Comparative Analysis of Muscle Atrophy During Spaceflight, Nutritional Deficiency and Disuse in the Nematode Caenorhabditis elegans

While spaceflight is becoming more common than before, the hazards spaceflight and space microgravity pose to the human body remain relatively unexplored. Astronauts experience muscle atrophy after spaceflight, but the exact reasons for this and solutions are unknown. Here, we take advantage of the nematode C. elegans to understand the effects of space microgravity on worm body wall muscle. We found that space microgravity induces muscle atrophy in C. elegans from two independent spaceflight missions. As a comparison to spaceflight-induced muscle atrophy, we assessed the effects of acute nutritional deprivation and muscle disuse on C. elegans muscle cells. We found that these two factors also induce muscle atrophy in the nematode. Finally, we identified clp-4, which encodes a calpain protease that promotes muscle atrophy. Mutants of clp-4 suppress starvation-induced muscle atrophy. Such comparative analyses of different factors causing muscle atrophy in C. elegans could provide a way to identify novel genetic factors regulating space microgravity-induced muscle atrophy.


Introduction
As more countries and companies advance spaceflight technology, space travel, habitation and tourism are becoming more common. Whereas space travel was reserved for astronauts in the past, today, private citizens can experience the space environment. However, with the increasing number of spaceflights and deeper ventures into space, understanding the changes and dangers that the space environment and microgravity pose to the human body has become more pertinent. Several studies have demonstrated that microgravity induces muscle atrophy in space [1,2], which is a serious concern for 2. Results 2.1. Muscle Atrophy in Space-Microgravity-Exposed C. elegans To better understand how space microgravity affects C. elegans muscle, we collaborated with the European Space Agency (ESA) to send C. elegans to the ISS. Young adult worms from the parental generation arrived on the ISS and remained in microgravity conditions for approximately 5 days, laying eggs to generate an F1 generation that was born aboard the ISS ( Figure 1A). Thus, the majority of the worms from these samples spent their entire life in space microgravity conditions. We analyzed a standard wild-type (WT) C. elegans strain N2 [12], and the unc-70 (e524) mutant strain, which is a weak mutant of the β-spectrin gene. unc-70 (e524) mutants were reported to have a smaller longitudinal muscle area ( Figure 2) than the wild type (WT) [21], but the overall structure of the muscle is normal, unlike that of stronger β-spectrin null mutants [22,23]. In addition, ground controls were maintained in similar conditions to those maintained for the space samples at NASA Kennedy Space Center.
bundle greatly vary in size across the longitudinal axis [21]. Thus, we set out to analyze a total of seven body wall muscle cells per animal: cells 10, 12, 14, 16, 18, 20 and 22 ( Figure  2A). An accurate measurement of muscle cell size would require measurement of the longitudinal area as well as cell height. However, since the height of the cells is much smaller than the longitudinal area [25] and accurate measurements of cell height would require electron microscopy analysis, we approximated muscle cell size by measuring only the longitudinal area of the cell, as previously shown [21].   After staining samples with phalloidin, we observed that the actin structure and sarcomere patterns of the muscle cells in the WT and unc-70 were indistinguishable in the space and control samples ( Figure 3A-D). Since the wild-type ground control samples we received were not in an adequate state for analysis and quantification, we instead generated laboratory control samples (lab WT) using the same flight conditions as the MME mission ( Figures 1A and 3A). Illustration of body wall muscles in C. elegans. (A) Dorsal lateral body wall muscles are arranged as staggered pairs. Seven dorsal lateral body wall muscle cells numbered 10, 12, 14, 16, 18, 20 and 22 are highlighted with a red border. (B) Cross-sectional illustration of the body of C. elegans. Four longitudinal bundles which consist of 23 or 24 paired muscles are located in four quadrants. Figure 3. Stained muscles of MME and NIS samples. (A-D) MME mission samples stained with Alexa Fluor™ 488 phalloidin. WT (MME) is N2; juIs76 (unc-25p::GFP), that is, a wild-type N2 strain expressing GFP transgenically in the DD/VD motor neurons. unc-70 is unc-70; juIs76 (unc-25p::GFP), that is, an unc-70 (e524) mutant strain expressing GFP transgenically in the DD/VD motor neurons. DD/VD neurons expressing GFP can be seen in (C,D). (E,F) NIS mission samples stained with phalloidin. WT (NIS) is the N2 strain. Body wall muscle #16 is highlighted with a white border in all pictures. The bar in (A) (=100 µm) is applied to (A-D), and the bar in (E) (=100 µm) is applied to (E,F).
