Dynamic regulation of inter-organelle communication by ubiquitylation controls skeletal muscle development and disease onset

  1. Arian Mansur
  2. Remi Joseph
  3. Euri S Kim
  4. Pierre M Jean-Beltran
  5. Namrata D Udeshi
  6. Cadence Pearce
  7. Hanjie Jiang
  8. Reina Iwase
  9. Miroslav P Milev
  10. Hashem A Almousa
  11. Elyshia McNamara
  12. Jeffrey Widrick
  13. Claudio Perez
  14. Gianina Ravenscroft
  15. Michael Sacher
  16. Philip A Cole
  17. Steven A Carr
  18. Vandana A Gupta  Is a corresponding author
  1. Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, United States
  2. Proteomics Platform, Broad Institute of MIT and Harvard, United States
  3. Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, United States
  4. Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, United States
  5. Department of Biology, Concordia University of Edmonton, Canada
  6. Faculty of Health and Medical Sciences, Centre of Medical Research, Harry Perkins Institute of Medical Research, University of Western Australia, Australia
  7. Division of Genetics, Boston Children’s Hospital, Harvard Medical School, United States
  8. Department of Anesthesiology, Brigham and Women’s Hospital, Harvard Medical School, United States
  9. Department of Anatomy and Cell Biology, McGill University, Canada

Abstract

Ubiquitin-proteasome system (UPS) dysfunction is associated with the pathology of a wide range of human diseases, including myopathies and muscular atrophy. However, the mechanistic understanding of specific components of the regulation of protein turnover during development and disease progression in skeletal muscle is unclear. Mutations in KLHL40, an E3 ubiquitin ligase cullin3 (CUL3) substrate-specific adapter protein, result in severe congenital nemaline myopathy, but the events that initiate the pathology and the mechanism through which it becomes pervasive remain poorly understood. To characterize the KLHL40-regulated ubiquitin-modified proteome during skeletal muscle development and disease onset, we used global, quantitative mass spectrometry-based ubiquitylome and global proteome analyses of klhl40a mutant zebrafish during disease progression. Global proteomics during skeletal muscle development revealed extensive remodeling of functional modules linked with sarcomere formation, energy, biosynthetic metabolic processes, and vesicle trafficking. Combined analysis of klh40 mutant muscle proteome and ubiquitylome identified thin filament proteins, metabolic enzymes, and ER-Golgi vesicle trafficking pathway proteins regulated by ubiquitylation during muscle development. Our studies identified a role for KLHL40 as a regulator of ER-Golgi anterograde trafficking through ubiquitin-mediated protein degradation of secretion-associated Ras-related GTPase1a (Sar1a). In KLHL40-deficient muscle, defects in ER exit site vesicle formation and downstream transport of extracellular cargo proteins result in structural and functional abnormalities. Our work reveals that the muscle proteome is dynamically fine-tuned by ubiquitylation to regulate skeletal muscle development and uncovers new disease mechanisms for therapeutic development in patients.

Editor's evaluation

This important study utilizes a model organism, zebrafish, to explore the roles of KLHL40, a component of the ubiquitin-proteasome system (UPS), in the development of skeletal muscle disease. Monitoring changes in transcriptome, proteome and ubiquitylome, the study finds a selective role for proteome remodeling in muscle development and monitors how KLHL40-deficiency leads to disease onset. A specific role for CUL3-KLHL40 in regulating the expression of Sar1a, a key component of biosynthetic secretion is described where abnormal Sar1a levels culminate in procollagen secretion defects. The compelling data on proteome remodeling and UPS-regulation of biosynthetic secretion make this work interesting to biologists who study the UPS, muscle development and intracellular traffic.

https://doi.org/10.7554/eLife.81966.sa0

Introduction

Fetal akinesia, arthrogryposis, and severe congenital myopathies are heterogeneous conditions of reduced fetal movement, usually presenting at birth (Beecroft et al., 2018; Langston and Chu, 2020). More than 50% of all causes of fetal akinesia are of neuromuscular origin, involving all points along the neuromuscular axis (motor neurons, peripheral nerves, neuromuscular junction, and the skeletal muscle regulatory and contractile apparatus; Ravenscroft et al., 2013; Oates et al., 2013; Vogt et al., 2008; Nowak et al., 1999; Pelin et al., 1999). These diseases exhibit a high clinical heterogeneity with a severe congenital onset with fetal akinesia to milder forms, often with a late childhood or adult-onset. At least 30 causative genes have been identified in these conditions (Ravenscroft et al., 2018; Ravenscroft et al., 2021). However, the origin and temporal ordering of molecular events that drive the disease pathology remains poorly understood.

Skeletal muscle is made up of myofibers highly specialized for contraction. To achieve this function, each myofiber contains myofibrils, which consist of a repetition of sarcomeres. After myoblast fusion, sarcomeres are assembled through the interaction of protein complexes that form complex supramolecular structures to form functional myofibers. This requires precisely controlled dynamic turn-over of proteins without perturbing the structure of assembling sarcomeres. The ubiquitin-proteasome system (UPS) regulates the relative abundance and functional modifications of proteins during multiple stages of myogenesis (Hnia et al., 2019; Piccirillo et al., 2014; Jirka et al., 2019). The UPS is a critical process that controls protein degradation and plays a key role in protein homeostasis. RING E3 ligases play key roles through the recognition of specific protein substrates and the transfer of ubiquitin to the substrate. Mutations in KLHL40, a CUL3 family E3 substrate adaptor protein, have been reported to result in server congenital nemaline myopathy (NM). The importance of UPS in skeletal muscle development has also been identified in human diseases where mutations in genes regulating ubiquitination and protein turnover processes result in sarcomeric disarray and functional deficits (Ravenscroft et al., 2013; Gupta et al., 2013; Frosk et al., 2002; Olivé et al., 2015).

Sarcomeres are present in close proximity to the triad system that is formed of T-tubules and sarcoplasmic reticulum, a modified endoplasmic compartment. In addition, different mitochondrial populations are also present in close juxtaposition to the sarcomere (Henderson et al., 2017). Mutations or deletions of sarcomeric genes affect the structure and function of surrounding organelles, and similarly, defects in other organelles in myofibers also affect sarcomere structure and function (Reimann et al., 2003; Voit et al., 2017; Fatkin et al., 2000; De Gasperi et al., 2022). This suggests that the sarcomere and surrounding organelles act as interconnected hubs that engage in extensive communication during skeletal muscle development and maintenance. Despite this, mechanistic insight into inter-organelle communication between different membrane compartments in protein trafficking and regulation of this process in skeletal muscle development remains largely unknown. Finally, how this communication is perturbed in disease states and contributes to disease pathology is not clear.

We have identified that inter-organelle communication is critical for vesicle trafficking and skeletal muscle development by ubiquitination signaling in the sarcomere. We performed global proteomic and ubiquitylome profiling of skeletal muscle during development and disease progression in a Klhl40 deficient zebrafish model of congenital nemaline myopathy. We identified that KLHL40 acts as a regulator of membrane vesicle trafficking through ubiquitylation and subsequent protein degradation of Secretion-associated Ras-related GTPase1a (SAR1A). In the absence of this negative feedback mechanism in KLHL40 deficiency, SAR1A is abnormally localized to the ER and contributes to membrane tubulation defects and disruption of the trafficking of collagen. Our work demonstrates that inter-organelle communication between sarcomeric and endomembrane compartments is dynamically regulated by ubiquitylation and is critical for skeletal muscle development, and defects in this process underlie pathology in skeletal muscle diseases.

Results

KLHL40 is required for skeletal muscle development

KLHL40 deficiency in humans results in a severe form of nemaline myopathy associated with neonatal lethality (Ravenscroft et al., 2013). Deleting Klhl40 in mice also results in an extensive structural damage in myofibers and neonatal lethality 2 to 3 weeks after birth (Garg et al., 2014). As zebrafish grow ex vivo, skeletal muscle development and disease progression can be visualized in the context of a living organism. We generated loss-of-function klhl40 alleles in zebrafish using the CRISPR/Cas9 gene-editing tool (Figure 1A–B). The human orthologue of the KLHL40 gene is duplicated in zebrafish as klhl40a and klhl40b and mapped on chromosome 2 and chromosome 24, respectively. klhl40a alleles created include klhl40abwg200 with insertion of one base (c.250_251insA; p.Val84Aspfs*36) and klhl40abwg201 with a two base pair deletion in exon 1 (c.251_252insTC;p.Val84Asnfs*36). These alleles result in frameshift mutations and are predicted to result in truncations in the N terminal BTB domain of the Klhl40a protein. For klhl40b, a CRISPR-edited allele (klhl40bbwg202) had insertion of one base (c.674_675insC; p.Arg225Profs*14) in exon 1 (Figure 1B). klhl40bbwg202 allele is predicted to result in an frameshift mutation and truncation of the Klhl40b protein in the BACK domain (Figure 1A). As klhl40abwg200 and klhl40bbwg202 alleles were predicted to produce the smallest truncated Klhl40 proteins, the rest of the analyses presented in this work were performed on these fish lines obtained after F3 generation and referred to as klhl40a and klhl40b in the rest of this work. The effect of different mutations on klhl40 mRNA levels was evaluated by RT-PCR (Figure 1—figure supplement 1). Both klhl40a and klhl40b alleles exhibited similar klhl40a and klhl40b mRNA levels, respectively, in comparison to +/+ siblings. As truncated mutant proteins could result in a dominant gain of function, a western blot was performed on the skeletal muscle extracts obtained from klhl40a and klhl40b mutant fish to evaluate the effect of these mutations on the klhl40 protein. Western blot analysis with a KLHL40 antibody that recognizes both Klhl40a and Klhl40b proteins revealed a 50% decrease in Klhl40 protein levels in both alleles compared to control and complete absence of Klhl40 protein in klhl40a/klhl40b double knockout fish (Figure 1—figure supplement 1). This suggests that klhl40a and klhl40b alleles result in the loss of Klhl40 protein in the mutant zebrafish. klhl40a and klhl40b mutant embryonic (2dpf) and larval fish (5dpf) did not exhibit gross morphological defects in any of the mutants examined. Previous studies have shown that the knockdown of klhl40 by morpholino results in myopathy in zebrafish embryos (Ravenscroft et al., 2013). The discrepancy between the morphants and mutants could be due to genetic compensatory mechanisms by other Kelch protein-coding genes or other modifier genes as described by several studies (El-Brolosy et al., 2019; Sztal et al., 2018). During the developmental transition from juvenile (1.5 months) to adult stage (3.0 months), the klhl40a mutants developed a myopathic phenotype with leaner bodies, whereas klhl40b were phenotypically indistinguishable from +/+ control siblings at this age (Figure 1C). No other obvious morphological defects were observed. klhl40a/klhl40b double mutants appeared phenotypically similar to klhl40a fish. klhl40a and klhl40b exhibit overlapping expression in skeletal muscle, and a lack of phenotype in klhl40b mutants suggested functional redundancy similar to a large number of duplicated genes (Ravenscroft et al., 2013; Taylor and Raes, 2004). To identify any defects in skeletal muscle function, the swimming performance of klhl40a mutants and +/+ siblings were analyzed by the flume tunnel assay to obtain maximum swimming speed (Umax) (Figure 1D; Widrick et al., 2018). No differences in the Umax values were observed between control and klhl40a mutants at the juvenile stage (1.5 months). The Umax values showed a significant decrease in klhl40a mutants compared to +/+ control siblings at the adult stage (3 months), indicating reduced endurance capacity of the klhl40a deficient fish. As klhl40a mutant fish at 1.5 months are phenotypically and functionally similar to control fish, this age group was termed ‘pre-symptomatic stage’, whereas klhl40a mutant fish at 3.0 months was termed as ‘symptomatic stage’ due to myopathic features. Control fish survived to 24 months of age, whereas most of the klhl40a mutant fish died between 9 and 12 months. These data show that loss of klhl40a leads to a myopathic phenotype, as observed in patients with KLHL40 variants (Figure 1E).

Figure 1 with 1 supplement see all
klhl40 is essential for vertebrate skeletal muscle development.

(A) Schematic diagram depicting the position of CRISPR-mediated mutant alleles and truncated proteins on the Kelch protein domain in klhl40abwg200 and klhl40bbwg202 knockout zebrafish. CRISPR-induced mutations in klhl40abwg200 and klhl40bbwg202 knockout zebrafish result in premature termination codons in BTB and BACK domain coding exons, respectively. (B) Sanger sequencing pherograms for control and klhl40a bwg200 and klhl40b bwg202 mutant zebrafish with an insertion of A in klhl40abwg200 and an insertion of C in klhl40bbwg202 coding regions. (C) Lateral view of the juvenile and adult zebrafish. klhl40a mutant zebrafish develop leaner bodies from transition to juvenile (1.5 months old) to the onset of the adult stage (3 months old) and exhibit reduced body length and body diameter. No obvious skeletal muscle phenotype is observed in the klhl40b allele compared to control (+/+) siblings. klhl40a/ klhl40b double mutant fish exhibit similar skeletal phenotype as observed in the klhl40a allele. (D) Endurance swimming behavior of klhl40a allele at juvenile state (1.5 months) and adult stage (3 months) (n=7–8). (E) The Kaplan-Meier survival curve of the different zebrafish groups was analyzed for 20 months (n=20 in each group). Data are mean ± S.E.M (unpaired t-test, parametric) for each experiment. Note: the survival curve of klhl40b mutant fish overlaps with the control fish.

KLHL40 plays pleiotropic roles in regulating skeletal muscle structure

KLHL40 deficient muscles in patients exhibit extensive myofiber damage and extensive sarcomeric disarray in many myofibers (Ravenscroft et al., 2013). As patient muscle biopsies are typically collected after the disease diagnosis, disease processes are usually already established. This prevents an understanding of pathological changes resulting in extensive muscle damage. Therefore, to understand how Klhl40a deficiency affects skeletal muscle structure during disease onset and progression, ultra-structure was evaluated by transmission electron microscopy (TEM) in both juvenile (pre-symptomatic) and adult stages (symptomatic) in control and klhl40a mutant fish (Figure 2 and Figure 2—figure supplement 1). No significant ultrastructural changes in the sarcomere or SR-ER region were observed during the juvenile stage in the klhl40a skeletal muscle (Figure 2—figure supplement 1). Both sarcomere width (w) and height (h) were significantly reduced in the klhl40a in comparison to +/+controls at the adult stage (Figure 2A–B). SR-ER vesicles in the mutant skeletal muscle were dilated compared to the control muscle and displayed an accumulation of membrane-bound structures (10–100 nm) in close proximity (Figure 2A–B). Mitochondria in the skeletal muscle of the klhl40a mutant muscle were rounder and displayed an electron dense-matrix compared to the controls (Figure 2C–D). Such abnormal mitochondrial are also seen in other muscle diseases such as Duchenne muscular dystrophy and polymyositis and are secondary consequences of muscle damage. Mutant muscle also exhibited abnormalities in the extracellular matrix (ECM) structure (Figure 2E–F). Compared to the controls, mutant muscle showed large gaps in the adjacent myofibers in the ECM region. These data indicate that Klhl40a is required to regulate sarcomere size, intracellular membrane homeostasis, and ECM stability in skeletal muscle.

Figure 2 with 1 supplement see all
klhl40a allele displays reduced sarcomere size and abnormal membrane organelles in skeletal muscle.

Transmission electron microscopy (TEM) showing the ultrastructure of control (+/+) and klhl40a KO in 3 months animals. (A–B) Longitudinal muscle section of control and klhl40a KO mutant muscle showing accumulation of membranous structures in SR-ER region (arrow) and reduced sarcomere width (w) and height (h). (C–D) Cross-section view showing mitochondrial in klhl40a KO mutant muscle contain electron-dense matrix (arrow) compared to control muscle (normal mitochondria). (E–F) The longitudinal view of skeletal muscle shows structural damage to the extracellular matrix (arrow) in the klhl40a KO mutant compared to the control. Electron microscopy was performed in three different control and klhl40a KO mutant fish. N=150–175 sarcomeres analyzed in each sample for quantification. N=75–100 mitochondria and 200–250 triads analyzed in each sample for quantification of the ER. Data are mean ± S.E.M; with one-way analysis of variance (ANOVA) and Tukey’s HSD test (****p<0.001).

Dynamic remodeling of the proteome during skeletal muscle development and disease progression in nemaline myopathy

KLHL40 is a substrate-specific adaptor of the E3 ubiquitin ligase CUL3, and the KLHL40-CUL3 ubiquitin ligase complex has previously been shown to stabilize the sarcomeric thin filament proteins such as leimodin3 (LMOD3) and nebulin (NEB) by ubiquitylation through in vitro studies (Garg et al., 2014), however, the in vivo relevance remains unknown. Protein complexes are changed dynamically during development to meet the constantly changing demands of differentiating cells. Subtle protein changes may significantly affect downstream processes, which can be hard to identify and require highly quantitative in vivo approaches. Identifying low-abundance proteins with critical roles may be difficult. These issues become particularly significant in human diseases as disease processes are mainly investigated during the pathological states when atrophic processes are prevalent. Still, our understanding can benefit by analyzing disease trajectories from a pre-symptomatic state to clinically symptomatic states.

To comprehensively quantify proteome remodeling during skeletal muscle growth, disease onset, and progression, global proteomic changes in skeletal muscle from control (+/+) and klhl40a zebrafish at pre-symptomatic (1.5 months) and symptomatic (3 months) stages of disease progression were analyzed. Deep-scale quantitative liquid chromatography-tandem mass spectrometry (LC-MS/MS) based proteomics was performed in skeletal muscle in these fish (Figure 3) (Supplementary file 1). Muscle samples from wild-type (WT) and klhl40a mutant (KO) were collected and analyzed in quadruplicate using tandem mass tags (TMT) for multiplexing and quantification (Figure 3A; Mertins et al., 2018). In parallel, we performed deep ubiquitylation profiling via enrichment of the lysine di-glycine remnant (KGG) from ubiquitin trypsinization from the exact same tissues (Udeshi et al., 2020). This allows the determination of the contribution of ubiquitylation in remodeling skeletal muscle proteome during muscle development and disease onset by the CUL3 E3 ubiquitin ligase-KLHL40 complex (Supplementary file 2; Figure 3A).

