Intrinsic muscles are a group of muscles deep inside the hands and feet. They help to control the precise movements required, for example, for a pianist to play their instrument or for certain animals to climb with remarkable agility.

Some animals, such as horses and deer, have evolved in such a way that they no longer grasp objects with hands and feet. Where intrinsic muscles were once present in the hands and feet of their ancestors, these animals now have strong ligaments that prevent over-extension of the wrist and ankle joints during hard landings.

Given their size, it is difficult to study horses and deer in the laboratory and understand how they lost their intrinsic muscles during evolution. Tran et al. therefore focused on a small rodent called the lesser Egyptian jerboa, which also displays long legs with strong ligaments and no intrinsic muscles.

Newborn jerboas have foot muscles that look very much like the intrinsic muscles found in mice, but these muscles disappear within 4 days of birth. A mechanism called programmed cell death is often responsible for specific tissues disappearing during development, but the experiments of Tran et al. revealed that this was not the case in jerboas. Instead, their intrinsic muscles were degraded by processes triggered by genes that disassemble underused muscles. In mice and humans, fasting, nerve injuries, or immobility trigger this type of muscle degradation, but in jerboas these processes appear to be a normal part of development.

This unexpected discovery shows that development and disease-like processes are linked, and that more studies of nontraditional research animals may help scientists better understand these connections.

Muscles in the feet of birds, reptiles, and mammals were lost multiple times in the course of limb evolution, usually coinciding with the loss of associated digits and elongation of remaining skeletal elements (Hudson, 1937; Raikow, 1987; Pavaux and Lignereux, 1995; Botelho et al., 2014; Abdala et al., 2015; Berman, 1985; Cunningham, 1883; Souza et al., 2010). Despite its frequent occurrence, the developmental mechanisms that lead to the natural absence of adult limb muscle are not known. We focus here on a representative example of distal limb muscle loss in the bipedal three-toed jerboa (Jaculus jaculus), a small laboratory rodent model for evolutionary developmental biology, to determine if evolutionary muscle loss conforms to expectations based on what was previously known about muscle cell biology.

The hindlimb architecture of the adult jerboa is strikingly similar by convergence to the more familiar hooved animals, like horses and deer, including the disproportionately elongated foot that lacks all intrinsic muscle (Berman, 1985; Cunningham, 1883). The tendons were retained and expanded in each of the anatomical positions where flexor muscles are absent (Figure 1A,B and Figure 1—figure supplement 1A,B) and serve to resist hyperextension when the terminal phalanx contacts the ground during locomotion (Lochner et al., 1980; Moore et al., 2017). The evolutionary origin of jerboa intrinsic foot muscle loss lies deep in the phylogenetic tree of Dipodoid rodents. Compared to the ancestral state, the number of intrinsic foot muscles are reduced from sixteen to six in pygmy jerboas (Stein, 1990) which diverged from the three-toed jerboa lineage more than 20 million years ago (Wu et al., 2012; Pisano et al., 2015).

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The mechanisms of limb muscle development have been extensively studied in traditional model systems, and its degeneration has been studied after injury and during disease. Briefly, limb muscle progenitors are specified from mesodermal cells at the ventrolateral edge of the dermomyotome in somites aligned with the prospective limb. These cells delaminate and migrate into the limb bud as dorsal and ventral muscle masses that proliferate and initiate a myoblast specification program (Chevallier et al., 1977; Christ et al., 1977; Hayashi and Ozawa, 1991; Murphy and Kardon, 2011). The muscle masses are then subdivided into individual muscle groups in response to cues from the developing muscle connective tissue, which is derived from limb field lateral plate mesoderm (Hayashi and Ozawa, 1991; Kardon, 1998; Kardon et al., 2003; Wortham, 1948). They then initiate a differentiation program, which includes cell fusion to form aligned multinucleated myofibers (Abmayr and Pavlath, 2012; Kelly and Zacks, 1969).

