Loss of mitochondrial protein CHCHD10 in skeletal muscle causes neuromuscular junction impairment (original) (raw)

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

Department of Neurobiology

, Key laboratory of Medical Neurobiology of Zhejiang Province, School of Medicine, Zhejiang University, Zhejiang, China 310058

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, School of Medicine, Zhejiang University, Zhejiang, China 310003

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The First Affiliated Hospital

, Institute of Translational Medicine, School of Medicine, Zhejiang University, Zhejiang, China 310003

Department of Neurobiology

, Key laboratory of Medical Neurobiology of Zhejiang Province, School of Medicine, Zhejiang University, Zhejiang, China 310058

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11 June 2019

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Yatao Xiao, Jianmin Zhang, Xiaoqiu Shu, Lei Bai, Wentao Xu, Ailian Wang, Aizhong Chen, Wen-Yo Tu, Jianwen Wang, Kejing Zhang, Benyan Luo, Chengyong Shen, Loss of mitochondrial protein CHCHD10 in skeletal muscle causes neuromuscular junction impairment, Human Molecular Genetics, Volume 29, Issue 11, 1 June 2020, Pages 1784–1796, https://doi.org/10.1093/hmg/ddz154
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Abstract

The neuromuscular junction (NMJ) is a synapse between motoneurons and skeletal muscles to control motor behavior. Acetylcholine receptors (AChRs) are restricted at the synaptic region for proper neurotransmission. Mutations in the mitochondrial CHCHD10 protein have been identified in multiple neuromuscular disorders; however, the physiological roles of CHCHD10 at NMJs remain elusive. Here, we report that CHCHD10 is highly expressed at the postsynapse of NMJs in skeletal muscles. Muscle conditional knockout CHCHD10 mice showed motor defects, abnormal neuromuscular transmission and NMJ structure. Mechanistically, we found that mitochondrial CHCHD10 is required for ATP production, which facilitates AChR expression and promotes agrin-induced AChR clustering. Importantly, ATP could effectively rescue the reduction of AChR clusters in the CHCHD10-ablated muscles. Our study elucidates a novel physiological role of CHCHD10 at the peripheral synapse. It suggests that mitochondria dysfunction contributes to neuromuscular pathogenesis.

Introduction

The neuromuscular junction (NMJ) is a peripheral synaptic connection between the motoneuron and skeletal muscle, controlling body movement. NMJ defects cause multiple neuromuscular disorders such as myasthenia gravis and amyotrophic lateral sclerosis (ALS) (1,2). Acetylcholine receptors (AChRs) are restricted at the synaptic region (density: >10 000 μm−2), compared with those in the non-synaptic region (density: <10 μm−2) (2). The agrin signal is well known to cluster the AChR proteins at the postsynapse, which is critical for NMJ development and maintenance (3–6). Interestingly, AChR mRNA is also highly enriched at the synaptic region, suggesting that in addition to agrin signal, there are other mechanisms that activate AChR transcription for NMJ assembly (2,7).

Mitochondria play important roles in the ATP synthesis, chromatin remodeling, intracellular calcium homeostasis and cell apoptosis (8,9). Mitochondria are enriched at NMJs, including at the presynapse (motor nerve terminals) and postsynapse (skeletal muscles) (10,11). Presynaptic mitochondria play roles in actin assembly, synaptic vesicle transportation and release (10). However, the roles of postsynaptic mitochondria remain largely elusive. Overexpression of mitochondrial uncoupling protein UCP1 in skeletal muscle caused NMJ destabilization and motoneuron degeneration (12). Overexpressing PGC1α in skeletal muscles enhanced mitochondria biogenesis, AChR clusters and motor performance (13,14). These reports indicate that muscle mitochondria regulate AChR clustering and NMJ function.

CHCHD10 (coiled-coil-helix-coiled-coil-helix domain containing 10) is a mitochondrial intermembrane protein that is enriched at cristae junctions (15,16). To date, at least 30 heterozygous mutations in the CHCHD10 gene have been identified in human patients, especially affecting neuromuscular system such as ALS, late-onset spinal motor neuropathy, Charcot–Marie–Tooth and mitochondrial myopathy (16–18). CHCHD10 is indicated to play a regulatory role in the maintenance of mitochondrial structure and function (19). Silencing of CHCHD10 or CHCHD10 mutation leads to a decrease in the ATP level, cytochrome C oxidase complex activity and respiratory capacity (15,19–22). The physiological functions of CHCHD10 in motor system in vivo are elusive. Loss of har-1, a CHCHD10 homology gene in Caenorhabditis elegans, showed impaired movement and a shortened lifespan (22). Injection of morpholinos targeting CHCHD10 into zebrafish caused motility deficit (23). Whether and how CHCHD10 physiologically regulates the peripheral synapse remains largely unknown.

Here, we report that CHCHD10 locates at postsynaptic NMJs. Loss of CHCHD10 in muscle caused motor defects, neurotransmission impairment and abnormal NMJ structure. Mechanistically, we found that CHCHD10 is required for ATP production at NMJs by promoting AChRs gene expression. ATP could rescue the NMJ defects in CHCHD10-deleted muscles. Our study elucidates a novel role of CHCHD10 in regulating NMJ integrity and suggests that postsynaptic mitochondria dysfunction might contribute to neuromuscular pathogenesis.

Results

Postsynaptic location of CHCHD10 at NMJs

The NMJ is a tripartite synapse consisting of muscle fibers, Schwann cells and motoneuron terminals (24). We isolated spinal cord, sciatic nerve and skeletal muscle from 2-month-old mice to examine CHCHD10 expression. Consistent with previous reports (20), we found that CHCHD10 (~15 kD) was highly expressed in skeletal muscles. Unexpectedly, there was very low expression of CHCHD10 in the spinal cord and sciatic nerve (Fig. 1A), suggesting that muscle CHCHD10 might play a predominant role at the peripheral synapse. We further examined CHCHD10 expression in the skeletal muscles of different aged mice and found that CHCHD10 was gradually increased after birth and reached a peak after 2 weeks (Fig. 1B), a time-window of NMJ maturation in mice (25).