Figure 3 with 2 supplements see all
Proteome and ubiquitylome disruption by Klhl40a deficiency.

(A) Experimental workflow for proteome and ubiquitylome quantification in klhl40a allele. (B) Heatmap showing protein abundances (log2 TMT ratios) across experimental samples. Only proteins with a significant differential response between control and klhl40a KO samples are shown (adjusted P-value <0.05). Proteins (rows) were clustered to show abundance patterns across experimental groups. (C) Heatmap showing ubiquitin sites following trypsin digestion (KGG)-site abundances (log2 TMT ratios) across experimental samples. Only proteins with a significant differential response between control and klhl40a KO samples are shown (adjusted P-value <0.05). KGG sites (rows) were clustered to show abundance patterns across experimental groups. (D) Correlation of protein response to klhl40a KO across juvenile (1.5 months) and adult animals (3 months). Plots show log2 fold changes for proteins quantified at both ages. Proteins are colored if they show differential abundance (adjusted p-value <0.05) at one age only (yellow), both ages with the same direction (green), and both ages with opposite directions (purple). (E) Correlation of ubiquitylome response to klhl40a KO across juvenile (1.5 months) and adult animals (3 months). Plots show log2 fold-changes for KGG-sites quantified at both ages. KGG sites are colored if they show differential abundance (adjusted p-value <0.05) at one age only (yellow), both ages with the same direction (green), and both ages with opposite directions (purple).

A total of 8,268 proteins were quantified across these 16 samples, and PCA showed the grouping of the different replicates from each age and experimental genotype group (Figure 3—figure supplement 1). To investigate changes in the proteome that are primarily regulated through the transcriptome during the disease state, we integrated our proteomic data with RNA sequencing (RNA-seq) results obtained on the same tissue samples (3 months; Supplementary file 3). The proteome-transcriptome correlation analyses revealed a high degree of discordance (78%) between transcript-protein pairs (Figure 3—figure supplement 2). This reflects extensive post-translational regulation of skeletal muscle development in vivo, therefore, we focused on the proteome dataset. Proteins with a significant differential response between control and mutant samples at either stage (moderated t-test, adjusted p-value <0.05) were clustered using hierarchical clustering to reveal proteome abundance patterns across experimental groups (Figure 3B). This revealed four distinct clusters defining critical trajectories of changes in the proteome during normal and disease states. Cluster 1 represented proteins that exhibited low abundance in the control muscle and high abundance in the mutant muscle at both the pre-symptomatic and symptomatic stages. Clusters 2 and 4 represented proteins with high levels in the juvenile stage but a significant reduction at the adult stage in the normal muscle and represented proteins in lipid catabolic process (e.g. Pck1, Pck2), vesicle trafficking (e.g. Sec16b, Srp14, Timm10), and the UDP-N-acetylglucosamine biosynthetic process indicating the involvement of vesicular trafficking pathway. Interestingly, these clusters in the mutant fish showed reduction at both juvenile (pre-symptomatic) and adult (symptomatic) stages. Finally, cluster 3 exhibited protein levels that were elevated in control muscle at both juvenile and adult stages but reduced in mutant muscle at both stages (Figure 3D). These data show extensive and dynamic remodeling of the cellular proteome during normal skeletal muscle development. Most differential proteomic changes in the Klhl40a mutant muscle emerge during the juvenile (pre-symptomatic) state. These changes are primarily static during disease onset and progression. This indicates that gene expression is subject to complex post-translational regulation in vivo, resulting in dynamic remodeling in normal skeletal muscle development.

Changes in proteome reveal delayed sarcomere maturation in Klhl40a deficiency

To investigate the biological pathway associated with Klhl40a deficiency, pathway enrichment analysis was performed on proteins significantly increasing or decreasing in the mutant at each stage (FDR p<0.05; Figure 4A and Figure 4—figure supplement 1). The skeletal muscle developmental process exhibited enrichment with increased abundance at the pre-symptomatic state in the klhl40a mutant muscle compared to controls. Many of these proteins were expressed in differentiating myotubes during early sarcomere assembly. They were either absent or exhibited low levels in the terminally differentiated mature skeletal muscle (Obscn, Nexn, Ilk, Vcl, Fxr1, Synpo2l, Tnnt2, Flnc, Pdlim5, Smyd1, Unc45, Mybpc1, Lamb2, Cav1, Alpk3, Pleca, and TnnT1) (Supplementary file 4). Hierarchical clustering also revealed that sarcomeric proteins associated with mature myofibers exhibited significantly less abundance in mutant muscle at both stages. Pathway enrichment showed that proteins in this cluster include proteins of the actin cytoskeleton (e.g. Tmod, Tnnt3, Tnni2, Tpma, Rock1), sarcomere assembly (e.g. Myom, Actn3, Capz) intermediate filaments (e.g. Plec, Dsp, Mtm1) and microtubule transport (e.g. KIif5b, Mfn2, Dync2h1) (Figure 4D, blue and cyan nodes). These proteins are required for the formation and maintenance of mature sarcomeres. This suggests that Klhl40-deficient skeletal muscle exhibit a defect in the terminal differentiation of skeletal muscle with increased levels of early sarcomeric and reduced abundance of late sarcomeric proteins.

Figure 4 with 1 supplement see all
Pathways regulated by changes in proteome and ubiquitylome mediated by Klhl40a.

(A) Network visualization of pathway enrichment results from klhl40a KO differential proteins compared to controls. Nodes (circles) indicate pathways significantly enriched in proteins that increase at 1.5 months, decrease at 1.5 months, increase at 3 months, or decrease at 3 months. Edges (connections) show nodes with overlapping genes. Clusters of nodes summarize pathways with similar biological functions. (B) Network visualization of pathway enrichment results from klhl40a KO differential KGG-sites compared to control. Nodes (circles) indicate pathways significantly enriched in KGG sites that increase at 1.5 months, decrease at 1.5 months, increase at 3 months, or decrease at 3 months. Edges (connections) indicate nodes with overlapping genes. Clusters of nodes summarize pathways with similar biological functions. (C) Fold-changes of KGG-sites and their cognate protein in response to klhl40a KO in 1.5 months animals compared to controls. Data points are colored if both the KGG-site and the cognate protein show differential abundance in KO vs. (+/+) control and have the same direction (green) or opposite directions (purple). (D) Fold-changes of KGG-sites and their cognate protein in response to klhl40a KO in 3 months animals compared to controls. Data points are colored if both the KGG-site and the cognate protein show differential abundance in KO vs. (+/+) control and have the same direction (green) or opposite directions (purple).

Bioenergetic and biosynthetic metabolic changes in proteomics precede structural changes in skeletal muscle in Klhl40 deficiency

The proteomics analysis additionally revealed significant changes in the metabolic processes in the absence of Klhl40a. Glucose uptake (hexokinase) and glycolytic pathway enzymes (Pygma, Pygmb, Eno3, Pfkm, Aldoa, Aldob, Aldoc, Pgk1, Pgam1, Pgam2, and Pkmb) (Supplementary file 4) were increased in the mutant muscle during the juvenile stage compared to controls (pre-symptomatic stage). Categories associated with cellular metabolism also showed amino acid and lipid metabolism enrichment, mitochondrial respiration, and nucleotide metabolism in mutant muscle at both stages (Figure 4A red and yellow nodes, Supplementary file 4). This suggests that pathways regulating bioenergetics balance in skeletal muscle are altered in the mutant muscle (Supplementary files 1 and 4). Glycolysis and mitochondrial respiration (oxidative phosphorylation, OXPHOS) are the primary regulators of cellular bioenergetics during development. While this metabolic shift of increased glycolytic and biosynthetic proteins is reminiscent of the Warburg effect (Oginuma et al., 2020; Tarazona and Pourquié, 2020; Tixier et al., 2013), a concurrent abundance of mitochondrial respiration enzymes indicates stress or disease-induced changes in the metabolic processes. Sarcomere remodeling in stress or disease states is associated with altered metabolic states by increasing glycolytic and mitochondrial proteins (Toepfer et al., 2020; Liu et al., 2022). Together, these studies link changes in proteome in Klhl40a deficiency to sarcomere structure and function, increased mitochondrial content, and altered metabolic state of the mutant muscle.

Quantitative KEGG proteome regulation in skeletal muscle is required for vesicle trafficking, glycolysis, and sarcomeric proteins

Deep ubiquitylome profiling illuminated changes in ubiquitylation dynamics during skeletal muscle development and disease onset in Klhl40 deficiency. Similar to the dynamics of changes observed in the proteome, changes in the ubiquitylome of mutant muscle were established before functional and structural changes were observed in skeletal muscle (Supplementary file 2). A heat map of Hierarchical cluster analysis of ubiquitin sites (Ub-sites) with differential abundance between wild-type and mutants in the ubiquitylome data showed four different clusters classified into two broad categories (Figure 3C and Supplementary file 5). Clusters 1 and 2 exhibited proteins with decreased Ub-sites in both pre-symptomatic and symptomatic mutant states. We expect many of these proteins to be direct targets of ubiquitylation by the Klhl40-Cul3 complex. Previous studies have shown nebulin is a direct ubiquitination target of Klhl40-Cul3 complex and was identified in cluster 1, validating our hypothesis (Garg et al., 2014). Pathway enrichment analysis showed that the most significantly downregulated Ub-sites nodes in the mutant muscle were the peroxisome and sarcomere proteins (Figure 4B, blue and cyan nodes; e.g., Ttn, Myha, Myhb, Tpma, Myhc4, Myom1). Most proteins exhibited changes in Ub-sites that correlated positively at the pre-symptomatic and symptomatic stages. (Figure 3C and E and Supplementary files 1–6). We did not identify any protein showing significant enrichment of Ub-sites in different directions (Figure 3E). This suggests that ubiquitylation marks by the Klhl40-Cul3 complex and potentially other ubiquitylation enzymes are robust and unidirectional in the disease state. Clusters 3 and 4 represented proteins that exhibited increased Ub-sites in the klhl40a mutant muscle compared to the control. As many ubiquitin ligases and deubiquitylases showed differential expression between control and mutant muscle, these proteins could be direct targets of many of these enzymes. Nodes that exhibited upregulated ubiquitylated peptides included proteins in muscle development and muscle fibers formation (Obscn, Tnni1, Tmod4, Tpm2, Myom2, Tnni2, Cfl2, Ldb3, Des) and ubiquitin-mediated proteolysis (Figure 4B, red and yellow nodes). Integration with the proteome data revealed that these highly ubiquitylated proteins had decreased abundance in mutant muscle (Tmod4, Tnni2, Tpm2, Cfl2, Myom2). Many of these highly ubiquitylated proteins are localized to thin filaments and contribute to nemaline myopathy, indicating other components of the ubiquitin proteasomal pathways may cause increased ubiquitylation and abnormal degradation of sarcomere proteins in mutant muscle and affect thin filaments stability (Figure 4C–D, Supplementary files 8-9). Finally, to identify potential targets of the Klhl40-Cul3 complex, analysis of fold changes of reduced Ub-sites and abundance of their cognate proteins in response to Klhl40a deficiency in opposite directions identified Sar1a (vesicle trafficking protein), glycolytic proteins (Pkmb, Aldo, Aldob) and sarcomeric proteins (Ttn, Tnnt2, Nckipsd). While the proteomic analysis indicated altered sarcomeric and glycolytic proteins might contribute to disease pathology in Klhl40a deficiency, reduced ubiquitylation of these proteins, suggest these may be directly regulated through ubiquitylation by the Klhl40-Cul3 complex.

Sar1a upregulation is associated with increased accumulation of ER-derived membrane-bound structures

The vesicle trafficking pathway is central in cells for transporting cargo and secretory proteins. However, the role of this process in normal and disease muscle is not completely clear. SAR1A is a small GTPase required to assemble COPII vesicles at endoplasmic reticulum exit sites (ERES) by recruiting the SEC23/24 complex for the protein trafficking (Lee et al., 2005). SAR1 is also involved in large cargo trafficking through large COPII structures that require TANGO1, cTAGE5, SEC23/24, and SEC13/31. While Sar1a protein levels were significantly increased, inner coat proteins of COPII vesicles Sec23b and Sec23d and Golga2 were reduced in the mutant muscle suggesting that they are co-regulated in Klhl40a deficiency. No changes were observed in the outer coat proteins of COPII vesicles (Sec13 and 31; Figure 5A). Proteomics analysis also showed no significant changes in the ER resident protein Sec12 or other COPII proteins, Tango1, cTAGE5, and Sec13/31 levels in Klhl40a deficiency. These changes to the vesicular trafficking proteins were regulated post-transcriptionally, as no differences were seen in these proteins at the mRNA level (Supplementary file 3). To investigate the possible role of Sar1a upregulation on the disease pathology in Klhl40a deficiency in skeletal muscle, western blot analysis was performed to validate the findings of the proteomics data. Western blot analysis in control and mutant klhl40a mutant skeletal muscle protein extracts at the adult stage (3 months) confirmed increased Sar1a protein levels in the mutant skeletal muscle compared to the WT control (Figure 5B). Moreover, Western blot also validated reduced Sec24d and Golga2 in Klhl40a deficient skeletal muscle. (Supplementary file 1, Figure 5B–C). Sar1a is expressed at low levels in normal skeletal muscle. However, the physiological roles and effects of Sar1a perturbations in skeletal muscle are unknown. To understand the implications of Sar1a upregulation on skeletal muscle pathology, human SAR1A mRNA was overexpressed in wild-type zebrafish. Ultrastructural examination of zebrafish larvae by electron microscopy showed that SAR1A mRNA overexpression (50–100 ng) resulted in abnormal membrane-bound structures in the SR-ER region similar to klhl40a mutant fish (Figure 5D). Immunofluorescence analysis of myofibers from control and SAR1A overexpressing zebrafish revealed SAR1A-positive punctate structures on SAR1A overexpression co-stained with the ER marker, protein disulphide isomerase (PDI) (Figure 5E). This suggests that increased SAR1A protein levels in skeletal muscle result in the abnormal accumulation of ER-derived membrane-bound structures.

Klhl40a loss results in perturbation of the ER-Golgi vesicle trafficking through secretion-associated Ras GTPase (Sar1a).

(A) ER-Golgi vesicle trafficking proteins exhibit altered levels in klhl40a mutant muscle compared to control (+/+) in proteome analysis; ns indicates no significant difference (B) Western blot showing ER-exit site protein Sar1a is upregulated in klhl40a mutant muscle, and downstream COPII and Golgi proteins are downregulated in mutant muscle (3mo) (C) Quantification of the protein by Western blot in klhl40a and control zebrafish. N=3 in each group. Data are mean ± S.E.M; with one-way ANOVA and Tukey’s HSD test (****p<0.001). (D) Transmission electron microscopy (TEM) of zebrafish larva (4 dpf) with SAR1A mRNA overexpression demonstrating abnormal membrane structures in the SR-ER region. (E) Immunofluorescence of control and SAR1A overexpressing myofibers (5 dpf); n=25–30 myofibers in each group.

Vesicle trafficking components are perturbed in Klhl40a deficiency

Klhl40a deficiency resulted in altered levels of several vesicle trafficking proteins. Therefore, we examined the morphology of the components of the protein trafficking process in myofibers from klhl40a and control zebrafish by immunofluorescence (3 months). Klhl40a deficient myofibers exhibited increased accumulation of Sar1a in the ER compared to controls (Figure 6). Moreover, an increased number of Sar1a and PDI-positive foci were observed in mutant muscle. The ultrastructure of abnormal membranous structures observed in the klhl40a mutant muscle was similar to ER and lacked the organization of the COPII-coated vesicles shown previously (Figure 2B; Barlowe et al., 1994; Matsuoka et al., 1998). This suggests that abnormal membranous structures in the ER-SR region in the mutant muscle originated from the ER. Immunofluorescence with Tango1 antibody revealed reduced ER exit sites (ERES) in Klhl40-deficient myofibers. Although the size of ERES sites showed variability in both control and mutant myofibers (50–500 nm), mutant myofibers showed an increased number of Tango1-positive enlarged ERES sites (~200–500 nm) (5.2 ± 1.91 %) compared to control myofibers (0.70 ± 0.39%). Examination of COPII vesicle protein Sec23B showed fewer COPII vesicles in Klhl40a deficiencyFinally, examination of the Golgi apparatus in Klha40a showed normal immunoreactivity in most of the myofibers (63 ± 14%) with varying amounts of GOLGA2-positive aggregates in other myofibers. These morphological changes are not overserved during the pre-symptomatic stage (Figure 2SB). As dysregulation of autophagy is associated with vesicle trafficking, we examined different autophagy markers (Atd5, Atg16l1, Atg4b, Atg9a, beclin, Lc3b, Lamp1, and Lamp2) in the proteomic data or by western blot (LC3) (Figure 6—figure supplement 1). We did not observe any altered autophagy markers in Klhl40a deficiency. ER stress can also trigger unfolded protein response (UPR) to restore ER proteostasis. No differences were observed in Bip, Calnexin, Ero1, Ireα1, CHOP, and PERK between control and klhl40a skeletal muscle (1.5 and 3 months) by proteome or RNA-seq analysis (3 months). xbp1 mRNA spliced during UPR in the ER also showed no change in the klhl40a mutant muscle (Figure 6—figure supplement 1). Western blot analysis of PERK showed no differences in the total PERK levels or phospho-PERK levels in Klhl40a deficiency compared to the control muscle indicating a lack of UPR activation in Klhl40a deficiency (Figure 6—figure supplement 1). These studies provide evidence that abnormally increased amounts of Sar1a do not result in the formation of productive COPII vesicles and contribute to abnormal vesicle formation associated with changes in crucial protein regulators of ER-Golgi vesicle trafficking.