Each differentiated myofiber produces an assemblage of Z-body proteins, Actin filaments, and non-muscle Myosin that form premyofibrils (Ono, 2010; Rhee et al., 1994; Sanger and Sanger, 2008; Sanger et al., 2002). Desmin, α-Actinin, and the Z-body portion of Titin also begin to organize (Furst et al., 1989; Sanger et al., 2002). Subsequent uncoiling of Titin increases Z-body spacing, and integration of embryonic skeletal muscle Myosin results in formation of nascent myofibrils (Ono, 2010; Sanger et al., 2010). Further maturation of the nascent myofibril into a mature myofibril involves incorporation of additional proteins that are important for sarcomere structure and function, and Z-lines are aligned and properly spaced to bring sarcomeres into register (Ehler and Gautel, 2008; Sanger et al., 2010). Failure at any point of myoblast specification, migration, myofiber differentiation, or myofibril maturation compromises muscle function and manifests as muscle degenerative disease in humans (Bönnemann and Laing, 2004; Laing and Nowak, 2005; Morita et al., 2005).

Working backward in time from the adult jerboa phenotype, we found that two of the three flexor muscle groups differentiate as multinucleated myofibers that initiate sarcomere assembly, as in other species. However, almost all jerboa intrinsic foot muscle is lost within a few days shortly after birth. Despite the rapid and near complete loss of myofibers, we found no molecular or ultrastructural evidence of apoptotic or necrotic cell death, no accumulation of autophagic vesicles, and no macrophage infiltration. Instead, we observed evidence of ordered sarcomere disassembly and upregulation of muscle-specific ubiquitin ligases, MuRF1 and Atrogin-1. Although the ultimate fate of intrinsic foot myofibers after loss of muscle identity remains unknown, these data suggest that the mechanism of myofiber loss is similar to atrophy, which is typically considered a pathological response to injury or disease.

The absence of intrinsic foot muscle in the adult jerboa could be due to a failure of early myoblasts to migrate into and/or to differentiate in the distal limb. Alternatively, embryonic muscles may form but not persist through development to the adult. In transverse sections of newborn mouse feet, immunofluorescent detection of skeletal muscle myosin heavy chain reveals each intrinsic muscle group (Figure 1—figure supplement 1G). In newborn jerboas, we observed two of the three groups of flexor muscles. While the m. lumbricales never form, the jerboa has a single m. flexor digitorum brevis and three pinnate m. interossei that are not present in adults (Figure 1—figure supplement 1H).

Postnatal growth of vertebrate skeletal muscle typically involves an increase in myofiber number (hyperplasia) within the first week, followed by an increase in myofiber size (hypertrophy) (Chiakulas and Pauly, 1965; Gokhin et al., 2008; White et al., 2010). In order to understand the dynamics of muscle growth and loss, we quantified the rate of myofiber hyperplasia at 2-day intervals after birth of the mouse and jerboa, focusing on the representative interosseous muscle that is associated with the third metatarsal (Figure 1E,F). As expected in the mouse, we observed a steady increase in the average number of myofibers in cross section from birth to P8 (Figure 1C). In contrast, the number of myofibers in the third interosseous of the jerboa foot rapidly declines beginning at approximately P4, and few myofibers remain by P8 (Figure 1D).

It is possible that the rate of myofiber loss outpaces a typical rate of new cell addition such that muscles with the potential to grow are instead steadily diminished. Alternatively, myofiber loss may be accelerated by a compromised ability to form new myofibers and to add nuclei to growing myofibers. To distinguish these hypotheses, we analyzed cohorts of animals 2 days after intraperitoneal BrdU injection at P0, P2, or P4. Since multinucleated jerboa foot myofibers are postmitotic (Figure 2—figure supplement 1), we reasoned that BrdU+ nuclei present within Dystroglycan+ myofiber membranes were added by myocyte fusion during the 2-day window after they were labeled as myoblasts or myocytes in S-phase (Figure 2A). When normalized to the total number of myofiber nuclei, we found that myocytes fuse to form multinucleated myofibers in jerboa hand muscle at a consistent rate from P0 to P6. However, their incorporation into jerboa foot muscle decreased significantly after P2 (Figure 2B). These results suggest that myofiber loss, which begins at P4, is preceded by reduced myogenesis.