Figure 1

CHCHD10 loss in skeletal muscle causes motor defects. (A) Spinal cord, sciatic nerve and skeletal muscle from P60 mice were collected for immunoblotting. GAPDH was set as the loading control. SC, spinal cord; SN, sciatic nerve; SM, skeletal muscle. (B) Gastrocnemius samples from different aged mice were subjected to immunoblotting. GAPDH was set as the loading control. (C) Diagram of the isolation of the synaptic region and non-synaptic region from the hemi-diaphragm. (D) Immunoblotting of CHCHD10 in the synaptic region and non-synaptic region. LRP4 was used as a positive control of synaptic protein. GAPDH was used as a loading control. Syn, synaptic region; Non-syn, non-synaptic region. (E) Relative protein and mRNA levels of CHCHD10 in the synaptic region and non-synaptic region. (F) Cross sections of the gastrocnemius were immunostained with anti-CHCHD10 (green) and AChR (indicated by R-BTX; red). Nuclei were stained with DAPI (blue). The yellow arrow indicates co-localization between CHCHD10 and AChR clusters. The white arrow indicates the extra-synapse location of CHCHD10. The boxed area in the left bottom is enlarged in the right bottom. (G) Diagram of the construction of CHCHD10 floxp mice. (H) Immunoblotting of CHCHD10 in the gastrocnemius and brain of HSA-CHCHD10−/− mice (P60). (I) Reduced grip strength in HSA-CHCHD10−/− mice. n = 13 mice per group; t-test; ***P < 0.001. (J) Beam walking test showed that HSA-CHCHD10−/− mice need more time to traverse the beam. n = 9 mice per group; t-test; *P < 0.05. (K) Vertical pole test showed the mutant mice need more time to reach the base of the pole. n = 9 mice per group; t-test; *P < 0.05.

To check whether CHCHD10 is expressed in the synaptic region in skeletal muscles, we isolated synaptic and non-synaptic regions of the thin diaphragm in 2-month-old mice (Fig. 1C) (26). LRP4 was used as a positive control to indicate the proper separation of synaptic region. We found that the CHCHD10 protein level was higher in the synaptic region than that in the non-synaptic region (2.00 ± 0.16 in the synaptic region vs. 1.00 ± 0.09 in the non-synaptic region, P < 0.05; Fig. 1D and E). The enrichment of CHCHD10 expression in the synaptic region occurs at the transcription level, as evidenced by an abundance of CHCHD10 mRNA (3.92 ± 0.44 in the synaptic region vs. 1.02 ± 0.02 in the non-synaptic region, P < 0.05; Fig. 1E). To directly observe the location of CHCHD10 at the synapse, cross sections of the gastrocnemius were immunostained with anti-CHCHD10. R-BTX was used to label the postsynaptic AChR clusters. As shown in Fig. 1F, most of CHCHD10 protein was co-localized with or nearby R-BTX positive AChR clusters (yellow arrow; Fig. 1F), indicating the synaptic location of CHCHD10. Together, our data suggested that CHCHD10 expression is enriched at the postsynapse of NMJs.

CHCHD10 deletion in skeletal muscle causes motor defects and neurotransmission impairment.

To explore CHCHD10 functions in skeletal muscles, we generated CHCHD10-floxp mice (see Material and Methods for details; Fig. 1G) and crossed them with HSA-Cre mice, which express the Cre gene under the control of the human skeletal actin (HSA) promoter. Cre starts to be expressed from embryonic day (E) 9.5 in the myotomal regions of somites and is detectable in almost all skeletal muscle fibers in P0 (postnatal 0) day mice (27,28). Immunoblot analysis showed that most of CHCHD10 proteins were lost in skeletal muscle homogenates of HSA-Cre; CHCHD10f/f (HSA-CHCHD10−/−) mice. The reduction was specific for muscles and was not observed in other tissues such as brain tissues (Fig. 1H). Homozygote conditional knockout animals were born with Mendelian ratios and were viable. To examine whether muscle functions were affected in HSA-CHCHD10−/− mice, we measured grip strength and found it was reduced from 100 ± 1.05% in the control to 79.67 ± 2.48% in the mutant (P < 0.001, Fig. 1I). Beam walking test and vertical pole test were performed to analyze motor behaviors. We found that the mutant mice need more time to traverse the beam (4.44 ± 0.21 s in the control while 6.19 ± 0.64 s in the control, n = 9 per group, P < 0.05; Fig. 1J) and to reach the base of the pore (7.58 ± 1.17 s in the control and 13.87 ± 2.04 s in the mutant, n = 9 per group, P < 0.05; Fig. 1K). Taken together, these behavior data suggest that impaired muscle functions in HSA-CHCHD10−/− mice.

To investigate the underlying mechanisms of muscle weakness in HSA-CHCHD10−/− mice, we used electromyography to determine whether neuromuscular transmission is impaired. Compound muscle action potentials (CMAPs) were measured in the gastrocnemius from adult mice (P60) in response to repetitive nerve stimuli in the sciatic nerve (29,30). In control littermates, the CMAP amplitude showed little change after 10 consecutive nerve stimuli at different frequencies (Fig. 2A and B). By contrast, CMAPs in the mutant muscles cannot be sustained (Fig. 2A and 2B). In details, the CMAP amplitude was decreased significantly from the 2nd stimulus at 30 Hz stimulation; the decrement of CMAPs at the 10th stimulus was ~13.71 ± 2.12% (n = 4 per group, P < 0.05; Fig. 2C). The reduction of the CMAP amplitude in HSA-CHCHD10−/− muscles was frequency dependent and showed a significant reduction from 20 to 40 Hz (Fig. 2D), indicating a progressive loss of successful neuromuscular transmission after repeated stimulation. Together, these observations suggested that muscle CHCHD10 is required for normal neurotransmission between motoneuron and skeletal muscle fibers.

