Regenerating Myofibers after an Acute Muscle Injury: Comparison
View Latest Version
Please note this is a comparison between Version 2 by Jason Zhu and Version 1 by Francis Xavier Pizza.

Injury to skeletal muscle through trauma, physical activity, or disease initiates a process called muscle regeneration. When injured myofibers undergo necrosis, muscle regeneration gives rise to myofibers that have myonuclei in a central position, which contrasts the normal, peripheral position of myonuclei. Myofibers with central myonuclei are called regenerating myofibers and are the hallmark feature of muscle regeneration. An important and underappreciated aspect of muscle regeneration is the maturation of regenerating myofibers into a normal sized myofiber with peripheral myonuclei. Strikingly, very little is known about processes that govern regenerating myofiber maturation after muscle injury. 

  • muscle regeneration
  • muscle repair
  • embryonic myogenesis

1. Embryonic and Postnatal Myogenesis vs. Muscle Regeneration

There is universal agreement that proliferation of muscle progenitor cells and myoblast–myoblast fusion are required for de novo myofiber formation in the embryo, fetus, and during muscle regeneration. Given this agreement, it is often assumed that cellular and molecular processes that facilitate myofiber maturation during muscle regeneration are analogous to those operating in the embryo, fetus, and after birth. This is a reasonable assumption as myogenic cell proliferation, myonuclear accretion, and hypertrophy of myofibers occurs during embryonic myogenesis, postnatal myogenesis, and muscle regeneration. However, myofibers maturing in the embryo, fetus, and after birth are phenotypically different from regenerating myofibers formed after an acute injury. The most striking differences between them are the position, organization, and number of myonuclei. The attainment of a normal myofiber size and morphology is also different during the course of postnatal myogenesis and muscle regeneration. These differences indicate that the cellular and molecular processes that regulate regenerating myofiber maturation are not identical to those that mediate myofiber maturation during embryonic and postnatal myogenesis.
To provide a framework for delineating cellular and molecular processes that are unique to muscle regeneration, the literature pertaining to indices of myofiber maturation during embryonic myogenesis, postnatal myogenesis, and muscle regeneration in mammals is summarized below. As knowledge of embryonic and postnatal myogenesis provides a foundation for understanding muscle regeneration, myofiber maturation during embryonic and postnatal myogenesis is discussed separately from muscle regeneration. When discussing regenerating myofibers, evidence pertaining to their maturation is discussed in the context of the paradigm of muscle regeneration, and comparisons are made to embryonic myogenesis and/or postnatal myogenesis.

2. Myonuclear Positioning during Embryonic and Postnatal Myogenesis

During embryonic myogenesis, myonuclei in developing myotubes/myofibers are dispersed evenly within a relatively small cytoplasmic area that contains isolated sarcomeres and/or developing myofibrils [62,69,77,78,82,95][1][2][3][4][5][6]. Transverse and longitudinal planes of skeletal muscle in the embryo or fetus indicate that myonuclei are centrally located in developing myotubes/myofibers [62,69,77,78,82,95][1][2][3][4][5][6]. The central position of myonuclei persists after myotubes/myofibers are innervated by a motor neuron [101,102][7][8].
Central myonuclei in myotubes/myofibers in the embryo/fetus move towards the sarcolemma during the course of the development [69,70,77,78][2][3][4][9]. The time scale of this repositioning, however, is unclear and appears to be temporally associated with myofibrillogenesis [69,77,82,103,104][2][3][5][10][11]. This temporal relationship most likely serves as the basis for the widely held belief for myonuclear repositioning during muscle development and regeneration. That is, the formation of myofibrils and function of sarcomeres within them is thought to propel myonuclei from a central to a peripheral position during the course of myofiber maturation. This premise has been supported by evidence of myonuclear trafficking during in vitro myogenesis [105][12]. Specifically, Roman et al. [105][12] reported that myofibrillar contraction, as well as desmin-mediated linking of myofibrils together, facilitated the migration of myonuclei towards the sarcolemma.
The repositioning of myonuclei from a central to a peripheral position during embryonic myogenesis is profound, as few, if any, myofibers at birth have central myonuclei [70,77,106,107,108,109][3][9][13][14][15][16]. Central myonuclei are also not normally observed during postnatal development [78,106,107[4][13][14][15][17][18][19][20][21],108,110,111,112,113,114], which is consistent with the number of myofibers within muscles being fixed at birth or shortly after [28,29,30,31][22][23][24][25]. The absence of central myonuclei during postnatal development is striking as myofibers rapidly add myonuclei within days of birth and continue to add myonuclei until adulthood [106,107,108,110,111,112,113,114][13][14][15][17][18][19][20][21] (discussed below in Section 4.3).
In summary, central positioning of myonuclei is a phenomenon that is restricted to embryonic myogenesis, as myonuclei are in a peripheral position during postnatal myogenesis. Simply put, repositioning of myonuclei from a central to a peripheral position occurs at a very early stage of myofiber maturation—that is, well before myofibers have reached their adult size and a full complement of myonuclei, myofibrils, organelles, and other components of their cytoplasm. Why myonuclear repositioning occurs at a very early stage of myofiber maturation is unclear. Is the movement of myonuclei from a central to a peripheral position during embryonic myogenesis a consequence of the formation and function of myofibrils? Or does the peripheral positioning of myonuclei prior to birth optimize molecular and cellular processes that facilitate myofiber maturation? If not, why do myonuclei added during postnatal myogenesis take a peripheral, rather than central, position in myofibers? Is their path to a central position during postnatal development blocked by a progressive increase in the number of myofibrils? These fundamentally important questions remain unanswered.