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The reduced rate of myocyte incorporation could be due to reduced numbers of muscle progenitor cells or to an inability of these cells to mature and fuse. To distinguish these possibilities by quantifying proliferative muscle progenitor cells, we analyzed animals 2 hr after BrdU injection at P0, P2, and P4 and counted the number of BrdU+ nuclei located between the Dystroglycan+ myofiber membrane and the Laminin+ basal lamina (Figure 2C). Normalized to the total number of myofibers, we found that the number of proliferative progenitor cells in jerboa foot muscle significantly decreased from P0 to P4 compared to hand muscles that showed no change over time (Figure 2D). These results suggest that a reduced number of muscle progenitor cells might contribute to the reduced prevalence of myocyte fusion events.

We next tested whether compromised proliferation and differentiation of jerboa foot muscle progenitors is cell autonomous or non-cell autonomous. We isolated single cells, including myoblasts and myocytes but excluding myofibers, by mechanical trituration and enzymatic digestion of P1 jerboa and mouse lower leg and foot muscles (Danoviz and Yablonka-Reuveni, 2012). After 6 days and 9 days of culture, we detected Myogenin+ differentiating myocytes and Myosin+ fully differentiated myofibers in primary cell cultures isolated from each muscle (Figure 2E). We did not detect a significant decline in the number of differentiated cells over time (Figure 2—figure supplement 2). Jerboa foot muscle cell differentiation and survival in vitro days after cell number begins to decline in vivo suggests that loss of jerboa foot myofibers is initiated non-cell autonomously.

The rapid and almost complete loss of differentiated myofibers in vivo from P4 to P8 suggested these cells die, since individual cells or groups of cells are commonly eliminated by apoptosis during development (Brill et al., 1999; Fernández-Terán et al., 2006). We therefore tested the hypothesis that neonatal intrinsic foot muscles undergo apoptosis by implementing the TUNEL assay to detect DNA fragmentation and by immunofluorescent detection of cleaved Caspase-3, a key protein in the apoptotic program (Elmore, 2007). Each revealed keratinocyte apoptosis in hair follicles, which are known to undergo programmed cell death, as a positive control in the same tissue sections (Magerl et al., 2001). However, TUNEL or cleaved Caspase-3-positive jerboa foot myofibers or cells in their vicinity were an extreme rarity (0.25% of myofibers) in animals ranging from P0 to P8 and comparable to mouse myofibers suggesting muscle is not eliminated by apoptosis (Figure 3A,B and Figure 3—figure supplement 1).

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Alternatively, myofiber loss may occur through a cell death mechanism that is first characterized by plasma membrane permeability, such as necrosis (Vanden Berghe et al., 2014). To test this hypothesis, we injected Evans blue dye (EBD), a fluorescent molecule that accumulates in cells with compromised plasma membranes (Hamer et al., 2002; Matsuda et al., 1995), into the peritoneum of P3 and P4 neonatal jerboas 24 hr before euthanasia. Although we detected EBD in mechanically injured myofibers of the gastrocnemius as a control, we saw no EBD fluorescence in jerboa foot myofibers or in surrounding cells (Figure 3C and Figure 3—figure supplement 2). We also saw no Annexin V immunofluorescence on the surface of jerboa foot myofibers, another hallmark of dying cells that flip Annexin V to the outer plasma membrane (Figure 3D and Figure 3—figure supplement 2).

Since we observed no direct evidence of cell death, we asked whether there was an immune response that might be an indirect proxy for undetected death. Dying muscle cells frequently recruit phagocytic macrophages that engulf cellular debris (Arnold et al., 2007; Londhe and Guttridge, 2015; Tidball and Wehling-Henricks, 2007). We predicted that myofibers that die by any mechanism that produces cellular debris might recruit macrophages that are detectable by expression of the F4/80 glycoprotein. However, consistent with the lack of evidence of cell death in the jerboa foot, no F4/80⁺ macrophages were found among myofibers from birth to P7 (Figure 3—figure supplement 3). Since immune cells other than mature macrophages might be recruited to a site of cell death, we also assessed expression of CD45 and found no evidence of T-cells, B-cells, dendritic cells, natural killer cells, monocytes, or granulocytes near jerboa foot myofibers from P4 to P8 (Figure 3E and Figure 3—figure supplement 3).

