Klhl41b orchestrates sarcomere assembly and growth to drive skeletal muscle hypertrophy in vivo

Hyewon Han; Grace R Tate; Janet Shi; Alfredo H Bongiorno; Arian Mansur; Jeffrey Widrick; Sehajpreet Gill; Esmat Karimi; Henk Granzier; Jeffrey R Moore; Vandana A Gupta

doi:10.1186/s13395-026-00453-2

. 2026 Oct 8;16:38. doi: 10.1186/s13395-026-00453-2

Klhl41b orchestrates sarcomere assembly and growth to drive skeletal muscle hypertrophy in vivo

Hyewon Han ¹, Grace R Tate ¹, Janet Shi ¹, Alfredo H Bongiorno ², Arian Mansur ¹, Jeffrey Widrick ³, Sehajpreet Gill ¹, Esmat Karimi ⁴, Henk Granzier ⁴, Jeffrey R Moore ², Vandana A Gupta ^1,^✉

PMCID: PMC13659986 PMID: 42870309

Abstract

Background

Nemaline myopathy (NM) is a clinically and genetically heterogeneous disease caused by mutations in sarcomere-associated genes, with congenital forms representing the most severe end of the spectrum. Recessive mutations in KLHL41 underlie a severe congenital NM characterized by sarcomeric disorganization and impaired muscle growth. The in vivo mechanisms by which KLHL41 governs sarcomere assembly and growth remain poorly understood.

Methods

We generated zebrafish models of KLHL41 deficiency by the CRISPR-Cas9 approach to examine the temporal and molecular basis of sarcomere assembly defects in NM. The phenotypic and physiological characterization of skeletal muscle defects was investigated by birefringence, spontaneous coiling, spontaneous swimming behavior, and force generation assays. Structural analyses by electron microscopy and immunofluorescence assessed myofibril organization and sarcomere structures across developmental time points. Quantitative proteomics and biochemical assays were performed to identify perturbations in sarcomeric proteins and associated molecular pathways.

Results

We show that Klhl41b is essential for the formation and organization of sarcomeres, supporting both radial and longitudinal muscle growth. Loss of Klhl41b resulted in reduced sarcomere size, defective sarcomere assembly followed by progressive destabilization of sarcomeres, nemaline body formation, and reduced muscle function. Proteomic analysis in Klhl41b-deficient zebrafish revealed reduced abundance of the sarcomeric troponin complex proteins, altered abundance of components of the translational machinery, and increased abundance of components of atrophy-associated pathways.

Conclusions

These data show that Klhl41b is required for sarcomere assembly and growth, and its deficiency drives a cascade of structural and molecular changes underlying skeletal muscle hypotrophy and NM pathology. These findings advance our understanding of disease mechanisms and may inform targeted therapeutic strategies to restore sarcomere integrity and muscle function.

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1186/s13395-026-00453-2.

Keywords: Nemaline Myopathy, Zebrafish, Disease Modeling, Sarcomere, thin filaments, KLHL41, Klhl41a, Klhl41b, Kelch, Duplicated genes

Introduction

The sarcomere is the fundamental contractile unit of striated muscle, playing a central role in muscle structure and function [1, 2]. Sarcomere assembly and growth involve a complex interplay of structural proteins, mechanical forces, signaling, metabolic pathways, epigenetic processes, and molecular chaperones that coordinate the sequential organization of actin and myosin filaments into repeating sarcomeric units [3–12]. Defects in sarcomere structure and function play a pivotal role in the development of genetic and acquired myopathies, which are characterized by muscle weakness and dysfunction and early lethality in severe cases [13, 14]. Mutations in genes encoding key sarcomeric proteins or their regulators can disrupt the structural integrity and contractile function of muscle fibers, leading to hypotrophy, atrophy, and impaired force transmission [15–18]. Therefore, a deeper understanding of sarcomere formation and assembly into mature myofibers could provide critical insights into the mechanisms underlying sarcomeric defects that lead to human myopathies.

Nemaline myopathy (NM) is a congenital neuromuscular condition marked by muscle weakness, hypotonia, and the presence of rod-like nemaline bodies in muscle fibers [19]. NM arises from mutations in genes encoding sarcomeric thin-filament proteins or regulators of sarcomere structure [20–30]. Mouse and zebrafish animal models of NM recapitulate the disease pathology and exhibit shorter or disorganized sarcomeres with nemaline bodies similar to those of NM patients. Mutations in Kelch family members KLHL40 or KLHL41 result in a severe form of nemaline myopathy with extensive hypotonia and early lethality in most patients [27, 28]. While KLHL41 has been linked to sarcomeric protein stability, its direct role in sarcomere assembly and organization in vivo remains unresolved [9, 31]. Defining the origins of these structural abnormalities requires insights into how sarcomeres are assembled and organized during muscle development in vivo.

Sarcomere assembly is described by several models, primarily based on in vitro studies, leaving key aspects of vertebrate in vivo assembly unresolved [32–36]. The premyofibril model suggests that disorganized, short premyofibrils with punctate Z-bodies gradually develop through an intermediate nascent state into mature, aligned myofibrils (with well-organized Z-lines) through the replacement of nonmuscle myosin with muscle-specific myosins [37]. Alternatively, another model proposes that thin and thick filaments assemble independently before aligning with each other through interdigitation [38]. The stress fiber model demonstrates that muscle stress fibers are precursors of actin and myosin filaments that assemble to form sarcomeres [35]. Consistent with these models, our previous studies of iPSC-derived muscle fibers showed the presence of immature and transitional states of sarcomere assembly that lack the organized periodicity characteristic of mature sarcomeres [4]. While these models suggest different mechanisms of sarcomere assembly, how these distinct pathways operate in vivo and contribute to myopathic phenotypes remains unresolved.

To investigate in vivo sarcomere assembly and the pathogenic mechanisms of NM, we developed zebrafish models of NM. Our studies reveal that the zebrafish orthologue of KLHL41, Klhl41b, plays a crucial role in sarcomere size and assembly, thereby contributing to the formation of functional myofibers. Most importantly, our findings suggest that KLHL41-related nemaline myopathy arises from the formation of structurally defective sarcomeres that progressively destabilize, rather than from a primary defect in sarcomere maintenance. Notably, we show that TNNT3, mutations in which cause a severe form of NM, contributes to KLHL41-dependent muscle function, underscoring the importance of thin filament components in regulating sarcomere architecture. These findings suggest that early defects in sarcomere assembly contribute to weak skeletal muscle in the congenital forms of NM. This refined perspective provides a foundation for developing more effective therapeutic strategies to restore sarcomere structure and function and improve clinical outcomes for patients with NM.

Results

Klhl41b deficiency results in skeletal muscle dysfunction in zebrafish

Mutations in KLHL41 result in severe muscle weakness at birth, underscoring its essential role in normal skeletal muscle development [28]. To uncover the role of KLHL41 in normal and disease states in skeletal muscle, we focused on identifying the earliest defects that emerge before the onset of overt pathological features. Zebrafish are particularly well suited for this purpose due to their rapid and externally developing embryos, optical transparency, and highly conserved skeletal muscle architecture. Therefore, we created a series of loss-of-function klhl41 alleles for KLHL41 zebrafish orthologs using CRISPR/Cas9 gene editing, enabling early in vivo analysis of skeletal muscle development (Supplemental Table 1, Fig. 1A-B). The KLHL41 ortholog in zebrafish is duplicated as klhl41a and klhl41b. We created mutants for both genes using the CRISPR-Cas9 approach. klhl41a alleles created include klhl41a^bw212g with an insertion of a base pair (c.250_251insA; p.Val84Aspfs*36) and klhl41a^bw213g with a deletion of 5 base pairs (c.1109_1113del; Trp371Cysfs*16) (Fig. 1A). klhl41b alleles include klhl41b^bw214g with an insertion of a base pair (c.674_675insC); p.Arg225Profs*14 and klhl41b^bw215g with a deletion of 5 base pairs (c.833_837del; p.Asn278Argfs*13) (Fig. 1B).

Fig. 1 — *klhl41b* is required for early skeletal muscle development and function. A-B Zebrafish *klhl41* series of knockout mutant alleles were generated by the CRISPR-Cas9 approach and analyzed after outcrossing for 4–5 generations. The *klhl41* gene is duplicated in zebrafish, and knockout mutant alleles were created for both duplicated genes (*klhl41a* and *klhl41b*). The locations of the mutations are shown in the protein domains (top) and exons (bottom). C-D Phenotypic (left panel) and birefringence analysis (right panel) of control, *klhl41a*, *klhl41b*, and *klhl41ab* larval fish in normal (left panel) and polarized light (right panel), demonstrating normal birefringence in *klhl41a* mutants but reduced brightness in the *klhl41b* and *klhl41ab* mutants compared to controls (5 dpf, n = 6 in each group from three different clutches). E Survival curve of control, *klhl41a*, *klhl41b*, and *klhl41ab* zebrafish. *klhl41a* mutants exhibited normal life span compared to controls, whereas *klhl41b* and *klhl41ab* mutants died within 7 days post-fertilization (n = 100 in each group, and 3–4 independent clutches were analyzed). F Quantification of the spontaneous coiling at 20 hpf demonstrates early muscle weakness in *klhl41b* and *klhl41ab* embryos (n = 20–24 in each group from three different clutches). G Swimming analysis was performed to analyze muscle function. Representative tail curvature profile of a wild-type and a *klhl41b* mutant larva during an escape response (5 dpf). The escape response stimulus is represented by the red arrow. The vertical dashed lines delineate the onset of movement (movement time = 0) and the three subsequent escape response stages: the C-start (1), the power stroke (2), and burst swimming (3). The images show the larvae prior to movement, at the stage 1 to 2 transition, the stage 2 to 3 transition, and during each subsequent stroke of stage 3 burst swimming. H Trend of distance traveled for the single wild-type and mutant larval fish shown in (G). I The instantaneous speed trend of the single wild-type and mutant larval fish is shown in (G). Speed oscillates due to the undulatory nature of larval swimming. The speed response was smoothed for further analysis. J Cumulative distance covered in 60 ms of escape response swimming, peak instantaneous speed, and the mean tail stroke amplitude and frequency of stage 3 burst swimming for wild-type and *klhl41b* larvae. Each point represents a single larva (the mean of three separate trials). Data are mean ± SEM; unpaired, two-tailed t-test, parametric for two groups or one-way analysis of variance (ANOVA) and Tukey’s HSD test for multiple group comparisons. **** p < 0.0001; ns: not significant