3. Myonuclear Positioning during Trauma-Induced Muscle Regeneration

Nuclear chains are the most striking, as well as the defining feature of regenerating myofibers. Myonuclei in nuclear chains are very closely aligned and, in many cases, appear to be in contact with each other after chemical or physical trauma in mice [83,86,87,88,89,90,91,115][26][27][28][29][30][31][32][33]. In contrast, the majority of central myonuclei during embryonic myogenesis appear to be separated from each other [69,77,82][2][3][5]. The number of myonuclei in nuclear chains of regenerating myofibers is notable, as well as variable. Specifically, the length of individual nuclear chains, a readout of the number of myonuclei in a single nuclear chain, varies considerably (40–90% of myofiber length) and increases during the course of regeneration [86][27]. As discussed later, the length of individual nuclear chains and the total number of nuclear chains in a regenerating myofiber can influence the detection of regenerating myofibers in transverse muscle sections.
Nuclear chains often run in parallel with each other and, in some cases, appear to converge/diverge to produce a ‘Y’ shaped appearance [86,89,92,115][27][30][33][34]. Parallel rows of nuclear chains are indicated in transverse muscle sections by the central position of two or three myonuclei within a myofiber, which is a common occurrence after chemical trauma in rodents. These organizational patterns of central myonuclei are unique to muscle regeneration, as no discernible rows or organizational patterns of myonuclei have been noted during embryonic myogenesis.
Using electron microscopy, nuclear chains can be found between myofibrils that contain organized sarcomeres [83,84][26][35]. This can also be seen in single myofibers stained to delineate filamentous (F-) actin [90][31] or desmin, which links Z-lines of adjacent myofibrils together [117][36]. The position of nuclear chains between myofibrils is also indicated in transverse sections stained with hematoxylin and eosin or reagents to detect cytoskeletal proteins (e.g., F-actin or an isoform of myosin) [44,96,98,118][37][38][39][40]. These techniques reveal that central myonuclei are surrounded by an abundance of cytoskeletal proteins. Similar to regenerating myofibers, central myonuclei during embryonic myogenesis are surrounded by cytoskeletal proteins, including developmental isoforms of myosin [44,82,97][5][37][41].
An important aspect of the muscle regeneration paradigm is that central myonuclei move towards the sarcolemma as the radial size of regenerating myofibers increases during the course of regeneration. This tenet likely stems from an apparent temporal relationship between myofibrillogenesis, myofiber hypertrophy, and the repositioning of myonuclei from a central to a peripheral position during embryonic myogenesis [69,77,82,103,104][2][3][5][10][11]. Accordingly, the number/percentage of regenerating myofibers is predicted to decrease during the course of muscle regeneration.
A well-recognized phenomenon after chemical and physical trauma to rodents is that regenerating myofibers remain in skeletal muscle for an extended period of time (e.g., 1–6 months post-injury) [14,36,86,87,89,96,119,120,121,122][27][28][30][38][42][43][44][45][46][47]. Importantly, the extent to which the number/percentage of regenerating myofibers diminishes during the course of trauma-induced muscle regeneration is unclear. For example, the percentage of regenerating myofibers (75–90%) in transverse sections and in isolated myofibers has been reported to remain relatively constant for at least one month of recovery from chemical (barium chloride or notexin) trauma [86,119,122][27][44][47]. Wada et al. [89][30] reported that the percentage of isolated myofibers that contained nuclear chains (i.e., regenerating myofibers) remained high (~80%) and constant for at least 6 months after physical trauma (400 needle punctures) to skeletal muscle. In contrast, others have reported that the percentage of regenerating myofibers in transverse sections decreases from 3–4 (74–90%) to 60 (14–60%) days of recovery from chemical (bupivacaine or notexin) trauma [14,121][42][46]. Conflicting viewpoints and evidence on how long regenerating myofibers remain in skeletal muscle after trauma has created much confusion about myonuclear repositioning during muscle regeneration and its use as an indicator of regenerating myofiber maturation.