The absence of any clear indication of muscle cell death motivated us to re-evaluate muscle maturation at greater resolution in order to capture the earliest detectable signs of muscle cell loss. We collected transmission electron micrographs of jerboa hand and foot muscle at P0, P2, and P4. We identified criteria for three categories of maturation, as described previously (Borisov et al., 2008; Raeker et al., 2014; Sanger et al., 2006), and two categories of degeneration. Category A cells have pre-myofibrils with thick and thin filaments and poorly resolved Z-discs, but the M-lines and I-bands are not yet apparent (Figure 4A). In Category B, Z-discs of nascent myofibrils are better resolved, and M-lines and I-bands are apparent, but parallel sarcomeres are not in register (Figure 4B). The mature myofibrils of Category C have Z-lines that are aligned with one another (Figure 4C). In Category D, early degeneration, some sarcomeres appear similar to Category A, but other areas of the cell contain disorganized filaments (Figure 4D). Category E includes those in the worst condition where less than half of the cell has any recognizable sarcomeres, and much of the cytoplasm is filled with pools of disorganized filaments and Z-protein aggregates (Figure 4E). Additionally, Category D and E cells have membrane-enclosed vacuoles and large lipid droplets (Figure 4—figure supplement 1). However, consistent with a lack of evidence for cell death, none of these cells or their organelles appear swollen, nuclear morphology appears normal, plasma membranes seem to be contiguous, and we do not observe an accumulation of autophagic vesicles that typically characterize cell death associated with unregulated autophagy (Mizushima, 2007; Kroemer and Levine, 2008; Denton and Kumar, 2019).

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We then coded and pooled all images of hand and foot myofibers from P0, P2, and P4 jerboas and blindly assigned each cell to one of the five categories. Quantification of the percent of myofibers in each category after unblinding revealed the progressive maturation of jerboa hand myofibers and the progressive degeneration of jerboa foot myofibers (Figure 4G). Compared to later stages, there is little difference in the maturation state of hand and foot sarcomeres at birth. Loss of ultrastructural integrity is therefore initiated perinatally, prior to complete myofibril maturation in the jerboa foot.

Our analysis of transmission electron micrographs also revealed the presence of filamentous aggregates that we did not include in our quantifications because they are enucleate, lack all other recognizable organelles, and are not bounded by a plasma membrane. Although these aggregates do not appear to be cellular, they are always closely associated with cells of a fibroblast morphology, and most lie between remaining myofibers in a space we presume was also once occupied by a myofiber (Figure 4F,H). To determine if these unusual structures contain muscle protein, we performed immunofluorescence on sections of P4 jerboa foot muscle and found similar aggregates of intensely fluorescent immunoreactivity to skeletal muscle myosin heavy chain. We also found that the surrounding cells, which correlate with the positions of fibroblasts in electron micrographs, express the intracellular pro-peptide of Collagen I (Figure 4I), the major component of tendon and other fibrous connective tissues and of fibrotic tissue after injury (Mann et al., 2011).

Given the apparent deterioration of nascent sarcomeres, we asked whether individual sarcomere proteins are lost from myofibrils in a temporal order or if proteins disassemble simultaneously. We assessed the organization of sarcomere proteins by multicolor immunofluorescence at P0, P2, and P4. Alpha-Actinin, Desmin, Myomesin, Myosin, Titin, and Tropomyosin are each localized to an ordered series of striations in a subset of myofibers suggesting all are initially incorporated into immature sarcomeres (Figure 5A and Figure 5—source data 1–5). By assessing all combinations of immunologically compatible primary antibodies, we identified populations of cells where Desmin was no longer present in an ordered array, but each of the other proteins appeared properly localized to the sarcomere (Figure 5B and Figure 5—source data 2). Although we could not distinguish such clear categories of mislocalization for each protein relative to all others, we inferred a relative timeline whereby Desmin disorganization is followed together by Myosin and Tropomyosin, then Titin, and lastly Myomesin and α-Actinin (Figure 5B–F and Figure 5—source data 1–5).