Quantitative analysis of the mRNA showed reduced amounts of klhl41 mRNA in the respective mutant alleles, suggesting impaired transcript stability (Figure S1A-B). Transcript analysis also showed no upregulation of klhl41a in klhl41b mutants, indicating no evidence of transcriptional compensation (Figure S1A-B). Western blot analysis of the protein extracts from klhl41a and klhl41b mutants showed a reduced amount of the Klhl41 protein, while klhl41a/klhl41b showed almost complete absence of protein. No truncated Klhl41 protein was detected, indicating reduced stability of the mutant protein as seen previously in human patients (Figure S1C) [27, 28, 39]. Phenotypic analysis of the wild-type control and mutant groups showed no obvious differences by light microscopy (Fig. 1C, left panel). Examination of skeletal muscle organization by polarized light showed no significant difference in birefringence between the control and klhl41a mutants. However, birefringence was reduced in the myotome of klhl41b and klhl41ab mutants compared to controls, indicating disrupted sarcomere organization (5 dpf) (Fig. 1C-D). Klhl41a deficiency did not affect survival in zebrafish, while Klhl41b and Klhl41ab deficiency resulted in reduced survival and death of mutants by 7 dpf (Fig. 1E).

Primary myogenesis leads to the formation of primary axial musculature during embryonic development in zebrafish that can contract to produce spontaneous coiling in the muscle as the first measure of motor function between 20 and 28 h post-fertilization (hpf). The frequency of spontaneous body coiling was reduced in klhl41b and klhl41ab embryos compared with the control siblings at 20 hpf, suggesting a defect in primary skeletal muscle formation (Fig. 1F). Because klhl41b and klhl41ab double mutants exhibited similar early muscle function, muscle structure, and survival phenotypes, whereas klhl41a mutants were indistinguishable from controls, all subsequent studies were performed using klhl41b mutants.

To evaluate the effect of Klhl41b deficiency on motor function, the escape swimming response of control and klhl41b mutant larvae was studied at 5 dpf using high-speed videography [40]. Escape responses were partitioned into an initiating C-start (stage 1) where the larva curves to bring the head and tail together, a counter bend or power stroke (stage 2), and a concluding burst of fast undulatory tail strokes (stage 3) (Fig. 1G). klhl41b mutants attained a significantly slower peak instantaneous speed and covered significantly less overall distance than wild-type larvae (Fig. 1H-J). Importantly, klhl41b larvae failed to achieve the extreme bending angles observed in wild-type larvae during the C-start (Figure S2, Stage 1: final tail curvature), power stroke (Figure S2, Stage 2: tail stroke amplitude), and stage 3 burst swimming (Fig. 1J). While the tail stroke frequency during stage 3 was similar to wild-type (Fig. 1G), the maximal tail stroke angular velocity was reduced in all three stages. Therefore, the impaired locomotion of the klhl41b mutants reflects their inability to bend the propulsive tail to the same extreme angles observed in wild-type controls, coupled with a failure to increase tail stroke angular velocity and stroke frequency to compensate, leading to reduced swimming compared to controls.

These results show that Klhl41b deficiency in zebrafish leads to reduced survival, changes in skeletal muscle organization, and reduced muscle function.

Klhl41b is required for early myofibril assembly but is dispensable for primary myoblast fusion

KLHL41 is expressed in developing mouse (E9.5) and zebrafish somites (11 h post-fertilization), suggesting a requirement in vertebrate skeletal muscle during early myogenesis [28, 41]. To identify the initial events regulated by Klhl41b during skeletal muscle development, we analyzed the myotome of control and klhl41b mutants at 24 h post-fertilization (hpf) when primary myogenesis is largely complete, for potential fusion defects (Figure S3). In zebrafish embryos, only fast muscle precursors undergo fusion to form multinucleated fibers, whereas slow muscle fibers remain mononucleated [42, 43]. Slow muscle is spatially segregated from fast muscle and forms a mononucleated outer layer in the myotome at 23–24 hpf. Analysis of the medial somite region showed no differences in the number of nuclei of control and Klhl41b mutant myotubes (somites 7–11), suggesting that Klhl41b is not required for primary myoblast fusion in vivo, and the defects observed in skeletal muscle potentially originated during sarcomere formation.

Next, we investigated sarcomere assembly in developing zebrafish embryos to determine whether the motility and birefringence defects observed in klhl41b mutants arise from impaired sarcomere formation during early myofibrillogenesis or from defects in the maintenance of mature sarcomeres. Ultrastructural examination of the anterior trunk region in control embryos showed that myotubes lacked sarcomeres at 16–17 hpf (Fig. 2A-B), whereas the earliest evidence of sarcomere formation was detected at ~ 18 hpf with a single row of myofibrils in control myotubes (Fig. 2C). The electron microscopy analysis of sarcomere assembly focused primarily on fast muscle fibers, as these can be identified by their multinucleated nature and constitute the majority of the zebrafish myotome. Compared to single rows of myofibrils in control myotubes, a significant number of the myotubes (~ 28%) in the klhl41b mutant embryos showed the absence of myofibrils at this stage (Fig. 2D, quantified in Fig. 2E) or the presence of much thinner myofibrils compared to controls (Fig. 2F-G, white arrowheads) at 18 hpf. Quantitative analysis of electron microscopy images demonstrated a significant reduction in sarcomere height and length in klhl41b mutants compared to controls at 18 hpf (Fig. 2H). Myotubes in klhl41b mutants also showed the presence of electron-dense aggregates in the vicinity of myosepta junctions (Fig. 2D, black arrows and Fig. 2D’ higher magnification). Within a few hours (24 hpf), control myotubes exhibit parallel, 3–4 organized rows of myofibrils along the longitudinal periphery in myotubes emanating from the myosepta in the proximity of the sarcolemma (Fig. 2I, L, and 4). In control myotubes, well-formed sarcomeres emanate from the myosepta (Fig. 2I), while klhl41b mutant myotubes exhibit only filamentous structures extending from the myosepta (Fig. 2J, white arrows), indicating a delay in sarcomere assembly. Compared to multiple rows of myofibrils in controls, klhl41b mutant myotubes predominantly displayed a single row of myofibrils at 24 hpf (Fig. 2L-M, quantification shown in 2N). As observed at earlier stages, klhl41b myotubes at 24 hpf also exhibited electron-dense aggregates near the myosepta region (Fig. 2J, black arrows). Whole-mount immunofluorescence with phalloidin in control myotubes at 24 hpf revealed a clear striated pattern consistent with organized sarcomeres. In contrast, klhl41b mutants lacked this striation, with increased phalloidin signal also localized at the myosepta region, suggesting that observed aggregates and filamentous structures may correspond to improperly organized F-actin (Fig. 2K, white arrows). Quantification of sarcomere number also showed fewer sarcomeres in klhl41b mutant myofibers compared to controls at 24 hpf (Fig. 2N). Sarcomere length and height in the klhl41b mutant muscle were reduced compared to controls (Fig. 2N). These data show that klhl41b is required for normal sarcomere assembly and its loss results in delayed and defective myofibrillogenesis accompanied by aberrant accumulation of F-actin at the myosepta.

Fig. 2 — Klhl41b regulates sarcomere size and assembly during primary myogenesis. A-B Transmission electron microscopy (16 hpf) showed the absence of sarcomeres in the control and *klhl41b* myotubes (n = 3 biological replicates in each group). C-D Transmission electron microscopy (18 hpf) revealed the presence of single rows of myofibrils in differentiating myotubes in the control myotome and the presence of electron-dense aggregates (shown at higher magnification in D’). A significant number of myotubes in the *klhl41b* mutants failed to show the presence of myofibrils. E Quantification of myotubes lacking myofibrils at 18 hpf. Each data point represents the mean value from an individual biological replicate (n = 4 biological replicates per group). Statistical comparisons were performed at the animal level using a two-tailed permutation test (70 labelings). F-G Compared to the control, myotubes in *klhl41b* mutants with myofibrils contained smaller sarcomeres, with reduced length and height. Sarcomere length and height were measured as indicated in the representative example shown at the top of the figure. H Quantification of sarcomere length and height at 18 hpf. Multiple sarcomeres were measured within each biological replicate and averaged to generate a single value per replicate; each data point represents one biological replicate (n = 4 biological replicates per group). Statistical comparisons were performed at the animal level using a two-tailed permutation test (70 labelings), *p < 0.05. I-J Controls showed the presence of well-developed sarcomeres attached to myosepta junctions, while the *klhl41b* mutant showed filamentous-like structures originating from the myosepta junctions (black arrow) (n = 3 biological replicates per group). K Confocal microscopy of the whole-mount staining with phalloidin (labeling F-actin) revealed the accumulation of F-actin in the vicinity of the myosepta junction in *klhl41b* mutants (white arrows) at 24 hpf (n = 15–20 in each group). L-M Transmission electron microscopy of the myotome at 24 hpf showed the presence of 3–4 rows of myofibrils in each myotube in controls compared to single rows of myofibrils in the *klhl41b* mutants (n = 3 biological replicates in each group). N Quantification of myofibril rows and sarcomere number at 24 hpf. Multiple myofibers and sarcomeres were measured within each biological replicate and averaged to generate a single value per replicate; each data point represents one biological replicate (n = 3 biological replicates per group). Statistical comparisons were performed at the animal level using a two-tailed permutation test (20 labelings), *p = 0.05. Data are presented as mean ± SEM. Somites 7–11 were analyzed at the medial depth of the trunk region