In an effort to gain insight into myonuclear positioning and repositioning during muscle regeneration, researchers analyzed single myofibers before and after chemical (barium chloride) trauma to gastrocnemius muscles of mice [90][31]. researchers found that the percentage of myonuclei in nuclear chains was high at 7 days post-injury (77%, range = 54–94%) and progressively decreased at 14 (70%, range = 60–84%) and 28 (60%, range = 35–80%) days post-injury [90][31]. The percentage of peripheral myonuclei was low at 7 days post-injury (16%, range = 3–43%) and progressively increased at 14 (24%, range = 12–37%) and 28 (29%, range = 13–55%) days post-injury [90][31].
Why regenerating myofibers remain in skeletal muscle for an extended period of time after chemical or physical trauma is unknown. Their sustained presence is either dismissed or interpreted to indicate that muscle regeneration has not yet completed. This interpretation is hard to reconcile with evidence of myofiber size (discussed in “Section 6”), as well as metabolic enzyme activity, mitochondrial respiration, and muscle function returning to control levels within four weeks of chemical trauma [14,15,36,86,96,122,123,124,125][27][38][42][43][47][48][49][50][51]. Maximum specific force of single myofibers (force normalized to myofiber cross-sectional area) after chemical (bupivacaine) trauma has also been reported to be similar to that of control myofibers [126][52]. Importantly, researchers are not aware of any study that has demonstrated that the central position of myonuclei or the presence of nuclear chains after muscle trauma impairs specific force or other contractile properties of single myofibers. The extent to which the metabolic profile of regenerating myofibers differs from that of normal myofibers also remains to be determined, as prior research targeting metabolism analyzed regenerating muscles [123[49][50][51],124,125], which normally contains both regenerating and non-regenerating/normal myofibers.
Myonuclear positioning during muscle regeneration is remarkably different from myonuclear positioning during embryonic and postnatal myogenesis. The alignment and organization of central myonuclei, as well as their sustained presence, are unique features of trauma-induced muscle regeneration. Why myonuclei are uniquely positioned during muscle regeneration is unknown. Is the central positioning of myonuclei during muscle regeneration a consequence of a high rate of myofibrillogenesis and a rapidly expanding cytoplasm? In other words, are myonuclei stuck in a central position because their path towards the periphery is blocked by a large number of myofibrils? Or is the central positioning of myonuclei optimal for orchestrating molecular processes (e.g., transcription) [90,91,92][31][32][34] that facilitate regenerating myofiber maturation? If true, why do central myonuclei persist after regenerating myofibers have reached an adult radial size? Do myonuclei remain in a central position because other aspects of regenerating myofiber maturation have yet to be completed? If yes, what aspect of their maturation lags behind? Clearly, additional research is needed to comprehend why the majority of myonuclei are centrally aligned in regenerating myofibers during trauma-induced muscle regeneration.