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Figure 5—source data 1

Information extracted from multicolor immunofluorescence of individual myofibers to infer the order of sarcomere protein disorganization in jerboa foot muscles.

‘Good’ represents striated localization of each protein to the sarcomere, and ‘bad’ refers to no distinguishable banded pattern of protein expression. In group 1, we saw myofibers with all three proteins properly localized suggesting disorganization follows an initial state of proper localization. When we compared the dRM and Drm categories, we saw loss of Desmin localization when Tropomyosin and Myosin were ‘good’ and almost no myofibers where Desmin was good and the others were bad. This suggests that Desmin is disorganized prior to Tropomyosin and Myosin. In group 2a, there were cells in the rmT category and almost none in the RMt category, suggesting Tropomyosin and Myosin are disorganized prior to Titin. Group 2b illustrates that both categories RmT and rMT appeared at similar frequency, suggesting it is unclear whether Tropomyosin or Myosin become disorganized prior to the other. In group 3, there were cells in the mtA category and none in the MTa category, suggesting Titin becomes disorganized before Alpha-actinin. Similarly, in group 4, there were cells in the mtY category but not in the MTy category suggesting Titin becomes disorganized before Myomesin. Due to shared antibody isotype for Alpha-actinin and Myomesin, the order of disorganization between these two could not be discerned. See Figure 5—source data 2–5 for full details of the percentage of myofibers in each category for each combination of multicolor immunofluorescence.

https://doi.org/10.7554/eLife.50645.016

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Figure 5—source data 2

Percentage of myofibers in each category for Desmin, Tropomyosin, Myosin, and Titin multicolor immunofluorescence of jerboa hand and foot muscles at three postnatal stages.

For the combination of Desmin, Tropomyosin and Myosin: P0 (two animals) hand n = 78 myofibers and foot n = 113 myofibers; P2 (two animals) hand n = 56 and foot n = 54 myofibers; P4 (two animals) hand n = 69 and foot n = 117 myofibers. For the combination of Desmin, Myosin, and Titin: P0 (two animals) hand n = 48 and foot n = 96 myofibers; P2 (two animals) hand n = 71 and foot n = 77 myofibers; P4 (two animals) hand n = 63 and foot n = 92 myofibers.

https://doi.org/10.7554/eLife.50645.017

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Figure 5—source data 3

Percentage of myofibers in each category for Tropomyosin, Myosin, and Titin multicolor immunofluorescence of jerboa hand and foot muscles at three postnatal stages.

P0 hand (three animals; n = 125 myofibers) and foot (four animals; n = 225 myofibers); P2 (four animals) hand n = 118 and foot n = 183 myofibers; P4 (three animals) hand n = 104 and foot n = 172 myofibers.

https://doi.org/10.7554/eLife.50645.018

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Figure 5—source data 4

Percentage of myofibers in each category for Myosin, Titin, and Alpha-actinin multicolor immunofluorescence of jerboa hand and foot muscles at three postnatal stages.

P0 hand (three animals) n = 156 and foot n = 182 myofibers; P2 (four animals) hand n = 189 and foot n = 203 myofibers; P4 hand (three animals; n = 104 myofibers) and foot (four animals; n = 172 myofibers).

https://doi.org/10.7554/eLife.50645.019

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Figure 5—source data 5

Percentage of myofibers in each category for Myosin, Titin, Myomesin multicolor immunofluorescence of jerboa hand and foot muscles at three postnatal stages.