Fig. 4 — Proteomic changes in Klhl41b deficiency in skeletal muscle. A Skeletal muscle tissue from control and *klhl41b* mutants was dissected (3 dpf), and the proteome was analyzed by quantitative mass spectrometry (n=3 replicates of 50 embryos in each group). B Volcano plot showing protein abundance in control versus *klhl41b* mutant groups (cutoff log2 fold change=1.25 and p < 0.05). C Reactome pathway enrichment in control or *klhl41b* mutants revealed alterations in sarcomeric contraction, metabolic, translational, and proteasomal pathway proteins in *klhl41b* mutants compared to controls. D Proteins that are downregulated (left) or upregulated (right) in the klhl41b mutants in the top enriched categories

Klhl41b is required for sarcomere structure and growth during secondary growth

Primary myogenesis in zebrafish skeletal muscle is complete by 24 hpf, establishing functional, contracting muscle with multiple rows of myofibrils within myotubes, contributing to the radial growth that continues during muscle growth. As development progresses, muscle fiber growth also proceeds via longitudinal growth, driven by the addition of serial sarcomeres at the ends of myofibers [44]. During skeletal muscle secondary growth, the number of sarcomeres in each myofiber was increased in controls by 3 dpf, while mutant myofibers contained significantly fewer sarcomeres along their length, indicating impaired longitudinal muscle growth due to defective serial sarcomere addition (Fig. 3A-D). Myofibers in the mutant muscle also showed the presence of overstretched sarcomeres, suggesting that reduced sarcomere addition in series during longitudinal growth places greater mechanical strain on the remaining sarcomeres (Fig. 3B, white arrows). Notably, we also observed an abnormal accumulation of F-actin at the myoseptal ends of mutant myofibers at 3 dpf similar to earlier stages (Fig. 3A-B). Ultrastructural examination showed a lack of sarcomeric organization and the presence of electron-dense aggregates in the proximity of myosepta in klhl41b mutants compared to controls (Fig. 3E-F, black arrows, inset higher magnification). Sarcomeres in the klhl41b mutant muscle also exhibited thickened or irregular Z-lines compared with controls (Fig. 3G-H, white arrows) as previously seen in the skeletal muscle of NM patients [45]. While the sarcomere length in klhl41b mutants was comparable to that of controls, the sarcomere height was significantly reduced compared to controls (Fig. 3I). Together, these observations are consistent with impaired incorporation or organization of actin within developing sarcomeres in klhl41b mutant muscle. The persistence of F-actin accumulation at myoseptal regions outside of organized sarcomeric structures in klhl41b mutants, together with reduced sarcomere number and height, points to an impairment in myofibril maturation that extends beyond primary myogenesis into the secondary growth phase.

Fig. 3 — Klhl41b deficiency results in reduced sarcomere size, number, and stability. A-B Whole-mount immunofluorescence of control and klhl41b mutants with phalloidin (green) and DAPI (blue) at 3 dpf. White arrows indicate overstretched sarcomeres at the myosepta junction. C Quantification of sarcomere number in control and *klhl41b* mutants at 3 dpf. Each data point represents the mean value from an individual biological replicate (n = 10 per group). Statistical comparison was performed at the animal level using a two-tailed Welch’s t-test. D Representative image illustrating the method used to quantify sarcomere number from phalloidin-stained myofibers; each segment along the dashed line corresponds to one sarcomere. E-F Transmission electron microscopy images showing lack of sarcomere organization and presence of electron-dense aggregates (inset, higher magnification) in the proximity of myosepta in *klhl41b* mutants (3 dpf). G-H *klhl41b* mutants exhibit the presence of irregular and thickened Z-lines (white arrows) at 3 dpf compared to controls, and quantification of sarcomere size showed reduced sarcomere height in *klhl41b* mutants compared to controls (I). Multiple sarcomeres were measured within each biological replicate and averaged to generate a single value per replicate; each data point represents one biological replicate (n = 3 biological replicates per group). Statistical comparisons were performed at the animal level using a two-tailed permutation test (20 labelings), *p = 0.05, ns = not significant. J-K Transmission electron microscopy of myotome at 5 dpf revealed the presence of electron-dense bodies in the skeletal muscle of *klhl41b* in the disorganized sarcomeric regions (white arrowheads) as well as on Z-lines (white arrows) (n = 3 biological replicates per group). L Quantification of sarcomere height and length at 5 dpf. Multiple sarcomeres were measured within each biological replicate and averaged to generate a single value per replicate; each data point represents one biological replicate (n = 3 biological replicates per group). Statistical comparisons were performed at the animal level using a two-tailed permutation test (20 labelings), *p = 0.05, ns = not significant. M-N Whole-mount immunofluorescence in zebrafish with phalloidin (F-actin, green), laminin (red), and DAPI (blue) revealed the accumulation of F-actin in the periphery of the myosepta junction (white arrows) and overstretched, broken myofibers (white arrowhead) (n = 3 biological replicates per group)

Klhl41b deficiency leads to the accumulation of nemaline bodies and sarcomeric dysfunction

To determine how these early defects in Klhl41b deficiency progress during later stages of muscle growth, we next examined skeletal muscle ultrastructure at 5 days post-fertilization (dpf). Transmission electron microscopy revealed striking differences in sarcomere organization between control and klhl41b mutant skeletal muscle at 5 dpf. In control myofibrils (Fig. 3J), sarcomeres displayed a highly regular and ordered architecture, with Z-discs aligned uniformly in parallel and evenly spaced across adjacent myofibrils. In contrast, klhl41b mutants (Fig. 3K) exhibited severe sarcomere disarray and regions of Z-lines showing the presence of sarcomeric aggregates (Fig. 3K; highlighted by white arrowheads). The mutant myofibrils showed deviation from their parallel orientation, with accumulation of electron-dense bodies in myofibrils, similar to nemaline bodies seen in NM patients [19, 27, 28]. To further characterize the structural defects, phalloidin staining was performed to visualize F-actin organization at 5 dpf. Control muscle (Fig. 3M) displayed uniform and well-ordered actin filament distribution along intact myofibers, with no evidence of structural disruption. Strikingly, klhl41b mutant muscle revealed focal accumulations of actin (Fig. 3N, white arrows) reflecting aberrant actin aggregation, as well as myofiber breakage (Fig. 3N, white arrowheads), indicative of compromised myofiber integrity and mechanical failure. Quantitative analysis corroborated these ultrastructural observations, demonstrating that while sarcomere length remained comparable between control and mutant muscle (Fig. 3L), sarcomere height was significantly reduced in klhl41b mutants compared to controls.

To investigate the effect of structural changes on force production, muscle physiological analysis was performed. We examined the kinetics of force production in control and klhl41b mutants. Compared with controls, the klhl41b mutant exhibited lower force production across all stimulation frequencies (1–200 Hz) (Figure S5). We further assessed twitch characteristics at 1 Hz. While no significant differences were observed in activation kinetics (time to peak force), mutant muscles showed a significant reduction in peak tension compared to controls, consistent with an overall deficit in force-generating capacity.

Together, these findings demonstrate that klhl41b deficiency leads to a combined pathology characterized by sarcomere disorganization, the presence of nemaline body-like aggregates, aberrant actin accumulation, myofiber fragility, and reduced force production by 5 dpf, underscoring a critical role for Klhl41b in maintaining the structural integrity of the contractile apparatus, myofiber architecture, and force generation capacity of the skeletal muscle.

Klhl41b deficiency results in changes in the sarcomeric, translational, and proteolytic proteome in skeletal muscle

KLHL41 is a substrate-specific adapter for CUL3 E3 ubiquitin ligase that targets specific proteins, and the KLHL41-CUL3 complex regulates the stability of skeletal muscle proteins by ubiquitination. To determine how the protein composition of skeletal muscle is altered in Klhl41b deficiency, we performed mass spectrometry (MS)-based quantitative proteome analysis of skeletal muscle from control and klhl41b mutant samples (log₂Fold change = 1.25, p < 0.05) (Fig. 4A), which were analyzed by the Reactome pathway analysis to identify enriched functional categories across experimental groups (Fig. 4B-C). Functional enrichment of proteins with a significant differential response between klhl41b mutants and controls revealed “striated muscle contraction” as the most significantly enriched functional category. Notably, several components of the skeletal muscle thin filament troponin complex, including Tnnt3a, Tnnt1, Tnnc2, Tnni1a, Tnni1b, Tnni2a, and Tnni2b, were among the most markedly decreased proteins, indicating broad disruption of troponin protein abundance in Klhl41b-deficient muscle (Fig. 4D). Consistent with the role of thin filament dysfunction in nemaline myopathy (NM), additional NM-associated proteins such as Neb, Tpm2, and Klhl40a were also reduced in mutants, whereas Acta1, Lmod3, Kbtbd13, and Cfl2 were unchanged. In contrast, Tpm3 and Mypn showed modest increases in abundance in klhl41b mutants (Table S3). In addition, protein groups associated with phospholipid metabolism were also reduced in abundance in klhl41b mutants compared to controls.

Interestingly, proteins involved in translational processes exhibited bidirectional changes; while some components were reduced, there was a concurrent enrichment of proteins associated with ribosome assembly and translational initiation, suggesting dysregulation rather than uniform suppression of translation. Conversely, mutant muscle showed a significant increase in proteasome-associated proteins, including skeletal muscle-specific E3 ubiquitin ligases implicated in atrophy, such as Murf1, Murf2, Murf3, Atrogin, and Mib (Fig. 4C–D). Together, these findings indicate that Klhl41b is required for maintaining thin filament protein homeostasis, with its loss associated with broad alterations in translational and proteostasis pathways in skeletal muscle. While some of these changes may result directly from Klhl41b deficiency, others likely reflect secondary consequences of disrupted sarcomere organization or impaired muscle function.

