Keywords

1.1 Introduction

As the building blocks for skeletal muscle during development, growth and regeneration (Anderson 2006; Mauro 1961; Mauro et al. 1970; Yablonka-Reuveni 2011), satellite cells play a fundamental role in muscle biology and animal function. They can respond dramatically and continuously to alterations in physiological demands on muscle, and to injuries and changes in nutritional constraints during adaptation. It is well-established that growth factors and cytokines have important roles in the repair of skeletal muscle (Karalaki et al. 2009), and that they have differential effects at various stages of myogenesis. Hepatocyte growth factor (HGF) for instance, is a protein made in myocytes and myoblasts (Charge and Rudnicki 2004; Sheehan et al. 2000; Tatsumi et al. 1998) and it promotes their proliferation while inhibiting myotube formation and differentiation in culture and regeneration.

HGF binds to the c-met receptor expressed on myoblasts and on satellite cells; that binding inhibits expression of the muscle regulatory genes, MyoD and myogenin, and the structural protein, myosin heavy chain (Anastasi et al. 1997; Gal-Levi et al. 1998; Leshem et al. 2000; Miller et al. 2000). By comparison, other growth factors also promote proliferation of myogenic cells, for example insulin-like growth factor (IGF-1) and fibroblast growth factor (FGF-2) and then promote myotube differentiation (Allen and Boxhorn 1989; Florini and Magri 1989; Hayashi et al. 2004).

Distinctions among growth factors in their activity in skeletal muscle were first explored in now-classic in vitro culture studies of proliferation and differentiation (Allen et al. 1995b; 1979) to understand the regulation of muscle growth. Studies of HGF, FGF and transforming growth factor beta (TGF-β) on proliferation and differentiation (Allen et al. 1995b; Allen and Boxhorn 1989; Allen and Rankin 1990; Johnson and Allen 1993; 1995) opened the field to studies of different stages in the myogenic regulatory program. As revealed at the time, the sequence of regulatory gene expression was delayed in cultures of satellite cells from muscle of old rats, pending a latent period between isolation of the quiescent satellite cells from muscle and their activation and proliferation in culture (Smith et al. 1994). Bischoff used a crushed-muscle extract (CME) preparation to induce activation of satellite cells on isolated fibers prepared for culture, and that extract was also shown to enhance muscle regeneration and satellite cell activity in vivo (Bischoff 1986a, b; Bischoff and Heintz 1994).

Early in situ hybridization experiments demonstrated that HGF was regulated during muscle development and then transiently expressed following ischemia-induced muscle injury and repair (Jennische et al. 1993). However, it was the notion that live and dead muscle fibers have distinctive interactions with their resident satellite cells and that living, intact fibers produce a factor that activates satellite cells and stimulates their transition from G0 to the cell cycle (Bischoff 1990). This was confirmed later, in studies of activation on fibers exposed to CME or the myotoxin, marcaine (Anderson and Pilipowicz 2002); those studies led to the identification of HGF as the activating factor in CME (Tatsumi et al. 1998). Subsequently, HGF-c-met interactions were examined in the context of paracrine signaling (Anastasi et al. 1997). More recently, an engineered high-affinity HGF with two c-met binding sites called Magic-Factor 1, has been shown to stimulate hypertrophy through anti-apoptotic pathways (Cassano et al. 2008). Current literature gives HGF a very prominent profile in processes mediating the balance between the activation of satellite cells and their return to quiescence (bou-Khalil et al. 2010; Chazaud 2010; Yamada et al. 2010), although other gene products and genetic background clearly impact satellite cell populations and their functionality in healthy and diseased muscle (Fukada et al. 2010; 2013).

Satellite cell activation is initiated through a cascade of calcium-dependent signals that release HGF from the extracellular matrix (Hara et al. 2012; Tatsumi et al. 2009b; Tatsumi 2010). Injury-induced activation leads to those processes involved in regeneration of muscle tissue, including angiogenesis, nerve sprouting and innervation, matrix remodelling and myogenesis, itself. All these processes impact directly on the outcome of tissue repair and the level of function attained by that tissue. Since the identification of nitric oxide as a potent mediator of satellite cell activation in mouse muscle (Anderson 2000; Wozniak and Anderson 2007; 2009), many roles of HGF and HGF-mediated signaling during the regeneration of muscle tissue have been reported. However, the scope of differences in HGF-mediated processes during activation and regeneration in different species, and many of the key aspects of chemical, mechanical, growth-factor- and chemokine-mediated signaling that affect muscle plasticity and function are not as well established across species, despite their importance in shaping muscle adaptation and the wide range of life-history challenges met by different species.