4. Myonuclear Accretion during Embryonic and Postnatal Myogenesis

Embryonic myogenesis is associated with a high rate of muscle progenitor cell proliferation and myonuclear accretion [61,62,63,70,127,128][1][9][53][54][55][56]. However, the kinetics of myonuclear accretion in developing myotubes/myofibers during embryonic myogenesis has yet to be quantitatively detailed.
Many studies have evaluated myonuclear accretion during postnatal myogenesis in rodents using myofibers isolated from several different skeletal muscles [106,107,108,110,111,112,113,114,129][13][14][15][17][18][19][20][21][57]. The kinetics of myonuclear accretion was first detailed in extensor digitorum longus (EDL) muscles of mice by White et al. [112][19]. Strikingly, they reported that the number of myonuclei/myofiber increased by ~3 fold from P3 to P7 [112][19] (P = postnatal age in days). Myonuclear accretion continued through P21, albeit at a slower rate [112][19]. Myonuclear accretion observed during the first three weeks of postnatal myogenesis was accompanied by a 3–4 fold increase in both myofiber length and cross-sectional area [112][19].
Others have analyzed myonuclear number in single myofibers isolated from mouse EDL muscles after three weeks of postnatal development [106,108,110,111,114][13][15][17][18][21]. These studies demonstrated that myonuclear number, expressed per myofiber or per mm of myofiber length, increased modestly (~10–35%) during the pre-puberty stage of development (P21 to P42) [106,110,111][13][17][18]. Increases in myonuclear number during this period were accompanied by an increase in myofiber length and notable increases in myofiber cross-sectional area [106,108,110,111,112][13][15][17][18][19]. Myonuclear accretion appears to continue to some degree after puberty in mice (P42/6 weeks of age) until 8–12 weeks of age [110,130][17][58]. During this period, myofiber length reaches adult levels [112,131][19][59] and myofiber cross-sectional area continues to increase until adulthood [30,106][13][24].
As detailed in a narrative review by Bachman et al. [132][60], the kinetics of myonuclear accretion during postnatal development in mice parallels the number and proliferative state of satellite cells/myoblasts. As a result, variability exists between muscles in myonuclear number, as well as the number of satellite cells associated with a myofiber during the course of postnatal myogenesis [110,112,130,133][17][19][58][61]. For example, satellite cell and myonuclear number are notably higher in myofibers isolated from soleus compared to EDL muscles throughout postnatal development and in adult mice [93,110,133][17][61][62]. Thus, myonuclear accretion varies between muscles during postnatal development in a manner that appears to reflect the number of myoblasts available for fusion [109][16]. Furthermore, as bone length increases during postnatal development [134,135][63][64] and myonuclear number scales with increases in myofiber length [106,112][13][19], differences in myonuclear number between muscles could also reflect differences in architectural (e.g., length and pennation angle) and/or functional features of myofibers [136][65]. Fundamentally, the genetic and physiological processes that dictate myonuclear number in myofibers during development are not fully understood.
From a physiology perspective, it is conceivable that increases in bone length drive the longitudinal growth of myofibers during postnatal development, which in turn creates a need for additional myonuclei to sustain both longitudinal and radial growth of myofibers. Upon myofibers reaching an adult length, the need for myonuclear accretion to sustain radial growth of myofibers could stem from increases in body mass and muscle use. Deciphering the cellular and molecular interplay between developing bone, satellite cells, and the needs of myofibers during postnatal myogenesis will undoubtedly be challenging. Equally challenging will be to determine why myoblasts during postnatal myogenesis fuse with existing myofibers and not with each other to form myotubes/myofibers. This is a fundamentally important question as myofiber number in a given muscle is widely believed to be fixed around the time of birth [28,29,30,31][22][23][24][25].