P0 (three animals) hand n = 130 and foot n = 159 myofibers; P2 (three animals) hand n = 126 and foot n = 142 myofibers; P4 hand (three animals; n = 98 myofibers) and foot (four animals; n = 164 myofibers).

https://doi.org/10.7554/eLife.50645.020

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Desmin forms a filamentous network that connects parallel sarcomeres to one another and coordinates myofibril contraction within cells and between neighboring cells (Bär et al., 2004; Capetanaki et al., 2015; Goldfarb et al., 2008). Mutations that cause desminopathies illustrate that Desmin is essential to maintain sarcomere integrity (Clemen et al., 2013). In mouse models of muscle atrophy triggered by fasting or denervation, phosphorylation of Desmin removes the protein from the sarcomere and targets it for ubiquitination and proteolytic degradation prior to degradation of other sarcomere proteins (Volodin et al., 2017). The observation that Desmin is the first of an ordered sarcomere disassembly in the jerboa foot may reflect targeted degradation of muscle proteins that is similar to muscle atrophy.

The ubiquitin-proteasome system is the main pathway through which cellular proteins are degraded during muscle atrophy, and MuRF1 and Atrogin-1 are E3 ubiquitin ligases among the ‘atrogenes’ that are highly upregulated (Bodine and Baehr, 2014; Schiaffino et al., 2013). To test the hypothesis that muscle loss in the jerboa foot exhibits molecular hallmarks of atrophy, we performed quantitative reverse transcriptase PCR (qRT-PCR) of MuRF1 and Atrogin-1 mRNA from intrinsic foot muscles and the flexor digitorum superficialis (FDS) of the mouse and jerboa. The FDS, which originates in the autopod during embryogenesis and translocates to the forearm (Huang et al., 2013), is the most easily dissected of the analogous forelimb muscles and serves as a control for typical muscle maturation in both species. When normalized to expression in the FDS at birth of each species, Atrogin-1 expression is 3.1-fold higher in the jerboa foot at P3 (Figure 5G). MuRF1 mRNA expression is already significantly elevated at birth and remains elevated at P3 (Figure 5H).

The NF-κB pathway is an upstream mediator of skeletal muscle atrophy (Li et al., 2008) and is both necessary and sufficient to induce MuRF1 expression (Cai et al., 2004; Wu et al., 2014). To lend further support to the hypothesis that jerboa foot muscle loss involves an ‘atrophy-like’ mechanism, we performed qRT-PCR of NF-κB2 and its binding partner, Relb. We observed that each gene is expressed greater than three-fold higher in jerboa foot at birth and at P3 (Figure 5—source data 1). The progressively disordered ultrastructure of the sarcomere that begins with loss of Desmin localization, the increased expression of multiple genes that are typically upregulated during atrophy, and the lack of evidence for cell death or macrophage infiltration are consistent with observations of atrophying muscle in mice and rats (Volodin et al., 2017; Bonaldo and Sandri, 2013; Sakuma et al., 2015; von Haehling et al., 2010).

Despite the similarities to muscle atrophy, myofiber loss in the jerboa foot does not seem to be simply explained by an atrophic response to denervation. First, and in contrast to the rapid rate of jerboa foot myofiber loss, chronic denervation in mice (100 days after nerve transection at P14) reduced the size but not the number of individual myofibers (Moschella and Ontell, 1987). Additionally, we found that the post-synaptic Acetylcholine Receptor (AchR) exclusively coincides with the presynaptic neuronal protein, Synaptophysin, in neonatal jerboa foot muscles (Figure 5—figure supplement 1). In the mouse, AchR clusters are present in a broad domain of fetal muscle prior to innervation and are refined to nerve terminals in response to chemical synapse activity before birth (Yang et al., 2001). The refinement of AchR clusters in jerboa foot muscles suggests that the muscles are not only innervated at birth but are also responsive to motor inputs.