We found that the average dorsal muscle longitudinal cell area ranged from 1135 µm 2 in cell 22 to 1821 µm 2 in cell 14 with an average size of 1501 µm 2 in the WT ( Figure 4A,B). Interestingly, the muscle cell sizes in the space samples were clearly smaller, ranging from 626 µm 2 in cell 22 to 1142 µm 2 in cell 14 with an average size of 891 µm 2 in the WT ( Figure  4A,B). The control unc-70 mutant muscle cells were smaller than the WT, as expected [21]. As in the WT, we also observed smaller-than-normal muscle cells in the space samples, with an average decrease of 37.3% in unc-70 mutant muscle cell area in space. The atrophy Figure 3. Stained muscles of MME and NIS samples. (A-D) MME mission samples stained with Alexa Fluor™ 488 phalloidin. WT (MME) is N2; juIs76 (unc-25p::GFP), that is, a wild-type N2 strain expressing GFP transgenically in the DD/VD motor neurons. unc-70 is unc-70; juIs76 (unc-25p::GFP), that is, an unc-70 (e524) mutant strain expressing GFP transgenically in the DD/VD motor neurons. DD/VD neurons expressing GFP can be seen in (C,D). (E,F) NIS mission samples stained with phalloidin. WT (NIS) is the N2 strain. Body wall muscle #16 is highlighted with a white border in all pictures. The bar in (A) (=100 µm) is applied to (A-D), and the bar in (E) (=100 µm) is applied to (E,F).
We noticed clear differences in the size of the muscle cells in space compared to the controls, so we measured the area of muscle cells in each of the worms. C. elegans body wall muscle comprises 95 rhomboid-shaped muscle cells arranged in four longitudinal bundles, two dorsally and two ventrally, spanning most of the anterior-posterior axis of the worm ( [24]; Figure 2). We did not observe any specific differences between muscle cells in the dorsal and ventral bundles in the space and control samples, so we focused our analysis on the dorsal lateral body wall muscle cells. The 24 muscle cells in a single bundle greatly vary in size across the longitudinal axis [21]. Thus, we set out to analyze a total of seven body wall muscle cells per animal: cells 10, 12, 14, 16, 18, 20 and 22 (Figure 2A). An accurate measurement of muscle cell size would require measurement of the longitudinal area as well as cell height. However, since the height of the cells is much smaller than the longitudinal area [25] and accurate measurements of cell height would require electron microscopy analysis, we approximated muscle cell size by measuring only the longitudinal area of the cell, as previously shown [21].
We found that the average dorsal muscle longitudinal cell area ranged from 1135 µm 2 in cell 22 to 1821 µm 2 in cell 14 with an average size of 1501 µm 2 in the WT ( Figure 4A,B). Interestingly, the muscle cell sizes in the space samples were clearly smaller, ranging from 626 µm 2 in cell 22 to 1142 µm 2 in cell 14 with an average size of 891 µm 2 in the WT ( Figure 4A,B). The control unc-70 mutant muscle cells were smaller than the WT, as expected [21]. As in the WT, we also observed smaller-than-normal muscle cells in the space samples, with an average decrease of 37.3% in unc-70 mutant muscle cell area in space. The atrophy in longitudinal muscle area was observed consistently in all seven dorsal lateral body wall muscle cells ( Figure 4A). Thus, the space environment causes atrophy of body wall muscle cells independently of unc-70 β-spectrin mutation.
length ratio (MBR). We confirmed the previous reports that body length is significan smaller in space in both the MME and NIS experiments and for both the WT and the u 70 mutant strain ( Figure 5A,B). Next, we calculated the muscle-to-body length ratio the MME and NIS experiments ( Figure 5C,D) and found a decrease in the MBR of 29. and 27.4% for the WT and unc-70, respectively, in the MME experiment, and a 19.8% d crease in the WT in the NIS experiment ( Figure 5E). This suggests that the level of mus shrinkage was much higher than the decrease in body length and indicates the occurren of microgravity-associated muscle atrophy.  We had an opportunity to confirm our results from the MME space mission during the Neural Integration System (NIS) mission. In contrast to the MME experiments, the NIS experiments included a 1 G centrifuge aboard the ISS, which generated a space 1 G control in addition to the space 0 G experimental samples. Similar to the MME experiments, we observed a consistent decrease in longitudinal muscle cell areas for all seven dorsal lateral body wall muscle cells, and an overall decrease in the average muscle cell area of 23.4% in space 0 G compared to space 1G ( Figure 4C,D). Taken together, exposure to space microgravity correlates with a decrease in body wall muscle cell size.