Capabilities of research using the single-fiber culture model to study HGF-mediated activation in mammalian and zebrafish muscle provide a strong platform for future exploration of HGF-induced signaling in development and muscle-tissue repair. This chapter provides a focused review of literature that relates to HGF-mediated mechanisms important in muscle growth and regeneration. Many studies demonstrate the interdependence of muscle structure, development and phenotype by modeling their combined impact on the activation of satellite cells. Evolutionary distinctions in activation by nitric oxide signaling (Anderson 2000) and possible species-dependent contributions of satellite cell regulation may contribute to determining the phenotypic pattern of muscle tissue expansion.

1.2 Satellite Cell Activation

The cytoskeletal transmembrane dystroglycan complex (DGC) anchors nitric oxide synthase (NOS)-1 close to the muscle fiber membrane. The DGC stabilizes the membrane of contracting muscle fibers by linking the actin cytoskeleton through the membrane to the surrounding matrix (Barresi and Campbell 2006; Campbell and Stull 2003; Durbeej and Campbell 2002; Ervasti and Campbell 1991; 1993; Matsumura et al. 1993). Changes in dystrophin, NOS-1 and transmembrane β-dystroglycan in the DGC, between muscles of wild-type, dystrophin-deficient and old mice accompany changes in satellite cell activation and cycling in muscle development, atrophy and regeneration (Janke et al. 2013; Leiter et al. 2012; Wozniak et al. 2003; Wozniak and Anderson 2007). We recently identified that the DGC develops in relation to muscle loading (Janke et al. 2013).

Satellite cells are precursor or stem cells on muscle fibers (Anderson 2006; Seale and Rudnicki 2000; Yablonka-Reuveni 2011) and are the source of cells for growth and regeneration (Collins et al. 2005; Collins and Partridge 2005). In adult mice, satellite cells are mitotically quiescent (G0); once activated to cycle, they form myoblasts that commit to the muscle lineage, divide, differentiate and fuse to extend existing fibers or form new ones. By asymmetrical division, some myoblasts return to a satellite position on fibers to renew the stem-cell niche (Collins et al. 2005; Kuang et al. 2007; Rudnicki et al. 2008). As fiber nuclei are post-mitotic, satellite cell activation is critical for muscle regeneration and plasticity.

Satellite cells are activated to cycle via signaling through the potent gaseous signal of nitric oxide release (Anderson 2000). Time-course studies of satellite cell activation using single muscle fibers that were mechanically stretched in culture revealed functional satellite-cell heterogeneity related to mRNA expression of c-met, the receptor for HGF (Fig. 1.1), and that loss of dystrophin appears to deplete particular subpopulations (Anderson and Wozniak 2004; Beauchamp et al. 1999; Collins et al. 2007; Heslop et al. 2000; Wozniak et al. 2003; Wozniak and Anderson 2007; 2009). Changes in NOS-1 expression or activity reduce or delay activation by injury, and then perturb the muscle-regeneration cascade (Anderson 2000; Wozniak and Anderson 2007). NOS-1 down-regulation in dystrophic muscle (Brenman et al. 1995) occurs with muscle loading in development and is not a direct result of the genetic loss of dystrophin (Janke et al. 2013). The mechano-sensitive NOS-1 protein releases nitric oxide after stretch; this liberates HGF from the fiber matrix. HGF binds to the c-met receptor on satellite cells, initiating satellite cell cycling and movement (Siegel et al. 2009; 2011; Tatsumi et al. 1998; 2002; Wozniak et al. 2003; 2005; Wozniak and Anderson 2007; 2009). Satellite cells also express dystrophin (Anderson et al. 1991), loss of which was shown recently to particularly impact satellite cell polarity in mitotic division and disease progression and regeneration in muscular dystrophy (Dumont et al. 2015).

Fig. 1.1
figure 1

Satellite cell heterogeneity on fibers. Zebrafish fibers were isolated, plated on collagen and incubated with or without cyclical stretching using a FlexCell vacuum-system apparatus. Fibers were fixed after 0 h, 3 of stretching or after 24 h (3 h stretching plus 21 h further incubation) before immunostaining for c-met protein. The number of c-met-positive satellite cells on each fiber was plotted as a frequency distribution for control fibers (without stretch) and fibers immediately or 21 h after stretch (Zhang and Anderson 2014). Graphs show significant heterogeneity for the distribution of c-met-positive satellite cells as a population on fibers (at least 67 fibers per group)