Figure 2

Muscle CHCHD10 ablation impaired neuromuscular transmission. (A) CMAPs were recorded in the gastrocnemius from P60 mice in response to a train of 10 submaximal stimuli at different frequencies. Representative CMAP traces between two genotypes at the 1st, 2nd and 10th stimuli. (B) Ten CMAP traces were stacked in succession for better comparison. With continuous stimulations, CMAP amplitudes were reduced in HSA-CHCHD10−/− muscles. Ten representative CMAP traces are shown as stacked in succession for better comparison. (C) Reduced CMAP amplitudes in HSA-CHCHD10−/− muscles at 10 stimulations at 30 Hz. n = 4 mice per group; t-test; *P < 0.05, **P < 0.01. (D) Reduction of the CMAP amplitude is stimulation frequency dependent. n = 4 mice per group; *P < 0.05, **P < 0.01. (E) Representative mEPP traces from P60 mice. mEPPs were recorded from hemidiaphragms. Traces underlined on the left are enlarged on the right. (F) Cumulative probability plot of mEPP amplitude distribution. (G) Reduced mEPP amplitudes in HSA-CHCHD10−/− mice. n = 4 mice per group, 5–6 muscle fibers per mouse; t-test; ***P < 0.001.

The decline in membrane potential could lead to a decline in excitability of muscle fibers, which would mimic the observed CMAP decline. We checked membrane potential but did not find significant differences between wild type and CHCHD10 mutants (Supplementary Material, Fig. 1B). To investigate whether the neurotransmission deficits result from pre- and/or postsynaptic impairment, we further measured miniature endplate potentials (mEPPs), events generated by spontaneous vesicle release (Fig. 2E). Electrophysiological recordings at diaphragm NMJs revealed that mEPP amplitudes were reduced by 29% compared with controls (1.140 ± 0.067 mV in the control vs. 0.806 ± 0.051 mV in the mutant, n = 5 per group, P < 0.001; Fig. 2F and G). These results were suggestive of reduced AChR density at the post-junctional membrane of HSA-CHCHD10−/− mice. Taken together, our findings are suggestive of postsynaptic deficits in neuromuscular transmission when the muscle CHCHD10 gene is ablated.

Muscle CHCHD10 is required for NMJ structural integrity.

The structural integrity of NMJs ensures proper neurotransmission from presynapse to postsynapse (4). To explain the neurotransmission defects and muscle weakness in HSA-CHCHD10−/− mutants, whole-mount gastrocnemius staining was performed to identify whether NMJ structural changes occur in the mutants. Z-serial images were collected using a confocal microscope and were collapsed into single images. We did not find any observable change in NMJs at the embryonic stage (E16.5) including NMJ size, number or endplate width (data not shown). This might be due to the relatively low levels of CHCHD10 before birth. In control muscles, AChR clusters were exhibited as plaques at P0 and matured at P60 with a characteristic pretzel-like morphology with complex continuous branches (Fig. 3A). The AChR cluster size and intensity was reduced at P0 (size: 91.68 ± 2.28% in the mutant vs. 100 ± 2.57% in the control, P < 0.05; Intensity: 82.04 ± 2.68% in the mutant vs. 100 ± 3.04% in the control, P < 0.001; Fig. 3A–C). At P60, we found that the pretzel-like morphology of NMJs was much simpler. The AChR cluster size and intensity was reduced than that in control. There was ~24% reduction in size (76.12 ± 5.43% in the mutant vs. 100.0 ± 6.62% in the control, P < 0.01) and ~23% reduction in intensity. At P300, there was ~34% reduction in size and ~18.4% reduction in intensity (size: 66.08 ± 5.20% in the mutant vs. 100 ± 7.38% in the control, P < 0.001; Intensity: 81.60 ± 5.20% in the mutant vs. 100 ± 6.40% in the control, P < 0.05; Fig. 3A–C). Strikingly, AChR clusters became fragmented at P300, indicating NMJ destabilization in the mutant mice (Fig. 3A and D). These results suggest that muscle CHCHD10 is required for postsynaptic NMJ maturation and maintenance both, which is consistent with the gradual increase in CHCHD10 expression after birth (Fig. 1B).

Figure 3

CHCHD10 regulates NMJ structural integrity. (A) Gastrocnemius muscles of control and HSA-CHCHD10−/− mice (P0, P60 and P300) were stained whole-mount with R-BTX (red) to label AChR. (B) Statistical results of the AChR cluster size in (A). n = 3 mice at each age per group; t-test; *P < 0.05, ***P < 0.001. (C) Statistical results of the AChR cluster intensity in (A). n = 3 mice per group; t-test; *P < 0.05, **P < 0.01, ***P < 0.001. (D) Statistical results of AChR cluster fragmentation in (A). n = 3 mice per group; t-test; ***P < 0.001. (E) Gastrocnemius muscles were stained with R-BTX (red) to label AChR and antibodies against NF and SV2 (NF/SV2; green) to label nerve branches and terminals. The arrows indicate degenerated axons in the mutant. Arrowheads indicate the fragmented NMJs without the coverage of nerve terminal staining. (F) Statistical results of the innervation of NMJs in control and HSA-CHCHD10−/− mice. n = 3 mice per group; two-way ANOVA; **P < 0.01.

To further examine whether motoneuron axon terminals are impaired in the mutants, we used antibodies against neurofilament (NF) and SV2 to label the nerve branches and terminals (26). In control adult mice, NF/SV2-labeled motoneuron axon terminals were smooth and aligned with postsynaptic AChR clusters at NMJs (Fig. 3E). In mutant mice, multiple NMJs showed fragmentation (arrowhead; Fig. 3A and E), and some motor neuron axons showed degeneration (arrow; Fig. 3E). Partial innervation of NMJs was increased ~1.7-fold, while full innervation was reduced by ~67% in the mutant (Fig. 3F). Axonal degeneration and fragmentation of synaptic AChR clusters are also observed at endplates regenerated from muscle fiber damage (31). We found that there were some central nuclei in mutant myotubes (Supplementary Material, Fig. 2A–C), suggesting that muscle regeneration occurred in CHCHD10 mutant mice. However, we did not find muscle defects at early stage (Supplementary Material, Fig. 2A and B), suggesting that loss of CHCHD10 preferentially damages NMJs rather than muscles. Taken together, these results suggest that muscle CHCHD10 is required for NMJ structural integrity.