Previous studies have shown that C. elegans overall body length is smaller in space [20,26]. Thus, one possibility is that the smaller muscle area observed in space may be due to growth limitations in the space environment and overall smaller body size. To ascertain this possibility, we determined the body length in the control and space samples, and then normalized our muscle cell area data with body length by calculating a muscle-to-body length ratio (MBR). We confirmed the previous reports that body length is significantly smaller in space in both the MME and NIS experiments and for both the WT and the unc-70 mutant strain ( Figure 5A,B). Next, we calculated the muscle-to-body length ratio for the MME and NIS experiments ( Figure 5C,D) and found a decrease in the MBR of 29.7% and 27.4% for the WT and unc-70, respectively, in the MME experiment, and a 19.8% decrease in the WT in the NIS experiment ( Figure 5E). This suggests that the level of muscle shrinkage was much higher than the decrease in body length and indicates the occurrence of microgravity-associated muscle atrophy. normalize muscle cell area with body length. ※ indicates MBR calculation which is indicated below the graphs. (E) Percent change in MBR between control and test samples to determine extent of muscle atrophy. Error bars indicate standard error. Statistical significance was determined via Student's t-test. * and ** and *** indicate statistical significance at p < 0.05, p < 0.005 and p < 0.001, respectively.

Muscle Atrophy in Nutritional Deprivation and Disuse Conditions in C. elegans
Genetic analysis using C. elegans affords us an opportunity to understand the cellular and molecular mechanism of how space microgravity induces muscle atrophy in the body wall muscle cells. However, sampling and other restrictions in space experiments, together with the few space mission opportunities, are limiting factors in conducting further genetic analysis in space. Muscle atrophy is not limited to space microgravity, and other factors can induce smaller muscle, including nutritional deprivation and muscle disuse [4][5][6]. Thus, we applied these two factors to determine whether they also induce muscle atrophy in C. elegans.
To determine whether nutritional deprivation is associated with muscle atrophy, we provided E. coli food to developmentally synchronized L1-stage larvae for 48 h, and then removed the food, starving the worms for 5 to 48 h. Normally, in the presence of ample nutrition, C. elegans continues to grow in both body length and longitudinal muscle area from 53 h to 96 h ( Figure 6A). However, upon starvation, body length growth slowed substantially, and muscle cell size stopped increasing ( Figure 6A) and appeared to shrink ( Figure 6B), eventually leading to the death of some worms after 48 h. After 30 h of starvation, muscle cell area and body length were greatly reduced in experimental worms compared to those undergoing continual feeding ( Figure 6C,D). More importantly, we found a large reduction in the MBR of over 40% by 30 h in starved animals ( Figure 6E). Error bars indicate standard error. Statistical significance was determined via Student's t-test. * and ** and *** indicate statistical significance at p < 0.05, p < 0.005 and p < 0.001, respectively.

Muscle Atrophy in Nutritional Deprivation and Disuse Conditions in C. elegans
Genetic analysis using C. elegans affords us an opportunity to understand the cellular and molecular mechanism of how space microgravity induces muscle atrophy in the body wall muscle cells. However, sampling and other restrictions in space experiments, together with the few space mission opportunities, are limiting factors in conducting further genetic analysis in space. Muscle atrophy is not limited to space microgravity, and other factors can induce smaller muscle, including nutritional deprivation and muscle disuse [4][5][6]. Thus, we applied these two factors to determine whether they also induce muscle atrophy in C. elegans.