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The level of c-met expression in muscle and the number of c-met expressing satellite cells on fibers increases after activating satellite cells in wild-type mouse muscle (Anderson and Wozniak 2004; Wozniak et al. 2003). Interestingly, HGF is expressed in many types of cells in muscle tissue including skeletal, smooth and cardiac muscle cells (Fig. 1.2). HGF is also expressed by infiltrating mononuclear inflammatory cells in regenerating mouse muscle (Sakaguchi et al. 2014) and in human muscle of patients with polymyositis/dermatomyositis (Sugiura et al. 2010). Interestingly, macrophages were reported to activate myogenic cells as early as 1999 (Merly et al. 1999). In cultures of CD56-expressing myoblasts from those patients, the level of HGF was reduced by treatment with TGFβ and anti-inflammatory treatment, and increased by treatment with interleukin-1α (Sugiura et al. 2010). HGF treatment of those cells in culture reduced their synthesis of pro-collagen 1, while interferon-gamma increased the expression of c-met, suggesting the use of HGF to promote regeneration of muscle in inflammatory conditions (Sugiura et al. 2010). HGF-mediated muscle cell proliferation was also suggested as an approach to promoting muscle growth for treatment of urinary incontinence (Sumino et al. 2007). Molecules including miRNA489 (Brack et al. 2009), FoxO1, TGFβ3 and myostatin promote quiescence and inhibit growth and cycling (Argiles et al. 2012; Huang et al. 2012; Reed et al. 2012; Thomas et al. 2000). The balance between quiescence and activation states in myogenic cells is influenced by epigenetic effects of the surrounding matrix that are mediated by signaling through the Wnt pathway and the methylation state of chromatin that is mediated in part by mixed lineage leukemia 5 (MLL5) (Dhawan and Rando 2005; Sebastian et al. 2009; Sellathurai et al. 2013; Srivastava et al. 2010; Subramaniam et al. 2013).

Fig. 1.2
figure 2

Hepatocyte growth factor (HGF) expression in control and dystrophic muscle. Muscle histology viewed by hematoxylin and eosin staining (a, e) and in situ hybridization for HGF) using digoxigenin-labeled riboprobe to identify mRNA (b–d, f–h) from wild-type C57BL10 mice (a–d) and dystrophin-deficient mdx mice (e–f). (a) Muscle fibers in control diaphragm. (b) Two satellite Fig. 1.2 (continued) cells (arrows) in tibialis anterior muscle contain cytoplasm densely stained for HGF transcript; less intense staining for HGF transcripts is visible around the periphery of myofibers. A nerve bundle is indicated by the crossed vertical arrow. (c) HGF expression is demonstrated in intrafusal fibers of a muscle spindle (arrow) and in smooth muscle around a blood vessel (crossed vertical arrow). (d) A small myotube (vertical arrow) is regenerating between intact fibers in the diaphragm of a control mouse. (e) Regenerating fibers (arrow) within sections of dystrophic tibialis anterior muscle. (f and g) Small regenerating fibers (arrows) are intensely stained for HGF mRNA in an inter-nuclear distribution in longitudinal section (g). (h) Cardiac myocytes from the atrium of a dystrophic mouse are stained for HGF transcripts (arrow). Bar = 10 μm (original magnification X200); bar in panel b applies to b–h (original magnification X400)

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1.3 Myogenic Cell Motility

Syndecans, especially syndecan 3 among the heparan sulphate proteoglycans (Casar et al. 2004; Cornelison et al. 2001; 2004; Pisconti et al. 2010; Rapraeger 2000), CD34, Eph-ephrin and Notch-delta interactions, Wnt, FGF2 and possibly Hedgehog (Xie et al. 2013) promote motility and cycling after activation (Alfaro et al. 2011; Ieronimakis et al. 2010; Luo et al. 2005; O’Brien et al. 2004; Stark et al. 2011). Syndecan-3 and dermatan sulfate are required for muscle regeneration and activation signaling (Casar et al. 2004; Villena and Brandan 2004). Interestingly, HGF induces the formation of lamellipodia and cell migration in cultures of C2C12 cells in a 3-dimensional assay. This is reported to occur through a balance of PI3-kinase-dependent signals in the WAVE2 N-WASP pathway that involve the formation of branched actin filaments and polarization of the plasma membrane (Kawamura et al. 2004). Such regulation likely plays a role in the fascinating HGF-dependent travel by activated satellite cells along muscle fibers that can be so beautifully demonstrated in fiber cultures (Siegel et al. 2009; 2011). This HGF-mediated migration may also interact with CD44-dependent regulation of the speed of satellite cell migration (Mylona et al. 2006). Involvement of glypican-1 in stabilizing HGF-c-met interactions via mechanisms related to lipid-raft formation is required for the optimal response of mouse C2C12 cells to HGF signaling and to stabilize CD44 (Gutierrez et al. 2014). HGF also induces expression of caveolin-1 which inhibits cell proliferation, and is proposed to regulate the activity of the ERK pathway in satellite cell mobility and proliferation during regeneration (Volonte et al. 2005).