KLHL41 stabilizes TNNT3 in skeletal muscle

Our previous studies have shown that KLHL41 regulates the dynamics of protein complexes by modulating their stability through ubiquitination by the CUL3 E3 ligase KLHL41 complex [9]. In the klhl41b mutant skeletal muscle, all the members of the troponin protein complex showed reduced abundance. The troponin complex is the Ca2+-regulating complex for skeletal muscle contraction and is localized to the thin filaments in sarcomeres (Fig. 5A). Mutations in skeletal muscle troponin complex proteins are associated with severe myopathy [25, 46–50]. Therefore, we investigated whether KLHL41 regulates the stability of the fast skeletal muscle troponin proteins. While all skeletal muscle troponin proteins were downregulated in the mutants, the highest downregulation in the klhl41b muscle was observed for the fast-muscle troponin protein complex (TNNT3, TNNI2, and TNNC2). Therefore, we focused on understanding the effect of KLHL41 on the fast-muscle-specific troponin proteins. To identify if KLHL41 also associates with different troponin protein members, KLHL41-FLAG was overexpressed in the C2C12 muscle cells, and co-immunoprecipitation with anti-FLAG antibody was performed in the differentiating myotubes (day 2; two days in the differentiation media). Western blot analysis of the immunoprecipitated complex revealed that KLHL41 associates with endogenous TNNT3 but not with the TNNC2 or TNNI2 (Fig. 5B). To confirm this association under endogenous conditions, we performed co-immunoprecipitation using a KLHL41 antibody from mouse skeletal muscle protein extracts. TNNT3 co-immunoprecipitated with endogenous KLHL41, further supporting an association between KLHL41 and TNNT3 in skeletal muscle (Fig. 5C).

We further analyzed if TNNT3 undergoes ubiquitination in the presence of KLHL41 and CUL3 ligase in C2C12 myotubes (Fig. 5D). No ubiquitination was observed for TNNT3 in the presence of KLHL41 and CUL3. MURF1 was used as a positive control, as TNNT3 has previously been reported to be a target of MURF1 E3 ubiquitin ligase. Immunoprecipitation of TNNT3 from C2C12 myotubes showed that TNNT3 is ubiquitinated in the presence of MURF1. These results indicate that KLHL41 association with TNNT3 does not promote CUL3-dependent ubiquitination. Next, we evaluated the effect of the KLHL41 protein on TNNT3 protein stability. Increasing amounts of the KLHL41-FLAG plasmid were overexpressed in C2C12 myotubes, and western blotting was performed to quantify the level of the troponin protein complex proteins (TNNT3, TNNI2, and TNNC2) (Fig. 5E). Increasing KLHL41 protein levels led to an increase in TNNT3 protein levels in differentiating myotubes. Similarly, an increase in KLHL41 also increased TNNC and TNNI levels (Fig. 5E-F). Together, these findings support a model in which KLHL41 associates with TNNT3, with increased TNNT3 abundance accompanied by increased levels of other components of the fast skeletal muscle troponin complex.

Klhl41b deficiency alters calcium sensitivity and cooperativity of thin filament activation

The troponin complex is the primary regulator of skeletal muscle contraction, where calcium binding to troponin C induces conformational changes in the thin filament that enable actin-myosin interactions and cross-bridge formation. To determine whether reduced abundance of troponin complex components affects thin filament activation, we performed in vitro motility assays using thin filaments purified from control and klhl41b mutant zebrafish (Fig. 5G–H). Calcium sensitivity was quantified by the pCa₅₀ value, which represents the calcium concentration required for half-maximal filament speed, and cooperativity was assessed using the Hill coefficient (nH). In control filaments, motility was effectively inhibited at low calcium concentrations and increased progressively with rising calcium levels, as expected (Fig. 5G). Thin filaments from klhl41b mutants also exhibited calcium-dependent activation and reached similar maximal filament speeds at high calcium, indicating preserved maximal activation capacity (Fig. 5H). However, mutant thin filaments failed to fully suppress motility at low calcium concentrations and exhibited a significant leftward shift in calcium sensitivity (increased pCa₅₀). In addition, klhl41b mutant filaments showed reduced cooperativity, as reflected by a significantly lower Hill coefficient, indicating a more gradual activation response to increasing calcium levels (Fig. 5). Together, these findings suggest that klhl41b deficiency disrupts the regulatory function of the thin filament, leading to increased calcium sensitivity and reduced cooperativity of activation, consistent with impaired control of actomyosin interactions.

Tnnt3a is required for myofibril architecture and sarcomere organization and partially rescues muscle function in Klhl41b deficiency

Proteomic analysis revealed that Tnnt3a protein levels are reduced in Klhl41-deficient muscle, suggesting that TNNT3 may act downstream of KLHL41 in maintaining thin filament integrity. This observation raises the possibility that defects in TNNT3 not only disrupt contractile regulation but may also contribute to sarcomere structural defects observed in KLHL41 deficiency. TNNT3 mutations have been previously reported in arthrogryposis with neonatal lethality and congenital nemaline myopathy [51], but suitable vertebrate animal models are unavailable to investigate the effect of TNNT3 in skeletal muscle development. To determine whether TNNT3 deficiency contributes to sarcomere assembly and stability, we analyzed a zebrafish knockout model of the human TNNT3 orthologue (tnnt3a).

tnnt3a mutant fish (c.A529T; NM_131565.1 and p.K177*) were obtained from the Zebrafish International Resource Center (Fig. 6A). Transcript analysis showed no differences in the tnnt3a mRNA levels between control and tnnt3a mutant zebrafish at 3 dpf (Figure S6A). Western blot analysis with a TNNT3 antibody revealed a decrease in the Tnnt3 protein in zebrafish, with no truncated protein detected, suggesting a loss of Tnnt3a protein in the tnnt3a mutant (Figure S6B). Compared to controls, tnnt3a mutants die between 3 and 4 dpf. Skeletal muscle analysis by birefringence assay showed no significant differences in the birefringence between control and tnnt3a mutants (3 dpf; Fig. 6B-C). However, ultrastructure analysis of the skeletal muscle of tnnt3a mutants (2 dpf) by electron microscopy revealed a reduced number of myofibril rows in myotubes compared to control siblings, suggesting that Tnnt3a is required for myofibril formation during skeletal muscle development (Fig. 6D-E, J). Control myotubes displayed well-organized sarcomeres while tnnt3a mutants exhibited sarcomeres with irregular and thickened Z-lines compared to controls (Fig. 6F-G). Quantification of the sarcomere size revealed that sarcomere height was normal, but sarcomere length was significantly reduced in tnnt3a mutants compared to controls (Fig. 6K-L). Moreover, Z-lines were irregular and thicker in tnnt3a mutants compared to controls (Fig. 6F-G and M). At the myosepta ends, control myofibers contained well-organized mature sarcomeres while tnnt3a mutant myofibers showed regions devoid of mature sarcomeres, with small and stretched sarcomeres at the myofiber ends (Fig. 6H-I). These data show that tnnt3a contributes to sarcomere growth, assembly into myofibrils, and stability during skeletal muscle development. The phenotypic convergence between tnnt3a and klhl41b mutants, together with the association between KLHL41 and TNNT3, supports a model in which the KLHL41-dependent regulation of TNNT3 contributes to proper sarcomere assembly and growth during skeletal muscle development.

To test whether Tnnt3a functions downstream of Klhl41b, we tested if restoring Tnnt3a expression could rescue muscle function in klhl41b mutants. Increasing doses of tnnt3a mRNA (0-100 pg) were injected into tnnt3a or klhl41b mutants or wild-type control embryos, and swimming performance was quantified by automated locomotor tracking (Fig. 6N). To confirm that the injected mRNA is translated into protein, we performed immunoblotting for Tnnt3 in wild-type embryos injected with tnnt3a mRNA, which showed dose-dependent production of Tnnt3 protein following injection (Figure S6C). In tnnt3a mutants, overexpression of the tnnt3a mRNA led to a dose-dependent improvement in average distance traveled, confirming the functional activity of the injected mRNA. Overexpression of tnnt3a mRNA in klhl41b mutants also improved swimming in a dose-dependent manner, with a significant increase over uninjected klhl41b mutants at the 50 pg dose. However, the rescue was incomplete, and the differences in swimming activity remained significant at all doses tested. Together, these results support a model in which Tnnt3a functions downstream of Klhl41b to promote muscle function, while the incomplete rescue indicates that Tnnt3a is unlikely to be the sole effector of Klhl41b-dependent muscle function and may act in concert with additional Klhl41b-regulated factors.

Early atrophy-associated gene activation accompanies primary sarcomeric defects

Proteomic analysis of klhl41b mutant muscle revealed enrichment of proteins involved in proteolytic pathways, suggesting activation of muscle atrophy programs. Although muscular atrophy is a common feature of myopathies and dystrophies, the initiating triggers of atrophic signaling in skeletal muscle remain poorly understood. Notably, many proteolytic enzymes and muscle-specific E3 ubiquitin ligases localize to the sarcomere, raising the possibility that disruption of sarcomere architecture may be associated with activation of these pathways.

To explore this possibility, we examined the expression of atrophy-associated genes across zebrafish models with distinct structural phenotypes: mutants with primary sarcomeric defects (klhl41b, tnnt3a) and dystrophic mutants (dmd, dag1) that retain normal sarcomere organization during early development (2 dpf) (Fig. 7). RT-PCR analysis demonstrated a significant upregulation of key atrophy-related transcripts, including murf1, murf2, murf3, atrogin, and mib1, in both klhl41b and tnnt3a mutants, indicating transcriptional activation of atrophy pathways. In contrast, these transcripts were not significantly altered in dmd (sapje) or dag1 mutants at the same developmental stages, despite eventual muscle degeneration in these mutants. Together, these findings suggest that early activation of atrophy-associated genes accompanies primary sarcomeric defects during zebrafish muscle development.