To determine whether nutritional deprivation is associated with muscle atrophy, we provided E. coli food to developmentally synchronized L1-stage larvae for 48 h, and then removed the food, starving the worms for 5 to 48 h. Normally, in the presence of ample nutrition, C. elegans continues to grow in both body length and longitudinal muscle area from 53 h to 96 h ( Figure 6A). However, upon starvation, body length growth slowed substantially, and muscle cell size stopped increasing ( Figure 6A) and appeared to shrink ( Figure 6B), eventually leading to the death of some worms after 48 h. After 30 h of starvation, muscle cell area and body length were greatly reduced in experimental worms compared to those undergoing continual feeding ( Figure 6C,D). More importantly, we found a large reduction in the MBR of over 40% by 30 h in starved animals ( Figure 6E).
Hence, nutritional deprivation correlates with muscle atrophy in C. elegans, as in most other animals [5,6,27]. Hence, nutritional deprivation correlates with muscle atrophy in C. elegans, as in most other animals [5,6,27]. Muscle disuse, whether due to disease, denervation experiments or hindlimb suspension experiments, is known to cause muscle atrophy [4,9,28,29]. In C. elegans, we used unc-13 strain mutants to replicate muscle disuse. The unc-13 gene encodes the C. elegans orthologue of the MUNC-13 mammalian protein, which serves a function in pre-synaptic neurotransmitter synaptic vesicle release [30,31]. The severe unc-13 (s69) mutant does not display any evoked synaptic activity, including any neuromuscular synaptic activity, and is almost completely paralyzed but overall can eat, develop and survive rather normally [12,30,32], unlike other mutants of key genes involved in the C. elegans neuromuscular junction. We found that the body wall muscle of unc-13 mutants was smaller than the muscle of well-fed WT strain worms, displaying an average area of 665 µm 2 , which is comparable in size to the area of muscle in starved animals at 741 µm 2 ( Figure 6B,C). The body length of unc-13 mutants was also smaller than that of well-fed WT C. elegans ( Figure  6D), but the MBR decreased nearly 40% in unc-13 mutants compared to WT animals, indicating substantial muscle atrophy due to disuse comparable to atrophy during nutritional deprivation ( Figure 6E).

clp-4 Promotes Muscle Atrophy during Nutritional Deprivation
To identify genetic factors that induce muscle atrophy, we conducted a candidate genetic screening using C. elegans mutants. Among the three experimental conditions of Muscle disuse, whether due to disease, denervation experiments or hindlimb suspension experiments, is known to cause muscle atrophy [4,9,28,29]. In C. elegans, we used unc-13 strain mutants to replicate muscle disuse. The unc-13 gene encodes the C. elegans orthologue of the MUNC-13 mammalian protein, which serves a function in pre-synaptic neurotransmitter synaptic vesicle release [30,31]. The severe unc-13 (s69) mutant does not display any evoked synaptic activity, including any neuromuscular synaptic activity, and is almost completely paralyzed but overall can eat, develop and survive rather normally [12,30,32], unlike other mutants of key genes involved in the C. elegans neuromuscular junction. We found that the body wall muscle of unc-13 mutants was smaller than the muscle of well-fed WT strain worms, displaying an average area of 665 µm 2 , which is comparable in size to the area of muscle in starved animals at 741 µm 2 ( Figure 6B,C). The body length of unc-13 mutants was also smaller than that of well-fed WT C. elegans ( Figure 6D), but the MBR decreased nearly 40% in unc-13 mutants compared to WT animals, indicating substantial muscle atrophy due to disuse comparable to atrophy during nutritional deprivation ( Figure 6E).