Another distinction among different growth factor-mediated processes is that HGF appears not to influence fibrinolysis required for invasion of myogenic cells into a matrix, although the migration process is dependent on urokinase plasminogen activator activity which is in turn, regulated by FGF-2 and TGFβ (Fibbi et al. 2002). Possible similarities among different types of cells (e.g., myogenic cells and keratinocytes) suggest that many features of migration, may show distinctions based on the surrounding matrix or particular features of how factors are applied in an experiment (e.g., separately or in combination), as illustrated by use of a wound-assay culture system (Peplow and Chatterjee 2013). In a review of heparanase and syndecan-1 in tumours, the up-regulation of HGF expression by heparanase, and subsequent binding of HGF to syndecan-1 for degradation, was highlighted as a mechanism regulating growth factor activity (Ramani et al. 2013). It is possible that similar interactions during HGF-mediated responses by satellite cells play a role in the timing or dose-response of activation and migration during growth and repair of muscle and could point the way to designing novel therapeutics to promote those processes.

1.4 Zebrafish Single-Fiber Studies of Satellite Cell Activation

Temperature, environment and life-history events such as migratory behaviour in fish and birds and hibernation patterns in mammals, influence muscle plasticity and metabolic adaptation (Corrigan et al. 2011; Johnston 2006; Johnston and Hall 2004; Meyerrochow and Ingram 1993; Price et al. 2011; Wund et al. 2008). For instance, hibernation affects muscle mass or growth in some vertebrates and not in others. Interestingly, hibernation can protect muscle from atrophy that ensues with starvation, disuse/immobilization or even denervation. Hibernation apparently changes the neural regulation of catabolic pathways, such that decreases in muscle mass and fiber cross-sectional area in hibernating bears after denervation are only one third to one half as pronounced as in non-hibernating bears (Lin et al. 2012). Hibernation is also accompanied by down-regulation of autophagic and proteolytic pathways mediated by serum- and glucocorticoid-inducible kinase-1 (GSK1) (Andres-Mateos et al. 2013). Moreover, in studies of the 13-striped ground squirrel, while the level of myostatin, which typically represses satellite cell proliferation in non-hibernating animals and typically increases in starvation and disuse, it is surprising that in hibernating animals, myostatin levels do not change. Myostatin stays at normal pre-hibernation levels well into hibernation, and the marked increase only happens when the animal is beginning to arouse from topor, toward the end of hibernation (Brooks et al. 2011). Regulation of muscle atrophy is clearly overlain by complex metabolic changes somewhat separable from neural and other activity and/or endocrine influences. Such fascinating observations encourage in-depth comparative studies of muscle satellite-cell biology and the impact of various life-history events.

A key distinction in muscle-growth capacity between mammals and teleosts is the regulation of fiber number. In mammals, fiber number does not increase after birth; by comparison, in teleost fish, the number of fibers, especially fast fibers, rises nearly continuously with body size and muscle mass, as part of the process termed indeterminate growth (Hall et al. 2004; Johnston 2006; Johnston and Hall 2004). A leader in the field, IA Johnston, notes the major gap in our understanding of the impact of indeterminate growth and ectothermy on the regulation of muscle growth in teleosts versus mammals (Johnston et al. 2011).

On the basis of the growing body of work on mammalian muscle fibers, and long-standing questions about the regulation of muscle growth in fish, we modified the protocol for isolating mouse-muscle fibers to enable culture studies of zebrafish muscle fibers (Anderson et al. 2012) prepared with careful attention to Bischoff’s technique (Bischoff 1986b). Using single-fiber cultures from zebrafish, experiments revealed that satellite cell activation on zebrafish fibers is mediated by nitric oxide, HGF and mechanical-stretch signals (Zhang and Anderson 2014) (Fig. 1.3a, b), similar to fiber cultures from mouse (Anderson and Pilipowicz 2002; Wozniak et al. 2003; Wozniak and Anderson 2007). However, although the cascade of activation involved similar signals, peak activation in zebrafish fiber cultures occurred at a distinctly higher concentration of nitric oxide and a lower concentration of HGF, than in our earlier findings on activation in mouse-fiber cultures; values in fish vs. mouse fiber experiments were: 1 mM isosorbide dinitrate (2 molar equivalents of nitric oxide released) vs. 100 nM of the NOS substrate L-arginine (1 molar equivalent of nitric oxide produced); and 10 ng/mL vs. 25 ng/mL HGF in medium, respectively (Anderson and Pilipowicz 2002; Wozniak and Anderson 2007; Zhang and Anderson 2014). [Unfortunately, the direction of the comparative fish-mouse difference for response to nitric oxide is reversed in Zhang and Anderson 2014.] While the levels of nitric oxide and HGF extant in the two types of fibers is not known, the relative differences indicate there is a differential “tuning” of activation by the HGF-mediated cascade in mouse and fish fibers. The dose-response of HGF-induced activation is shifted to a lower peak concentration of exogenous HGF by stretching in culture (Fig. 1.3c), due to release of HGF from the low-affinity receptors in the extracellular matrix surrounding fibers (Tatsumi et al. 2001; 2002; Tatsumi and Allen 2004). Activation by exposure to HGF and by mechanical stretching in the fish fibers is additionally mediated by temperature. Results of studies using inhibition of HGF-c-met binding and NOS activity demonstrated that activation of satellite cells on fibers in culture is reduced at 21 °C compared to the level observed at 27 °C, the typical temperature for rearing zebrafish (Zhang and Anderson 2014).