Fig. 7 — Early atrophy-gene activation accompanies sarcomeric defects in klhl41b and tnnt3a mutants. Quantitative RT-PCR analysis of control and klhl41b, tnnt3a, dmd and dag1 mutants for skeletal muscle atrophy transcripts (2 dpf) (n=20 in each group). Data are mean ± SEM. Statistical comparisons were performed with one-way analysis of variance (ANOVA) followed by Tukey’s HSD test. **** p < 0.0001; ** p < 0.01; ns: not significant

Discussion

Mutations in approximately 200 genes contribute to the diverse array of sarcomeric diseases affecting sarcomere assembly, maintenance, or growth (https://www.musclegenetable.fr/). However, no therapies are available for most of these disorders because the origins of the specific sarcomeric defects that contribute to skeletal muscle weakness remain poorly understood. Therapeutic development for sarcomeric diseases remains inherently challenging due to fundamental biological constraints: many causal genes encode exceptionally large structural proteins that are difficult to target with current gene delivery approaches, while others underlie rare or ultra-rare myopathies, limiting opportunities for scalable therapeutic development. By understanding the specific sarcomeric defects in vertebrate animal models of human myopathies, we may devise more broadly applicable therapeutic strategies to restore sarcomere structure and function in these diseases.

Nemaline myopathy is a sarcomeric disease in which mutations in 12 genes, including KLHL41, result in disorganized sarcomeres, nemaline bodies, and hypotonia in affected patients. However, it remains unclear whether the sarcomeric defects observed in patients result from defects in sarcomere assembly, growth, or maintenance. In this study, we demonstrate that Klhl41b is required for the initiation of sarcomere assembly and plays a key role in regulating sarcomere size and stability in vivo. Consistent with these structural defects, mutants exhibit reduced spontaneous coiling during early development, a period when nascent sarcomeres first become functionally competent to support muscle contraction. Similarly, patients with KLHL41 mutations, as well as those with congenital nemaline myopathy caused by other genes, exhibit fetal akinesia and reduced fetal movement, suggesting that muscle weakness originates during fetal development [28].

Klhl41 is duplicated in zebrafish, and both klhl41a and klhl41b are expressed during somitogenesis. Morpholino-based knockdown of either gene or both resulted in myopathy in zebrafish [28]. While klhl41b mutants recapitulated the myopathy observed in klhl41b morphants, klhl41a mutants did not exhibit the overt phenotype observed with morpholinos. This discrepancy may be attributed to genetic compensation by other genes in klhl41a mutants or off-target effects of morpholinos, as previously seen in other zebrafish models of skeletal muscle diseases [17, 52].

Using the zebrafish model, our findings provide in vivo insight into the earliest stages of sarcomere assembly in vertebrate skeletal muscle. The earliest detectable sarcomeres appear as single rows of myofibrils along the sarcolemmal membrane, anchored to the myosepta at both ends. Importantly, although previous immunofluorescence studies have identified the presence of sarcomeric proteins near the plasma membrane during the onset of myofibrillogenesis, they lacked the resolution to resolve the structural integrity or assembly state of individual sarcomeres [53]. In contrast, our EM analyses address this gap by providing ultrastructural resolution on myofibril organization, revealing that even at this earliest stage, sarcomeres exhibited well-defined Z-lines, organized thick and thin filaments, and distinct A-bands, I-bands, and H-zones, indicating that key structural features were already established at the time of detection. Notably, we did not observe isolated sarcomeres or intermediate assemblies resembling pre-myofibrillar structures described in previous models of myofibrillogenesis [54, 55]. This difference may reflect methodological differences, including the higher structural resolution afforded by electron microscopy compared to earlier imaging approaches. Following initial assembly, skeletal muscle growth is characterized by coordinated increases in myofibril number and sarcomere length. Specifically, we observed radial expansion through the addition of myofibril rows and longitudinal growth through serial addition of sarcomeres within existing myofibrils, thereby supporting muscle growth and function. Consistent with our observations, recent work in zebrafish demonstrated that sarcomere growth involves progressive expansion of myofibrils and that the abundance of the fast-muscle myosin light chain Mylpf regulates this process by promoting myofilament localization and myofibril growth [56]. Together, these findings highlight the importance of appropriate sarcomeric protein abundance and organization in supporting myofibril growth during muscle development.

Nemaline myopathy is classically defined by the presence of nemaline bodies, electron-dense protein aggregates derived from Z-line and other sarcomeric proteins, yet the sequence of pathogenic events leading to their formation remains incompletely understood [57]. In this study, using a zebrafish model of Klhl41b deficiency, we identify early disease-associated alterations that precede the appearance of nemaline bodies and provide insight into the temporal progression of pathology. Our data reveal that Klhl41b loss disrupts early myofibrillogenesis, as evidenced by a delay in the formation of organized myofibrillar rows and reduced sarcomere size. These findings are consistent with impaired radial growth of developing myofibrils contributing to hypotrophy, rather than a defect limited to maintenance of mature sarcomeres. With disease progression, we observe a reduction in the number of sarcomeres, limiting longitudinal myofiber growth, accompanied by irregular and thickened Z-lines, a structural abnormality reported across multiple forms of NM [19, 58]. Importantly, these early structural abnormalities are followed by the emergence of nemaline bodies and widespread sarcomeric disorganization, recapitulating the histopathological features observed in patient muscle at advanced stages [28]. Together, these data indicate that defects in sarcomere assembly and growth occur prior to overt protein aggregation and are associated with the progression of muscle pathology in KLHL41-associated NM (Fig. 8).

Fig. 8 — Proposed role of Klhl41b in normal and nemaline myopathy (NM) skeletal muscle. Klhl41b is required for (A) early sarcomere assembly and (B) subsequent skeletal muscle growth through the addition of new myofibril rows during radial growth and terminal sarcomere addition during longitudinal growth. In Klhl41b deficiency, myofibrils form with reduced sarcomere size (A) and later exhibit defective sarcomere addition during growth, accompanied by Z-line thickening (blue) (B). These abnormalities ultimately progress to sarcomere disorganization and protein aggregate accumulation resembling nemaline bodies in patients (C). At the molecular level, altered sarcomeric protein turnover and stability, along with disrupted translational and atrophy-associated pathways, contribute to disease pathogenesis. Together, these defects lead to skeletal muscle hypotrophy, weakness, and reduced force production

Proteomic and biochemical analyses demonstrate that loss of Klhl41b leads to coordinated downregulation of multiple troponin subunits, with TNNT3 emerging as a key KLHL41-associated protein whose stability is enhanced by KLHL41. These data support a role for KLHL41 in regulating TNNT3 stability, with associated effects on the abundance of other troponin complex proteins and thin filament composition. Consistent with this, we and others have previously demonstrated that KLHL41 is critical for thin filament integrity by regulating the stability of thin filament proteins, including nebulin and NRAP [9, 59]. Mechanistically, KLHL41 associates with TNNT3 and promotes its stability, whereas MURF1 ubiquitinates TNNT3 and reduces its protein abundance. Consistent with these molecular changes, functional analysis of thin filaments revealed increased calcium sensitivity and reduced cooperativity of activation, indicating impaired regulation of contraction. This altered calcium sensitivity in klhl41b mutants is consistent with impaired relaxation dynamics, which provides a mechanistic explanation for the reduced body curvature observed during swimming. Although TNNT3 is classically defined as a regulator of Ca²⁺-mediated contraction [60], its coordinated loss in KLHL41-deficient muscle, together with its integral position within the thin filament, prompted us to investigate whether TNNT3 plays a role beyond contractile regulation, specifically in sarcomere assembly and organization. Analysis of tnnt3a mutant zebrafish revealed reduced myofibril content, shortened sarcomeres, and disrupted Z-line architecture, indicating an additional requirement for TNNT3 in sarcomere assembly and organization during development. The phenotypic convergence between klhl41b and tnnt3a mutants, together with the observed alterations in thin filament regulation, supports a model in which KLHL41-dependent regulation of TNNT3 contributes to coordinated sarcomere assembly and functional regulation. Consistent with this model, overexpression of tnnt3a mRNA in klhl41b mutants partially rescued swimming activity, suggesting that Tnnt3a acts downstream of Klhl41b. However, the incomplete rescue, together with the distinct structural phenotypes of the two mutants, indicates that TNNT3 is not the sole mediator of KLHL41 function and that additional KLHL41-regulated thin-filament proteins likely contribute to the phenotype, as seen in our proteomics data (Neb, Tpm2) and previously described [31].

Beyond its role in regulating thin filament stability, our data indicate that Klhl41b also influences broader cellular processes required for muscle growth and maintenance. Proteomic analysis showed altered abundance of proteins associated with the translational machinery, suggesting disruptions in ribosome biogenesis and translation initiation in Klhl41b-deficient muscle. Such alterations may affect the synthesis of sarcomeric and regulatory proteins required for efficient myofibril assembly and muscle growth. Consistent with this, β2-adrenergic–mediated muscle hypertrophy, an anabolic process dependent on increased ribosomal protein production, is significantly impaired in KLHL41-deficient cellular models [61]. In addition, we observed activation of atrophy-associated pathways in both Klhl41b- and Tnnt3a-deficient muscle, whereas similar responses were not detected in Dag1- or Dmd-deficient muscle at the same developmental stage, when these mutants lack structural sarcomeric abnormalities. These findings indicate that early activation of atrophy-associated pathways accompanies primary sarcomeric defects during muscle development. Such activation of proteolytic pathways may further exacerbate sarcomeric instability, thereby contributing to progressive muscle weakness. Together, these findings suggest that Klhl41b has broader roles in protein stability, translational capacity, and proteolytic regulation that collectively support sarcomere assembly and muscle integrity (Fig. 9).

Overall, our findings provide a framework linking early defects in myofibril assembly to clinical manifestations of nemaline myopathy. Impaired radial growth of myofibrils is expected to limit muscle fiber cross-sectional area, contributing to muscle hypotrophy and reduced force generation, while defects in the addition of sarcomeres in series may impair longitudinal muscle growth and elasticity, predisposing to contractures. Notably, both hypotrophy and contractures are prominent features of KLHL41-associated NM and are also observed in other severe forms of NM caused by mutations in distinct genes, suggesting convergence on shared pathogenic mechanisms [25, 27, 28, 51, 62, 63]. While clinical presentations can vary across different genetic forms and mutation types in NM, our data support the idea that disruptions in sarcomere assembly and growth represent common disease drivers, particularly in severe forms such as KLHL41 or TNNT3 deficiency. Further comparative studies across genetic subtypes will be important to define the extent to which these mechanisms are conserved versus mutation-specific. Ultimately, these insights provide a foundation for identifying shared therapeutic targets and guiding the development of interventions to improve muscle growth and function across multiple forms of nemaline myopathy and other sarcomeric disorders.