clp-4 Promotes Muscle Atrophy during Nutritional Deprivation
To identify genetic factors that induce muscle atrophy, we conducted a candidate genetic screening using C. elegans mutants. Among the three experimental conditions of muscle atrophy that we have identified, starvation was the most straightforward experimentally. Therefore, we based our genetic screening on the nutritional deprivation method and evaluated the muscle atrophy of various mutant lines by determining the change in the MBR after 30 h of starvation. We first tested mutants of dim-1, which encodes a muscle-expressed immunoglobin protein. DIM-1 is known to be involved in the maintenance of muscle structure, and loss of dim-1 protects against disruption of attachment complexes and associated phenotypes [33,34]. In addition, dim-1 expression was decreased during spaceflight in two independent space experiments [20]. However, in dim-1 starved animals, we observed smaller muscle and a decreased MBR similar to that of WT animals, indicating that the loss of dim-1 does not alter muscle atrophy during nutritional deficiency ( Figure 7A,B). We next tested genes encoding members of the calpain family of calcium-dependent cysteine proteases. Various calpain proteins have differential functions in muscle sarcomere and muscle mitochondrial development as well as attachment complex formation [34]. We found that mutation of the calpain genes clp-1 and tra-3 did not alter starvation-induced muscle atrophy, and clp-7 mutation had a mild effect ( Figure 7A,B). However, disruption of clp-4 had a profound effect, decreasing the MBR of WT animals from 43.6% to only 21% in clp-4 mutants ( Figure 7A,B). Finally, we also observed muscle atrophy in daf-12and tbh-1-deficient animals. DAF-12 and TBH-1 are known to protect C. elegans from starvation-induced death [35]. However, both daf-12 and tbh-1 mutants showed a similar decrease in the MBR during starvation, indicating that starvation-induced muscle atrophy may be independent of starvation tolerance pathways ( Figure 7A,B). Overall, we found that the calpain gene clp-4 promotes muscle atrophy, and inhibition of clp-4 can protect C. elegans muscle from complete starvation-induced muscle atrophy. muscle atrophy that we have identified, starvation was the most straightforward experimentally. Therefore, we based our genetic screening on the nutritional deprivation method and evaluated the muscle atrophy of various mutant lines by determining the change in the MBR after 30 h of starvation. We first tested mutants of dim-1, which encodes a muscle-expressed immunoglobin protein. DIM-1 is known to be involved in the maintenance of muscle structure, and loss of dim-1 protects against disruption of attachment complexes and associated phenotypes [33,34]. In addition, dim-1 expression was decreased during spaceflight in two independent space experiments [20]. However, in dim-1 starved animals, we observed smaller muscle and a decreased MBR similar to that of WT animals, indicating that the loss of dim-1 does not alter muscle atrophy during nutritional deficiency ( Figure 7A,B). We next tested genes encoding members of the calpain family of calcium-dependent cysteine proteases. Various calpain proteins have differential functions in muscle sarcomere and muscle mitochondrial development as well as attachment complex formation [34]. We found that mutation of the calpain genes clp-1 and tra-3 did not alter starvation-induced muscle atrophy, and clp-7 mutation had a mild effect (Figure 7A,B). However, disruption of clp-4 had a profound effect, decreasing the MBR of WT animals from 43.6% to only 21% in clp-4 mutants ( Figure 7A,B). Finally, we also observed muscle atrophy in daf-12-and tbh-1-deficient animals. DAF-12 and TBH-1 are known to protect C. elegans from starvation-induced death [35]. However, both daf-12 and tbh-1 mutants showed a similar decrease in the MBR during starvation, indicating that starvationinduced muscle atrophy may be independent of starvation tolerance pathways ( Figure  7A,B). Overall, we found that the calpain gene clp-4 promotes muscle atrophy, and inhibition of clp-4 can protect C. elegans muscle from complete starvation-induced muscle atrophy.

Discussion
Long-term space travel, tourism and habitation are on the horizon for humankind, but microgravity poses significant health and biological problems that must be overcome. Space-induced muscle atrophy has been demonstrated in a range of organisms including humans, rodents and primates, and now includes the nematode C. elegans [36]. However, identifying the molecular and genetic underpinnings of microgravity-induced muscle atrophy and developing therapies to prevent it are hampered by the difficulty of conducting practical and effective experiments in space. Using C. elegans as a model animal for muscle atrophy affords us the opportunity to conduct experiments in space and the laboratory,

Discussion
Long-term space travel, tourism and habitation are on the horizon for humankind, but microgravity poses significant health and biological problems that must be overcome. Space-induced muscle atrophy has been demonstrated in a range of organisms including humans, rodents and primates, and now includes the nematode C. elegans [36]. However, identifying the molecular and genetic underpinnings of microgravity-induced muscle atrophy and developing therapies to prevent it are hampered by the difficulty of conducting practical and effective experiments in space. Using C. elegans as a model animal for muscle atrophy affords us the opportunity to conduct experiments in space and the laboratory, and to study muscle atrophy from a broader perspective with multiple useful genetic tools. Here, we showed C. elegans muscle cell area decreased in two independent experiments aboard the ISS. The physiological phenotype we observed in muscle is consistent with previous reports showing gene expression declines in key muscle genes in C. elegans, including myo-3 and unc-54 myosin heavy chain genes, and the unc-15 paramyosin gene, from two independent space experiments [19,20,37]. Indeed, the downregulation of myosin heavy chain and paramyosin genes was consistent with observed decreases in protein levels in the same spaceflight experiments [20] as well as previous spaceflight experiments [15,38]. These data together with our observations suggest that space microgravity regulates C. elegans muscle cell size by altering muscle gene expression [39,40].