Figure 6 with 1 supplement see all
Morphological changes in vesicular trafficking compartments in Klhl40a deficient myofibers.

Sar1a is increased and co-localized with PDI in Klhl40a deficiency. The number of PDI-positive foci is also increased in the absence of Klhl40a. The fraction of Sec23 and Tango1 positive foci is decreased in klhl40a mutant myofibers. Disruption of the Golgi architecture was observed in a fraction of Klhl40a deficient myofibers. Data are mean ± S.E.M (unpaired t-test, parametric) for each quantification.

KLHL40-CUL3 regulates SAR1A levels through ubiquitylation

KLHL40 is a substrate-specific adaptor protein for the E3 ubiquitin ligase CUL3 that targets particular protein substrates for ubiquitination, affecting the target protein stability. To test if SAR1A is a direct substrate of KLHL40, we performed co-immunoprecipitation assays in C2C12 cells that showed SAR1A is a direct interactor of KLHL40 (Figure 7A). As SAR1A protein increases in KLHL40 deficiency, we evaluated if there is a direct reciprocal interaction between KLHL40 and SAR1A proteins by overexpression assays in C2C12 cells. A gradual decrease in KLHL40 protein levels resulted in a concomitant increase in SAR1A protein levels in muscle cells (Figure 7B). There was no evidence of promoter competition with KLHL40 and SAR1A plasmids (Figure 7—figure supplement 1). To understand if the interaction between KLHL40 and SAR1A results in reduced levels of SAR1A through ubiquitylation-mediated protein degradation, we evaluated the effect of KLHL40 protein on SAR1A stability in the presence of the proteasome inhibitor MG132 (Figure 7C). In the presence of MG132, increased stability of SAR1A was observed, suggesting that KLHL40 targets SAR1A for degradation through proteasomes. Analysis of vertebrate SAR1A protein sequences revealed that the SAR1A ubiquitylation site identified by ubiquitylome analysis is highly conserved in vertebrates suggesting that SAR1A ubiquitylation in skeletal muscle may be conserved in all vertebrates (Figure 7D). Finally, to examine whether the KLHL40-CUL3 complex can directly promote SAR1A ubiquitination, an in vitro ubiquitination assay was performed with neddylated CUL3 and recombinant SAR1A protein with wild type or disease-causing KLHL40-GST mutant proteins (Figure 7E–F). Western blot analysis showed increased SAR1A ubiquitylation as a function of time in the presence of wild-type KLHL40 but not GST-only control (Figure 7F, Figure 7—figure supplement 1). To understand the role of disease-causing KLHL40 missense variants in disease pathology through SAR1A-mediated pathways, ubiquitylation of SAR1A by NM-causing KLHL40 missense variants was also studied. Variants in the N-terminal BTB domain of KLHL40 (L86P) and BACK domain (W201L) showed similar SAR1A ubiquitylation as the wild-type protein. Still, variants in the Kelch domains (R311L and E528K) resulted in a significant reduction in SAR1A ubiquitylation (Figure 7F–G). As Kelch proteins bind their targets through the C-terminal Kelch domains, this suggests that patients with loss of function or missense variants in the Kelch domains in KLHL40 may exhibit reduced SAR1A ubiquitylation. Finally, SAR1A ubiquitylation was evaluated in C2C12 cells in the presence or absence of KLHL40 with CUL3 by overexpression and immunoprecipitation assays. This showed that KLHL40 is required for SAR1A ubiquitination in the context of muscle cells by CUL3, as no ubiquitylation was observed in the absence of KLHL40 (Figure 7H). Together, these results demonstrate that KLHL40-CUL3 is a regulator of SAR1A in skeletal muscle under normal conditions.

Figure 7 with 1 supplement see all
SAR1A is a direct ubiquitylation target of the KLHL40-CUL3 complex and is differently ubiquitylated by a disease-causing mutation in KLHL40.

(A) Coimmunoprecipitation in C2C12 cells showing KLHL40 directly interacts with SAR1A. (B) Co-overexpression of decreasing KLHL40-FLAG and constant SAR1A-V5 in C2C12 myoblasts demonstrates that KLHL40 is a regulator of Sar1A protein. (C) Co-overexpression of decreasing amounts of KLHL40-FLAG and constant amount of SAR1A-V5 in C2C12 myoblasts in the presence of UPS inhibitor MG132 increases the SAR1A protein levels in comparison to MG132- condition. (D) Alignment of the amino acid sequence of the SAR1A ubiquitylation site demonstrates high conservation in vertebrates (K182 in all species, marked by the asterisk). (E) Localization of different disease-causing variants in KLHL40 in the protein domains. (F) In vitro ubiquitylation of human SAR1A by CUL3 protein complex in the presence of wild-type and disease-causing KLHL40 proteins. (G) Quantifying the relative human SAR1A ubiquitylation by wild-type and disease-causing KLHL40-CUL3 complex. (H) Ubiquitylation of overexpressed SAR1A in the presence of KLHL40 in C2C12 myoblasts. Data are mean ± S.E.M; with one-way analysis of variance (ANOVA) with Dunnett’s multiple comparisons test and Brown-Forsythe test (****p<0.001; n.s. non significant) n=3.

Figure 7—source data 1

Full unedited 7A immunoblot with FLAG antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data1-v2.zip
Figure 7—source data 2

Full unedited 7A immunoblot with V5 antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data2-v2.zip
Figure 7—source data 3

Full unedited 7B immunoblot with FLAG antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data3-v2.zip
Figure 7—source data 4

Full unedited 7B immunoblot with V5 antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data4-v2.zip
Figure 7—source data 5

Full unedited 7B immunoblot with GAPDH antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data5-v2.zip
Figure 7—source data 6

Full unedited 7C immunoblot with FLAG antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data6-v2.zip
Figure 7—source data 7

Full unedited 7C immunoblot with V5 antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data7-v2.zip
Figure 7—source data 8

Full unedited 7C immunoblot with GAPDH antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data8-v2.zip
Figure 7—source data 9

Full unedited 7F immunoblot with SAR1A antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data9-v2.zip
Figure 7—source data 10

Full unedited 7F protein gel.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data10-v2.zip
Figure 7—source data 11

Full unedited 7H immunoblot with FLAG antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data11-v2.zip
Figure 7—source data 12

Full unedited 7H immunoblot with SAR1A antibody.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data12-v2.zip
Figure 7—source data 13

Annotated immunoblots and gel.

https://cdn.elifesciences.org/articles/81966/elife-81966-fig7-data13-v2.pdf

Trafficking of extracellular proteins is perturbed in klhl40a mutant muscle

Defects in ER-Golgi trafficking underlie many skeletal muscle diseases, but the role of COPII vesicles and the trafficking of specific proteins is not known in skeletal muscle. Tango1 is essential for transporting large cargo proteins such as procollagens. As Klhl40a deficient muscle (3 months) showed extensive disruption in the ECM region, we examined the procollagen trafficking in the skeletal muscle of klhl40a mutant and control zebrafish. Immunofluorescence analysis with intracellular muscle protein α-actinin showed extensive immunoreactivity of procollagens intracellularly in the klhl40a mutant muscle compared to the control muscle (Figure 8A). Collagen staining was also reduced in the klhl40a mutant muscle and no collagen immunoreactivity was detected intracellularly (Figure 8A arrow). While no immunostaining of another ECM protein, integrin β–1, was observed intracellularly, significantly reduced levels were seen in ECM in the klhl40a mutant muscle compared to the control muscle. We tested if Klhl40a deficiency affects the ER-Golgi transport of procollagens in MB135 human myoblasts. We employed a selective hook (RUSH) assay with procollagen type1α1 (large cargo protein) and Golgi-localized enzyme sialyl transferase (ST ,small cargo protein). We observed a reduced rate of ER-export of procollagen in human KLHL40 knockout MB135 myoblasts compared to controls and small cargo sialyl transferase (Figure 8B–C). This suggests that KLHL40 is critical for the trafficking of large cargo proteins in the skeletal muscle.

Abnormal ER-Golgi trafficking of procollagen contributes to reduced collagen in the extracellular matrix in Klhl40a deficiency.

(A) Immunofluorescence of control and klhl40a KO zebrafish muscle (3 months). Mutant muscle displays the intracellular accumulation of procollagen (as seen with co-labeling with sarcomeric α-actinin; white arrow) in klhl40a KO muscle compared to +/+control. Mutant muscle showed reduced levels of collagen compared to controls (white arrow). Integrinβ1 level is also reduced in the ECM in the mutant muscle. (B) Retention using selective hooks (RUSH) assay for ER-Golgi trafficking of procollagens and (C) sialyltransferase (ST) in control and KLHL40 knockout human myoblasts. The Golgi apparatus is marked with a yellow dashed line. N=33–45 cells in each group; data are mean ± S.E.M; and Tukey’s HSD test (p<0.01).

KLHL40 human patients exhibit vesicle accumulation and ECM defects

To investigate if SAR1A upregulation and collagen accumulation is associated with disease pathology in KLHL40 deficiency, the skeletal muscle of NM patient KLHL40 patient (c.46C>T, p.Gln16*) was examined by immunofluorescence. Immunofluorescence analysis of the skeletal muscle of the KLHL40 NM muscle showed increased immunoreactivity for SAR1A protein in most of the myofibers. Many myofibers exhibited a very high level of SAR1A immunoreactivity (Figure 9A, white arrows) compared to control muscle. KLHL40 deficient skeletal muscle also showed increased collagen within the muscle fibers, similar to klhl40a fish (Figure 9A, arrowheads). This increased expression of SAR1A is specific to KLHL40-deficient skeletal muscle as analysis of skeletal muscle from centronuclear myopathy patients (RYR1 or MTM1 disease-causing mutations) showed no changes in the SAR1A protein levels compared to the control (Figure 9—figure supplement 1). Analysis of the ultrastructure of skeletal muscle from KLHL40 patient (c.[932G>T];[1516A>C] p.[Ag311Leu];[Thr506Pro]) revealed that in addition to nemaline bodies, extensive vesicle accumulation (arrows) and aberrant ECM structures with reduced collagen fibers (arrowhead) similar to Klhl40 deficient zebrafish were observed (Figure 9B). This suggests that defects in vesicle trafficking contribute to disease onset and pathology in KLHL40 deficiency in patient skeletal muscles.

Figure 9 with 1 supplement see all
KLHL40-NM patients exhibit increased SAR1A protein and vesicle accumulation with ECM defects in skeletal muscle.

(A) Immunofluorescence in control and a KLHL40 patient muscle biopsy showing increased SAR1A protein in the patient muscle (white arrows). Moreover, collagen accumulation is seen in the patient muscle (white arrowhead). (B) Transmission electron microscopy of KLHL40 patient muscle showed vesicle accumulation (arrows) and disorganized and damaged extracellular matrix between myofibers (arrows).

Discussion

Skeletal muscle development is a highly coordinated process involving the differentiation of muscle stem cells, fusion of myoblasts, the formation of multinucleated myotubes, and the development of sarcomeres. In parallel, an extensive intracellular membrane network is established in juxtaposition to sarcomeres to produce force-generating myofibers. Rapid fine-tuning of cellular phenotypes to support the dynamic transitions are accomplished through post-transcriptional and post-translational processes. These control protein synthesis rates and modify protein functions and dynamic degradation through the UPS. The protein degradation process during skeletal muscle growth and disease onset is highly selective, as evident from the identification of mutations in components in the UPS pathway in human myopathies that perturb specific stages of muscle development and growth and result in impaired motor function (Olivé et al., 2015; Carrasco-Rando and Ruiz-Gómez, 2008; Cirak et al., 2010). In particular, KLHL40 and KLHL41, that functions as substrate-specific adaptors for E3 ubiquitin ligase CUL3, contribute to severe forms of congenital myopathy with neonatal lethality with extensive sarcomeric disarray and contractures (Ravenscroft et al., 2013; Gupta et al., 2013).

To address the gaps in the understanding of disease development and identify in vivo events that initiate the pathology and the mechanisms through which these events become pervasive in Klhl40a deficiency, we performed quantitative global proteome and ubiquitylome analysis in skeletal muscle from non-symptomatic stages to symptomatic stages of disease progression in the zebrafish model (Mertins et al., 2018; Udeshi et al., 2020; Satpathy et al., 2021). We identified that normal skeletal muscle proteome exhibited plasticity and was dynamically changed during growth. In contrast, Klhl40a-deficient skeletal muscle showed a highly altered proteome during the early stages of skeletal muscle growth (i.e. during pre-symptomatic stages) compared to controls that remained primarily static during the transition from juvenile to adult stages and throughout disease progression. In addition, the proteome data identified early preclinical signatures suggesting disease-causing processes are established during pre-symptomatic stages before structural and functional deficits are observed in skeletal muscle. Moreover, most of the proteome (78%) that exhibited changes during the juvenile-to-adult transition did not show significant differences in gene expression by RNA sequencing. In addition, no changes were detected at the transcriptome levels for proteins that showed significant changes in the ubiquitylation and their cognate proteins suggesting that regulatory processes such as post-transcription, post-translation, and protein degradation impact protein abundance after mRNA is made (Zeng et al., 2022).

Skeletal muscle depends on anaerobic glycolysis and oxidative phosphorylation for its bioenergetic demands (Hargreaves and Spriet, 2020). During skeletal muscle development, aerobic glycolysis is the primary source of bioenergetics in proliferative cells, but this changes to oxidative phosphorylation (OXPHOS) during differentiation. Our combined analysis of the ubiquitylome and proteome identified many glycolytic enzymes that exhibited reduced ubiquitylation and increased protein levels in Klhl40a deficiency. High glycolytic enzyme pyruvate kinase (Pkm2) levels lead to defects in energy metabolism and skeletal muscle atrophy in the myotonic dystrophy (Gao and Cooper, 2013). This suggests that Klhl40a directly or indirectly acts as a regulator of glycolysis in skeletal muscle, and a deficiency may contribute to a perturbed bioenergetic state and muscle defects. We also observed the upregulation of OXPHOS proteins in Klhl40a deficiency and associated changes in mitochondrial morphology. While no changes in the ubiquitylation of these proteins were seen, this dysregulation of mitochondrial proteins could be a response to altered glycolysis or structural abnormalities in skeletal muscle. The upregulation of mitochondrial proteins is associated with increased energetic demands of the diseased muscle and may further exacerbate the disease pathology through increased oxidative stress (Toepfer et al., 2020; Pant et al., 2015). Klhl40a-deficient muscle also showed reduced ubiquitylation and increased abundance of early sarcomeric proteins that are normally expressed in the early differentiation of myofibers and are decreased or absent during terminal myofiber differentiation. We have previously shown that reduced ubiquitylation and upregulated protein level of an early sarcomeric protein, NRAP, in KLHL41 deficient skeletal muscle contributes to myopathy by abnormal sequestration of the late sarcomeric proteins and preventing their localization to mature sarcomeres (Jirka et al., 2019). This suggests a similar mechanism may contribute to smaller sarcomeres and myopathy observed in Klhl40a deficiency. Nemaline myopathy is associated with defects in sarcomere structure, and it is not clear if the formation or growth of the sarcomere is affected in NM patients. Our studies demonstrate that in KLHL40-NM, sarcomeres are initially formed normally, but further sarcomere growth is perturbed and provides mechanistic insights into disease pathology.

Our studies identified that Klhl40a regulates skeletal muscle growth and function through regulation of vesicle trafficking, a poorly understood pathway in skeletal muscle. Vesicle trafficking is mediated by multiple organelles that coordinate the transport of proteins synthesized in the ER to the extracellular space or other endomembrane compartments through the Golgi apparatus (Barlowe and Schekman, 1993). Human genetic studies have identified pathogenic variants in critical proteins in the vesicular trafficking pathway, resulting in myopathies in affected patients such as GOLGA2, BIDC2 and BET1 (Oates et al., 2013; Kotecha et al., 2021; Shamseldin et al., 2016; Neveling et al., 2013; Unger et al., 2016; Shomron et al., 2021; Donkervoort et al., 2021). Variants in many genes associated with congenital muscular dystrophies (POMT1, POMT2, TRAPPC11, GOSR2) encode proteins that are localized to different membrane compartments of the vesicle trafficking pathway (Beltrán-Valero de Bernabé et al., 2002; van Reeuwijk et al., 2005; Bögershausen et al., 2013; Larson et al., 2018). How defects in specific components in the trafficking pathway affect the distribution of secretory and extracellular proteins locally or globally and contribute to muscle diseases is not well understood. Nevertheless, our studies provide in vivo insights into the requirement of vesicle trafficking to transport ECM proteins to maintain healthy skeletal muscle. KLHL40-CUL3 acts as a regulator of this process through ubiquitylation-mediated protein degradation of SAR1A, which is required for budding COPII vesicles from ER to transport proteins. CUL3-dependent ubiquitylation is previously shown to dynamically regulate the trafficking of large COPII carriers (Jin et al., 2012). KLHL12, another substrate-specific adapter for CUL3 E3 ubiquitin ligase forms a complex with CUL3 which ubiquitylates SEC31 leading to an increase in COPII vesicle size to accommodate large procollagen molecules for secretion in mouse embryonic stem cells (mESC; Jin et al., 2012). While most procollagens are trafficked through the secretory pathway, a subset is directed towards lysosomal degradation to remove excess procollagen from cells through the autophagy pathway (Omari et al., 2018). Recent studies in skin fibroblasts have shown that the CUL3-KLHL12 complex is involved in routing procollagens to lysosomes to regulate intracellular collagen levels (Moretti et al., 2023). Moreover, inhibition of CUL3 neddylation which is critical for the ubiquitylation activity still led to the formation of large COPII vesicles by the CUL3-KLHL12 complex, which is required for the secretion of procollagens. As CUL3-mediated ubiquitylation also regulates KLHL12 protein stability, further studies are needed to understand the ubiquitylation-dependent and independent roles of the CUL3-KLHL12 complex on procollagens secretion in different cellular contexts and physiological conditions. KLHL12 is expressed at very low levels in normal skeletal muscle compared to other cells and tissue types (http://gtexportal.org). We did not identify any differential changes in Klhl12 and the target protein Sec31 in Klhl40a deficiency.