Materials and methods

Zebrafish maintenance and husbandry

Zebrafish were maintained and bred using standard methods as described [64].

All experiments and procedures were approved by the Institutional Animal Care and Use Committee at Brigham and Women’s Hospital. Wild-type fish were obtained from the Tübingen (TU) line and staged by hours (h) or days (d) post-fertilization at 28.5 °C. Zebrafish embryonic, larval, juvenile, and adult stages of development were classified as described previously [65]. In all experiments, biological replicates were used.

Gene and protein nomenclature

Gene, mRNA, and protein nomenclature follows species-specific conventions established by the Zebrafish Information Network (ZFIN; www.zfin.org), Mouse Genome Informatics (MGI; www.informatics.jax.org), and the HUGO Gene Nomenclature Committee (HGNC; www.genenames.org). For zebrafish, gene symbols and mRNAs are written in lowercase italics (klhl41a, klhl41b, and tnnt3a), whereas the corresponding proteins are written in non-italicized Roman type with an initial capital letter (Klhl41a, Klhl41b, and Tnnt3a). For mouse, gene symbols and mRNAs are written in italics with an initial capital letter (Klhl41 and Tnnt3), whereas the corresponding proteins are written in non-italicized uppercase (KLHL41 and TNNT3). For human, gene symbols and mRNAs are written in italicized uppercase (KLHL41 and TNNT3), whereas the corresponding proteins are written in non-italicized uppercase (KLHL41 and TNNT3)

Creation of zebrafish lines

sgRNAs were designed using the web-based ZiFiT Targeter program (http://zifit.partners.org/) to target distinct sites in exon 1 or exon 2 of the zebrafish klhl41a and klhl41b genes. The first 2 bp GG sequences at the 5’ end of the target site are a constraint imposed by T7 promoter sequence requirements in addition to the NGG protospacer adjacent motif (PAM) sequence requirement immediately 3’ to the target site. Two oligonucleotides, each of 22 nucleotides in length, were used to construct the guide RNA for each target site. Forward and reverse primers were annealed to create a sgRNA oligonucleotide duplex. Primer sequences are summarized in Table S2. The guide RNA expression vector pDR274 was used to create the sgRNA expression system using the T7 promoter followed by in vitro transcription as previously described [66]. sgRNAs and Cas9 protein (Thermo Scientific, CA) were co-injected into the yolk sac of one- and two-celled stage zebrafish embryos. Each embryo was injected with 1–2 nL of the injection stock solution containing 2 uL of 400 ng/nL Cas9 protein and 3 uL of 100 ng/uL sgRNA. Injected embryos were inspected under the microscope for 3 days and classified as dead, deformed, or normal. Embryos displaying normal phenotypes were analyzed to assess the efficacy of sgRNAs by identifying target-site mutations. To analyze injected fish, genomic DNA was extracted from 6 to 8 pooled embryos at 3–5 days post-fertilization (dpf), PCR amplified, cloned into the pCR-TOPO vector, and subjected to Sanger sequencing to assess the efficacy of sgRNA-induced mutagenesis. tnnt3a mutant line (tnnt3a^sa24709) was obtained from the Zebrafish International Resource Center and contains a premature stop codon (c.A529T; NM_131565.1 and p.K177*). Heterozygous fish were outcrossed with the TU fish, and embryos obtained from heterozygous incrosses after the F3 generation were analyzed for all experiments in this work.

Identification of founder fish and generation of stable mutant fish

Because a fertilized zebrafish embryo develops quickly, direct delivery of sgRNA-Cas9 protein via injection results in chimeric embryos. Founder fish were determined by genotyping tail fin clips of the F0 generation. The mosaic F0 generation was outcrossed to wild-type TU fish for at least three generations for the studies presented in this work. Genomic DNA was extracted from the caudal fin tissue of F0 adult zebrafish and genotyped by Sanger sequencing. Mutation type and frequency were assessed using the ICE analysis tool (Synthego). F0 founders with a high proportion of predicted deleterious mutations were outcrossed with wild-type TU zebrafish. The presence of a single klhl41a or klhl41b mutation was confirmed by Sanger sequencing and ICE analysis. Three F3 founder zebrafish carrying the same mutation were selected for each allele, and outcrossed embryos from these individuals were pooled and raised to adulthood. For all experiments, at least 3–4 independent clutches from different parental pairs were analyzed. The sequences of sgRNAs are listed in Table S1, and sequencing primers are listed in Table S2. Wild-type control and mutant zebrafish (siblings) used in experimental procedures were genotyped either before the experimental procedures (proteomics, Western blotting, RT-PCR) or after experiments were performed (electron microscopy, immunofluorescence, activity analysis, survival curve). Head tissues (for imaging and proteomics studies) or the whole body were used for the DNA extraction for genotyping. All animal experiments and image analyses were performed in a blinded manner using coded sample identifiers. For most experiments, data acquisition and analysis were completed prior to genotyping, which was performed afterward to assign WT, heterozygous, and mutant groups. For experiments requiring prior genotype identification (e.g., proteomics), samples were coded after genotyping to maintain blinding during downstream analyses.

tnnt3a mRNA rescue and swimming analysis

Full-length zebrafish tnnt3a coding sequence (ENSDART00000103395) was cloned into a PCSDest destination vector and in vitro transcription was performed as previously described [67]. 25–100 pg of tnnt3a mRNA was injected into 1-cell embryos obtained from tnnt3a or klhl41b heterozygous crosses. Automated activity analysis was performed on larvae (4 dpf) as previously described [66].

Kinematic measurements in zebrafish larvae

The methods used to analyze escape response kinematics have been previously described in detail [40]. Briefly, a single larva was placed in a small arena, and an escape response was evoked using a brief electrical field pulse. The response was captured at 1000 frames/s by a high-speed camera. Recording began 10 ms prior to the stimulus and continued until the initial 60 ms of escape response movement was recorded. A minimum interval of 60 s was maintained between successive trials, four times longer than the inter-escape response period used in previous studies to prevent habituation. No evidence of response attenuation was observed [40]. A deep-learning neural network developed using the open-source toolkit DeepLabCut identified seven anatomical key points along the body and tail on each video frame [68, 69]. The x- and y-coordinates of these key points were used to calculate the linear kinematics of the larvae’s putative center of mass (COM, defined by a key point at the posterior edge of the swim bladder) and the angular kinematics of the propulsive tail. The average of three separate escape responses obtained per larva was used for analysis. Instantaneous speed fluctuates during swimming because of the oscillatory action of the propulsive tail. To obtain an estimate of peak swimming speed, the raw speed response was smoothed with LOESS (span = 0.7), as previously described [40].

Sample preparation for mass spectrometry

Samples for protein analysis were prepared essentially as previously described [70, 71]. Proteomes were extracted using a buffer containing 200 mM EPPS pH 8.5, 8 M urea, 0.1% SDS, and protease inhibitors. Following lysis, 50 µg of each proteome was reduced with 5 mM TCEP. Cysteine residues were alkylated using 10 mM iodoacetamide for 20 min at RT in the dark. Excess iodoacetamide was quenched with 10 mM DTT. A buffer exchange was carried out using a modified SP3 protocol [72]. Briefly, ~ 250 µg of Cytiva SpeedBead Magnetic Carboxylate Modified Particles mixed at a 1:1 ratio were added to each sample. 100% ethanol was added to each sample to achieve a final ethanol concentration of at least 50%. Samples were incubated with gentle shaking for 15 min. Samples were washed three times with 80% ethanol. Protein was eluted from SP3 beads using 200 mM EPPS pH 8.5 containing Lys-C (Wako, 129–02541). Samples were digested overnight at room temperature with vigorous shaking. The next morning, trypsin was added to each sample and further incubated for 6 h at 37 °C. Acetonitrile was added to each sample to achieve a final concentration of ~ 33%. Each sample was labeled, in the presence of SP3 beads, with ~ 125 µg of TMT reagents (ThermoFisher Scientific). Following confirmation of satisfactory labeling (> 97%), excess TMT was quenched by adding hydroxylamine to a final concentration of 0.3%. The full volume from each sample was pooled, and acetonitrile was removed by vacuum centrifugation for 1 h. The pooled sample was acidified, and peptides were desalted using a Sep-Pak 50 mg tC18 cartridge (Waters). Peptides were eluted in 70% acetonitrile, 1% formic acid, and dried by vacuum centrifugation.

Basic pH Reversed-Phase Separation (BPRP)

TMT-labeled peptides were solubilized in 5% acetonitrile/10 mM ammonium bicarbonate, pH 8.0, and ~ 300 µg of TMT-labeled peptides were separated by an Agilent 300 Extend C18 column (3.5 μm particles, 4.6 mm ID, and 250 mm in length). An Agilent 1260 binary pump coupled with a photodiode array (PDA) detector (ThermoFisher Scientific) was used to separate the peptides. A 45-minute linear gradient from 10% to 40% acetonitrile in 10 mM ammonium bicarbonate, pH 8.0 (flow rate of 0.6 mL/min), separated the peptide mixtures into 96 fractions (36 s). A total of 96 fractions were consolidated into 24 samples in a checkerboard fashion and vacuum-dried to completion. Each sample was desalted using StageTips and redissolved in 5% formic acid/5% acetonitrile for LC-MS3 analysis.