Our methodology to assess the size of the dorsal lateral muscle cells is based on phalloidin fluorescence staining of muscle actin and determining the approximate borders of each muscle cell. In addition, we measured the maximal longitudinal area of each rhomboid-shaped muscle cell, ignoring the height dimension of each cell. Our methodology gives an approximate measurement of muscle cell volume and atrophy observed during spaceflight. Experimental limitations with the space samples prevented us from conducting more accurate methods, such as EM experiments, to assess muscle cell volume. In addition, C. elegans body length differences in space, as reported previously [20,41], can indirectly contribute to smaller muscle. Instead, we normalized muscle area with body length by calculating the MBR in all our experiments.
Although our initial MME mission samples showed a decline in muscle cell area in space, this experiment was hampered by low-quality ground controls for the WT, and we produced a laboratory control that mimicked spaceflight conditions. However, we confirmed our results from MME in the NIS experiments, which included a 1 G control sample aboard the ISS in addition to a 0 G sample. In these experiments, both the control and experimental samples experienced the same spaceflight launch, handling by astronauts and timing and conditions aboard the ISS. However, some of the conditions between MME and NIS were different. This included the cultivation time aboard the ISS (5 days in total for MME, 12 days in total for NIS; see Figure 1), the conditions of the medium (the chemical FUdR was added at 7.5 days to prevent egg laying of the F1 generation; see Methods) and the fixation conditions (−80 • C for MME, PFA fixation and 4 • C for NIS). Overall, these differences did not change the developmental time or quality of muscle and phalloidin staining, and we have confidence in the differences in muscle area and body length in both MME and NIS experiments.
Nutritional deprivation results in muscle atrophy in humans, rodents and flies [5,6,27]. Starvation in C. elegans is a large source of stress for adult animals, resulting in multiple physiological and behavior changes [42]. Interestingly, starvation does not result in death but rather an increase in C. elegans' lifespan [43,44], presumably through a starvation resistance pathway that alters metabolism [42]. However, the effects of starvation on muscle and body length have not been assessed. Here, we show that longitudinal muscle cell area and body length decreased substantially within 5 h of starvation at the L4 stage, and muscle cell size continued to decline after 48 h of starvation ( Figure 6A). Octopamine regulates starvation resistance in adult animals as well as other physiological and behavioral aspects of starvation, and the nuclear receptor daf-12 and the octopamine biosynthesis gene tbh-1 promote starvation resistance [35,42,45,46]. We wondered whether tbh-1 and daf-12 could also mediate nutritional deprivation-induced muscle atrophy, but tbh-1 and daf-12 mutants showed a similar starvation-induced MBR to the WT, demonstrating that another starvation-resistance pathway may be regulating these changes ( Figure 7A,B).
Disuse of muscle, as observed in bedridden patients, patients with spinal cord injury and spinal cord denervation and hind-limb suspension experiments in rodents, results in muscle atrophy [4,9,28,29]. We used a C. elegans mutant, unc-13 (s69), to simulate muscle disuse experiments in the nematode. UNC-13 protein is necessary for synaptic vesicle release, and unc-13 (s69) mutants are almost completely paralyzed because no evoked synaptic activity occurs at the neuromuscular junction [32]. Mutations in genes more specific to the neuromuscular junction, such as acetylcholine receptor genes unc-17 and cha-1, may be more appropriate; however, null mutants of these genes are so severe that they induce larval developmental arrest [47][48][49]. unc-13 mutants had a smaller body wall muscle area and decreased MBR compared to WT animals ( Figure 7A,B), according to results obtained using the standard method.
However, since unc-13 (s69) mutants have a complete loss of synaptic activity throughout the body and have some neurosecretory defects [50], we cannot completely rule out the possibility that other neural factors may cause smaller muscle cell size.