CHCHD10 is required for agrin-induced AChR clustering in cultured myotubes

To further explore the underlying cellular mechanisms of NMJ defects in the HSA-CHCHD10−/− mutant, we first checked endogenous CHCHD10 expression during C2C12 differentiation. We found that CHCHD10 was expressed in myotubes but was undetectable in myoblasts (Fig. 4A), consistent with the increased expression in skeletal muscles after birth (Fig. 1). To examine the endogenous role of CHCHD10, we designed sgRNA of CHCHD10 (with Cas9 cDNA in the same vector) to efficiently knockdown its expression (Fig. 4B). C2C12 cells were transfected with sgRNA of CHCHD10 and then stimulated with agrin (1 nm; 16 h), an inducer of AChR clustering and NMJ formation and maintenance (32). In non-transfected myotubes, agrin effectively promoted AChR clusters; however, in the sgRNA of CHCHD10 transfected myotubes, agrin-induced AChR clusters were significantly reduced (17.550 ± 1.066 in the control versus 7.414 ± 0.669 in the sgRNA group, P < 0.001; Fig. 4C and D).

Figure 4

CHCHD10 is required for agrin-induced AChR clustering in myotubes. (A) Immunoblotting of CHCHD10 protein in C2C12 myoblasts and myotubes. D, day. (B) Immunoblotting showed that endogenous CHCHD10 expression is knocked down by CRISPR-cas9. (C) C2C12 myoblasts were transfected with Cas9 and sgRNA of CHCHD10. Agrin-induced AChR clusters were reduced in the CHCHD10 sgRNA transfected myotubes. (D) Quantification results of AChR clusters in C. Three independent experiments were performed; t-test; ***P < 0.001. (E) Primary myotubes were cultured from CHCHD10f/f mice (P0) and were infected with Ad-GFP (Ad-Ctrl) or Ad-Cre-GFP (Ad-Cre). After 4 days, primary myotubes were treated with agrin (1 nm; 16 h) and then stained with R-BTX. Note that the agrin-induced AChR cluster (white arrow) is reduced in Ad-Cre-infected CHCHD10f/f myotubes (green) compared with that in Ad-Ctrl-infected ones (white arrow; green). The non-infected myotubes in the two groups showed no difference (yellow arrow). (F) Quantification results in E. Three independent experiments were performed; t-test; **P < 0.01.

To further verify the role of CHCHD10 in agrin-induced AChR clustering in C2C12 myotubes, we cultured primary myotubes from CHCHD10f/f pups. Primary myotubes were infected with adenovirus expressing GFP (Ad-Ctrl) or GFP-IRES-Cre (Ad-Cre) for 4 days and were stimulated with agrin followed by R-BTX staining. Virus-infected myotubes were indicated with a green fluorescence signal. As shown in Figure 4E and F, agrin-induced AChR clusters were significantly reduced in Ad-Cre-infected CHCHD10f/f myotubes (white arrow; Ad-Cre) compared with those in Ad-Ctrl-infected myotubes (white arrow; Ad-Ctrl). There was no detectable difference in the intensity of AChR clusters between Ad-Ctrl infected myotubes and non-infected ones (Fig. 4E). Taken together, our data suggested that muscle CHCHD10 is required for agrin-induced AChR clustering in a cell-autonomous manner.

CHCHD10 is required for mitochondria structure and ATP production.

CHCHD10 was originally identified as a mitochondrial cristae protein (15,20). We co-transfected Flag-CHCHD10 and mito-RFP (an indicator of mitochondria) into C2C12 cells. Immunostaining confirmed that Flag-CHCHD10 proteins are highly co-localized with mitochondria (Fig. 5A). Although accumulated evidence has indicated that CHCHD10 plays roles in mitochondria structure and function, it remains controversial in vivo (22,23,33). Thus, we used electron microscope analysis to examine mitochondria morphology in the adult diaphragm of wild-type and HSA-CHCHD10−/− mice. Consistent with the previous report (33), we found that there were many large lysosome-like vesicles (red arrow, high-dense round vesicles) in the mutant mitochondria that were rarely shown in the mitochondria in control animals (13.45 ± 1.80% in the mutant vs. 1.50 ± 0.43% in the control, P < 0.001; Fig. 5B and C), indicating active degradation of unhealthy mitochondria in the mutant. We further found that the mRNA levels of mitochondria-related transcripts including those of ATP6, Cytochrome C, PGC1α and GABPα were downregulated, suggesting the impaired mitochondrial activities (Fig. 5D).

Figure 5

CHCHD10 regulates mitochondria structure and ATP production. (A) Immunostaining of Flag-CHCHD10 in C2C12 cells. Green: Flag-CHCHD10; Red: Mito-RFP; Blue: DAPI. (B) EM analysis of mitochondria in control and HSA-CHCHD10−/− muscles (P60). The red arrows indicate the high-density round vesicles in the mutant. (C) Quantitative results in B; t-test; ***P < 0.001. (D) mRNA expression of mitochondria-related genes in two genotypes. Note that most of the mitochondria-related gene expression is reduced in the mutant. Three independent experiments were performed; t-test; **P < 0.01; ***P < 0.001. (E) Reduction of ATP levels in CHCHD10-ablated primary myotubes (Ad-Cre virus infected) and synaptic regions of muscles in HSA-CHCHD10−/− mice. At least three independent experiments were performed; t-test; *P < 0.05; ***P < 0.001. SR, synaptic region.

To further examine whether mitochondria function in skeletal muscle fibers is impaired without CHCHD10, we checked the ATP levels in Ad-Ctrl and Ad-Cre infected CHCHD10f/f primary myotubes and found that the total ATP levels were reduced by ~30% (69.92 ± 2.38% in Ad-Cre-infected myotubes and 100 ± 2.07% in the control, P < 0.001; Fig. 5E). We next asked whether CHCHD10 regulates ATP levels in synaptic regions in vivo, which were isolated as previously described (Fig. 1C). We found that ATP levels in the synaptic region were reduced by ~46% in the HSA-CHCHD10−/− compared with those in the control (control: 100 ± 4.65% vs. mutant: 54.42 ± 13.95%, n = 3, P < 0.05; Fig. 5E). Taken together, our data suggested that muscle CHCHD10 is required for normal mitochondria morphology and function at NMJs.