Moreover, no changes in the protein levels of autophagy markers were observed in Klhl40a deficiency at the disease states examined. CUL3 interacts with many Kelch proteins in a tissue-specific context and therefore, may regulate specific aspects of secretory and degradative pathways in response to different stimuli and disease states. Klhl40a deficient skeletal muscle showed an increased number of enlarged ERES sites compared to controls, while the overall number of ERES sites was reduced in the mutant muscle. ERES functions as an inter-organelle transport apparatus that actively modulates its shape and size while directing diverse cargo types to Golgi and increases in size during active transport (Weigel et al., 2021). As Klhl40a deficiency resulted in reduced procollagen trafficking from ER, this increase in the size of ERES sites could be a compensatory mechanism to enhance the secretory flux. Sar1 is associated with the ERES sites and regulates the formation of COPII vesicles. Despite the presence of increased Sar1A levels in Klhl40a-deficient myofibers, reduced or limited amounts of other proteins, such as Sec16b and COPII proteins in the mutant myofibers may underlie a reduced number of ERES sites and COPII vesicles.

While the Sar1a level was increased in the klhl40a mutant, no differences were observed in closely related family member Sar1b in Klhl40a deficiency (Supplementary file 1) showing a decrease in Sar1a and Sar1b ratio. SAR1A is 90% identical to SAR1B, and these proteins exhibit overlapping and unique functions with different biochemical properties in COPII assembly (Melville et al., 2020; Georges et al., 2011; Jones et al., 2003). While SAR1B specifically regulates chylomicron trafficking in the small intestine, both proteins are required for the trafficking of cardiac sodium channel Nav1.5 protein and efficient ER export of procollagens and may be able to compensate for each other function for the trafficking of common proteins (Jones et al., 2003; Levic et al., 2015; Kim et al., 2012; Cutrona et al., 2013). In Klhl40a deficiency, increased amounts of Sar1a resulted in the formation of abnormal ER-derived membrane-bound structures and decreased procollagens trafficking; therefore, downregulation of Sar1a levels in skeletal muscle may be able to restore the procollagen trafficking defects. Klhl40a deficiency muscle showed extensive ER dilation that can lead to activation of the UPR caused by the ER stress. However, most of the markers of UPR activation showed no significant difference in the Klhl40a deficiency in skeletal muscle, suggesting a lack of UPR activation in vivo.

A key clinical feature of KLHL40 NM is the presence of contractures in most individuals. Defects in ECM are directly associated with contracture in many neuromuscular diseases. Vesicle trafficking dysregulation is associated with neurodegenerative and skeletal disorders, and our findings open new avenues on this pathway in neuromuscular diseases. Our studies identified structural defects in skeletal muscle in Klhl40a deficiency and showed that Klhl40a directly regulates vesicle trafficking and contributes to disease pathology.

Overall, we provide a comprehensive temporal proteomic landscape during skeletal muscle growth and disease development, and dynamic fine-tuning of the cellular proteome by ubiquitylation is critical for muscle function. While comprehensive studies are needed to define the specific role of each of the different processes identified in skeletal muscle defects in KLHL40 deficiency, these studies suggest these altered molecular processes contribute to specific pathological defects observed in Klhl40a deficiency. Given the several pathways in which KLHL40 is involved, approaches aiming to inactivate pro-disease pathways and activate protective pathways may be a promising therapeutic strategy for at least this form of NM.

Methods

Zebrafish maintenance and husbandry

Zebrafish were maintained and bred using standard methods as described (Westerfield, 2000). The Institutional Animal Care and Use Committee approved all experiments and procedures at Brigham and Women’s Hospital (2016000304). Wild-type fish were obtained from Tubingen (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 have been described previously (Melville et al., 2020).

Creation of zebrafish lines

sgRNAs were designed using the web-based ZiFiT Targeter program (http://zifit.partners.org/) and targeting specific sites in exon 1 or exon 2 of zebrafish Kelch genes. The first two bases (GG) 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 22 bases 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 Supplementary file 10. The zebrafish 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 (Hwang et al., 2013). sgRNA and Cas9 protein (Thermo Scientific, CA) were co-injected into the yolk sac of one- and two-cell stage zebrafish embryos. Each embryo was injected with a 5 μl solution containing 2 μl of 400 ng/nl Cas9 protein and 3 μl of 100 ng/μl sgRNA. Injected embryos were inspected under the microscope for three days and were classified as dead, deformed, or normal phenotypes. Embryos displaying normal phenotypes were analyzed to test 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) and used for DNA sequencing experiments by Topo cloning.

Identification of founder fish and generation of isogenic 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 and observing mosaicism at the target site. The mosaic F0 generation was outcrossed to wild-type TU fish for at least three generations for studies presented in this work. The sequences of sgRNAs are listed in Supplementary file 10.

In-solution digestion

Muscle samples frozen in liquid nitrogen were cryofractured using the cryoPREP tissue disruption system on setting 4 (Covaris). Samples were then lysed for 30 min at 4 °C in urea lysis buffer (8 M urea, 50 mM Tris-HCl pH 8.0, 75 mM NaCl, 1 mM EDTA, 2 µg/µl aprotinin (Sigma-Aldrich), 10 µg/µl leupeptin (Roche), and 1 mM phenylmethylsulfonyl fluoride (PMSF) (Sigma-Aldrich)) and cleared by centrifugation at 20,000xg. Protein concentrations were determined by bicinchoninic acid (BCA) protein assay (Pierce), and samples were diluted to a protein concentration of 2 µg/μl. Samples were reduced with 5 mM dithiothreitol (DTT) for 1 hr at 21 °C, followed by alkylation with 10 mM iodoacetamide for 45 min at 21 °C. Samples were diluted with 50 mM Tris-HCl pH 8.0 to a final urea concentration of 2 M prior to enzymatic digestion. Proteins were digested with the endoproteinase LysC (Wako Laboratories) for 2 hr at 25 °C followed by overnight digestion with sequencing-grade trypsin (Promega) at 25 °C (enzyme-to-substrate ratios of 1:50). Following digestion, samples were acidified to a concentration of 1% formic acid (FA) and cleared by centrifugation at 20,000xg. The remaining soluble peptides were desalted using a 100 mg reverse phase tC18 SepPak cartridge (Waters). Cartridges were conditioned with 1 ml 100% acetonitrile (MeCN) and 1 ml 50% MeCN/0.1% FA, then equilibrated with 4X1 ml 0.1% trifluoroacetic acid (TFA). Samples were loaded onto the cartridge and washed 3 X with 1 ml 0.1% TFA and 1 X with 1 ml 1% FA, then eluted two times with 600 µl 50% MeCN/0.1% FA per elution. Peptide concentration of desalted samples was again estimated by BCA assay and dried by vacuum centrifugation.

TMT labeling of peptides

Tandem mass tag (TMT) labeling was performed as previously described (Zecha et al., 2019). Briefly, 100 µg peptides per sample were resuspended in 50 mM HEPES pH 8.5 at a concentration of 5 mg/ml. Dried Tandem Mass Tag (TMT) pro 16-plex reagent (ThermoFisher Scientific) was reconstituted at 20 µg/µl in 100% anhydrous MeCN and added to samples at a 2:1 TMT to peptide mass ratio. The reaction was incubated for 1 hr at 25 °C while shaking and quenched with 5% hydroxylamine to a final concentration of 0.2% for 15 min at 25 °C while shaking. The TMT-labeled samples were then combined, dried to completion by vacuum centrifugation, reconstituted in 1 ml 0.1% FA, and desalted with a 100 mg SepPak cartridge as described above.

Basic reverse phase (bRP) fractionation

TMT-labeled peptides were fractionated via offline basic reverse-phase (bRP) chromatography as previously described (Mertins et al., 2018). Chromatography was performed with a Zorbax 300 Extend-C18 column (4.6x250 mm, 3.5 µm, Agilent) on an Agilent 1100 high-pressure liquid chromatography (HPLC) system. Samples were reconstituted in 900 µl of bRP solvent A (5 mM ammonium formate, pH 10.0 in 2% vol/vol MeCN). Peptides were separated at a flow rate of 1 ml/min in a 96 min gradient with the following concentrations of solvent B (5 mM ammonium formate, pH 10.0 in 90% vol/vol MeCN) 16%B at 13 min, 40%B at 73 min, 44%B at 77 min, 60%B at 82 min, 60%B at 96 min. A total of 96 fractions were collected and concatenated non-sequentially into 24 fractions for proteomic analysis. Fractions were dried via vacuum centrifugation, and an equivalent of 1 µg of the peptide was injected for LC-MS/MS analysis.

Liquid chromatography and mass spectrometry for global proteome analysis

Dried fractions were reconstituted in 3% MeCN/0.1% FA to an estimated peptide concentration of 1 µg/µl and analyzed via coupled nanoflow liquid chromatography and tandem mass spectrometry (LC-MS/MS) using a Proxeon Easy-nLC 1200 (Thermo Fisher Scientific) coupled to an Orbitrap Exploris 480 Mass Spectrometer (Thermo Fisher Scientific). A sample load of 1 µg for each fraction was separated on a capillary column (360x75 µm, 50 °C) containing an integrated emitter tip packed to a length of approximately 25 cm with ReproSil-Pur C18-AQ 1.9 μm beads (Dr. Maisch GmbH). Chromatography was performed with a 110 min gradient of solvent A (3% MeCN/0.1% FA) and solvent B (90% MeCN/0.1% FA). The gradient profile, described as min:% solvent B, was 0:2, 1:6, 85:30, 94:60, 95:90, 100:90, 101:50, 110:50. Ion acquisition was performed in data-dependent mode with the following relevant parameters: MS1 orbitrap acquisition (60,000 resolution, 350–1800 scan range (m/z), 300% normalized AGC target, 25ms max injection time) and MS2 orbitrap acquisition (20 scans per cycle, 0.7 m/z isolation window, 32% HCD collision energy, 45,000 resolution, 50% normalized AGC target, 50ms max injection time, 15 s dynamic exclusion, 50% fit threshold, and 1.2 m/z fit window).

K-GG enrichment for ubiquitylome analysis

Ubiquitin enrichment was performed based on the UbiFast protocol (Udeshi et al., 2020). Anti-K-e-GG bead-bound antibodies from the PTM-Scan ubiquitin remnant motif kit (Cell Signaling Technologies #5562) were cross-linked as follows. Beads were washed 3 X with 100 mM sodium borate (pH 9.0) and incubated with 20 mM DMP for 30 min at RT. Beads were then washed 2 X with 200 mM ethanolamine and incubated overnight at 4 °C in 200 mM ethanolamine with end-over-end rotation. Following incubation, beads were washed 3 times with IAP buffer and stored at 4 °C at a concentration of 0.5 µg/μL. For each 11-plex experiment, 31.25 µg of cross-linked anti-K-GG bead-bound antibody at 0.5 µg/μL in IAP per channel was aliquoted into 1.5 mL Eppendorf tubes on ice. 1 mg peptide per sample was reconstituted to 0.5 mg/mL concentration in IAP buffer and vortexed for 10 min. Peptides were then centrifuged for 5 min at 5000 g. Each peptide solution was added to a tube of antibody and gently rotated end-over-end at 4 °C for 1 hr. Following enrichment, samples were centrifuged (1 min, 2000xg), and the supernatant was removed. Beads were washed with 1.5 mL ice-cold IAP followed by 1.5 mL ice-cold PBS (30 s, 2000xg) and reconstituted in 200 μL 100 mM HEPES buffer. A total of 400 µg of TMTpro 16-plex labeling reagent in 10 μL acetonitrile was added for each sample. Peptides were TMT labeled on-beads while shaking vigorously (1400 rpm) at 20 °C for 10 min, then quenched with 8 μL 5% hydroxylamine and shaken vigorously for another 5 min washed once with 1.3 mL cold IAP, and again with 1.5 mL cold IAP. Each channel was resuspended and transferred to a combination tube with 130 μL cold IAP. Following the combination, each now-empty tube was serially washed with 1.5 mL cold IAP to remove the remaining beads, and this 1.5 mL IAP was added to the combination tube and used to wash the combined beads. Combined beads were washed one final time with 1.5 mL ice-cold PBS. Once the channels were combined and washed, peptides were eluted twice from the beads by resuspending with 150 μL room temperature 0.15% TFA and incubated for 5 min at RT. Each round of acid-eluted K-GG-modified peptides was desalted on an equilibrated two-punch C18 stage tip. Both elutions of K-GG peptides were loaded sequentially, washed twice with 100 μL 0.1% FA, and eluted into an MS vial with 50 μL 50% ACN/0.1% FA. The eluted peptides were frozen, lyophilized, and reconstituted in 9 μL 3% ACN/0.1% FA, with 4 μL injected twice for two consecutive LC-MS/MS runs.

Liquid chromatography and mass spectrometry for global proteome analysis

Reconstituted K-GG enriched peptides were analyzed via coupled nanoflow liquid chromatography and tandem mass spectrometry (LC-MS/MS) using a Proxeon Easy-nLC 1200 (Thermo Fisher Scientific) coupled to an Orbitrap Exploris 480 Mass Spectrometer (Thermo Fisher Scientific) equipped with a FAIMS interphase. Four out of 9 μl of total eluted material was separated on a capillary column (360x75 µm, 50 °C) containing an integrated emitter tip packed to a length of approximately 25 cm with ReproSil-Pur C18-AQ 1.9 μm beads (Dr. Maisch GmbH). Chromatography was performed with a 154 min gradient of solvent A (3% MeCN/0.1% FA) and solvent B (90% MeCN/0.1% FA). The gradient profile, described as min:% solvent B, was 0:2, 2:6, 122:35, 130:60, 133:90, 143:90, 144:50, 154:50. Ion acquisition was performed in data-dependent mode with the following relevant parameters: three FAIMS CV settings (–45 V, –50 V, and –70 V), MS1 orbitrap acquisition (60,000 resolution, 350–1800 scan range (m/z), 100% normalized AGC target, 10ms max injection time) and MS2 orbitrap acquisition (10 scans per cycle, 0.7 m/z isolation window, 32% HCD collision energy, 45,000 resolution, 50% normalized AGC target, 120ms max injection time, 20 s dynamic exclusion, 50% fit threshold, and 1.4 m/z fit window).

Data analysis

Raw MS/MS data from heart and liver samples were processed using Spectrum Mill v.7.09.215 (Proteomics.broadinstitute.org). MS2 spectra were extracted from RAW files and merged if originating from the same precursor or within a retention time window of +/-60 s and m/z range of +/-1.4, followed by filtering for precursor mass range of 750–6000 Da and sequence tag length >0. MS/MS search was performed against the UniProt Danio rerio protein database downloaded in November 2020 and common contaminants, with digestion enzyme conditions set to “Trypsin allow P,”<5 missed cleavages, fixed modifications (cysteine carbamidomethylation and TMTpro on N-term and lysine), and variable modifications (oxidized methionine, acetylation of the protein N-terminus, pyroglutamic acid on N-term Q, and pyro carbamidomethyl on N-term C). Additional variable modifications were added for ubiquitylome (di-glycine residual in K). Matching criteria included a 30% minimum matched peak intensity and a precursor and product mass tolerance of +/-20 ppm. Peptide-level matches were validated if found to be below the 1.0% false discovery rate (FDR) threshold and within a precursor charge range of 2–6. A second round of validation was performed for protein-level matches for proteome datasets, requiring a minimum protein score of 13. Ubiquitylome site-centric and protein-centric data, including TMT intensity values and ratio to the median of all samples, was extracted and summarized in a table. Raw mass spectrometry data will be made publicly available in MassIVE upon acceptance of the manuscript.

Statistical analysis of proteomics data

Statistical analysis was performed in the R environment for statistical computing. Sample log2 TMT ratios were median centered. Proteins with less than 2 unique peptides were removed from downstream analysis. One sample (4 month knockout replicate 1) was identified as an outlier by PCA and removed from the dataset. To identify proteins and KGG-sites with differential abundance between WT and KO groups, a linear model was fit with the age and genetic background as experimental factors, and moderated T-tests were performed using the limma package (Ritchie et al., 2015). Multiple hypothesis testing correction was performed using the BH method.

Pathway enrichment and network visualization

Proteins and KGG-sites showing a differential abundance in response to klhl40a knockout at both ages were used for downstream pathway analysis (adjusted p-value <0.05). Pathway enrichment analysis was performed for features increasing or decreasing in abundance in the KO stain at each of the two ages using the g: profiler tool (Raudvere et al., 2019). The background list of proteins was set to all detected in the proteomics analysis. The list of enriched pathways and genes contained in each pathway were exported to Cytoscape (Shannon et al., 2003). The EnrichmentMap application was used to generate a network of enriched pathways with the following parameters (pathway FDR p-value <0.05; Jaccard index >0.35) (Merico et al., 2010).