5. Myonuclear Accretion during Trauma-Induced Muscle Regeneration

The paradigm for muscle regeneration proposes that myonuclear accretion in myotubes facilitates their longitudinal and radial growth, which in turn gives rise to small regenerating myofibers. Although the distinction between a mature myotube and a small regenerating myofiber is ambiguous (discussed above in “Section 3.3”), myonuclear accretion is thought to continue until regenerating myofibers obtain the normal complement of myonuclei. The attainment of a normal number of myonuclei is believed to be necessary for regenerating myofibers to achieve a radial size that is comparable to a normal myofiber. Surprisingly, little research has directed towards detailing myonuclear accretion during regeneration [86,90,92][27][31][34].
researchers have evaluated myonuclear accretion during muscle regeneration by analyzing single myofibers before and after chemical (barium chloride) trauma to tibialis anterior and gastrocnemius muscles of mice [86,90,92][27][31][34]. The magnitude and rate of myonuclear accretion observed during the course of muscle regeneration was surprising. Specifically, the number of myonuclei in regenerating myofibers, expressed per unit myofiber length (myonuclei/100 μm), was ~2 fold higher at 7, 14, and 28 days post-injury compared to non-regenerating myofibers in control muscles (0 days post-injury) [86,90][27][31]. Furthermore, myonuclear number (myonuclei/100 μm) in regenerating myofibers was similar at 7, 14, and 28 days post-injury [86,90][27][31]. The magnitude and rate of myonuclear accretion that researchers observed in regenerating myofibers paralleled the robust satellite cell/myoblast proliferation that occurs within 7 days of chemical trauma in mice [36,37,38][43][66][67]. The profile of myonuclear accretion, however, was not temporally related to regenerating myofiber hypertrophy, as cross-sectional area of regenerating myofibers increased by 2.2 fold from 7 to 14 days post-injury and by 17% from 14 to 28 days post-injury [90][31] (discussed below in “Section 6”). Simply put, myonuclear accretion preceded the robust hypertrophy of regenerating myofibers.
Although reporting the myonuclear number relative to myofiber length (e.g., myonuclei/100 μm) is informative and widely done, it is not a measure of the absolute number of myonuclei in a myofiber. This is an important issue to consider when myofiber length is changing, such as during postnatal myogenesis [112,131][19][59]. Unfortunately, changes in the length of myotubes or regenerating myofibers during regeneration have not been described in the literature. The measurement of myotube/regenerating myofiber length requires that they remain intact after being enzymatically and physically liberated from a whole muscle, procedures that can fracture small, fragile myofibers. Nevertheless, further work is needed to detail changes in regenerating myofiber length and the absolute number of myonuclei in regenerating myofibers to fully appreciate the kinetics of myonuclear accretion during muscle regeneration.
Why regenerating myofibers rapidly add more than the normal complement of myonuclei is unknown. Intuitively, the number of myonuclei in a regenerating myofiber reflects both extrinsic and intrinsic regulation of myonuclear accretion. Extrinsic factors, such as molecules (e.g., cytokines) and cell types (e.g., macrophages) within injured muscle, increase the number of myoblasts available for fusion by augmenting satellite cell/myoblast proliferation [23,26,32,33][68][69][70][71]. The local environment of injured muscle could also enhance the ability of myoblast to fuse to myotubes/regenerating myofibers by augmenting the expression and/or function of molecules that facilitate cell-to-cell adhesion [25,51,52][72][73][74] and/or membrane dynamics during fusion [24,53,54][75][76][77]. Independent of the extrinsic regulation of myonuclear accretion, myotubes/regenerating myofibers likely have an intrinsic need to add myonuclei to support their sustained maturation. However, such a need could only be satisfied if satellite cells proliferate, and myoblasts are available for fusion.
To reconcile the above logic, researchers propose that myotubes/regenerating myofibers obtain a sufficient number of myonuclei to sustain their maturation through an exquisite interplay between their needs and the local environment of injured muscle. The details of such an interplay are difficult to fathom as they likely involve paracrine signaling in myotubes/regenerating myofibers, enhanced function of molecules that mediate cell-to-cell adhesion and fusion, and synergistic interactions with multiples cytokines and cell types. Alternatively, the myonuclear number in regenerating myofibers could be solely dependent on the extrinsic regulation of satellite cell/myoblast proliferation and fusion. Such a scenario is not favored because it implies that developing myotubes/regenerating myofibers after muscle trauma have no role in determining how many myonuclei they house. In other words, it does not seem prudent to ignore the intrinsic needs of myotubes/regenerating myofibers, particularly when their longitudinal and radial growth demands an increase in transcription.
Given the proposed regulation of myonuclear number in regenerating myofibers, researchers speculate that the higher-than-normal number of myonuclei in regenerating myofibers, which was obtained within 7 days of muscle trauma [90[31][34],92], is a consequence of extrinsic regulation of myogenic cell proliferation and fusion. In other words, researchers think that the pro-proliferative and pro-fusion environment of severely injured muscle caused the myonuclear number to exceed levels that are required for regenerating myofiber maturation.
Superficially, myonuclear accretion during muscle regeneration is different from that occurring during postnatal myogenesis. Regenerating myofibers obtain a higher-than-normal number of myonuclei within 7 days of muscle trauma [90,92][31][34]; whereas myofibers developing after birth take at least a couple of months to obtain adult levels of myonuclei [106,110,130][13][17][58]. As cells of the myogenic lineage (e.g., myoblasts) are the principal source of myonuclei for all myofibers [37[37][66][78][79],43,44,45], it is likely that differences in the magnitude and kinetics of myonuclear accretion between muscle regeneration and postnatal myogenesis parallel differences in satellite cell/myoblast proliferation. Specifically, researchers suspect that satellite cell/myoblast proliferation after chemical trauma is notably higher for a shorter period of time compared to postnatal myogenesis. If true, this could reflect a differential profile of cell types (e.g., macrophages) and cytokines during muscle regeneration and postnatal myogenesis. This speculation is reasonable as a large number of macrophages accumulate in injured muscle and their secretome augments myogenic cell proliferation after chemical trauma [26,33,137][69][71][80]. However, the macrophage accumulation profile during postnatal myogenesis, as well as the extent to which their secretome contributes to myogenic cell proliferation and/or fusion during postnatal myogenesis, remains to be determined. Independent of the extrinsic regulation of myonuclear accretion, it is also possible that myotubes/regenerating myofibers have an intrinsic need to quickly obtain a large number of myonuclei, a need that may not be germane to postnatal myogenesis.