ATP promotes AChR expression and rescues NMJ defects in HSA-CHCHD10−/− mice

Agrin binds LRP4 to activate MuSK, thus stimulating AChR clustering in muscle cells, which is critical for NMJ development and maintenance (4–6). Having demonstrated the defects in NMJ structure and function in HSA-CHCHD10−/− mice (Figs 1–3), we next ask whether CHCHD10-mediated mitochondria function regulates agrin-induced AChR clustering. ATP was reported to enhance agrin-induced AChR clustering (34,35). As shown in Figure 6A, AChR clusters were increased in response to agrin stimulation, while pretreatment with ATP could amplify the effects of agrin-induced AChR clustering, in a dose-dependent manner (Fig. 6A and B). ATP alone showed no effect on inducing AChR clustering (Fig. 6A), but increased the R-BTX staining signal in the background, indicating enhanced expression of AChR. NaN3 is an inhibitor of complex IV in the mitochondrial respiratory chain (36), and we found that NaN3 could inhibit ATP levels (Supplementary Material, Fig. 3). It reduced agrin-induced AChR clustering in C2C12 myotubes in a dose-dependent manner (Fig. 6C). More importantly, NaN3 administration in skeletal muscles of wild-type mice impaired NMJ structure in vivo (arrowheads; Fig. 6D).

Figure 6

ATP promotes AChR expression and rescues NMJ defects in HSA-CHCHD10−/− mice. (A) C2C12 myotubes were treated with agrin (1 nm; 16 h) with or without ATP (50 μm; 16 h) and were stained with R-BTX. The arrowhead indicates agrin-induced AChR clusters. Note that ATP alone has no effect on AChR clustering but increases the BTX (bungarotoxin) staining signal. (B) Statistical results of agrin-induced AChR cluster with different doses of ATP (0, 0.5, 2.5 and 12.5 μm). Three independent experiments were performed. **P < 0.01, ***P < 0.001. (C) Statistical results of agrin-induced AChR cluster following pretreatment with different dose of NaN3 (0, 0.16, 0.8, 4 and 20 mM; 8 h). Three independent experiments were performed. **P < 0.01, ***P < 0.001. (D) Gastrocnemius muscles were stained with R-BTX to label AChR cluster after NaN3 (0.16 mm) injection. (E) Immunoblotting of pMuSK in C2C12 myotubes. C2C12 myotubes were treated with agrin (1 nm; 16 h) with or without ATP (50 μm; 16 h). GAPDH was set as the loading control. (F) Immunoblotting of cell surface AChRs proteins. HEK293T cells were co-transfected with indicated GFP and AChR subunit plasmids. After 48 h, cells were incubated with CHX (10 μg/ml) and following with or without ATP (50 μm) treatment. Cell membrane was isolated and subjected to immunoblot. ATP1A1 was set as cell membrane control. (G) The indicated gene expression in ATP-treated C2C12 myotubes (50 μm; 16 h). ***P < 0.001. (H) ATP administration rescued the reduction of NMJ size in CHCHD10-deleted muscles. (I) Statistical results of H. One-way ANOVA; n = 3; ***P < 0.001.

We speculate that ATP amplifies agrin-induced AChR clustering through three possible mechanisms: (1) promoting agrin-induced activation of the receptor kinase MuSK; (2) promoting the transportation of new synthesized AChR proteins from cytoplasm to cell membrane and (3) promoting AChR gene expression. To distinguish these possibilities, C2C12 myotubes pretreated with ATP and agrin were subjected to MuSK activation analysis. We found that ATP had no effect on agrin-elicited MuSK tyrosine phosphorylation at the Tyr755 site (Fig. 6E). Next we examined the AChR protein levels on the cell surface. HEK293T cells were transfected with AChRs subunits for 48 h. Cells were pretreated with cycloheximide (CHX) to inhibit the AChR expression and were followed by ATP treatment. HEK293T cells were subjected to stain with R-BTX (without permeabilization) to label the cell surface protein as previously described (37). Cell membrane was isolated and subjected to western blotting. Immunostaining (Supplementary Material, Fig. 4A and B) and western blotting (Fig. 6F) showed a similar amount of AChRs on cell membrane, suggesting that ATP treatment has little effect on AChR transportation. We next checked the effect of ATP on mRNA levels of AChR subunits. Real-time polymerase chain reaction (PCR) results showed that the mRNA levels of AChRα, AChRβ, AChRδ and AChRγ were significantly increased after ATP incubation (P < 0.001; Fig. 6G). This is consistent with the increase in the R-BTX staining signal but not in the number of AChR clusters when stimulated with ATP alone (Fig. 6A). Interestingly, PGC1α, COXIII and CHCHD10 genes had no response to ATP stimulation (Fig. 6G), suggesting that ATP promotes agrin-induced AChR clustering through specifically regulating AChR subunit gene expression.

Considering the reduction of ATP levels in HSA-CHCHD10−/− mice, we next examine whether ATP could rescue the NMJ defects in the mutant muscles. Ad-GFP or Ad-Cre virus was injected into gastrocnemius muscles of CHCHD10f/f mice (Fig. 6H). ATP administration could effectively rescue the NMJ defects in the CHCHD10-deleted muscles, compared with control (Fig. 6H and I). Taken together, our data support that CHCHD10-mediated ATP could promote AChR gene expression and NMJ development.