Beginning only with knowledge of adult jerboa foot anatomy, we reached evidence for a cellular mechanism of intrinsic muscle loss during neonatal development that is surprising in the context of what is known about muscle development and pathology in more traditional laboratory species. Although we found multinucleated myofibers in the feet of neonatal jerboas, all muscle protein expression rapidly disappears from the jerboa foot shortly after birth. We were perplexed to find no evidence of apoptotic or necrotic cell death by a variety of assays and throughout the time when muscle cells are lost, nor did we observe immune cells that are commonly recruited to clear the remains of dying cells. Instead, we saw structural and molecular similarities to muscle atrophy, although atrophy in young mice leads to reduced myofiber size rather than number as in the jerboa (Bruusgaard and Gundersen, 2008; Moschella and Ontell, 1987).

In an effort to functionally connect the atrogenes, MuRF1 and Atrogin-1, to a mechanism of muscle loss, we endeavored to determine whether knocking down both genes in jerboa foot muscle in vivo could be sufficient to rescue myofibers or to delay their loss. Although we developed a system to validate shRNAs targeting each jerboa gene after first inducing MuRF1 and Atrogin-1 expression in cultured primary jerboa myofibers, shRNA delivery into rodent neonatal foot muscle was not feasible (not shown). Lentiviral injection bathes the ensheathing muscle connective tissue, and infection rarely reaches myofibers of these small muscles in either mouse or jerboa. Plasmid injection and electroporation, which is feasible in adult rodent feet (DiFranco et al., 2009), is not efficient in neonatal feet of either species. This is likely because the neonatal foot lacks a cavity to contain injected DNA, provided by separation of the overlying skin in mature animals, which is required for efficient plasmid uptake by electroporation (Krull, 2004).

Even if genetic manipulations were technically feasible, intrinsic foot muscle loss first appeared in the jerboa lineage more than 20 million years ago. For phenotypes that diverged over such long evolutionary distances, a large set of genes and mechanical acommodation of integrated tissues are likely at play, which were collectively honed by millions of years of evolution; thus, manipulation of one or two genes may not be sufficient to rescue or delay muscle loss. It is therefore important to consider that a standard of ‘molecular mechanism’ applied to the experimental manipulations of traditional laboratory species may not be appropriate in the context of understanding complex macroevolutionary processes.

As for why the phenotype is limited to the distal limb, it is possible that disuse contributes to jerboa foot muscle loss, since jerboas and ungulates each fuse metatarsals into a single cannon bone, which would be expected to physically impair lateral motion of the digit elements (Cunningham, 1883; Moore et al., 2015). However, the rapid and complete loss of myofibers in the neonatal jerboa foot does not appear to simply reflect a species-level difference in the animal’s generalized response to disuse atrophy, since hindlimb denervation or immobilization in adult jerboas causes gradual loss of muscle mass, primarily through a significant reduction in the diameter of individual myofibers (AlWohaib and Alnaqeeb, 1997; Aryan and Alnaqeeb, 2002). These observations are very similar to what has been shown in disuse atrophy models in mice and in rats (Bonaldo and Sandri, 2013; Moschella and Ontell, 1987) and differ from what we see in the neonatal foot.

Why would an embryo expend energy to form muscles that are almost immediately lost? The formation and subsequent loss of intrinsic foot muscles in jerboas and hooved animals may simply reflect a series of chance events in each lineage with no fitness cost, or these similarities in multiple species may reveal true developmental constraints. Muscle is not required for autopod tendon formation or maintenance in mice, but the tendons that develop in a muscle-less or a paralyzed mouse are thinner and less well organized (Huang et al., 2015). It is therefore possible that muscle is initially required in the fetus and neonate for tendons to establish sufficient architecture from origin to insertion so that the tendon, after further growth, can withstand high locomotor forces in the adult (Lochner et al., 1980; Moore et al., 2017).

Regardless of whether these nascent muscles serve an essential purpose, we are left wondering what is the ultimate fate of jerboa foot myofibers. If these cells do indeed die, perhaps death is too rapid for detection. However, programmed cell death is thought to occur over hours or even days from the initial trigger to the final corpse (Green, 2005). Alternatively, death may result from a mechanism that does not proceed through DNA fragmentation, plasma membrane permeability, macrophage recruitment, or stereotyped ultrastructural changes, and yet this would seem to eliminate most known forms of regulated cell death (Galluzzi et al., 2007).