6. Myofiber Hypertrophy during Postnatal Myogenesis

Myofibers at birth undergo sustained hypertrophy until adulthood. This hypertrophy has been attributed to a progressive increase in the number and size of myofibrils, which normally occupy nearly all of the cytoplasm of myofibers [103,104][10][11]. Because sustained hypertrophy during postnatal myogenesis demands a net accumulation of myofibrillar proteins [138[81][82],139], there has been a long standing interest in deciphering the contribution of myonuclear accretion and downstream processes (e.g., transcription and translation) to myofiber hypertrophy during postnatal development.
Prior research has reported statistically significant positive correlations between myonuclear number and myofiber cross-sectional area or volume at selected time points of postnatal development [108,110,112,129][15][17][19][57]. These findings indicated a role of myonuclear accretion in facilitating myofiber hypertrophy during postnatal myogenesis.
Two studies have directly tested the relationship between myonuclear number and the radial size of myofibers during postnatal myogenesis in mice [106,110][13][17]. Cramer et al. [106][13] inhibited myoblast fusion by conditionally deleting myomaker during the first, second, or third week of postnatal development in mice [106][13], which corresponded to a timeframe when myofibers are rapidly adding myonuclei [112][19]. Deleting myomaker during the first of week of postnatal development substantially reduced myonuclear number and myofiber volume in myofibers isolated from EDL muscles at 4 weeks of postnatal development [106][13]. Notable reductions in myofiber cross-sectional area were also observed in other hindlimb muscles at 4 weeks of postnatal development [106][13]. Interestingly, deleting myomaker during the first of week of postnatal development resulted in premature death, which was interpreted to indicate a finite ability of myonuclei to compensate for severe impairments in myonuclear accretion [106][13]. Deleting myomaker during the second week of postnatal development reduced myonuclear number in EDL myofibers by ~35–55% throughout 5 months of postnatal development [106][13]. Importantly, sustained reductions in myonuclei number did not impair the ability of myofibers to undergo hypertrophy. Rather, sustained reductions in myonuclei number prevented myofibers from reaching a radial size that is comparable to controls [106][13]. Bachman et al. [110][17] also reported that reductions in myonuclear number prevented myofibers from reaching a normal size during postnatal myogenesis. This was demonstrated by depleting satellite cells (Pax7+ cells) after 4 weeks of postnatal development in mice. The near absence of satellite cells in EDL and soleus muscles reduced myonuclear number by 10–15% and myofiber cross-sectional area by 15–19% at 8 weeks of postnatal development [110][17]. The findings of Cramer et al. [106][13] and Bachman et al. [110][17] demonstrate that myonuclear number is a critical determinant of the radial size of myofibers during postnatal myogenesis. In other words, if the intrinsic need for additional myonuclei during postnatal myogenesis is not met via myogenic cell proliferation and subsequent fusion, myofibers fail to reach their normal radial size [106,110][13][17].
Additional research is needed to understand how the need for myofibrillar protein synthesis to sustain myofiber hypertrophy during postnatal myogenesis is coordinated amongst the myonuclei, and how such a need triggers myonuclear accretion. This will be a challenging endeavor as the transcriptome of individual myonuclei during postnatal myogenesis is heterogeneous [140][83] and the translational regulation of protein synthesis is complex [141,142][84][85]. Interestingly, Cramer et al. [106][13] proposed that individual myonuclei have a finite transcriptional capacity during postnatal myogenesis. If true, myonuclear accretion during postnatal myogenesis could be initiated when individual myonuclei are approaching their maximal transcriptional capacity.