Discussion

The roles of postsynaptic mitochondria in NMJ homeostasis remain unclear. Here we reported that the mitochondria CHCHD10 protein is highly expressed at the postsynapse in skeletal muscles. Muscle conditional knockout of the CHCHD10 gene caused motor defects, impaired neuromuscular transmission and NMJ structure. Mechanistically, we found that CHCHD10 is required for mitochondrial ATP production to promote agrin-induced AChR clustering by enhancing AChR gene expression. Finally, ATP could effectively rescue NMJ defects in the CHCHD10 mutant mice. Our study first established CHCHD10 conditional knockout mice to demonstrate that muscle CHCHD10 is required for normal NMJ transmission through regulating AChR gene expression, linking mitochondria dysfunction and neuromuscular pathogenesis.

Agrin signal is well known to induce AChR clustering (at the protein level) which is critical for both NMJs development and maintenance (4–6,30). In addition, AChRs transcripts (at the mRNA level) are also highly enriched at NMJs (2,7). The underlying mechanisms of controlling the synaptic gene expression remain elusive. Our study first provides that such a molecule, CHCHD10, is enriched at the postsynaptic mitochondria to promote AChR mRNA expression, which synergistically regulates agrin-induced AChR clustering for NMJ structure and function. In mammals, each muscle fiber has multiple nuclei, but they do not evenly distribute along the whole myotube. Interestingly, some are clustered around NMJs, and these so-called synaptic nuclei have different chromatin conformation compared with extrasynaptic ones (38). Considering that ATP is important for chromatin remodeling (9,39), we speculate that CHCHD10-mediated ATP could promote chromatin remodeling in the synaptic nucleus and enable transcript factors (such as GABP) to access the AChR gene promoter at NMJs. Thus, it may explain the high enrichment of AChR transcripts at the synaptic region of NMJs in vivo (2).

To date, at least 30 mutations in the CHCHD10 gene have been recently identified in human patients. Interestingly, most of them affect motor systems such as ALS (16,18). CHCHD10 has been reported to regulate mitochondria structure and function in vitro (15,20). The physiological role of CHCHD10 in motor system in vivo remains elusive. Woo et al. (22) found impaired movement and a shortened lifespan in the CHCHD10 mutant C. elegans, and CHCHD10 shRNA could reduce Synaptophysin and Drebrin puncta in primary cultured neurons. In zebrafish, the injection of morpholinos targeting CHCHD10 caused a shortened motoneuron axon length, abnormal myofibrillar structure and motility deficit (23). In mice, Burstein et al. introduced a single nucleotide in exon 2 of CHCHD10 gene resulting in a frameshift mutation and premature stop. Although the mutant mice were viable and gross normal until 6 months of age, they did show some defects in mitochondria in different tissues: decreased ADP-stimulated respiration, increased mitochondrial iron levels and the presence of numerous lysosome-like structures in close proximity to heart mitochondria (33), which is consistent with our finding (Fig. 5). Different from homogenizing the whole muscle mass for ATP analysis (33), we speculate that mitochondria at extrasynaptic regions in myotubes could also contribute to ATP production. To solve this problem, we isolated CHCHD10-enriched synaptic regions (Fig. 1) and found that ATP levels were significantly reduced at NMJs in HSA-CHCHD10−/− mice compared with control (Fig. 5). This result suggested that CHCHD10-mediated ATP production is more restricted at NMJs and plays a fine regulative role in synaptic gene expression. We found that mEPP amplitude reduction and CMAP decline in mutant mice (Fig. 2). CMAP is a very complicated event which could be affected by many biological factors ranging from anatomical and physiological aspects to architectural properties of the muscle, such as the spreading of the neuromuscular and the fiber–tendon junctions, number of muscle fibers in each innervation unit, fiber shortening and tendon stretching (40). In our study, CHCHD10 is ablated in mitochondria of skeletal muscles including synaptic region and non-synaptic region, which might cause muscle dysfunction (Fig. 5 and Supplementary Material, Fig. S2) and contribute to the reduction of CMAPs (Fig. 2). Noticeably, NMJ defects in HSA-CHCHD10−/− were more severe with an age-dependent manner. NMJ defects in mutants were observed at the early developmental stage, and postsynaptic AChR clusters were frequently fragmented at P300, indicating roles of CHCHD10 in both NMJ development and maintenance. The disassembly of NMJs in CHCHD10-deleted mice could be caused by the slow decline of NMJs and eventually motoneuron degeneration (5), consistent with the fact that ALS patients with CHCHD10 mutations are more frequently late onset with slowly progressive symptoms (41,42). Interestingly, during our manuscript preparation, two groups recently reported that severe NMJ degeneration and motor behavior defects in CHCHD10S59L knockin mice, supporting the concept that CHCHD10 plays critical roles at NMJs (43,44).

ALS is a fatal motoneuron degenerative disease in which spinal motoneurons gradually lose control of skeletal muscles. Clinical symptoms include muscle atrophy, muscle weakness, respiratory failure and eventual death. Degeneration of NMJs is reported to occur at the initial stage of ALS, earlier than motoneuron cell body loss and clinical symptoms begin (1, 45–47). There is no effective treatment for ALS in clinic, at least in part, due to ALS patients are not being treated early enough when the motoneuron cell body is starting to degenerate (48). Our studies suggest that targeting NMJs could be an early therapeutic intervention for ALS and other neuromuscular disorders.

Material and Methods

Animals

CHCHD10-floxp mice, in which the loxp sites were inserted into the flank of exons 1 and 2 of the CHCHD10 gene, were generated using a CRISPR-cas9 strategy in the C57/B6 background (Fig. 1G, Beijing Biocytogen Co., Ltd, China). The insertion of the loxp site was verified by sequencing and southern blotting. HSA-Cre mice were purchased from The Jackson Laboratory (#006149) (28). Unless otherwise indicated, control mice were either relevant floxed or Cre littermates. All mice were raised on standard conditions and a 12 h light and 12 h dark cycle with free access to food and water. All animal experimental procedures were approved by the Institutional Animal Care and Use Committee of Zhejiang University.