Liquid Chromatography Separation and Tandem Mass Spectrometry (LC-MS3)

Proteome data were collected on an Orbitrap Eclipse mass spectrometer (ThermoFisher Scientific) coupled to a Proxeon EASY-nLC 1000 LC pump (ThermoFisher Scientific). Fractionated peptides were separated using a 120 min gradient at 550 nL/min on a 35 cm column (i.d. 100 μm, Accucore, 2.6 μm, 150 Å) packed in-house. MS1 data were collected in the Orbitrap (60,000 resolution; maximum injection time 50 ms; AGC 4 × 10⁵). Charge states between 2 and 5 were required for MS2 analysis in the ion trap, and a 120 s dynamic exclusion window was used. Top 10 MS2 scans were performed in the ion trap with CID fragmentation (isolation window 0.5 Da; Turbo; NCE 35%; maximum injection time 35 ms; AGC 1 × 10⁴). Real-time search was used to trigger MS3 scans for quantification [73]. MS3 scans were collected in the Orbitrap using a resolution of 50,000, NCE of 55%, maximum injection time of 250 ms, and AGC of 1.25 × 10⁵. The close out was set at two peptides per protein per fraction (73).

Data analysis for mass spectrometry

Raw files were converted to mzXML, and monoisotopic peaks were re-assigned using Monocle [74]. Searches were performed using the Comet search algorithm against the Uniprot zebrafish (Danio rerio) database. We used a 50 ppm precursor ion tolerance, 1.0005 fragment ion tolerance, and 0.4 fragment bin offset for MS2 scans collected in the ion trap. TMT on lysine residues and peptide N-termini (+ 229.1629 Da) and carbamidomethylation of cysteine residues (+ 57.0215 Da) were set as static modifications, while oxidation of methionine residues (+ 15.9949 Da) was set as a variable modification. Each run was filtered separately to 1% False Discovery Rate (FDR) on the peptide-spectrum match (PSM) level. Then, the proteins were filtered to the target 1% FDR level across the entire combined data set. For reporter ion quantification, a 0.003 Da window around the theoretical m/z of each reporter ion was scanned, and the most intense m/z was used. Reporter ion intensities were adjusted to correct for isotopic impurities of the different TMT reagents according to manufacturer specifications. Peptides were filtered to include only those with a summed signal-to-noise (SN) ≥ 60 across all TMT channels. For each protein, the filtered peptide TMT SN values were summed to generate protein quantification values. The signal-to-noise (S/N) measurements of peptides assigned to each protein were summed (for a given protein). These values were normalized so that the sum of the signal for all proteins in each channel was equivalent, thereby accounting for equal protein loading.

Native thin filament preparation

Buffers were prepared for native thin filament (NTF) isolation as described [75]. The relaxing solution consisted of 50 mM NaCl, 5 mM MgCl₂, 2 mM EGTA, 1 mM dithiothreitol (DTT), 2.5 mM ATP, and 2.5 mg/mL phosphocreatine disodium salt hydrate, with the pH adjusted to 7.0 using phosphate buffer. The homogenization buffer contained 25 mM imidazole, 1 mM EGTA, 100 mM KCl, 5 mM MgCl₂, 5 mM ATP, 10 mM DTT, and 0.1% Triton X-100, adjusted to a pH of 6.45. The suspension buffer consisted of 25 mM imidazole, 1 mM EGTA, 100 mM KCl, 5 mM MgCl₂, and 5 mM ATP, adjusted to pH 7.9 with 0.55 mM Tris. The storage buffer consisted of a homogenization buffer without ATP.

Euthanized zebrafish larvae (5 dpf) were centrifuged at 10,000 g to remove excess water. Zebrafish (< 10 mg) were dissected in 500 µL of relaxing solution and teased apart with Dumont tweezers over the course of an hour to isolate single myofibers. Further dissection was performed in 500 µL of relaxing solution containing 0.5% Triton X-100. Subsequently, homogenization was performed in 150 µL of homogenization buffer using a 0.2 mL glass homogenizer. Samples were centrifuged for 30 min at 40,000 g, and the pellet was discarded. The supernatant was centrifuged for 45 min at 200,000 g to isolate NTFs. The resulting pellet was resuspended in 75–100 µL of suspension buffer and centrifuged for 30 min at 40,000 g to remove residual debris. The final supernatant was centrifuged again at 200,000 g for 45 min to obtain the purified NTF pellet. The NTF pellet was solubilized overnight in 40–60 µL of storage buffer.

Protein concentration was determined using the Bradford assay and samples were labeled with TRITC-phalloidin (tetramethyl rhodamine) 1 µM by overnight incubation at 4 °C. All centrifugation steps were performed at 4 °C. Dissections were conducted either on ice or in a 4 °C room. A TLA-100 rotor was used for centrifugation at 40,000 g (32,198 rpm), while centrifugation at 200,000 g was performed at 71,996 rpm.

In vitro motility assay

The in vitro motility assays were performed as previously described [76], with modifications to accommodate NTFs. In vitro motility assays were performed using actin filaments prepared from embryos pooled from three different clutches from different mating pairs for klhl41b mutant fish and WT control fish. For each genotype, three independent technical replicates were performed. Each technical replicate involved a separate assay chamber and a separate day with independently prepared myosin and corresponding motility buffers. At least fifteen filaments were analyzed per chamber to ensure robust measurement of motility parameters. Flow cells were assembled by fastening a nitrocellulose-coated coverslip to a glass slide with double-sided tape.

Chicken skeletal muscle myosin was purified as previously described and diluted to 50 µg/mL in high-salt buffer (300 mM KCl, 25 mM BES, 5 mM EGTA, 4 mM MgCl₂, and 10 mM DTT, pH 7.4) and added to the flow cell [77]. Unbound or weakly bound myosin was removed by washing the flow cells with actin buffer (55 mM KCl, 25 mM BES, 5mM EGTA, 4 mM MgCl₂, 10 mM DTT, pH 7.4), followed by the addition of 1 mg/mL bovine serum albumin (BSA) diluted in high-salt buffer to prevent nonspecific binding of thin filament proteins to the nitrocellulose membrane.

Unlabeled NTFs sourced from control or klhl41b zebrafish larvae (1 µM) were diluted in low-salt buffer (55 mM KCl, 25 mM BES, 5 mM EGTA, 4 mM MgCl₂, and 10 mM DTT, pH 7.4) and added into the myosin-coated flow cell and incubated for 2 min. A wash step was performed with 1 mM ATP to block any remaining non-functional myosin, followed by four additional low-salt buffer washes to remove excess ATP. Finally, regulated movement of labeled NTFs was initiated by adding motility buffer (low-salt buffer with 10 mM DTT, dextrose, and 0.5% methylcellulose) containing 1 mM ATP and calcium buffers at varying concentrations (pCa 4.0 to pCa 10.0). A scavenger mix containing glucose oxidase and catalase was also included to prevent photobleaching. The sliding velocity of the labeled NTFs and the percentage of motile filaments were analyzed using the ImageJ plug-in MTrackJ (https://imagescience.org/meijering/software/mtrackj/).

Sample preparation and western blotting analysis

C2C12 cells (10–15 mg) were placed in RIPA Lysis and Extraction Buffer (Thermo Fisher Scientific) with a cocktail of protease inhibitors and homogenized (2 × 15 s) using the TissueMiser homogenizer (Thermo Fisher Scientific). 30–50 zebrafish larvae were homogenized in T-per tissue extraction buffer (Thermo Fisher Scientific), centrifuged (13,000 x g, 4 °C, 15 min), and the supernatant was collected.

Samples were separated on 4–12% Bis-Tris plus mini gels (Thermo Fisher Scientific) and blotted using Trans-Blot Turbo Mini 0.2 μm Nitrocellulose (Bio-Rad). The membranes were blocked using 5% non-fat milk powder in 1X Tris Buffered Saline (Boston Bioproducts, MA) and 0.1% TWEEN 20 (TBST) for 1 h at room temperature and incubated with primary antibodies overnight at 4 °C. The membranes were subsequently washed and incubated with polyclonal anti-rabbit or anti-mouse IgG antibody conjugated to horseradish peroxidase. The antibody pairings and associated dilutions are: Anti-FLAG M2, 1: 250 (F1804, Millipore Sigma); TNNT3, 1:500 (Millipore Sigma, AV51287), TNNI2, 1:500 (Thermo Fisher Scientific, TA807853), TNNC2, 1:500 (Millipore Sigma, HPA043174), GAPDH (Cell Signaling Technology, 2118 S) and KLHL41, 1:250 (Millipore Sigma, AV38732). Secondary antibodies were anti-rabbit 1:1000 (170–6515, Bio-Rad) and anti-mouse, 1:1000 (170–6516, Bio-Rad). The quantification of protein bands was performed using ImageJ.

Whole-mount immunofluorescence

Whole-mount immunofluorescence on zebrafish was performed as previously described [59]. 1:100 Laminin (Millipore MAB1922) and 1:40 FITC-phalloidin (Thermo Fisher Scientific) were used. The secondary antibody was anti-mouse, 1:250 (A11005, Thermo Fisher Scientific).

C2C12 cell culture studies

Coimmunoprecipitation of KLHL41 and troponins was performed using the previously described method [9]. To study the association between KLHL41 and troponins, C2C12 cells (ATCC #CRL-1772) were transfected with different amounts of KLHL41-pEZYFLAG plasmid. Cells were differentiated for two days, after which cell lysates were prepared in RIPA buffer, and proteins were analyzed by Western blotting.

Coimmunoprecipitation in Mouse Skeletal Muscle Extract

Mouse skeletal muscle tissue (500 mg) was minced on ice and homogenized in ice-cold lysis buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% NP-40, 5% glycerol, supplemented with protease and phosphatase inhibitors) using a Dounce homogenizer. Lysates were pre-cleared with Protein A/G agarose beads (Thermo Fisher Scientific) for 1 h at 4 °C, then incubated with KLHL41 antibody-crosslinked Protein A/G beads overnight at 4 °C. Beads were washed five times with lysis buffer, and bound proteins were eluted with 0.1 M glycine-HCl (pH 2.5) for 10 min at room temperature, and the eluate was immediately neutralized with 1/10 volume of 1 M Tris-HCl (pH 9.0). Immunoprecipitated proteins were resolved by SDS-PAGE alongside input lysate and IgG control and analyzed by Western blot.