RT-PCR

Total RNA from control and klhl40a zebrafish muscle was isolated and cDNA was synthesized as performed previously (Bennett et al., 2018). xbp1 splicing was analyzed by RT-PCR as reported (Li et al., 2015).

Tissue sample preparation and western blotting analysis

Zebrafish muscle tissue (10–15 mg) was placed in RIPA Lysis and Extraction Buffer (Thermo Fisher Scientific) with a cocktail of protease inhibitors and homogenized (2X15 s) using the Tissuemiser homogenizer (Thermo Fisher Scientific). Samples were separated by an SDS-PAGE and blotted onto polyvinylidene difluoride (PVDF) membranes. The membranes were blocked using 5% non-fat milk powder in 1 X Tris Buffered Saline (Boston Bioproducts, MA) and 0.1% TWEEN 20 (Sigma-Aldrich, cat. no. P9416) (TBST) for 1 hr at room temperature and incubated with primary antibodies overnight at 4 °C. The membranes were subsequently washed and incubated with polyclonal anti-mouse-IgG antibody conjugated to horseradish peroxidase. To isolate protein from the human skeletal muscle biopsies, 50-μM-thick frozen sections were resuspended in tissue protein extraction buffer (T-PER, Thermofisher Scientific) with inhibitors and homogenized (1X15 s) using the Tissuemiser homogenizer (Thermo Fisher Scientific). The antibody used and associated dilutions are: Anti-KBTBD5 for KLHL40, 1:100 dilution (sc-99943, Santa Cruz Biotechnology); anti-α-Tubulin,1:500 (ab18251, Abcam); anti-Sar1a,1:100 (ab125871, Abcam); anti-Sec24d, 1:100 (14687, Cell Signaling technology); anti-Golga2, 1:100 (ab30637, Abcam); anti-FLAGM2, 1: 250 (F1804, Sigma-Aldrich); anti-V5, 1:500 (R960-25, Thermo Fisher Scientific), LC3B,1:100 (3868, Cell Signaling), PERK, 1:100 (3192, Cell Signaling) and phospho-PERK, 1:100 (3179, Cell Signaling). 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 Image J.

Immunofluorescence

Zebrafish or human frozen skeletal muscle tissue were cryosectioned (8 μm) or myofibers were used for immunofluorescence as previously described (Gupta et al., 2013). Myofibers were isolated from control or klhl40a zebrafish (1.5 or 3 months) as described previously with minor modifications (Ganassi et al., 2021). Skinned zebrafish muscle samples were treated with collagenase for 90 min and triturated to release the myofibers. Myofibers were centrifuged at 1000 g for 60 s, washed and resuspended in DMEM media. Myofibers were plated on laminin coated 8 chamber permanox slides (Thermo Fisher Scientific) for further analysis. Fixed cells were blocked in 10% goat serum/0.3% Triton, incubated in primary antibody overnight at 4 °C, washed in PBS, incubated in secondary antibody for 1 hr at room temperature, washed in PBS, then mounted with Vectashield Mounting Medium (Vector Laboratories, Burlingame, CA, USA).

Antibodies used for immunofluorescence were anti-Sar1a,1:100 (ab125871, Abcam); anti-RYR1,1:250 (R129, Sigma-Aldrich); anti-procollagen, 1:100 (MAB1912, Millipore Sigma); Integrin, 1:25 (clone8c8, DSHB); PDI, 1:100 (ab2792, Abcam); Tango1, 1:100 (17481–1-AP, Proteintech); Sec23B, 1:100 (Sigma, HAP069974). Secondary antibodies were anti-rabbit,1:250 (A11008, Thermo Fisher Scientific); anti-mouse,1:250 (A11005, Thermo Fisher Scientific). Imaging was performed using a Nikon Ti2 spinning disk confocal microscope.

SAR1A mRNA overexpression in zebrafish

Human SAR1A cDNA was subcloned from a pDEST40-SAR1-V5-His6 plasmid (a gift from Richard Kahn; Addgene plasmid # 67451; http://n2t.net/addgene:67451; RRID: Addgene_67451) into pCSDest vector. mRNA was synthesized in vitro using mMessage kits (Ambion, Austin, TX, USA). 50–100 pg of mRNA was injected into embryos at the one-cell stage.

C2C12 cell culture studies

Coimmunoprecipitation of KLHL40 and SAR1A (from pDEST40-SAR1-V5-His6) was performed using the previously described method (Jirka et al., 2019). To study the reciprocal interaction between KLHL40 and SAR1A, C2C12 cells (ATCC, #CRL-1772) were transfected with different amounts of KLHL40-pEZYFLAG and pDEST40-SAR1-V5-His6 plasmids. MG132 (10 μM) was added at 40 hr post-transfection, and cells were harvested 48 hr post-transfection. Cell lysates were prepared in RIPA buffer, and proteins were analyzed by western blot analysis. The cells were routinely checked for mycoplasma and tested negative.

Creation of KLHL40 knockout human myoblasts

sgRNAs were designed to target exon 1 of the human KLHL40 gene using the Broad Institute’s CRISPick tool (https://portals.broadinstitute.org/gppx/crispick/public). The oligonucleotides for the sgRNAs were cloned into lentiCRISPRv2 plasmid; Exon 1–1: (F: 5’-CACCGATGGTGAAGGATGCACACGA –3’ R: 5’-AAACTCGTGTGCATCCTTCACCATC –3’) and Exon 1–2 (F: 5’-CACCGGGAAGCACAGTAGCACTCGT –3’ R: 5’-AAACACGAGTGCTACTGTGCTTCCC-3’). The sgRNAlentiCRISPRv2 DNA was cotransfected with pCMVVSVG and psPAX2 into HEK293 cells and the supernatant containing lentivirus was collected at 48 hr post-transfection. Transduction with the sgRNA lentiviruses was performed in human MB135 myoblasts (source PMID:28171552), followed by antibiotic selection (Puromycin, 10 µg/μl) of the positive clones. Single cells were subsequently plated for clonal expansion, and Sanger sequencing was performed with the genomic DNA to identify KLHL40 knockout clones. Control and mutant cells were validated by Sanger sequencing for all experiments. The cells were routinely checked for mycoplasma and tested negative.

Retention using selective hooks assay

The retention using selective hooks (RUSH) assay was performed as previously described (Boncompain and Perez, 2012). Briefly, wild-type and KLHL40 knockout myoblasts (c.1010_1011insA;p.Cys337Valfs) were co-transfected by electroporation with two plasmids (pLVX-SBP-mGFP-COL1A1; Addgene plasmid: 110726 and Str-KDEL_ST-SBP-mCherry; Addgene plasmid: 65265). The first plasmid expresses an engineered human procollagen type I alpha 1 with streptavidin binding protein and mGFP between the prosequence and triple helical region. The second plasmid co-expresses the Golgi-localized enzyme sialyl transferase-mCherry (ST-mCherry) fused to streptavidin binding protein and also KDEL-tagged streptavidin, necessary for the ER retention of both cargo proteins (mGFP-COL1A1 and ST-mCherry). Simultaneous release of both reporters from the ER was accomplished by the addition of 40 μM biotin, and live cells were monitored by fluorescence microscopy every 2 min. Images were obtained on a Nikon Livescan sweptfield confocal microscope with a×40 objective lens (NA 0.95), and the resulting movies used for quantitative fluorescence analysis were not subjected to processing. Integrated fluorescence intensity of mGFP-COL1A1 at the Golgi region (defined by the region of perinuclear intensity seen 20–30 min after biotin addition) and from the whole cell was measured using ImageJ. The ratio between fluorescent intensities within the Golgi region and the whole cell was generated for each time point. The ratio at the 0 min time point, representing ER background signal at the Golgi region, was subtracted from the corresponding ones at each time point and then normalized to the maximum value. The kinetics of mGFP-COL1A1 trafficking represents a change in the ratio over time (0–30 min).

Electron microscopy

Muscle tissue was dissected from juvenile and adult zebrafish (1.5 and 3 months), deskinned, and fixed in formaldehyde–glutaraldehyde– picric acid in cacodylate buffer overnight at 4 °C, followed by osmication and uranyl acetate staining. Subsequently, muscle tissue samples were dehydrated in a series of ethanol washes and finally embedded in Taab epon (Marivac Ltd., Nova Scotia, Canada). We dissected a single animal at a time, collected 50–100 mg of skeletal muscle biopsy, and fixed it immediately to prevent any contracting state artifacts which can result from the uneven fixation of thick samples. Moreover, blinded sample processing and image analysis, and quantification of muscle biopsies from multiple fish tissues was performed to examine if the differences between control and experimental groups were reproducible. Ninety-five nanometer sections were cut with a Leica ultra cut microtome, picked up on 100 m formvar-coated copper grids, and stained with 0.2% lead citrate. Sections were viewed and imaged on a Joel 1200EX Transmission Electron Microscope (Electron Microscopy Core, Harvard Medical School).

Expression and purification of SAR1A and KLHL40 proteins

SAR1A cDNA (addgene, #67451) or KLHL40 (WT or mutant cDNAs) were cloned into the pDEST15 vector by gateway cloning. The SAR1A-pDEST15 or KLHL40-pDEST15 vectors were transformed into BL21-Codon Plus (DE3) E. coli cells and cultured in LB media supplemented with 100 μg/mL ampicillin at a 1 L scale. The cells were grown at 37℃ until O.D600=0.4 and induced with 0.5 mM IPTG for 18 hr at 18 °C. The harvested cells were resuspended in 25 mM HEPES (pH 7.4), 130 mM NaCl, 20 mM MgCl2, 1 mM TCEP, 1 mM PMSF, and 1 tablet of protease inhibitor cocktail (Pierce). Following lysis by the French press, cell debris was pelleted by centrifuging at 27,000xg for 40 min, and the soluble lysate was loaded onto glutathione-agarose resin (MCLAB) for affinity purification. The resin was washed with wash buffer containing 25 mM HEPES (pH 7.4), 130 mM NaCl, 20 mM MgCl2, 1 mM TCEP, 0.1% Triton X-100, and then washed with the same buffer without Triton X-100. GST-tagged proteins were eluted with 50 mM reduced glutathione in 25 mM HEPES (pH7.4), 130 mM NaCl, 20 mM MgCl2, 1 mM TCEP, and dialyzed into 50 mM HEPES (pH7.4), 150 mM NaCl, 1 mM TCEP, 10% glycerol. The purified protein was concentrated to 5 mg/ml, flash-frozen, and stored at 18 °C.

In vitro ubiquitination assays

The in vitro ubiquitination assays for SAR1A were conducted at 37 °C in a total volume of 20 μL. The reaction mixture containing 5 mM ATP, 100 μM wild-type ubiquitin, 100 nM E1 protein, 2 μM E2 (UbcH5b), 0.38 μM CUL3-NEDD8-RBX2 (BostonBiochem, USA), 0.3 μM KLHL40-GST(WT or mutants) or GST and 5 μM SAR1A, with 40 mM Tris-HCl (pH 7.5), 50 mM NaCl, 0.5 mM TCEP and 5 mM MgCl2 as the reaction buffer. Substrate SAR1A was preincubated with everything in the reaction mixture except E1 at 37 °C for 20 min before E1 was added to the reaction system to initiate the reactions. Reactions were quenched at the indicated time points (0, 30, and 90 min) by adding an SDS loading buffer containing the reducing agent dithiothreitol (DTT). The reaction samples were then resolved by SDS-PAGE gels and analyzed by either the Colloidal Blue Staining kit (Thermo Fisher Scientific, USA) or western blot analysis. Assays were repeated on at least three independent occasions revealing results similar to the data presented in the figures.

Western blotting for in vitro ubiquitination assays

After SDS-PAGE, the proteins were transferred to nitrocellulose membranes using an iBlot blotting system (Thermo Fisher Scientific, USA). The membranes were then blocked with 5% BSA in phosphate buffer saline tween (0.5%) (PBST) buffer for 1 hr and then incubated with the anti-SAR1A antibody (1:500) at 4 °C overnight. After this, the membranes were washed with PBST and probed with horseradish peroxidase-conjugated anti-Rabbit secondary antibody. The bands were detected by chemiluminescence using a Clarity Western ECL substrate (Bio-Rad, USA).

Data analysis for in vitro ubiquitination assays

To quantify the reaction rate of the SAR1A ubiquitination reactions, the ubiquitinated SAR1A bands detected by western blot were quantified by densitometric analysis using Image J (version 1.53 a). The relative ubiquitination rate of the KLHL40 mutant proteins group versus the WT KLHL40 group was calculated from three biological replicates. The average values and standard deviations (presented as error bars) were calculated and shown in the Figure 7. The statistical significance and p values (or non-significant, n.s.) between groups were calculated using GraphPad Prism 9 using one-way ANOVA and reported in the Figure 7.

Materials availability

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

Data availability

The data is publicly available via Sequence Read Archive (SRA) (Accession Number: PRJNA861969) and MassIVE (https://massive.ucsd.edu) and are accessible at ftp://MSV000090018@massive.ucsd.edu.

The following previously published data sets were used
    1. Mansur A
    2. Joseph R
    3. Kim ES
    4. Jean-Beltran PM
    5. Udeshi ND
    6. Pearce C
    7. Jiang H
    8. Iwase R
    9. Milev MP
    10. Almousa HA
    11. McNamara E
    12. Widrick J
    13. Perez C
    14. Ravenscroft G
    15. Sacher M
    16. Cole PA
    17. Carr SA
    18. Gupta VA
    (2022) MassIVE
    ID MSV000090018. Dynamic regulation of inter-organelle communication by ubiquitylation controls skeletal muscle development and disease onset in nemaline myopathy.

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Decision letter

  1. Meir Aridor
    Reviewing Editor; University of Pittsburgh, United States
  2. Vivek Malhotra
    Senior Editor; Barcelona Institute for Science and Technology, Spain

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Dynamic regulation of interorganelle communication by ubiquitylation controls skeletal muscle development and disease onset" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Vivek Malhotra as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1) Overall myopathic effects and changes on a cellular level to the morphology of organelles and sorting sites including ER exit sites and Golgi need further analysis and quantification.

2) Analysis of biosynthetic secretion and in particular collagen mobilization in the secretory pathway or degradation that may lead to ECM deposition defects are required.

3) Further analysis of the findings on Sar1 with added controls and analysis of all COPII and ER exit site components is required to provide a clear view of the basis for secretion defects that may lead to disease onset.

4) Attention should be given to the complexity of effects leading to the overall disease phenotypes.

Reviewer #1 (Recommendations for the authors):

Specific points.

1. Although the authors demonstrate that selective effects are derived from the deletion of KLHL40a, both a and b forms are expressed to almost the same levels (Figure S1A-E). What is the explanation for the differential effect? Is it localization? Is it a lack of different activities of the b isoform and why? Could it be that a possible intact BTB domain in the klhl40b deletion/ truncation vs the truncated BTB in the A isoform is significant here (Figure 1A-B, S1A-B)? Could it be that the klhl40a truncation is generating a dominant negative effect? This should be explored and at least discussed.

2. Although it is clear that morphological effects are observed in KLHL40a deletion muscle, these are largely undefined. Are there morphology changes to the SR-ER and Golgi at 1.5 months in KO, particularly since proteome changes are already observed at that stage? Key to understanding the effects of KLHL40a deletion are the morphological changes depicted by EM in figure 2. A-H and these will truly benefit from better analysis. The authors should utilize organelle markers and immunofluorescence, in order to define the identity and morphology of affected organelles in control and KO states (in particular the SR-ER, some shown in Figure 6 but deserves more markers and better analysis and importantly also Golgi compartments). Is the ER dilated? What is the extent of Golgi fragmentation? The presentation of the EM images will also greatly benefit from the enlargements of individual examples of organelle images and better morphological descriptions. Similar analysis will benefit Figure 5D in particular following Golgi morphology thus defining if it is the increased Sar1a levels that lead to Golgi fragmentation.

3. It has been demonstrated that the activities of biosynthetic secretion are adjusted during developmental processes by unfolded protein response (UPR) signaling. It would be an informative and important addition to the work if the authors examine their data and expand their analysis (transcriptome and proteome) to describe potential UPR activity (selected targets) both during development and in particular in KLHL40a KO where UPR is likely induced (added to Figure 4). This should provide an important control to the study and current conclusions that suggest perturbed secretion from the ER (the alternative where there is no UPR activation suggesting perhaps a subtle secretion defect or a very selective defect in collagen secretion, although the latter may not agree with Golgi fragmentation).

4. The authors describe the regulation of Sar1a yet this isoform-specific Sar1 effect is not discussed or examined. This is not trivial as Sar1b is selectively implicated in supporting traffic of large lipid particles as was also established in zebrafish by the Knapik group and was also implicated in the packaging of large-size cargo from the ER in early work from the Schekman group. Is it the ratio between the two Sar1 isoforms that are perturbed to give the observed collagen and ECM secretion/deposition defects? Is it just the excess of Sar1 proteins regardless of the isoform nature that is leading to ER tubulation and inhibition of general secretion? Given the authors finding and the body of previous work on Sar1a and Sar1b, it would markedly benefit this work to address these points. The authors should also better define when Sar1a or all isoforms are followed in their experiments.

5. The results showing elevated levels of Sar1a in KLHL40a deletion are very interesting. The authors further show that the levels of other COPII subunits are mostly reduced (Figure 5 A-C) but provide an incomplete analysis. It is known that COPII proteins including inner and outer layer components undergo ubiquitination and de-ubiquitylation cycles. The authors should specifically examine if overall Sec23 and Sec24 levels are co-regulated as these form Sec23 -Sec24 complexes. Similarly, and more importantly, the analysis ignores Sec13 and Sec31 subunits of the coat outer layer and this should be explored. In accordance, key regulatory proteins including TANGO1, cTAGE5, and in particular Sec12 should also be highlighted (all is in the data and if interesting, perhaps deserve western blot analysis).