Western blotting

Western blotting was performed as previously described (30). Fresh tissue and cell lysates were resolved by Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and were transferred to 0.4 μm nitrocellulose membranes. The membranes were incubated in 5% dry non-fat milk in TBS buffer for 1 h, and with the following primary antibodies for western blotting: anti-CHCHD10 (1:1000; 25671-1-AP; Proteintech), anti-pMuSK (1:1000; D151396; Sangon Biotech), anti-LRP4 (1:1000; N207/27; NeuroMab), anti-ATP1A1 (1:1000; 14 418-1-AP; Proteintech), anti-α-tubulin (1:2000; sc-23948; Santa Cruz), anti-GAPDH (1:5000; 5174S; Cell Signaling Technology) and anti-Flag (1:1000; F1804; Sigma-Aldrich). HRP-conjugated goat anti-mouse and Rabbit IgG were from Thermo (1:5000; 31430 and 31460). Immunoreactive bands were visualized using Super Signal West Femto Maximum Sensitivity Substrate (34095; Thermo). Autoradiographic films were scanned with a Bio-Red scanner (1708371; Bio-Rad ChemiDoc Touch), and captured images were analyzed with ImageJ.

Quantitative reverse transcription polymerase chain reaction (qRT-PCR) analysis

RNA was isolated with Trizol as previously described (49). Two micrograms of RNA were reverse-transcribed using the iScript cDNA synthesis kit (Bio-Rad). Real-time PCR was performed using qPCR SYBR Green Master Mix (YEASEN). The RNA levels were normalized with internal controls (GAPDH) that were assayed simultaneously on the same reaction plates. Each sample was measured in triplicate. PCR primers will be provided if requested.

Immunostaining

Whole-mount staining of muscles was performed as described previously (30). Briefly, the gastrocnemius was fixed with 4% paraformaldehyde (PFA) at 4°C overnight, rinsed with PBS and incubated with 0.1 M glycine for 20 min at room temperature, permeabilized with 1% Triton X-100 for 3 h and then blocked in blocking buffer (1% BSA, 0.5% goat serum, 0.15 M NaCl, 0.1% Triton X-100 and PBS) for 1 h, followed by incubation with the primary antibodies and rhodamine-conjugated α-bungarotoxin (R-BTX; 1:2000; Life Sciences) in the blocking buffer. The following primary antibodies were used: anti-CHCHD10 (1:500; A81854; Sigma-Aldrich or 1:500; 5671-1-AP; Proteintech), anti-neurofilament (1:1000; 2837S; Cell Signaling Technology), anti-SV2 (1:1000; SV2; Developmental Studies Hybridoma Bank) and anti-laminin (1:500; L9393; Sigma). Muscle samples were washed three times with 1% Triton X-100 in PBS. The samples were then incubated with goat anti-mouse/rabbit IgG conjugated with Alexa Fluor 488 (1:500; A-11029/A-11034; Life Sciences) or Alexa Fluor 594 (1:500; A-11029/A-11034; Life Sciences) for 1 h at room temperature. Other primary antibodies for immunostaining were anti-Flag (1:1000; F7425; Sigma-Aldrich); Z-serial images were collected using a Nikon confocal laser scanning microscope and were analyzed with ImageJ.

Hematoxylin and eosin staining

Gastrocnemius muscles were excised and dehydrated quickly in isopentane precooled with liquid nitrogen and cut with cross section (10 μm thickness) using freezing microtome (Leica CM1950). The samples were stained with hematoxylin and eosin (C0105; Beyotime Biotechnology) and imaged using upright microscope (Leica DM4000).

Grip strength measurement

Grip strength was performed as previously described (28). Briefly, mice have the tendency to grasp a horizontal metal grid while being suspended by their tails. To measure the grip strength, 2-month-old mice were subjected to grip strength by a grasping force measuring instrument (YLS-13A; Shandong Academy of Medical Sciences, China). When the mice grasp a metal grid that is connected to a force transducer, they are gently pulled horizontally to produce a force until the grip is released.

Beam walking test

The beam walking test was performed as previously reported (50,51). Briefly, mice were trained at a width of 4 cm and a length of 100 cm beam on the first day and trained at a diameter of 1 cm and a length of 100 cm beam the next day. On the third day, mice were tested at a diameter of 1 cm and a length of 100 cm beam. A bright light was projected at the start of the beam to urge the moving from the start line to the home cage of mice which was at the end point. The total traveled distance was 90 cm, and the time for the entire process was recorded. Each mouse was tested three times. The investigator was blind to mouse genotype in the process of experiments.

Vertical pole test

The vertical pole test was performed as previously reported (52). Before the test, mice were trained in the behavioral procedure for 2 days. Mice were placed facing up at the top (5 cm from the top of the pole) of a gauze-covered vertical pole with a diameter of 1 cm and a length of 50 cm. The total time was recorded when fore limbs of mice reach the base of the pole. Each mouse was tested three times. The investigator was blind to mice genotype during the experiments.

Electromyography

CMAP was measured as previously described (53). The mice were anesthetized with isoflurane (R510–22; RWD Life Science). The stimulation needle electrode (092-DMF25-S; TECA) was inserted near the sciatic nerve in the left leg thigh. The reference needle electrode was inserted near the Achilles tendon, and the recording needle electrode was inserted into the middle of the gastrocnemius muscle of the left leg. The reference and recording electrodes were connected to an Axopatch 200B amplifier (Molecular Devices). Supramaximal stimulation was applied to the sciatic nerve with trains of 10 stimuli at 1, 2, 5, 10, 20, 30 and 40 Hz (with a 30 s pause between trains). CMAPs were collected using Digidata 1322A (Molecular Devices). Peak-to-peak amplitudes were analyzed in Clampfit 9.2 (Molecular Devices).

Electrophysiological recording

Electrophysiological recording of neuromuscular transmission was performed as described previously (28,54). For the analysis of neuromuscular transmission, mice with hemidiaphragms with ribs and phrenic nerve distal endings were dissected and then pinned on Sylgard gel in oxygenated (95% O2, 5% CO2), 26–28°C Ringer’s solution (136.8 mm NaCl, 5 mm KCl, 12 mm NaHCO3, 1 mm NaH2PO4, 1 mm MgCl2, 2 mm CaCl2, 11 mm d-glucose; pH 7.3). Microelectrodes (20–40 MΩ when filled with 3 M KCl) were pierced into myotubes just lateral of the main intramuscular phrenic nerve of mouse left diaphragm which could be seen under the light microscope. Fluorescence microscopical studies showed that R-BTX labeled NMJs are strictly localized in that area (Supplementary Material, Fig. S1A). The resting membrane potentials remained stable throughout the experiment. From each hemidiaphragm, ≥5 muscle fibers were recorded for a >3 min period. The data were collected using an Axopatch 200B amplifier, digitized (10 kHz low-pass filtered) with Digidata 1322A and analyzed in Clampfit 9.2.