Fig. 1.3
figure 3

Dose-response curves for satellite cell activation on fibers in culture. Muscle fibers were isolated from zebrafish (a, b) and mouse (c) and cultured in basal growth medium containing bromodeoxyuridine (BrdU) for 24 h in the presence of: (a) the NO donor isosorbide dinitrate, ISDN (0–2.5 mM); (b) HGF (0–25 ng/mL); and (c) HGF (0–30 ng/mL) with or without cyclical stretching. After 24 h incubation, fibers were fixed and immunostained for BrdU, the dose-response of satellite cell activation was determined as the number of BrdU-positive satellite cells per fiber. Note that peak activation by HGF occurred at a lower concentration in unstretched zebrafish than mouse fiber cultures (c vs. b), and that HGF-induced activation on mouse fibers shifts to a lower concentration of HGF in the presence of 3 h of stretching

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The interesting finding of a novel temperature-sensitive pattern of satellite cell activation on zebrafish fibers was accompanied by observation that the population of satellite cells identified by immunostaining studies for the c-met receptor protein, increased in size only 3 h after stretch (Zhang and Anderson 2014). This finding indicated that c-met is acting as an immediate early gene in satellite cell activation on zebrafish fibers, consistent with findings from mouse-fiber cultures (Wozniak et al. 2003). As well, there were more c-met-positive satellite cells on fish fibers, and they cycled faster than in mouse (Zhang and Anderson 2014). Very recent extension of these fiber-culture studies is now examining developmental aspects of interactions between fibers and their resident satellite cells.

The study demonstrated the sensitivity of the fiber-model system for testing the impact of brief or subtle changes in the fiber environment, on muscle growth and adaptive capacity. While there is clear conservation of regulatory pathways between zebrafish and mouse muscle, at least in the fiber-culture model system, these recent studies provide fresh insights on possible mechanisms of indeterminate muscle growth in fish that utilize fiber hyperplasia and hypertrophy, in comparison to post-natal muscle growth by hypertrophy of fibers in mammalian species. Such information could be a strong asset to aquaculture and food-meat production industries, and assist in further studies of the epigenetic interactions of the fiber cytoskeleton with satellite cell activity as they relate to understanding how the growth and repair of muscle tissue shape functional muscle plasticity.

1.5 Re-defining the Impact of Age on Satellite Cell Activation and Muscle-Growth Capacity

Previous research in this laboratory tracked the concurrent loss of muscle mass, strength, proliferation and fiber size in aging mice, with changes in NOS-1 and β-dystroglycan in the DGC and membrane function. Exercise increased levels of the protein NOS-1 in skeletal muscle and increased the proliferation of satellite cells when the intervention was given at eight but not 18 months-of-age, and the response was muscle-specific rather generalized in all muscles (Leiter et al. 2011). We also found that satellite cells were not activated in old muscle by the level of stretching that effectively activates them when younger muscle is stretched in culture, and this refractoriness to stretching increased with greater age (Leiter and Anderson 2010). The novel finding was that in muscles from old mice, proteins of the DGC were relocated and reduced in concentration (Leiter et al. 2012), and that remarkably, combined treatment with voluntary-running exercise and the nitric oxide-donor compound called isosorbide dinitrate, partly restored the beneficial muscle-building effects of exercise in the old mice, toward effects that are more typical of exercise effects on muscle in younger animals. This was despite the significant decline in the number of satellite cells expressing transcripts for myoD, myogenin or pax7 found in the gastrocnemius muscle of animals at 18 months-of-age, the age at which the combined treatment with ISDN plus exercise had started, compared to muscle at 3, 8 or 12 months-of-age (Fig. 1.4, Leiter & Anderson, unpublished data). Treatment also shifted the subcellular pattern of the β-dystroglycan and NOS-1 in muscle from the old mice, toward the localization pattern seen in muscle from much younger mice. This was concurrent with decreased permeability to serum immunoglobulins in the muscle of old mice; overall, the combined treatment of exercise and the nitric oxide donor resulted in a 25 % increase in muscle mass and increased fiber diameter in the old mice (Leiter et al. 2012). Without the increase in nitric oxide provided by the NO-donor treatment, exercised old mice displayed the typical loss of muscle mass and the distribution of the two proteins in the DGC had the pattern that was observed in muscle of old mice. There may be potential for application of therapy based on nitric oxide, to improve the functional state of muscle growth capacity in aging or to assist in “pre-habilitation” of muscle prior to surgical procedures. This study shows the growth capacity of satellite cells in old muscle is retained. A comprehensive examination of the capacity for muscle regeneration in response to three different types of injury (freeze, crush and myotoxin) in very old mice, showed that the capacity of satellite cells in muscle of old or very old mice to provide fully functional regenerated muscle tissue can be retained if muscle damage has not affected the neurovascular supply (Lee et al. 2013). The possibility of progressive changes in HGF expression and HGF-mediated signaling in satellite cell activation has not been fully explored through the aging process.