7. Regenerating Myofiber Hypertrophy after Trauma-Induced Muscle Regeneration

According to the muscle regeneration paradigm, regenerating myofibers undergo hypertrophy until they reach a radial size that is comparable to normal myofibers. Furthermore, the radial size of a regenerating myofiber is thought to be closely coupled to myonuclear number. The logic behind the coupling is that myonuclear accretion increases the capacity of regenerating myofibers to transcribe genes, which, in theory, optimizes their hypertrophy. The transcriptional activity in regenerating myofibers and the division of transcriptional labor amongst myonuclei within them is poorly understood [90,91,92,143][31][32][34][86].
A large number of studies have quantified the radial size of myofibers before and after muscle trauma in rodents. It is difficult to construct a composite profile of regenerating myofiber hypertrophy after trauma for several reasons. One, temporal variability in indices of muscle regeneration exists between experimental models of inducing muscle trauma [36,144][43][87]. Two, the radial size of myofibers after trauma is routinely measured at one or two recovery time points. Lastly, in some cases, it is unclear if the reported data after trauma is representative of all myofibers (non-regenerating and regenerating) or just regenerating myofibers. The reason for raising this issue is not to question the appropriateness of data reporting in prior studies, as there are valid reasons for reporting the radial size of all myofibers after trauma. Rather, the intent is to highlight belief that regenerating myofiber hypertrophy cannot be accurately described from changes in the radial size of all myofibers. This belief is rooted in the fact that the number/percentage of regenerating myofibers in transverse sections after chemical or physical trauma can vary between muscles at any recovery time point (e.g., 46–93%) [86][27]. This variability means that the number/percentage of non-regenerating myofibers in transverse sections after muscle trauma also varies at each recovery time point (e.g., 7–54%) [86][27]. A secondary analysis of published data [86][27] revealed that the cross-sectional area of non-regenerating myofibers after chemical (barium chloride) trauma is 30–55% lower compared to non-regenerating myofibers in control muscle.
Although regenerating myofibers are discernible 4 days after chemical trauma in mice, it is difficult to accurately measure their radial size in transverse muscle sections and to isolate them from whole muscles. On the other hand, regenerating myofibers at 7 days post-injury and beyond can be measured in transverse sections, as well as isolated from whole muscles of mice. researchers found that regenerating myofiber cross-sectional area in transverse sections [86][27] and in isolation [90][31] increased by 1.7–2.6 fold between 7 and 14 days after chemical (barium chloride) trauma in mice. Others have also reported a robust hypertrophy of regenerating myofibers over a similar time frame of recovery from chemical (barium chloride or notexin) trauma in mice [96,99,145,146][38][88][89][90]. Rapid hypertrophy during early recovery from chemical trauma is preceded by myonuclear accretion [90][31] and is temporally associated with innervation of regenerating myofibers [96,120][38][45] and a high rate of muscle protein synthesis [86][27]. Regenerating myofiber cross-sectional area continued to increase between 14 and 21 days post-injury, albeit at a slower rate [145][89]. This slower rate of hypertrophy was temporally associated with a gradual return of protein synthesis rates to levels that were observed in control muscles [86][27]. No further increase in regenerating myofiber cross-sectional area was observed between 21 and 28 days post-injury. Interestingly, cross-sectional area of regenerating myofibers at 14 days post-injury was comparable to the size of normal (non-regenerating) myofibers in control muscles. Despite reaching control levels, regenerating myofiber cross-sectional area continued to increase to levels that were 25% and 18% higher than control myofibers at 21 and 28 days post-injury, respectively. Others have reported that the radial size of myofibers exceeded control levels within 4 weeks of chemical (notexin) trauma [122][47] and remained elevated for at least 6 months after chemical (cardiotoxin, barium chloride, and notexin) or physical (cold exposure) trauma to skeletal muscle [36][43]. The hypertrophy of regenerating myofibers beyond control levels is not consistent with a central tenet of the muscle regeneration paradigm. That is, hypertrophy of regenerating myofibers is thought to cease when they achieve a size that is comparable to normal myofibers.