6. It is interesting to note that the COPII protein Sec31 was previously shown to be developmentally regulated by Cul3-KLHL12 mediated Sec31 ubiquitylation to support collagen secretion. It is possible that Cul3-driven tight control over COPII subunit levels and ratios is generally at play here. Indeed, the authors suggest that levels of multiple ubiquitin ligases and deubiquitylases are modified leading to the observed overall complex effects on the proteome. However special focus should be given to known regulators of COPII. Is KLHL12 expression modified in development here? Is it modified by KLHL40a deletion? The authors should examine the potential adjustments made by Cul3-KLHL12 that may control Sec31 levels. This analysis will provide a required overview of Cul3-regulated secretion proposed here and would suggest that selective control of the levels of individual COPII components (and not just Sar1a) may direct coat composition, a potentially highly significant outcome with broad implications.

7. The authors localize Sar1 in control and KLHL40a deletion mutants (Figure 6) and in KLHL40 nm patient samples (Figure 9A) but should complement this with analysis with a similar analysis of other COPII components and of ER exit site proteins such as TANGO1 or Sec16. This analysis is required to define the effects of Sar1a levels on COPII assembly and ER exit site assembly. The analysis is critical for the work, generating some mechanistic insights on the roles of KLHL40a in regulating secretion, and monitoring outcomes of elevated Sar1a levels on its downstream effectors.

Reviewer #2 (Recommendations for the authors):

The phenotypes look like they could have arisen from alterations in several cellular processes, from mitochondrial activity to secretion. It seems challenging to be able to distinguish between the relative contributions from each of these processes. It would be instructive to test whether autophagy is affected as well, and include a section in the discussion to highlight as a caveat, the diverse pathways that might contribute to the phenotype.

The authors show a clear lack of correlation between the proteome and transcriptome, but in order to focus on protein ubiquitylation of specific proteins, the authors need to present a lack of correlation, between the proteomic and RNAseq data for the individual proteins studied in the manuscript.

Altered Golgi architecture and an increase in vesicular structures could arise from a number of reasons. In order to argue for a direct role of secretion, the authors need to carry out a more direct secretion assay to confirm the altered secretory rate.

From figure 7, it is challenging to distinguish between secreted (extracellular) collagen between control and mutant fish. Could the authors visualise and quantify extracellular collagen specifically? Imaging collagen fibres should show only extracellular collagen.

It is unclear why the levels of total collagen appear very different, is this due to altered collagen degradation? The authors need to show whether the observed increase in collagen is KLHL40 arises, in part, from reduced intracellular collagen degradation.

Co-localise collagen and other cellular structures including the ER, to see where collagen is retained.

The authors should test whether the excess vesicular structures in the KLHL40 fish are secretory or degradative, for instance, do they have LC3?

The authors have used C2C12 cells that have been used for immunoprecipitation analysis, the Golgi apparatus, and assay for collagen secretion.

Figure 9 is unclear. The control and affected tissues look to be from different areas, could this be resolved?

There are related published results on CUL3 and KLHL12, which could play a related role in cellular responses to collagen secretion. The results in this manuscript could suggest a common theme of proteostatic control over collagen and COPII degradation, rather than secretion, thereby resolving confusing data on the KLHL12/Cul3 model. Could the authors include a brief discussion about links between these two studies?

Reviewer #3 (Recommendations for the authors):

The manuscript could benefit from more experiments and quantitation. Specifically, The finding that KLH40A knockout leads to smaller fish is important but whether this is solely related to a skeletal muscle phenotype is less clear. For example, where are these orthologs expressed (which tissues?)? What other morphologic issues are present? The image just shows a smaller fish. I would also like to see a Kaplan Meier curve for the death of the mutant fish.

Figure 2 relies only on EM images and would be helpful to have other pathologic images to get a sense of muscle morphology. Specifically, statements such as myopathic phenotype are not clearly demonstrated. regarding sarcomere width, how does one control for the contractile state? In addition, an assay of mitochondrial function would be helpful seahorse or evidence of oxidative damage, changes in ETS components. The EM descriptions of the Golgi and basal lamina seem qualitative, can we quantify them?

The large amount of data from the RNAseq, ubiquitin remnant, and proteome data is laudable and described in extensive detail with minimal validation and lots of speculation. Is there a decrease/increase of total proteins on Western blot, the most interesting subset is the proteins that are enriched but decreased in ubiquitin remnants?

The focus on Sar1a is premature in vivo and figure 5 shows vesiculation is not convincing and needs quantitation. The altered localization is challenging to assess in Figure 6 because of the difference in morphology of the sections.

Figure 7 needs many controls for the biochemical experiments. Control vectors with just V5 or Flag. total input for the KLHL40 flag.

Control vectors/proteins for 7B are needed to assess promoter competition that can occur when expressing two different proteins at different levels. 7C could benefit from a stability assay rather than steady-state levels of the proteins. Figure 7F needs controls without the addition of the Kelch protein and just GST lysates. The ubiquitination could be occurring just with the addition of contaminating proteins.

Figure 9 needs disease controls. Hard to know if sar1 is upregulated in many myopathies. the same is true for ECM defects.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Dynamic regulation of interorganelle communication by ubiquitylation controls skeletal muscle development and disease onset" for further consideration by eLife. Your revised article has been evaluated by Vivek Malhotra (Senior Editor) and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

1. It is suggested that the provided western blots (Figure S1) negate the potential for dominant negative effects of expressed truncated KLHL40 fragments, but can the antibody listed (directed to the C-terminus) recognize the remaining and potentially expressed N-terminal fragments in edited fish where the C-terminus is truncated (Figure 1A, Figure S1E)? Alternatively, other experiments that may address the potential (or lack of) dominant negative effects (rescue experiments, RNAi with similar phenotypes or otherwise) can be discussed.

2. It is not clear how ECM- collagen is defined in the added experiments (Figure 8 collagen panel) and how it is differentiated from intracellular accumulated procollagen 1. The procollagen antibody listed in methods may not stain mature collagen fibers. This should be addressed and the intracellular localization of accumulated procollagen 1 in KO conditions should be defined.

3. The outcomes of the RUSH-traffic experiments in KO cells are interesting and suggest a delay in PC traffic in KO vs WT cells with comparable traffic rates at later time points. Corresponding images depicting the quantified data should be added for clarity. The effects on the traffic of other cargoes should be examined to define if we are looking at a selective effect on large-size cargo such as procollagen or a more global effect on traffic from the ER. In the methods and Results sections, the authors state that an added cargo is being expressed (ST-mCherry) and its traffic (or that of another small-size cargo) from the ER in WT and KO cells should be compared to that of procollagen. This would be highly valuable and important.

4. It is suggested that the stabilization of Sar1 with MG132 negates the possibility of promoter competition in the transfection-based Sar1 expression level analysis yet there is a visible decrease in Sar1 levels in both conditions (+/- MG123). Simple normalized quantification of the presented data (Figure 7 B and C) should be added to define the selective contributions of proteasome-mediated degradation.

5. The added IF of PDI (IF, Figure 6) supports the EM analysis showing a substantive dilation of the ER in KLHL40 deleted cells. Yet the added cursory analysis of UPR (omics of Bip calnexin, CHOP, Ero1) suggests that UPR is not induced but no alternative mechanism for ER-expansion is discussed. A closer look at the UPR (phospho-Ire1, phospho-PERK, spliced XBP1) should address if subtle effects on secretion from the ER lead to a more limited UPR outcome. The added new data showing mostly intact Golgi (as opposed to previous conclusions) may support this alternative.

6. In Figure 6 S1, increase in levels of autophagy markers may (or may not) report on autophagy levels as this may also depend on other factors. Similarly, added blots provided for LC3 I/II ratios require ratio quantification. Overall, the limitations of both analyses should be discussed.

7. The definitions of TANGO1 localization as "large" or bigger "vesicles" should be clarified (both for TANGO1 and COPII staining) with TANGO1 likely not marking vesicles but instead marking ER exit sites (ERES). Increased Sar1a may lead in some undefined manner to a decrease in ERES assembly and this point should be discussed (see below).

8. The added info on Sar1b levels (not modified), suggests that indeed the ratio between Sar1a and Sar1b is modified in KLHL40 KO. If so, co-expression of Sar1a and b may nullify the effects of Sar1a overexpression. This point can be addressed and discussed, given the selective roles of Sar1 proteins in procollagen traffic.

9. The discussion on KLHL12-Cul3 should highlight both potential contributions in procollagen degradation in lysosomes and on traffic, given recent work on degradation of procollagen in KLHL12 expressing cells from the Kim and Lippincott-Schwartz labs. Also, note the recent paper by Jinoh Kim and colleagues (MBoC) showing the involvement of maintaining KLHL12 in collagen levels rather than secretion

Other points:

1. KLHL40 is defined in the text as a "negative regulator" of traffic, yet its deletion inhibits procollagen traffic. A more appropriate definition may be simply "regulator".

2. In the abstract and in the title the authors use the term "interorganelle communication" to describe effects on multiple organelles (mitochondria ER) but this term is not clear as the work describes developmental control over organelle morphology which may be independent (Mitochondria, ER) with one explored functional outcome (biosynthetic secretion or intracellular traffic rather than communication).

https://doi.org/10.7554/eLife.81966.sa1

Author response

Essential revisions:

1) Overall myopathic effects and changes on a cellular level to the morphology of organelles and sorting sites including ER exit sites and Golgi need further analysis and quantification.

We thank the reviewers for raising this critical point and apologize for not being clear previously. We have acquired new electron microscopy (EM) data, reanalyzed the previous EM data, and performed detailed immunofluorescence analysis for a number of antibodies (PDI, Sar1a, Sec23, Sec24, Sec31, Tango1, and Golga2). As many of these antibodies previously failed to resolve well in frozen sections, we have performed these studies in isolated myofibers from control and klhl40a mutant zebrafish. While Sec31 and Sec24 antibodies didn’t work in zebrafish by IF, the results from all other antibodies are provided in Figure 6. The following are the findings from these studies (Please see new Figure 6 and main text pages 15 and 16):

  1. Overall Myopathic effects: klhl40a mutant fish exhibit leaner bodies with reduced sarcomere size and decreased swimming behavior suggesting structural and functional deficits in skeletal muscle. Previous studies in zebrafish have shown that the expression of klhl40a and klhl40b is restricted to skeletal muscle (References 23 and 26). Analysis of Gtex data in human tissues also showed that KLHL40 is specifically expressed in skeletal muscle (https://gtexportal.org/home/gene/KLHL40). Weak skeletal muscle can result in secondary morphological defects in other interacting tissues, such as scoliosis. No signs of scoliosis or other morphological defects were observed in klhl40a mutants, as shown in Figure 1A, suggesting klhl40a deficiency results in a primary myopathy (added to text on page 6).

  2. klhl40a mutant muscle exhibits an accumulation of membrane-bound structures near the ER-SR region. To understand the origin of these membrane-bound structures, we performed immunofluorescence analysis in control and klhl40a mutant myofibers. These studies showed a significantly increased number of PDI-positive foci in the mutant muscle suggesting the increased number of membrane-bound structures are derived from ER. We also observed increased Sar1a immunoreactivity in the mutant myofibers co-localized with PDI. Moreover, the ultrastructure of these membrane-bound structures shows a similarity to ER and lacks the organization of CopII-coated vesicles shown previously (Barlowe et al., 1994 and; Matsuoka et al., 1998). This suggests that the absence of Klhl40a results in an increased number of Sar1A-positive ER-derived vesicles that do not form CopII vesicles (text added to page 16).

  3. In normal conditions, Sec23/24 and Sec13/31 assemble with Sar1A the ER exit sites to form CopII vesicles. Similarly, Tango1, Sec23, and Sec24 assemble at Sar1A ER exit sites to form large cargo containing CopII vesicles. We observed a decrease in Sec23 and Tango1 positive foci in klhl40a deficient myofibers compared to controls. This suggests that increased Sar1A level does not result in the formation of productive CopII vesicles (text added to page 16).

  4. Immunofluorescence with Golga2 antibody revealed normal Golgi structure in most myofibers (63 ± 14%) with varying amounts of GOLGA2-positive aggregates in other myofibers. Based on our previous EM results, we previously concluded that the Golgi apparatus appeared to be fragmented in the klhl40a mutant muscle. However, with Golga2 specific antibody in a large number of myofibers, we did not observe any extensive defect in the Golgi apparatus (New Figure 6 and text on pages 15-16).

2) Analysis of biosynthetic secretion and in particular collagen mobilization in the secretory pathway or degradation that may lead to ECM deposition defects are required.

We have created human KLHL40 knockout muscle cell lines and performed intracellular collagen trafficking assays to investigate collagen mobilization in KLHL40 deficiency. We analyzed the ER-Golgi trafficking of procollagens by the RUSH assay and identified delayed ER-Golgi trafficking of procollagens in KLHL40 knockout cells compared to control myoblasts (Figure 8, pages 18-19). We also analyzed the proteomics data for changes in the UPR or autophagy proteins, and no significant changes were observed between the control and klhl40a mutant muscle. Finally, western blot analysis with the LC3 antibody did not reveal any differences between the control and klhl40a mutant muscle, indicating a defect in secretory pathways and not in the degradation process. These results are added as Figure 6—figure supplement 1 and text on page 16.

3) Further analysis of the findings on Sar1 with added controls and analysis of all COPII and ER exit site components is required to provide a clear view of the basis for secretion defects that may lead to disease onset.

Please see response 1 to the essential revision. In addition, to evaluate if SAR1A overexpression results in abnormal membrane-bound structures in skeletal muscle, we performed immunofluorescence in myofibers from control and SAR1A overexpressing zebrafish (5 dpf). SAR1A overexpression increased foci in skeletal muscle that co-stained with both PDI and SAR1A, further proving that increased amounts of SAR1A contribute to abnormal membrane-bound structures (Added as Figure 5D and text on page 15).

4) Attention should be given to the complexity of effects leading to the overall disease phenotypes.

Thanks for raising this point. We have reorganized our data to reflect the major disease phenotypes and associated molecular changes. Our morphological studies point to complexity in the skeletal muscle defects in KLHL40 deficiency with reduced sarcomere size, mitochondrial changes, and abnormal vesicle trafficking. Our proteomics studies have identified reduced Ub-sites enrichment and increased abundance of their cognate proteins as potential KLHL40-CUL3 targets for vesicle trafficking (Sar1a), glycolytic (Pkm, Aldo), and early sarcomeric proteins (Ttn, Tnnt2, and Nckipsd).

Abnormal upregulation of early sarcomeric protein in mature skeletal muscle is associated with structural defects in sarcomeres, and increased rates of glycolysis in differentiated skeletal muscle lead to atrophy. While no changes in the ubiquitylation of mitochondrial proteins were seen, this dysregulation of mitochondrial protein could be a response to altered glycolysis or structural abnormalities in skeletal muscle. Finally, we have performed a detailed analysis of the vesicle trafficking process in skeletal muscle and show that alteration of this process results in ECM defects through reduced trafficking of collagens.

While comprehensive studies are needed to define the specific role of each of these processes in skeletal muscle defects in KLHL40 deficiency, these studies show these altered molecular processes contribute to specific pathological changes observed in Klhl40 deficiency in skeletal muscle. (Discussed on pages 22-23).

Reviewer #1 (Recommendations for the authors):

Specific points.

1. Although the authors demonstrate that selective effects are derived from the deletion of KLHL40a, both a and b forms are expressed to almost the same levels (Figure S1A-E). What is the explanation for the differential effect? Is it localization? Is it a lack of different activities of the b isoform and why? Could it be that a possible intact BTB domain in the klhl40b deletion/ truncation vs the truncated BTB in the A isoform is significant here (Figure 1A-B, S1A-B)? Could it be that the klhl40a truncation is generating a dominant negative effect? This should be explored and at least discussed.

Genome duplication events occurred in the ancestor of all vertebrates. After duplication, one of the duplicates most frequently loses its function, and the other retains all the original functions. This redundancy in zebrafish klhl40 genes is evident from a previous study that showed a similar expression pattern of klhl40a and klhl40b in zebrafish skeletal muscle but different severity of muscle phenotypes of the morphant fish. While klhl40a knockdown by morpholinos resulted in a severe phenotype, klhl40b morphant fish exhibited a relatively mild phenotype suggesting functional redundancy similar to our studies (Ravenscroft et al., 2013). klhl40a and klhl40b mutations are predicted to encode for truncated proteins. However, western blot analysis (Figure 1 and Figure 1—figure supplement 1) showed a complete absence of klhl40 proteins in the mutant fish lines. It thus ruled out a dominant negative effect of the truncated protein. These changes are incorporated in the main text on pages 5 and 6.

2. Although it is clear that morphological effects are observed in KLHL40a deletion muscle, these are largely undefined. Are there morphology changes to the SR-ER and Golgi at 1.5 months in KO, particularly since proteome changes are already observed at that stage?

Thank you for asking this critical question. We have performed a detailed analysis of klhl40a and control zebrafish skeletal muscle at 1.5 months (added Figure 2—figure supplement 1 and text on page 7). Evaluation of the ultrastructure by electron microscopy did not reveal any significant changes in sarcomeres or SR/ER region (Figure 2SA). We looked specifically at the ER and Golgi membranes by immunofluorescence in cultured myofibers isolated from klhl40a and control zebrafish skeletal muscle at 1.5 months. No changes were observed in the localization of the ER and Golgi markers (Figure 2—figure supplement 1). Finally, we have looked at different vesicles in the control and klhl40a myofibers to define specific defects. We included that data in updated figure 6 (Please see responses 1 and 3 to the editor’s questions).