Electron microscopy

3 different control and 3 mutant fish from three different mating pairs were analyzed by electron microscopy, and different myofibers were analyzed from somites 7–11 (area near the end of yolk sac). Sections were analyzed from the middle area of the somite along the longitudinal axis (primarily fast muscle fibers). Zebrafish embryos and larvae were fixed in formaldehyde–glutaraldehyde–picric acid in cacodylate buffer overnight at 4 °C, followed by osmication and uranyl acetate staining. Subsequently, tissue samples were dehydrated in a series of ethanol washes and finally embedded in TAAB Epon (Marivac Ltd., Nova Scotia, Canada). Ninety-five nanometer sections were cut with a Leica ultra cut microtome, picked up on 100-mesh formvar-coated Cu grids, and stained with 0.2% lead citrate. Sections were viewed and imaged by JEOL 1200EX Transmission Electron Microscope (Electron Microscopy Core, Harvard Medical School). Images were taken from somites located in the mid-trunk region, and the same somite positions were used for all samples to maintain consistency. A total of at least n = 3 fish were examined for each genotype by EM for this analysis. Sarcomere length was measured across all samples and found to be consistent within replicates in each genotype and time point, indicating comparable contractile states at the time of fixation. The maximal width of a structure is inferred when the section passes through the central longitudinal plane rather than a tangential or oblique plane. For sarcomeres or myofibrils, this is identified by the simultaneous visualization of continuous and well-aligned structural landmarks, including distinct Z-lines, A-bands, I-bands, and consistent filament organization. Therefore, measurements were restricted to regions displaying the most complete and symmetric ultrastructural organization. Moreover, consistent measurements across multiple samples and independent sections further supported that these dimensions represent near-maximal structural width. All tissue was transferred to fixative within the same time window, minimizing pre-fixation variability. While chemical fixation cannot guarantee identical contractile states, the uniformity of sarcomere lengths across samples provides empirical evidence that the fibers were in a comparable state.

qRT-PCR

Total RNA was extracted from the control or mutant zebrafish embryos using RNeasy mini kit (Qiagen) and cDNA was synthesized using Superscript IV Reverse Transcriptase kit using random hexamers. RT-PCR was performed as previously described [59].

Data analysis

Statistical analyses were performed in GraphPad Prism 10. The biological replicate was defined according to the experimental design and is specified in the corresponding figure legends. For morphometric analyses in which multiple myofibers or sarcomeres were measured from the same animal, measurements were averaged within each animal to generate a single animal-level value, and these animal-level values were used for statistical comparisons. For datasets with small numbers of independent biological replicates and for which parametric assumptions could not be justified, differences between groups were evaluated using an exact two-tailed permutation test. For datasets in which continuous measurements were approximately normally distributed, comparisons between two groups were performed using an unpaired two-tailed Student’s t-test or Welch’s t-test, as appropriate, with Welch’s correction used when equal variances could not be assumed. Comparisons involving more than two groups were performed using one-way or two-way ANOVA followed by the multiple-comparison test specified in the corresponding figure legend. Exact statistical tests, sample sizes, and definitions of biological replicates are provided in the individual figure legends. Data are presented as mean ± SEM unless otherwise indicated. Statistical significance was defined as p < 0.05.

Supplementary Information

Supplementary Material 1.^{(27.3KB, pdf)}

Supplementary Material 2.^{(17.3KB, pdf)}

Supplementary Material 3.^{(279.7KB, pdf)}

Supplementary Material 4. ^{(203.4KB, xlsx)}

13395_2026_453_MOESM5_ESM.odt^{(8.9MB, odt)}

Supplementary Material 5: Figure S1: (A-B) qRT-PCR analysis of control, klhl41abw213g, and klhl41bbw215g zebrafish larvae at 5 dpf. n=10 fish in each group, and three biological replicates were tested. β-actin was used as the control. (C) Western blot analysis of protein extracts from control, klhl41abw213g, klhl41bbw215g, and klhl41abw213g/klhl41bbw215g zebrafish larvae at 5 dpf. n=30 fish in each group, and three biological replicates were tested. Gapdh was used as the loading control. Data are mean ± S.E.M. (unpaired t-test, parametric), **** p<0.001; * p<0.01; ns: not significant. (D) Alignment of the KLHL41 antibody epitope sequence with zebrafish Klhl41a and Klhl41b proteins, performed using ClustalW. The number 1 denotes the first amino acid position in zebrafish proteins. Figure S2: Effect size estimates for 3 pre-stimulus variables, 4 overall performance kinematic variables, and 27 additional kinematic variables classified by escape response stage. Data were obtained using a bootstrap procedure and indicate the estimated difference between genotypes (Klhl41b minus wild-type) with the 95% confidence interval. Figure S3: Klhl41b is dispensable for primary myoblast fusion. Confocal image of the whole mount fluorescence in control and klhl41b zebrafish embryos (24 hpf) labeled with phalloidin (F-actin green) and DAPI (pink) (Somite 7-11, medial somite depth). Quantification of the number of nuclei in each myofiber (n=20 embryos in each group) showed no significant differences between control and klhl41b mutants. Data are mean ± S.E.M. (unpaired t-test, parametric), ns: not significant. Figure S4: Myofibrils are initially formed near the sarcolemma during muscle development. Transmission electron microscopy of control and klhl41b mutants skeletal muscle (24 hpf) showing that nascent myofibrils (white arrows) are located at the peripheral regions of myofibers, adjacent to the sarcolemma (blue lines). Figure S5: Muscle physiology assay. (A) Force production in control and klhl41b mutants at all the frequency ranges of stimulation resulted in lower force production in the mutants. (B) Activation and (C) tension of a single twitch produced at 1 Hz frequency. Figure S6: (A) qRT-PCR analysis of control and tnnt3a zebrafish larvae at 3 dpf. n=10 fish in each group, and three biological replicates were tested. (B) Western blot analysis of protein extracts from control and tnnt3a zebrafish larvae at 3 dpf. n=30 fish in each group, and three biological replicates were tested. Gapdh was used as the loading control. Data are mean ± S.E.M. (unpaired t-test, parametric), ** p<0.01; ns: not significant.

Acknowledgements

This work was supported by the National Institutes of Health R56AR077017 (to VAG), A Foundation Building Strength grant (to VAG) and National Institutes of Health R01AR053897 (to HG). VAG is also supported by a grant from the Muscular Dystrophy Association. We thank Jonathan Van Vranken at the Thermofisher Center for Multiplexed Proteomics at Harvard Medical School for consultation on sample preparation and performing mass spectrometry. We thank Louise Trakimas (HMS Electron Microscopy Core) for her excellent help in the sample preparation and microscopy resources on the North Quad (MicRon) core at Harvard Medical School for confocal microscopy.

Authors’ contributions

V.A.G. , J.W., and J.R.M. contributed to the main text. A.M., J.W., and H.H. contributed to the Fig. 1. J.S. contributed to Figs. 2 and 3. H.H. contributed to Figs. 4 and 5. E.K and H.G. contributed to Figure S4. A. H. B. contributed to Fig. 5. G.R.T. contributed to Fig. 6. H.H. contributed to Fig. 7. V.A.G. created Fig. 8. All authors reviewed the manuscript.

Data availability

Mass Spectrometry data have been deposited to Proteomics Identifications Database and can be accessed using the provided link: https://www.ebi.ac.uk/pride/archive/projects/PXD061858. The source file of all the raw data is provided.

Materials availability

Newly created zebrafish lines and plasmids generated in this work are available on request.

Declarations

Competing Interests

All authors declare no competing interests.

Footnotes

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Material 1.^{(27.3KB, pdf)}

Supplementary Material 2.^{(17.3KB, pdf)}

Supplementary Material 3.^{(279.7KB, pdf)}

Supplementary Material 4. ^{(203.4KB, xlsx)}

13395_2026_453_MOESM5_ESM.odt^{(8.9MB, odt)}

Data Availability Statement

Newly created zebrafish lines and plasmids generated in this work are available on request.

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Klhl41b orchestrates sarcomere assembly and growth to drive skeletal muscle hypertrophy in vivo

Hyewon Han

Grace R Tate

Janet Shi

Alfredo H Bongiorno

Arian Mansur

Jeffrey Widrick

Sehajpreet Gill

Esmat Karimi

Henk Granzier

Jeffrey R Moore

Vandana A Gupta

Abstract

Background

Methods

Results

Conclusions

Supplementary Information

Introduction

Results

Klhl41b deficiency results in skeletal muscle dysfunction in zebrafish

Fig. 1.

Klhl41b is required for early myofibril assembly but is dispensable for primary myoblast fusion

Fig. 2.

Fig. 4.

Klhl41b is required for sarcomere structure and growth during secondary growth

Fig. 3.

Klhl41b deficiency leads to the accumulation of nemaline bodies and sarcomeric dysfunction

Klhl41b deficiency results in changes in the sarcomeric, translational, and proteolytic proteome in skeletal muscle

KLHL41 stabilizes TNNT3 in skeletal muscle

Fig. 5.

Klhl41b deficiency alters calcium sensitivity and cooperativity of thin filament activation

Tnnt3a is required for myofibril architecture and sarcomere organization and partially rescues muscle function in Klhl41b deficiency

Fig. 6.

Early atrophy-associated gene activation accompanies primary sarcomeric defects

Fig. 7.

Discussion

Fig. 8.

Materials and methods

Zebrafish maintenance and husbandry

Gene and protein nomenclature

Creation of zebrafish lines

Identification of founder fish and generation of stable mutant fish

tnnt3a mRNA rescue and swimming analysis

Kinematic measurements in zebrafish larvae

Sample preparation for mass spectrometry

Basic pH Reversed-Phase Separation (BPRP)

Liquid Chromatography Separation and Tandem Mass Spectrometry (LC-MS3)

Data analysis for mass spectrometry

Native thin filament preparation

In vitro motility assay

Sample preparation and western blotting analysis

Whole-mount immunofluorescence

C2C12 cell culture studies

Coimmunoprecipitation in Mouse Skeletal Muscle Extract

Electron microscopy

qRT-PCR

Data analysis

Supplementary Information

Acknowledgements

Authors’ contributions

Data availability

Materials availability

Declarations

Competing Interests

Footnotes

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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