Muscle cell culture

C2C12 cells and primary myotubes were cultured as described previously (30). For primary myotube culture, the dissected muscle from CHCHD10f/f P0 pups was digested by 0.25% trypsin-EDTA (Invitrogen) for 30–45 min at 37°C and was plated onto 30% gelatin-coated 12 well plates in the growth medium (10% horse serum, 10% Fetal bovine serum (FBS), 1% penicillin-streptomycin, in Dulbecco’s Modified Eagle Medium (DMEM). After 24 h, the growth medium was replaced with differentiation medium (2% horse serum, 1% penicillin-streptomycin, in DMEM) for 48–72 h. Adenovirus (2 × 1010 PFU/ml, 0.5 μl) was added at the second day of culture. C2C12 myotubes or primary myotubes were treated with agrin (1 nm; 16 h) and were harvested for R-BTX staining. AChR clusters were analyzed with ImageJ.

Electron microscopy analysis

Electron microscopy (EM) analysis was performed as described previously (55). Briefly, diaphragm muscles were fixed in 2% glutaraldehyde in 0.1 M PBS at 4°C overnight, were washed 3 times with 0.1 M PBS and were further fixed in 1% osmium tetroxide in sodium cacodylate buffer (pH 7.3) for 1 h at room temperature. After washing 3 times with 0.1 M PBS, the tissues were dehydrated through a series of ethanol (50%, 70%, 90% and 100%) and were permeated 2 times in 100% acetone. After three rinses with 100% propylene oxide, the samples were embedded in plastic resin (EM-bed 812; EMSciences). Serial sections (1–2 μm thick) of tissue blocks were stained with 1% toluidine blue and then were cut into ultrathin sections, mounted on 200 mesh unsupported copper grids and stained with uranyl acetate (3% in 50% methanol) and lead citrate (2.6% lead nitrate and 3.5% sodium citrate; pH 12.0). Electron micrographs were taken using a Tecnai T10 operated at 100 KV.

Measurement of the ATP levels

ATP levels were measured using a bioluminescence detection kit (S0027; Beyotime) according to the manufacturer’s instructions. The ATP levels of muscle synaptic region or cell lysate samples were measured in a luciferase reaction based on the production of light caused by the reaction of ATP with added luciferase and D-luciferin. Briefly, tissue or cell lysate samples were incubated with the ectonucleotidase inhibitor ARL 67165 trisodium salt hydrates (A265; Sigma-Aldrich) to inhibit ATP hydrolysis. ATP was measured by a luciferase reaction in which 560 nm light was emitted when D-luciferin was converted to oxyluciferin. Luminescence was measured using a luminometer (SpectraMax M5/M5e; Molecular Devices). ATP was calculated based on a calibration curve with standard samples. The total amount of cell or tissue proteins was used for normalization.

AChR transportation assay

HEK293T cells were co-transfected with GFP, AChRα, AChRβ, AChRδ and either AChRε or AChRγ subunits (0.3:1:1:1:1) as previously described (37). After 48 h, cells were preincubated with CHX (10 μg/ml), following with ATP (50 μM) for 8 h before harvest. For immunostaining, cells were fixed with 4% PFA and stained with R-BTX without permeabilization. For immunoblotting, cells were lysed in cell lysis buffer (150 mm NaCl, 5 mm EDTA, 1 mm PMSF and cocktails of protease inhibitors in 50 mm Tris-HCl; pH 7.4) and centrifuged at 2500 rpm for 5 min to get rid of the cell debris. The supernatant was centrifuged at 13 000 rpm for 10 min. The pellet was resolved in SDS buffer as cell membrane extract.

Pharmacological experiments

C57/B6 mice (P30) were anesthetized with isoflurane and injected NaN3 (10 μl, 0.16 mm) into gastrocnemius, every other day. After 15 days, animals were sacrificed for NMJ morphological analysis.

CHCHD10f/f mice at P3 were anesthetized with isoflurane and injected with Ad-Cre-GFP or Ad-GFP (5 μl, 2 × 1010 PFU/ml) into gastrocnemius muscles of both hind legs. After 1 day recovery, ATP (5 μl, 50 μm) was injected into gastrocnemius of the right hind leg. As control, same volume of PBS was injected into the left. Injections were given twice per day at the interval of 12 h for 15 days before NMJ morphological analysis.

Statistical analysis

The data were analyzed by two-tailed paired or unpaired Student’s t-test and were expressed as means ± SEM, and GraphPad Prism 5 software was used for statistical analyses unless otherwise indicated. Statistically significant differences were indicated as *P < 0.05, **P < 0.01 and ***P < 0.001.

Acknowledgments

We are grateful to the histology core at Zhejiang University for image analysis and Shen laboratory members for their suggestions. We are grateful to Dr Lin Mei, Dr Kai Zhao, Dr Yisheng Lu, and Dr Lei Li for materials and suggestions.

Conflict of Interest statement. None declared.

Funding

This work was supported, in part, by Zhejiang Provincial Natural Science Foundation of China (LR17H090001 to S.C.), the National Key Research and Development Program of China (2017YFA0104903 to S.C.) and National Natural Science Foundation of China (31671040, 31871203 to S.C., 31701036 to K.Z.).

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© The Author(s) 2019. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oup.com

Topic:

• neuromuscular diseases
• physical activity
• mutation
• mitochondria
• agrin
• mitochondrial proteins
• skeletal muscles
• neuromuscular junction
• cholinergic receptors
• synapses
• synaptic transmission
• mice
• neuromuscular transmission
• skeletal myocytes
• postsynaptic region

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