Key to understanding the effects of KLHL40a deletion are the morphological changes depicted by EM in figure 2. A-H and these will truly benefit from better analysis. The authors should utilize organelle markers and immunofluorescence, in order to define the identity and morphology of affected organelles in control and KO states (in particular the SR-ER, some shown in Figure 6 but deserves more markers and better analysis and importantly also Golgi compartments). Is the ER dilated? What is the extent of Golgi fragmentation? The presentation of the EM images will also greatly benefit from the enlargements of individual examples of organelle images and better morphological descriptions. Similar analysis will benefit Figure 5D in particular following Golgi morphology thus defining if it is the increased Sar1a levels that lead to Golgi fragmentation.

Thanks for these excellent suggestions, and we regret omitting this critical information in the previous version of the manuscript. We have performed a detailed analysis of klhl40a and control muscle by different vesicle markers by immunofluorescence. To improve the sensitivity of immunofluorescence and rule out any freezing artifacts, we have performed these analyses in the cultured myofibers from klhl40a and control zebrafish (3 months). No changes in the ER and Golgi are observed at 1.5 months in the Klhl40a mutants compared to the control (Figure 2S). The ER appear to be dilated by electron microscopy analysis, and quantification of this is now added to Figure 2. After careful reanalysis, we do not see large-scale changes in the Golgi in Klhl40a mutants. We have analyzed a large number of myofibers (n=10-12, 3 replicates) and found normal Golgi architecture in most of the myofibers. In a small percentage of myofibers, we observed Golga2 positive aggregates. These changes are quantified and added to figure 6, and the text is added to pages 15 and 16.

3. It has been demonstrated that the activities of biosynthetic secretion are adjusted during developmental processes by unfolded protein response (UPR) signaling. It would be an informative and important addition to the work if the authors examine their data and expand their analysis (transcriptome and proteome) to describe potential UPR activity (selected targets) both during development and in particular in KLHL40a KO where UPR is likely induced (added to Figure 4). This should provide an important control to the study and current conclusions that suggest perturbed secretion from the ER (the alternative where there is no UPR activation suggesting perhaps a subtle secretion defect or a very selective defect in collagen secretion, although the latter may not agree with Golgi fragmentation).

Thanks for asking about this critical point. We have examined the UPR activity by evaluating differential levels of Bip, Calnexin, Ero1, Irea1, CHOP, and PERK and did not identify any significant differences between control and klhl40a mutants at 1.5 and 3 months of age by proteome and RNA seq (3 months). We observed a defect in procollagen trafficking from ER to Golgi (Figure 8, text page 19). Moreover, new IF revealed that Golgi structure is preserved in most of the myofibers (Figure 6, text pages 15-16). Future studies may be able to provide if the vesicle trafficking defects in klhl40a deficiency specifically affect collagens or other proteins.

4. The authors describe the regulation of Sar1a yet this isoform-specific Sar1 effect is not discussed or examined. This is not trivial as Sar1b is selectively implicated in supporting traffic of large lipid particles as was also established in zebrafish by the Knapik group and was also implicated in the packaging of large-size cargo from the ER in early work from the Schekman group. Is it the ratio between the two Sar1 isoforms that are perturbed to give the observed collagen and ECM secretion/deposition defects? Is it just the excess of Sar1 proteins regardless of the isoform nature that is leading to ER tubulation and inhibition of general secretion? Given the authors finding and the body of previous work on Sar1a and Sar1b, it would markedly benefit this work to address these points. The authors should also better define when Sar1a or all isoforms are followed in their experiments.

This is an important point and we apologize for not addressing this earlier. While a significant increase was observed in Sar1a protein in klhl40a knockout muscle, no significant changes were observed in Sar1b, indicating the defects observed are isoform-specific. As no differences were observed in Sar1b, all the experiments were performed with reagents specific to Sar1a (cDNA for ubiquitination and cell culture studies and antibody for Sar1a protein). The following text to reflect this is added to the main text on page 24 “While the Sar1a level was increased in the Klhl40a mutant, no differences were observed in closely related family member Sar1b in Klhl40a deficiency (Table S1). SAR1B regulates chylomicron trafficking and is 90% identical to SAR1A; these proteins exhibit different biochemical properties in COPII assembly and do not compensate for each other function in vivo (58-60). Therefore, in Klhl40a deficiency, altered levels of Sar1a contribute to procollagen trafficking defects in skeletal muscle”.

5. The results showing elevated levels of Sar1a in KLHL40a deletion are very interesting. The authors further show that the levels of other COPII subunits are mostly reduced (Figure 5 A-C) but provide an incomplete analysis. It is known that COPII proteins including inner and outer layer components undergo ubiquitination and de-ubiquitylation cycles. The authors should specifically examine if overall Sec23 and Sec24 levels are co-regulated as these form Sec23 -Sec24 complexes. Similarly, and more importantly, the analysis ignores Sec13 and Sec31 subunits of the coat outer layer and this should be explored. In accordance, key regulatory proteins including TANGO1, cTAGE5, and in particular Sec12 should also be highlighted (all is in the data and if interesting, perhaps deserve western blot analysis).

We have performed a detailed analysis of COPII subunits and identified that inner layer proteins sec23 and sec24 are downregulated in KLhl40a deficiency, suggesting that they are co-regulated in klhl40a deficiency. No significant changes were observed in the ER-resident protein Sec12 or outer COPII membrane proteins Sec13 and Sec31 and TANGO1 and cTAGE5 levels (added to text on page 14). While Sec31 and Sec24 antibodies didn’t work by IF or western, we have performed IF to quantify the number of vesicles formed by these proteins and provided the data in Figure 6 ( text pages 15-16).

6. It is interesting to note that the COPII protein Sec31 was previously shown to be developmentally regulated by Cul3-KLHL12 mediated Sec31 ubiquitylation to support collagen secretion. It is possible that Cul3-driven tight control over COPII subunit levels and ratios is generally at play here. Indeed, the authors suggest that levels of multiple ubiquitin ligases and deubiquitylases are modified leading to the observed overall complex effects on the proteome. However special focus should be given to known regulators of COPII. Is KLHL12 expression modified in development here? Is it modified by KLHL40a deletion? The authors should examine the potential adjustments made by Cul3-KLHL12 that may control Sec31 levels. This analysis will provide a required overview of Cul3-regulated secretion proposed here and would suggest that selective control of the levels of individual COPII components (and not just Sar1a) may direct coat composition, a potentially highly significant outcome with broad implications.

Reviewer 1 has raised a very interesting point and we apologize for not addressing this earlier. Gene expression analysis showed that KLHL12 is expressed at very low levels in skeletal muscle compared to other cells and tissue types (http://gtexportal.org). KLHL12 and Sec31 expression was also not modified in Klhl40a deficiency at the protein level (Table S1). This suggests that there are tissue-specific roles of CUL3 where CUL3 interacts with different Kelch proteins and regulates different aspects of secretory pathways in different tissues. We have added this point to the discussion on page 24 in the following text.

“CUL3-dependent ubiquitylation is previously shown to dynamically regulate the trafficking of large COPII carriers (56). KLHL12, another substrate-specific adapter for CUL3 E3 ubiquitin ligase forms a complex with CUL3 which ubiquitylates SEC31 leading to an increase in COPII vesicle size to accommodate large procollagen molecules for secretion (56). KLHL12 is expressed at very low levels in normal skeletal muscle compared to other cells and tissue types (http://gtexportal.org). We did not identify any differential changes in Klhl12 and the target protein Sec31 in Klhl40a deficiency. CUL3 interacts with many Kelch proteins in a tissue specific context and therefore, may regulate specific aspects of secretory pathways in response to different stimuli”.

7. The authors localize Sar1 in control and KLHL40a deletion mutants (Figure 6) and in KLHL40 nm patient samples (Figure 9A) but should complement this with analysis with a similar analysis of other COPII components and of ER exit site proteins such as TANGO1 or Sec16. This analysis is required to define the effects of Sar1a levels on COPII assembly and ER exit site assembly. The analysis is critical for the work, generating some mechanistic insights on the roles of KLHL40a in regulating secretion, and monitoring outcomes of elevated Sar1a levels on its downstream effectors.

Thanks for raising this critical point. Please see response 1 to the editor’s questions. We have performed detailed experiments and analysis to address these points and have included the updated data in figures 2S, 5, and 6 and text on pages 15-16.

Reviewer #2 (Recommendations for the authors):

The phenotypes look like they could have arisen from alterations in several cellular processes, from mitochondrial activity to secretion. It seems challenging to be able to distinguish between the relative contributions from each of these processes. It would be instructive to test whether autophagy is affected as well, and include a section in the discussion to highlight as a caveat, the diverse pathways that might contribute to the phenotype.

We agree with the reviewer that the relative contribution of different processes on disease phenotype may be challenging to assess. Please see the answer to point 4 compiled by the editors. We also evaluated autophagy markers in the proteomics data and performed a western blot with LC3B antibody and found no differences in klhl40a deficiency. We have added Figure 6—figure supplement 1 to show these data and included the following description in the text (Page 16).

“As dysregulation of autophagy is associated with vesicle trafficking, we examined different autophagy markers (ATD5, ATG16L1, ATG4B, ATG9A, beclin, LC3B, LAMP1, and LAMP2) in proteomic data or by western blot (LC3) (Figure 6—figure supplement 1). We did not observe any altered autophagy markers in Klhl40 deficiency, suggesting autophagy is not changed in the mutant muscle at the stages analyzed.”

The authors show a clear lack of correlation between the proteome and transcriptome, but in order to focus on protein ubiquitylation of specific proteins, the authors need to present a lack of correlation, between the proteomic and RNAseq data for the individual proteins studied in the manuscript.

We analyzed the RNA-seq data for all the key proteins studied in this work and found no correlation with the RNA-seq data. The following sentence is added to the text (pages 21-22) to reflect his “In addition, no changes were detected at the transcriptome levels for proteins that showed significant changes in the ubiquitylation and their cognate proteins suggesting that regulatory processes such as post-transcription, post-translation, and protein degradation impact protein abundance after mRNA is made (41).”

Altered Golgi architecture and an increase in vesicular structures could arise from a number of reasons. In order to argue for a direct role of secretion, the authors need to carry out a more direct secretion assay to confirm the altered secretory rate.

To address this concern, we generated human KLHL40 knockout primary myoblast cell lines for performing collagen trafficking assay. Our results showed that rate of intracellular collagen trafficking is significantly reduced in the KLHL40 KO cells in comparison to control myoblasts. These results are now added to Figure 8 and text on pages 18-19.

From figure 7, it is challenging to distinguish between secreted (extracellular) collagen between control and mutant fish. Could the authors visualise and quantify extracellular collagen specifically? Imaging collagen fibres should show only extracellular collagen.

We have performed staining for mature collagens localized in the ECM region that show reduced ECM collagen staining in the klhl40a muscle. We have added a new panel of the staining to Figure 8 and added the following text in the manuscript (Page 19):

“Collagen staining is also reduced in the klhl40a mutant muscle with some myofibers also lacking collagen in the ECM region compared to the control muscle (Figure 8 arrow)”.

It is unclear why the levels of total collagen appear very different, is this due to altered collagen degradation? The authors need to show whether the observed increase in collagen is KLHL40 arises, in part, from reduced intracellular collagen degradation.

Co-localise collagen and other cellular structures including the ER, to see where collagen is retained.

The authors should test whether the excess vesicular structures in the KLHL40 fish are secretory or degradative, for instance, do they have LC3?

The authors have used C2C12 cells that have been used for immunoprecipitation analysis, the Golgi apparatus, and assay for collagen secretion.

Thanks for raising this critical point. We have performed additional experiments and have shown that increased procollagen accumulation in muscle fibers is due to reduced trafficking of collagens from ER (Figure 8). We could not resolve the precise localization of procollagens in the skeletal muscle by immunofluorescence. While future immunogold-EM studies may be able to resolve these differences, these results suggest that collagen is retained in the ER. We have also performed western blot analysis with LC3 (added as Figure 6—figure supplement-1 and text on page 16) and did not observe any significant differences in control versus mutants suggesting the excess vesicles in the Klhl40a mutant are potentially secretive and not degradative.

Figure 9 is unclear. The control and affected tissues look to be from different areas, could this be resolved?

For congenital muscle diseases, it is extremely difficult to obtain healthy control muscle biopsies from neonatal babies, and therefore, adult muscle biopsies are used as the control.

There are related published results on CUL3 and KLHL12, which could play a related role in cellular responses to collagen secretion. The results in this manuscript could suggest a common theme of proteostatic control over collagen and COPII degradation, rather than secretion, thereby resolving confusing data on the KLHL12/Cul3 model. Could the authors include a brief discussion about links between these two studies?

Thanks for suggesting including this important detail we neglected to address previously. We have now addressed this point and added the following information to the discussion (Page 24):

“CUL3-dependent ubiquitylation is shown to dynamically regulate the trafficking of large COPII carriers. KLHL12-CUL3 complex ubiquitylates SEC31 leading to an increase in COPII vesicle size to accommodate procollagen molecules for secretion (56). USP8 negatively regulates this process by deubiquitylation of SEC31 and inhibiting the formation of large COPII carriers (57). KLHL12 is expressed at very low levels in normal skeletal muscle compared to other cells and tissue types (http://gtexportal.org). KLHL12 and the target protein Sec31 are also not differentially regulated in KLHL40a deficiency. CUL3 interacts with many Kelch proteins in in a tissue-specific context and may regulate specific aspects of secretory pathways in response to different stimuli.”

Reviewer #3 (Recommendations for the authors):

The manuscript could benefit from more experiments and quantitation. Specifically, The finding that KLH40A knockout leads to smaller fish is important but whether this is solely related to a skeletal muscle phenotype is less clear. For example, where are these orthologs expressed (which tissues?)? What other morphologic issues are present? The image just shows a smaller fish. I would also like to see a Kaplan Meier curve for the death of the mutant fish.

Previous studies in zebrafish have shown that the expression of Klhl40a and Klhl40b is restricted to skeletal muscle (References 23 and 26). Analysis of Gtex data in human tissues also showed that KLHL40 is specifically expressed in skeletal muscle (https://gtexportal.org/home/gene/KLHL40). Weak skeletal muscle can result in secondary morphological defects in other interacting tissues, such as scoliosis. No signs of scoliosis or other morphological defects were observed (added this sentence to page 6). We have added the Kaplan Meier Curve for the death of the mutant fish (Figure 1E and text to pages 6-7).

Figure 2 relies only on EM images and would be helpful to have other pathologic images to get a sense of muscle morphology. Specifically, statements such as myopathic phenotype are not clearly demonstrated. regarding sarcomere width, how does one control for the contractile state? In addition, an assay of mitochondrial function would be helpful seahorse or evidence of oxidative damage, changes in ETS components. The EM descriptions of the Golgi and basal lamina seem qualitative, can we quantify them?

Thanks for raising this critical point regarding the specific changes to different compartments in skeletal muscle. We have included IF data to show different ER, COPII, and Golgi structures in control and klhl40a mutant myofibers and quantified them (Please see response 1 to editors).

We agree with reviewer 2 that artifacts in the skeletal muscle contracting state can lead to a misinterpretation of results. We dissect one animal at a time, collect 50-100 mg of skeletal muscle biopsy, and fix it immediately to prevent any contracting state artifacts which can result from the uneven fixation of thick samples. Moreover, blinded sample processing, image analysis, and quantification of muscle biopsies from multiple fish tissues are performed to examine whether the differences between the control and experimental groups are reproducible and statistically significant. These details are added to the Electron microscopy section in methods on page 39. Finally, a comprehensive detailed metabolic analysis (e.g. seahorse and potentially mass spectroscopy) will be required to understand the detailed effects of changes in the glycolytic and mitochondrial proteins. While this may provide additional insights, is beyond the scope of the current work.

The large amount of data from the RNAseq, ubiquitin remnant, and proteome data is laudable and described in extensive detail with minimal validation and lots of speculation. Is there a decrease/increase of total proteins on Western blot, the most interesting subset is the proteins that are enriched but decreased in ubiquitin remnants?

The focus on Sar1a is premature in vivo and figure 5 shows vesiculation is not convincing and needs quantitation. The altered localization is challenging to assess in Figure 6 because of the difference in morphology of the sections.

We have identified several potential pathways that are altered in klhl40a deficiency and kept the focus of this work on the sar1a-mediated vesicle trafficking. We are pursuing many novel candidates identified from these studies. However, that involves extensive characterization and generation of novel reagents, which is beyond the scope of the current study. We have now performed additional immunofluorescence on Sar1a and associated vesicles in klhl40a KO and have added that, as updated in figure 6. We have also performed IF analysis on zebrafish in figure 5d which showed increased Sar1a staining is colocalized with the ER marker PDI in zebrafish myofibers.

Figure 7 needs many controls for the biochemical experiments. Control vectors with just V5 or Flag. total input for the KLHL40 flag.

Control vectors/proteins for 7B are needed to assess promoter competition that can occur when expressing two different proteins at different levels. 7C could benefit from a stability assay rather than steady-state levels of the proteins. Figure 7F needs controls without the addition of the Kelch protein and just GST lysates. The ubiquitination could be occurring just with the addition of contaminating proteins.

We have performed additional experiments or added controls for previously missing experiments in Figure 7. Figure 7A: We have added the controls for control V5 and FLAG plasmids and the total input for KLHl40 FLAG as the new figure 7A.

Figure 7B: We have performed an additional experiment to assess the effect of promoter competition of KLHL40FLAG and Sar1V5 vectors. We observed that the expression of KLHL40FLAG was the same in the absence and the presence of Sar1V5. As observed previously, the expression of Sar1V5 was reduced in the presence of KLHL40FLAG. This suggests a lack of promoter competition between Sar1V5 and KLHL40 FLAG (Added as Figure 7—figure supplement 1 and text on page 17).