Alternatively, multinucleated myofibers may transform to another cellular identity after degrading all sarcomere proteins. Although a fate transformation would be surprising, it would not be without precedent. The electric organ of fish that can produce an electric field (e.g. knifefish and elephantfish) is thought to be derived from skeletal muscle. Electrocytes of Sternopygus macrurus express skeletal muscle Actin, Desmin, and α-Actinin, and electrocytes of Paramormyrops kingsleyae retain sarcomeres that are disarrayed and non-contractile (Gallant et al., 2014; Unguez and Zakon, 1998). If myofibers in the jerboa foot indeed transdifferentiate, it is possible that they transform into the pro-Collagen I positive fibroblasts that are entangled with the filamentous aggregates, although these could also be phagocytic fibroblasts recruited to consume the enucleate detritus without stimulating inflammation (Heredia et al., 2013; Joe et al., 2010; Monks et al., 2005; Schwegler et al., 2015). Unfortunately, the lineage labeling approaches required to track the ultimate fate of jerboa myofibers are exceptionally challenging in this non-traditional animal model.

It is clear, however, that regardless of the ultimate fate of jerboa foot myofibers, their path passes through a phase marked by cell biology that is typical of atrophy, including the ordered disassembly of sarcomeres and expression of the E3 ubiquitin ligases, MuRF1 and Atrogin-1. However, skeletal muscle atrophy is typically associated with pathology in the context of disuse, nerve injury, starvation, or disease. In this context, we were struck by a statement in the 1883 anatomical description of the fetal and adult suspensory ligament of four species of hooved mammals: ‘It is an instance of pathological change assisting a morphological process’ (emphasis his) (Cunningham, 1883). Indeed, there are remarkable similarities in the histology of jerboa and horse foot muscle compared to human clinical observations of tissue remodeling that follows rotator cuff tear characterized by muscle atrophy, myofiber loss, and fibrosis (Souza et al., 2010; Gibbons et al., 2017).

Foot muscle atrophy in the jerboa may be one of many cellular responses associated with injury or disease in humans that is utilized in the normal development and physiology of other species. These data suggest that there is less of a clear divide between natural and pathological states than typically thought. Studies of non-traditional species may not only reveal the mechanisms of evolutionary malleability, but may also advance our understanding of fundamental biological processes that are typically associated with pathological conditions.

All raw images and other associated data for this manuscript, including TEM, immunofluorescence, and qRT-PCR, have been curated and deposited with Zenodo. They can be found at the object identifier https://doi.org/10.5281/zenodo.3404257.

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Author details

1. Mai P Tran

   Division of Biological Sciences, Section of Cellular and Developmental Biology, University of California, San Diego, La Jolla, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-7127-4947

2. Rio Tsutsumi

   Division of Biological Sciences, Section of Cellular and Developmental Biology, University of California, San Diego, La Jolla, United States

   Contribution

   Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-9473-3923

3. Joel M Erberich

   Division of Biological Sciences, Section of Cellular and Developmental Biology, University of California, San Diego, La Jolla, United States

   Contribution

   Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

4. Kevin D Chen

   Division of Biological Sciences, Section of Cellular and Developmental Biology, University of California, San Diego, La Jolla, United States

   Contribution

   Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

5. Michelle D Flores

   Division of Biological Sciences, Section of Cellular and Developmental Biology, University of California, San Diego, La Jolla, United States

   Contribution

   Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

6. Kimberly L Cooper

   Division of Biological Sciences, Section of Cellular and Developmental Biology, University of California, San Diego, La Jolla, United States

   Contribution

   Conceptualization, Formal analysis, Supervision, Funding acquisition, Project administration, Writing—review and editing

   For correspondence

   kcooper@ucsd.edu

   Competing interests

   is on the science advisory board for Synbal, Inc, a company pursuing the use of active genetics technologies in laboratory rodents. This activity is unrelated to the work in this manuscript

   "This ORCID iD identifies the author of this article:" 0000-0001-5892-8838

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