Fig. 1.4
figure 4

Satellite cell populations decline with increasing age. (a) Gastrocnemius muscle from mice aged 3, 8, 12, and 18 months, showing satellite cells stained positive by combined in situ hybridization (ISH) using a cocktail of riboprobes for pax7 (a muscle-specification gene, expressed in quiescent and activated satellite cells), and myoD and myogenin RNA (muscle-regulatory genes Fig. 1.4 (continued) expressed by activated satellite cells). Satellite cells are directly juxtaposed to the sarcolemma of unstained fibers (arrows indicate satellite cells with a prominent nucleus). Insets show a negative-control section (arrow points to an unstained satellite cell) and a satellite cell on an atrophic fiber (at 3 and 18 months, respectively). Bar = 10 μm. (b) Satellite cell density (number of ISH-positive mononuclear cells in the satellite position per field, mean ± SEM, 247–303 fields per group) was highest at 12 months. Brackets indicate significant differences (p < 0.001). (c) The percentage of satellite cells detected by in situ hybridization for each age group, plotted against satellite cell density (the number of ISH+ cells per field). The density of satellite cells expressing at least one of myoD, myogenin or pax7 declined with increasing age (p < 0.01, Chi-square statistics); 18-mo gastrocnemius muscles had the highest frequency of fields without any satellite cells (Leiter and Anderson, unpublished)

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1.6 Integrating Growth, Satellite-Cell Behaviour and Cytoskeletal Development

Cytoskeletal NOS-1 and β-dystroglycan develop earlier in mouse diaphragm than limb muscle. This pattern changes with loss of dystrophin from the DGC (Janke et al. 2013). Such findings support the idea that differential muscle loading, which is higher and earlier in diaphragm than in limb muscle, and the structure of the DGC that differs in important but seemingly subtle ways from the DGC in the central nervous system, may explain muscle-specific variations in muscular dystrophy (Janke et al. 2013; Snow et al. 2013a; b). Previous reports on c-met regulation of cell functions, such as migration of invasive tumour cells, limb muscle precursors in development, and tubule formation by epithelial cells, in regeneration of liver and proliferation of liver and placental cells (Trusolino et al. 2010), and the outcome of muscle satellite-cell activation (Leiter et al. 2012; Webster and Fan 2013; Wozniak and Anderson 2007), clearly demonstrate the importance of the cytoskeleton in mediating that regulation.

Semaphorin3A (Sema3A) is a paracrine neuro-repellant protein, novel to muscle biology since 2009 (Tatsumi et al. 2009a). Sema3A is implicated in an axon-guidance role during the regeneration of skeletal muscle. HGF induces Sema3A secretion by activated satellite cells, but interestingly this does not occur through HGF binding to c-met. These findings suggest that satellite cells serve to collect and integrate activating and quiescence factors and the functional capacity for nerve-independent and nerve-dependent repair via Sema3A (Do et al. 2011; 2012; Suzuki et al. 2013). Sema3A is up-regulated by HGF, peaks early in myoblast differentiation and could mediate the timing of axonal ingrowth to regenerating fibers. It is intriguing that Sema3A may act through a receptor in the plexinA2-myogenin pathway that is particularly active in slow muscle (Suzuki et al. 2013; Tatsumi et al. 2009a). This field of study opens exciting new ways to study nerve-muscle interplay during muscle development. For example, it would be fascinating to examine the expression and behaviour of satellite cells on developing fibers at different life-stages from a number of species, as the stage that is particularly critical to organism survival may differ in different species. Satellite cell activity could be explored using an architectural approach, by examining the expression of HGF, Sema3A and other regulatory and structural proteins by satellite cells and their activation status, according to their distance from vessels or nerve-muscle junctions along a fiber in culture or in vivo. Such studies could help identify factors that contribute to producing a heterogeneous population of stem cells in a muscle, and how the behaviour of satellite cells during development and activation in growth, may influence the muscle phenotype that develops in different species.