Comparing muscle atrophy in space microgravity with nutritional deprivation and disuse, we found a decrease in the MBR of between 20% and 30% under space microgravity conditions, and a roughly 40% decline in both starvation and disuse conditions ( Figure 6E). In addition, body wall muscle shape was almost always preserved in the control under each of these atrophy-inducing conditions (Figures 3 and 6B). This is in contrast to the altered shape observed in the disruption of core proteins involved in muscle attachment complexes, which results in a complete collapse of the sarcomere structure into ball-like structures [34] that we never observed in our experiments. Although we occasionally observed spacing between the phalloidin-stained actin of adjacent muscle cells and slight alterations in shape in nutritionally deprived and disuse animals ( Figure 6B), these qualitative phenotypes were rare and inconsistent. Overall, the muscle atrophy observed in space microgravity was not greatly different from that induced by nutritional deprivation or disuse.
In our candidate genetic screening, we found that mutation in the clp-4 gene suppressed starvation-induced muscle atrophy from over 43% of the MBR to less than half at 21% of the MBR. Calpains are Ca 2+ -dependent proteases that exhibit limited proteolytic activity and are ubiquitously present [51,52]. They are known to be involved in apoptosis, muscle protein turnover and muscle maintenance in general [34,53,54]. Humans have three clp-4 orthologs, CAPN1 (calpain 1), CAPN2 (calpain 2) and CAPN3 (calpain 3), which have been implicated in limb-girdle muscular dystrophy [55,56]. Calpains play a role in muscle protein degradation when attachment complexes are disrupted, and CLP-4 is expressed in muscle cells and required to maintain arrayed attachment complexes [34,56]. How then can CLP-4 promote starvation-induced muscle atrophy? We speculate that nutritional deprivation leads to cell and ER stress, resulting in release of ER calcium stores and activation of the Ca 2+ -dependent protease, but further analysis will be required to substantiate this. Furthermore, we would like to eventually confirm whether clp-4 or calpain-mediated proteolysis also plays a role in promoting space-microgravity-induced muscle atrophy.

Conditions for MME Space Experiment
To determine a suitable liquid culture for the MME mission that would preserve the worms' integrity when frozen but not affect the worms' development and population growth, different freezing solutions with varying concentrations and combinations were tested. Synchronized L1 larva-stage worms were prepared and transferred into polyethylene bags containing LabTie E. coli OP50 (Biovirid, Veenendaal, The Netherlands) dissolved in buffer (S-basal + cholesterol, 2× final LabTie concentration) with or without cryoprotectant solution and incubated for 6 days at 12 • C (to mimic the storage temperature prior to arrival at the ISS) and another 6 days at 20 • C (actual incubation temperature aboard the ISS). Culture bags were then retrieved, and worms' developmental and growth states were checked under a dissecting microscope. Then, they were frozen at −80 • C. After several days, culture bags were thawed, and worms were again observed for GFP integrity and signals. We found that dimethyl sulfoxide (DMSO) and trehalose treatment optimally preserved the GFP quality from the samples compared to sorbitol treatment or no cryoprotectant treatment (Table S1). However, it does not provide ideal development and population growth for C. elegans. Thus, a lower concentration of trehalose solution was tested under our MME space sample conditions, and the worms' survival, development and population growth were observed regularly under a dissecting microscope. However, 0.08 mM trehalose treatment still conferred a slightly negative effect on the growth of the worms (Table S2). Thus, we proceeded with S-basal with cholesterol without any cryoprotectant as the appropriate culture condition for the MME space experiment.

MME Space Experiments
The MME space mission was conducted in collaboration with the European Space Agency (ESA). We initially determined the proper conditions for spaceflight, and the samples were prepared using a method similar to the one described previously by Laranjeiro et al. (2021) and Sudevan et al. (2022) [58,59]. The eggs were harvested by bleaching gravid worms following the standard protocol [60] on Day −1. On Day 0, approximately 500 synchronized L1 larvae were transferred to polyethylene flight culture bags containing S-basal (for 1 L S-basal: NaCl 5.85 g, K 2 HPO 4 1 g, KH 2 PO 4 6 g), cholesterol (5 mg/mL in EtOH stock; 1 ml for 1 L) and 2× concentration LabTie freeze-dried OP50 (Biovirid, Veenendaal, The Netherlands), totaling a volume of 6.5 mL per bag. These samples were then sealed and placed into experiment cassettes (ECs, Kayser, Toscana, Italy; [18]) and kept in cold stowage at temperatures ranging from 8 to 13 • C until they arrived at the International Space Station (ISS). After five days of incubation in the ISS, the samples were frozen at −80 • C until analysis. The control samples were prepared in the same manner as the space samples and handled and stored at NASA Kennedy Space Center (Cape Canaveral, FL, USA) or in the laboratory for lab WT.