Figure 7C: This experiment aims to determine the effect of the ubiquitin-mediated proteasomal degradation of SAR1A in the presence of KLHL40, and we show that MG132 prevents this degradation. This suggests that the reduced levels of SAR1A in the presence of KLHL40 are caused by ubiquitin-mediated proteasomal degradation, and a cycloheximide-based half-life stability assay may not be necessary.

Figure 7F: We have performed the control experiment regarding the in vitro ubiquitination reaction for SAR1A in the presence of GST. We found that GST itself did not result in the ubiquitination of SAR1A, suggesting that the ubiquitination of SAR1A requires KLHL40. This figure has been added as a supplemental figure (added Figure 7—figure supplement 1 and text on page 18).

Figure 9 needs disease controls. Hard to know if sar1 is upregulated in many myopathies. the same is true for ECM defects.

Thanks for asking this question. We have obtained skeletal muscle samples from centronuclear myopathy patients (RYR1 and MTM1 mutations) and quantified the amount of total SAR1A protein by western blotting (added as Figure 7—figure supplement 1). The amount of SAR1A was significantly upregulated in KLHL40 deficiency but not in other myopathies. This suggests that SAR1A upregulation is specific to KLHL40related myopathy. Due to the limited amount of muscle biopsies, we could not evaluate ECM structure. Previous studies have shown normal ECM structures in RYR1 and MTM centronuclear myopathies. Therefore, SAR1A is not associated with disease pathology in other forms of myopathies. This text is added to the text on page 20.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

1. It is suggested that the provided western blots (Figure S1) negate the potential for dominant negative effects of expressed truncated KLHL40 fragments, but can the antibody listed (directed to the C-terminus) recognize the remaining and potentially expressed N-terminal fragments in edited fish where the C-terminus is truncated (Figure 1A, Figure S1E)? Alternatively, other experiments that may address the potential (or lack of) dominant negative effects (rescue experiments, RNAi with similar phenotypes or otherwise) can be discussed.

We have repeated western blots with an antibody that recognizes 1-50 amino acids of the N-terminal of the KLHL40 proteins (upstream of the mutations in the different klhl40 fish lines). Western blot analysis showed the absence of the klhl40 mutant protein in the klhl40ab mutant and a reduction in protein amounts in klhl40a or klhl40b mutants compared to the wildtype control. Updated westerns are added to Figure S1.

2. It is not clear how ECM- collagen is defined in the added experiments (Figure 8 collagen panel) and how it is differentiated from intracellular accumulated procollagen 1. The procollagen antibody listed in methods may not stain mature collagen fibers. This should be addressed and the intracellular localization of accumulated procollagen 1 in KO conditions should be defined.

To distinguish between the ECM at the periphery of myofibers and the intracellular region, we have added the panels with the sarcomeric α-actin antibody (Figure 8), and the corresponding text is added on pages 19-20. Procollagens in the mutant muscle are predominantly intracellular, as seen with actinin co-labeling. Precise localization of procollagens, while an important question, will need high-resolution imaging such as immunoEM and extensive optimization with multiple antibodies and is the beyond the scope of current work.

3. The outcomes of the RUSH-traffic experiments in KO cells are interesting and suggest a delay in PC traffic in KO vs WT cells with comparable traffic rates at later time points. Corresponding images depicting the quantified data should be added for clarity. The effects on the traffic of other cargoes should be examined to define if we are looking at a selective effect on large-size cargo such as procollagen or a more global effect on traffic from the ER. In the methods and Results sections, the authors state that an added cargo is being expressed (ST-mCherry) and its traffic (or that of another small-size cargo) from the ER in WT and KO cells should be compared to that of procollagen. This would be highly valuable and important.

We have added the images depicting the quantified data for the RUSH assay for procollagen trafficking in Figure 8B. We have also added data on the trafficking of the small cargo, sialyltransferase-mCherry, which was not significantly affected in KLHL40 KO myoblasts compared to WT control (Figure 8C). This is also added to the main text (page 20). These data suggest that KLHL40 deficiency primarily affects the trafficking of large cargo such as procollagens.

4. It is suggested that the stabilization of Sar1 with MG132 negates the possibility of promoter competition in the transfection-based Sar1 expression level analysis yet there is a visible decrease in Sar1 levels in both conditions (+/- MG123). Simple normalized quantification of the presented data (Figure 7 B and C) should be added to define the selective contributions of proteasome-mediated degradation.

Thanks we have added the quantification of western blots in Figures 7 B and C, which shows that the difference in SAR1A level in the presence of MG135 at different amounts of KLHL40 is not significant.

5. The added IF of PDI (IF, Figure 6) supports the EM analysis showing a substantive dilation of the ER in KLHL40 deleted cells. Yet the added cursory analysis of UPR (omics of Bip calnexin, CHOP, Ero1) suggests that UPR is not induced but no alternative mechanism for ER-expansion is discussed. A closer look at the UPR (phospho-Ire1, phospho-PERK, spliced XBP1) should address if subtle effects on secretion from the ER lead to a more limited UPR outcome. The added new data showing mostly intact Golgi (as opposed to previous conclusions) may support this alternative.

While we didn’t see the differences in the protein levels of the ER stress markers, we performed additional experiments to analyze phospho-Ire1, phosphor-PERK, and spliced XBP1 (Figure 6S D-E). This analysis showed lack of the spliced XBP1 in the Klhl40a mutant muscle. Western blot analysis revealed no differences in the PERK and phosphor-PERK levels between control and Klhl40a deficient skeletal muscle. We also tried two different antibodies for Phospho-Ire1alpha that showed the highest similarity with the zebrafish protein, but they failed to yield specific signals by western blots. Our proteome analysis, as well as RT-PCR assay for XBP1 and western blot analysis of phospho-PERK, showed no evidence of the UPR activation in vivo skeletal muscle. These results are added in Figure 6S, and the text is added in the main manuscript on pages 17 and 26.

6. In Figure 6 S1, increase in levels of autophagy markers may (or may not) report on autophagy levels as this may also depend on other factors. Similarly, added blots provided for LC3 I/II ratios require ratio quantification. Overall, the limitations of both analyses should be discussed.

We have added quantification of the LC3II-LC3-1 ratio in Figure 6S and updated the corresponding figure legend.

7. The definitions of TANGO1 localization as "large" or bigger "vesicles" should be clarified (both for TANGO1 and COPII staining) with TANGO1 likely not marking vesicles but instead marking ER exit sites (ERES). Increased Sar1a may lead in some undefined manner to a decrease in ERES assembly and this point should be discussed (see below).

We have clarified TANGO1 localization as ERES (Main manuscript, page 16) and discussed changes in ERES sites (Main manuscript, page 25) as follows.

“Klhl40a deficient skeletal muscle showed an increased number of enlarged ERES sites compared to controls, while the overall number of ERES sites was reduced in the mutant muscle. ERES functions as an inter-organelle transport apparatus that actively modulates its shape and size while directing diverse cargo types to Golgi and increases in size during active transport. As Klhl40a deficiency resulted in reduced procollagen trafficking from ER, this increase in the size of ERES sites could be a compensatory mechanism to enhance the secretory flux. Sar1 is associated with the ERES sites and regulates the formation of COPII vesicles. Despite the presence of increased Sar1A levels in Klhl40a deficient myofibers, reduced or limited amounts of other proteins, such as Sec16b and COPII proteins in the mutant myofibers may underlie a reduced number of ERES sites and COPII vesicles.”

8. The added info on Sar1b levels (not modified), suggests that indeed the ratio between Sar1a and Sar1b is modified in KLHL40 KO. If so, co-expression of Sar1a and b may nullify the effects of Sar1a overexpression. This point can be addressed and discussed, given the selective roles of Sar1 proteins in procollagen traffic.

We agree that the ratio between the Sar1a and Sar1b is modified in Klhl40a deficiency and have added this to the discussion. Sar1a and Sar1b have been shown to regulate the trafficking of common proteins such as cardiac sodium channel Nav1.5 and procollagens, and therefore, co-expression of Sar1a and b may compensate for the deficiency of either of these Sar1 proteins. However, in Klhl40a deficiency, increased amounts of Sar1a resulted in the formation of abnormal ER-derived membrane bound-structures and, therefore, restoring the normal amount of Sar1a (than overexpression of Sar1b) may be able to restore the procollagen trafficking defects. This updated discussion is added to pages 25-26 in the main manuscript as follows:

“While the Sar1a level was increased in the klhl40a mutant, no differences were observed in closely related family member Sar1b in Klhl40a deficiency (Supplementary file 1) showing a decrease in Sar1a and Sar1b ratio. SAR1A is 90% identical to SAR1B and these proteins exhibit overlapping and unique functions with different biochemical properties in COPII assembly (60-62). While SAR1B specifically regulates chylomicron trafficking in the small intestine, both proteins are required for the trafficking of cardiac sodium channel Nav1.5 protein and efficient ER export of procollagens and may be able to compensate for each other function for the trafficking of common proteins (62-65). In Klhl40a deficiency, increased amounts of Sar1a resulted in the formation of abnormal ER-derived membrane-bound structures and decreased procollagens trafficking; therefore, downregulation of Sar1a levels in skeletal muscle may be able to restore the procollagen trafficking defects.”

9. The discussion on KLHL12-Cul3 should highlight both potential contributions in procollagen degradation in lysosomes and on traffic, given recent work on degradation of procollagen in KLHL12 expressing cells from the Kim and Lippincott-Schwartz labs. Also, note the recent paper by Jinoh Kim and colleagues (MBoC) showing the involvement of maintaining KLHL12 in collagen levels rather than secretion

Thanks, this is an important point, and we regret excluding this from the previous discussion. We have addressed these points in the main manuscript (Discussion, Pages 24-25). as follows.

“KLHL12, another substrate-specific adapter for CUL3 E3 ubiquitin ligase forms a complex with CUL3 which ubiquitylates SEC31 leading to an increase in COPII vesicle size to accommodate large procollagen molecules for secretion in mouse embryonic stem cells (mESC) (56). While most procollagens are trafficked through the secretory pathway, a subset is directed towards lysosomal degradation to remove excess procollagen from cells through the autophagy pathway (57). Recent studies in skin fibroblasts have shown that the CUL3-KLHL12 complex is involved in the routing of procollagens to lysosomes to regulate intracellular collagen levels (58). Moreover, inhibition of CUL3 neddylation which is critical for the ubiquitylation activity still led to the formation of large COPII vesicles by the CUL3-KLHL12 complex, which is required for the secretion of procollagens. As CUL3-mediated ubiquitylation also regulates KLHL12 protein stability, further studies are needed to understand the ubiquitylation-dependent and independent roles of CUL3-KLHL12 complex on procollagens secretion in different cellular contexts and physiological conditions. KLHL12 is expressed at very low levels in normal skeletal muscle compared to other cells and tissue types (http://gtexportal.org). We did not identify any differential changes in Klhl12 and the target protein Sec31 in Klhl40a deficiency. Moreover, no changes in the protein levels of autophagy markers were observed in Klhl40a deficiency at the disease states examined. CUL3 interacts with many Kelch proteins in a tissue-specific context and therefore, may regulate specific aspects of secretory and degradative pathways in response to different stimuli and disease states.”

Other points:

1. KLHL40 is defined in the text as a "negative regulator" of traffic, yet its deletion inhibits procollagen traffic. A more appropriate definition may be simply "regulator".

We have removed “negative” and now kept this as “regulator”.

2. In the abstract and in the title the authors use the term "interorganelle communication" to describe effects on multiple organelles (mitochondria ER) but this term is not clear as the work describes developmental control over organelle morphology which may be independent (Mitochondria, ER) with one explored functional outcome (biosynthetic secretion or intracellular traffic rather than communication).

Inter-organelle communication is used to describe the communication between different membrane compartments in protein trafficking and is clarified in the main text (page 4).

https://doi.org/10.7554/eLife.81966.sa2

Article and author information

Author details

  1. Arian Mansur

    Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  2. Remi Joseph

    Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    Contribution
    Data curation, Validation, Investigation, Methodology
    Competing interests
    No competing interests declared
  3. Euri S Kim

    Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    Contribution
    Data curation, Investigation, Methodology
    Competing interests
    No competing interests declared
  4. Pierre M Jean-Beltran

    Proteomics Platform, Broad Institute of MIT and Harvard, Cambridge, United States
    Contribution
    Data curation, Supervision, Validation, Investigation, Methodology, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5106-0992
  5. Namrata D Udeshi

    Proteomics Platform, Broad Institute of MIT and Harvard, Cambridge, United States
    Contribution
    Data curation, Formal analysis, Supervision, Investigation, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
  6. Cadence Pearce

    Proteomics Platform, Broad Institute of MIT and Harvard, Cambridge, United States
    Contribution
    Data curation, Validation, Investigation
    Competing interests
    No competing interests declared
  7. Hanjie Jiang

    1. Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    2. Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Boston, United States
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  8. Reina Iwase

    1. Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    2. Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, United States
    Contribution
    Data curation, Validation, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3703-2511
  9. Miroslav P Milev

    Department of Biology, Concordia University of Edmonton, Montreal, Canada
    Contribution
    Data curation, Formal analysis, Methodology
    Competing interests
    No competing interests declared
  10. Hashem A Almousa

    Department of Biology, Concordia University of Edmonton, Montreal, Canada
    Contribution
    Data curation, Investigation, Methodology
    Competing interests
    No competing interests declared
  11. Elyshia McNamara

    Faculty of Health and Medical Sciences, Centre of Medical Research, Harry Perkins Institute of Medical Research, University of Western Australia, Perth, Australia
    Contribution
    Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  12. Jeffrey Widrick

    Division of Genetics, Boston Children’s Hospital, Harvard Medical School, Boston, United States
    Contribution
    Data curation, Investigation, Methodology
    Competing interests
    No competing interests declared
  13. Claudio Perez

    Department of Anesthesiology, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    Contribution
    Investigation, Methodology
    Competing interests
    No competing interests declared
  14. Gianina Ravenscroft

    Faculty of Health and Medical Sciences, Centre of Medical Research, Harry Perkins Institute of Medical Research, University of Western Australia, Perth, Australia
    Contribution
    Supervision, Validation, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  15. Michael Sacher

    1. Department of Biology, Concordia University of Edmonton, Montreal, Canada
    2. Department of Anatomy and Cell Biology, McGill University, Montreal, Canada
    Contribution
    Formal analysis, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
  16. Philip A Cole

    Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    Contribution
    Supervision, Funding acquisition, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6873-7824
  17. Steven A Carr

    Proteomics Platform, Broad Institute of MIT and Harvard, Cambridge, United States
    Contribution
    Supervision, Investigation, Methodology, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
  18. Vandana A Gupta

    Division of Genetics, Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Methodology, Writing - original draft, Project administration, Writing – review and editing
    For correspondence
    vgupta@research.bwh.harvard.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4057-8451

Funding

A Foundation Building Strength

  • Vandana A Gupta

National Institute of Arthritis and Musculoskeletal and Skin Diseases (R56AR077017)

  • Vandana A Gupta

National Institutes of Health (R37GM62437)

  • Philip A Cole

National Cancer Institute (R01CA74305)

  • Philip A Cole

Brigham and Women's Hospital

  • Vandana A Gupta

National Heart, Lung, and Blood Institute (F32HL154711)

  • Pierre M Jean-Beltran

National Health and Medical Research Council (APP2002640)

  • Gianina Ravenscroft

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Dr. Nigel Laing for critically reading this manuscript and providing valuable suggestions. We also thank Dr. Michael Lawlor and Stacy Crossette (Congenital Muscle Disease Tissue Repository) for providing the centronuclear patients' muscle biopsies. This work was supported by NIH R56AR077017 (VAG), R37GM62437 and R01CA74305 (PAC), A Foundation Building Strength grant and Innovation Evergreen Fund Award (VAG), and NIH F32HL154711 (PMJB). GR is supported by an Australian NHMRC EL2 Investigator Grant (APP2007769). This work is also supported by an NHMRC Ideas Grant to GR and NL (APP2002640). The DSHB antibody (8c8) developed by (Hausen and Gawantka) was obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and maintained at the University of Iowa, Department of Biology, Iowa City, IA 52241.

Ethics

Human Research Ethics Committee of the University of Western Australia (RA/4/20/1008). Written informed consent was provided by all families.

Zebrafish were maintained and bred using standard methods as described (Westerfield,2000). All experiments and procedures were approved by the Institutional Animal Care and Use Committee at Brigham and Women's Hospital. (2016000304).

Senior Editor

  1. Vivek Malhotra, Barcelona Institute for Science and Technology, Spain

Reviewing Editor

  1. Meir Aridor, University of Pittsburgh, United States

Version history

  1. Received: July 18, 2022
  2. Preprint posted: July 22, 2022 (view preprint)
  3. Accepted: June 16, 2023
  4. Accepted Manuscript published: July 11, 2023 (version 1)
  5. Version of Record published: July 19, 2023 (version 2)

Copyright

© 2023, Mansur et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Arian Mansur
  2. Remi Joseph
  3. Euri S Kim
  4. Pierre M Jean-Beltran
  5. Namrata D Udeshi
  6. Cadence Pearce
  7. Hanjie Jiang
  8. Reina Iwase
  9. Miroslav P Milev
  10. Hashem A Almousa
  11. Elyshia McNamara
  12. Jeffrey Widrick
  13. Claudio Perez
  14. Gianina Ravenscroft
  15. Michael Sacher
  16. Philip A Cole
  17. Steven A Carr
  18. Vandana A Gupta
(2023)
Dynamic regulation of inter-organelle communication by ubiquitylation controls skeletal muscle development and disease onset
eLife 12:e81966.
https://doi.org/10.7554/eLife.81966

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https://doi.org/10.7554/eLife.81966

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