Characterization of the satellite cell populations in vivo on adult and developing muscle fibers on non-mammalian species is an important and achievable goal, given the acceleration of the satellite cell field that has resulted in the past three decades of focused research using rodent models of human disease. Understanding how satellite cell populations vary in size and activation state by species, fiber type and through development can now be examined in relation to differences in the environment, function and behaviour of an organism. A good “baseline” understanding is still missing information about cross-species comparisons of satellite cell populations in adult and developing muscle that explain the differential responses by muscle to activating stimuli and environmental change, and the distinctive capacity for nearly life-long formation of new fibers in some species. Studies of the differential growth by fish developing in cold versus warm water, and the genes expressed by muscle tissue or fibers (Johnston 2006; Johnston et al. 2009; Johnston and Hall 2004; Steinbacher et al. 2011) have not fully explained the important biology of muscle satellite cells, the proteins they make or bind and their response to changing conditions. That information is crucial to anticipating animal responses to climate change, toxic habitat or tissue injury.

By studying the regulation of satellite cell activation signaling on single fibers in vitro, experiments could be conducted to examine species-specific differences in the regulation of satellite cell activation and how they may relate to features of the DGC. A synthesis of information on how the DGC and internal cytoskeleton shape nuclear position and development is also missing from the literature, despite the critical roles of those proteins in regulating satellite cell activation (for example, see Janke et al. 2013; Wozniak and Anderson 2007) and muscle fiber integrity.

A further gap in our knowledge relates to understanding the basis for central nucleation in regenerated fibers. Typically in development, as myotubes differentiate and receive innervation, nuclei shift from the central region of myotubes toward the periphery of fibers (just inside the DGC). Nesprin1 may be involved in that relocation. Nesprins are part of a complex that links the outer nuclear envelope with the actin cytoskeleton, and preliminary observations showed nesprin1 distribution differed between the peripheral nuclei in normal adult fibers and those centrally located nuclei in regenerated fibers in the absence of dystrophin (Kaluzny and Anderson, unpublished). The positioning of satellite cells in proximity to in-growing nerves and blood vessels (Duxson and Sheard 1995; Harris et al. 1989), particularly at the nerve-muscle junction or endplate regions, is also reported to relate to the location of giant nesprinG proteins (>1000 kb) (Burke and Roux 2009; Crisp et al. 2006; Roux et al. 2009; Starr and Fischer 2005; Tzur et al. 2006; Zhang et al. 2007). It would be fascinating to examine nerve-muscle junctions on fibers in culture, in relation to the position and activity of the resident satellite cells and myonuclei at important stages in development and regeneration, and also the interaction of satellite cells with angiogenic processes and vascular endothelial growth factor through those processes.

Since muscle stem-cell activity on fibers is so strongly linked with the ultimate outcome of muscle growth, physiology and adaption to neurovascular regulation (Allen et al. 1995a), it will be important to take new advantage from the fiber-model system to interrogate the relationship between satellite cell activation, behaviour and muscle phenotype. Fiber cultures could be used to study whether growth factors such as HGF or cytokines derived from the many tissue types in a muscle belly, direct satellite cell populations toward regenerating fibers that take up a particular fast or slower phenotype after the repair. Alternatively, satellite cells may be the prime organizers of key stages in the regenerative response through their integration of incoming signals and subsequent release of factors that actively signal to the local or systemic environment. For example, an interesting report examined systemic interactions mediating HGF expression, and found that within an hour of a muscle injury, both liver and spleen had up-regulated HGF expression and production of the inactive 90 kDa form of HGF protein. Serum from rats undergoing muscle regeneration was also able to induce expression of HGF transcripts in cells cultured from rat spleen, suggesting a role for an endocrine-level regulation of satellite cell behaviour (Suzuki et al. 2002). Fiber cultures could be used to screen for myogenic or growth-promoting effects of a range of compounds or extracts from natural or engineered tissues. Another report noted the importance of HGF in recruiting myogenic cells into injured muscle by mediating their interactions with vascular endothelial cells (Corti et al. 2001). With the recent discovery that satellite cells make Sema3A during activation and regeneration (Do et al. 2011; Suzuki et al. 2013; Tatsumi et al. 2009a), it is time to open the window wide for study of this novel aspect of muscle biology and nerve-muscle interaction.