NIS Space Experiments
The NIS space mission was conducted in collaboration with the Japan Aerospace Exploration Agency (JAXA). Approximately 200 synchronized L1 larvae were prepared on Day −2, added to polyethylene flight culture bags with M9 buffer with cholesterol (KH 2 PO 4 3 g, Na 2 HPO 4 12H 2 O 15.1 g, NaCl 5 g, 1 M MgSO 4 1 mL and 1 mL of cholesterol (5 mg/mL in EtOH) for a total of 1 L) and stored at 12 • C until the launch. When the samples arrived at the International Space Station (ISS), they were transferred to 20 • C incubators. The astronaut responsible for handling the samples took half of each sample and fixed them with 0.7% paraformaldehyde (PFA) when the F1-generation worms reached Day 1 and Day 10 of adulthood. To prevent starvation and the mixing of generations, the astronaut added media M9 buffer with cholesterol and E. coli food with FUdR (Sigma-Aldrich F0503, St. Louis, MO, USA, 75 µM), which interferes with DNA synthesis and prevents reproduction, after extracting the portion of samples. In this paper, only Day-10 adult samples were used for analysis. Control samples were also sent to the ISS and placed in a centrifuge that simulated a 1 G gravity force.  [57]) with LabTIE freeze-dried OP50 (0.9 g/250 mL) was added to each well of a 96-well plate (30096, SPL). To prevent rapid evaporation and ensure normal gas exchange, the 96-well plate was covered with a gas-permeable film (A9224, Sigma-Aldrich) instead of its plastic cover. L1-synchronized larvae were grown on E. coli OP50-seeded NGM plates at 20 • C for 48 h. These worms were then washed with S-basal to remove as much OP50 as possible from their bodies and divided into two groups, one with food and the other without food, and transferred to a liquid culture plate. After an additional 30 h of incubation, the samples were used for analysis. Each well contained fewer than 20 worms.

Phalloidin Muscle Staining
A previously described method for muscle staining, as outlined in [61], was used in this study. All samples were fixed with 4%PFA prior to staining. Following PFA fixation, the samples were washed with 1× PBS, and acetone was added for 1 min to enhance the permeability of phalloidin within the worms. Subsequently, the samples were washed again and treated with phalloidin conjugated with a fluorescent molecule. After overnight incubation at 37 • C, the samples were mounted onto glass slides covered with a 2% dry agarose pad. Observation and image capture were conducted using an Olympus BX50 microscope (Tokyo, Japan). All samples were stained with Alexa Fluor™ 488 Phalloidin (A12379, Invitrogen™, Waltham, MA, USA), which emits a green fluorescent signal, except for the NIS samples, which were stained with Rhodamine Phalloidin (R415, Invitrogen™), which emits a red fluorescent signal.
An adult C. elegans possesses 95 body wall muscles distributed along its body. These muscles are organized into four longitudinal bundles, one in each quadrant. Each bundle consists of staggered pairs of muscles, and the muscle cells within a bundle are numbered from 1 to 23 or 24, as described previously [62]. For this study, we specifically focused on measuring seven dorsal lateral body wall muscles located in the mid-body region, which are numbered 10, 12, 14, 16, 18, 20 and 22 (Figure 1). To measure the longitudinal muscle cell area and body length, a previously published protocol was used [21]. Briefly, the ImageJ software V1.52 was utilized as a measurement tool. The muscle-to-body length ratio, MBR, was calculated by dividing individual worms' average muscle cell area by body length. Statistical analysis of the obtained data was conducted using Student's t-test for space missions and one-way ANOVA followed by Tukey's post hoc analysis (IBM SPSS, USA) for starvation and disuse experiments to determine the significance of the results.