Experiments on HGF-induced Sema3A production by satellite cells on single fibers, classified by immunostaining (Anderson et al. 1988; Merrifield and Atkinson 2000) and structure from mouse and zebrafish could be designed to manipulate satellite cell activation by HGF and nitric oxide, inhibit NOS-1 or block c-met and other cytokine and growth-factor receptors. Time-course studies such as those that tracked repair (Anderson et al. 1998; Grounds 1987; Grounds et al. 1992; Grounds and McGeachie 1987; 1992; McGeachie and Grounds 1987; McIntosh et al. 1998a, b, c; Moor et al. 2000; Smythe et al. 2008) may identify further fiber-type (Suzuki et al. 2013) or differential species effects on activation and Sema3A synthesis. Studies of larval muscle development in various fish species may identify species-specific patterns of HGF and Sema3A signaling that influence receptor clustering at nerve-muscle junctions (Lomo 2003), motor synapse elimination (Busetto et al. 2003; Jansen and Fladby 1990; Missias et al. 1996) and satellite cell positioning near synapses and vessels (Duxson et al. 1986; Duxson and Sheard 1995; Paul et al. 2004). HGF is known to increase blood vessel formation if it is incorporated in local-release acellular grafts after nerve sectioning (Li et al. 2008; Watanabe and Okada 1967). HGF also promotes angiogenesis and muscle-fascicle organization, as studied in a gel matrix around engrafted myoblasts (Barbero et al. 2001), and HGF expression increases during hyperbaric oxygen treatment to improve muscle repair after severe ischemia (Asano et al. 2007). Both findings are consistent with a recent report on the impact of oxygen levels on HGF-promoter activity and satellite cell-mediated angiogenesis (Flann et al. 2014).

Interestingly, acetylcholine receptors cluster on the muscle membrane before, not after developing fibers are innervated, so the pattern of fiber formation may influence the growth of motor axons and synapse formation (Arber et al. 2002). There are also distinct innervation patterns in parasitic and free-living lamprey (Atkins and Pezzementi 1993; Pezzementi and Chatonnet 2010) that have not been studied in relation to satellite cells on those muscle fibers. Such research could pursue the impact of satellite cell-derived factors such as HGF on driving angiogenesis (Ferrara et al. 2003; Williams and Annex 2004) or axon-guidance, and provide an exciting revision of classical thinking that is untested at the level of satellite-cell biology on fibers.

The precise state of satellite cells in a particular model system during assay, is critically important in interpreting findings. Satellite cells can be examined in many model systems: single-cell primary cultures, cell homogenates after expansion or differential plating, myoblasts derived from bulk cultures of mixed muscle cells, fibers cultured either free-floating or attached to a substrate, and in sections of normal, developing or regenerating muscle. In each system, the previous activities of those cells, the experimental conditions, and the way satellite cells or fibers are isolated as reagents for use in each assay all affect research findings and their interpretation. For instance, the mechanism of fiber isolation impacts the activation status of their resident satellite cells. Isolation to study satellite cell motility in response to HGF can effectively use cultures in which the preparation protocol activates the satellite cells (Siegel et al. 2009). By comparison, studies of the transition from quiescence to activation need preparations that avoid shaking or other activating stimuli (Anderson et al. 2012; Wozniak and Anderson 2005). The status of satellite cells, quiescent, activated, mobile and/or proliferative, needs to be verified in a control group under the same conditions (Wozniak et al. 2003; Wozniak and Anderson 2007). An understanding the status of satellite cells, such as their expression of muscle-regulatory genes (therefore activated), genes such as pax7 that specify muscle (Seale et al. 2000) or identify satellite cells (e.g., c-met, CD44, or CD56/NCAM) and their proliferation state, is as critical as experimental design in interpreting in vivo experiments. A recent report suggesting that HGF-c-met signaling may not be required for activation or proliferation of satellite cells (Webster and Fan 2013) did not control for a number of important features of experimental design: the interval between tamoxifen-knockdown of c-met gene activity and experimental assay; the possibility of residual c-met in satellite cells that were most stem-like or quiescent; satellite cell activation in different-aged mice at the time of injury; uncertain half-life of c-met; the activity of c-met as an immediate early gene (Wozniak and Anderson 2007; Zhang and Anderson 2014); and the complex implications of satellite-cell heterogeneity. Thus, the conclusions of that report regarding satellite cell activation are not well founded. Satellite cell behaviour following injury, stretching and isolation reflects the events of activation, including entry into the cell cycle and mobilization (Anderson 2000); the many molecular-genetic pathways and epigenetic and feedback mechanisms that regulate activation and/or are impacted by it, are yet to be fully established.

1.7 Conclusion

HGF activates satellite cells; its release from the muscle extracellular matrix is initially mediated via nitric oxide release, calcium-calmodulin signaling and matrix metalloproteinase activity after mechanical or injury-induced signals. Satellite cells respond by proliferating and making HGF and Sema3A; they also underpin the multi-directional signaling process between muscle fibers and proteins and cells in the large volume of tissue outside the endomysium. By using the single-fiber model system, single-cell physiology is accessible to comparative study and can give insights into the integral role of a heterogeneous population of satellite cells. Such research has potential to improve our ability to promote human muscle growth and repair, to prevent conditions such as age-related atrophy and to improve management and treatment of neuromuscular diseases. For animal and comparative physiologists, such basic muscle-cell biology studies could test new approaches to aquaculture and species conservation.