researchers have examined the statistical relationship between myonuclear number and the radial size of regenerating myofibers after chemical (barium chloride) trauma in mice [86,90,92][27][31][34]. Significant correlations were observed between myonuclear number and regenerating myofiber area (r = 0.67) [86][27] and volume (r = 0.77) [90][31]. Interestingly, regenerating myofiber volume was closely coupled to myonuclei number at 7 days post-injury (r = 0.95) and became more responsive to myonuclear number at 14 and 28 days post-injury, as indicated by a progressive increase in the slope of regression lines during the course of regeneration [90][31]. The increased responsiveness of myofiber volume to myonuclear number was temporally associated with an elevated level of transcription within regenerating myofibers [90][31].
No studies have directly tested the relationship between myonuclear number and the radial size of regenerating myofibers during regeneration. In contrast, prior studies have tested the contribution of myonuclear accretion to myofiber hypertrophy during postnatal myogenesis [106,110][13][17] and after resistance exercise [147,148][91][92] and muscle overload (synergist ablation) [47,149,150][93][94][95]. To be clear, investigators have demonstrated that genetically depleting satellite cells [38[37][67][93][96][97],44,46,47,48], myomaker [60][98], or myomixer [56][99] prior to chemical trauma in mice prevents regenerating myofiber formation during recovery. Because few, if any, regenerating myofibers formed in the absence of satellite cells, myomaker, or myomixer, these studies [38,44,46,47,48,56,60][37][67][93][96][97][98][99] do not provide insight into the relationship between myonuclear number and the radial size of regenerating myofiber during regeneration. Unfortunately, current genetic models are not well suited to test such a relationship as they require several days of Cre-mediated recombination to produce the desired outcome (e.g., depletion of satellite cells or absence of myomaker). This is problematic because myonuclear number in regenerating myofibers reaches a plateau within 7 days of muscle trauma [86,90,92][27][31][34]. In other words, initiating Cre-mediated recombination just prior to or immediately after muscle trauma may not reduce the number of satellite cells or myomaker expressing myoblasts to a level that compromises myonuclear accretion during recovery. Thus, testing the extent to which myonuclear number dictates the radial size of regenerating myofibers may require the development of new approaches that manipulate myonuclear number or myonuclear function in regenerating myofibers during the course of regeneration.
Regenerating myofibers during muscle regeneration and myofibers during postnatal myogenesis in mice are similar in that they both undergo robust hypertrophy. However, regenerating and developing myofibers differ in the time needed to reach an adult size, as well as the temporal relationship between myonuclear accretion and myofiber hypertrophy. Regenerating myofibers achieve an adult radial size much sooner (3–4 weeks) [36,86,90,122][27][31][43][47] than developing myofibers (8–12 weeks) [30,106][13][24]. During muscle regeneration, myonuclear accretion reaches a plateau within 7 days of chemical trauma and precedes notable increases in the radial size of regenerating myofibers [86,90][27][31]. In contrast, myonuclear accretion and myofiber hypertrophy during postnatal myogenesis are closely coupled, particularly when myofibers are rapidly undergoing hypertrophy [110,112][17][19]. Reconciling such disparities awaits a better understanding of the extrinsic and intrinsic regulators of myonuclear accretion and myofiber hypertrophy during muscle regeneration, as well as during postnatal myogenesis.

8. Myofiber Formation and Morphology during Embryonic Myogenesis

It is widely accepted that myofiber formation during embryonic myogenesis in mammals occurs in two successive stages [69,70,77,78][2][3][4][9]. Each stage has been attributed to the proliferation and fusion of distinct populations of progenitor cells, namely embryonic and fetal myoblasts [151,152,153][100][101][102]. Morphological, functional, and molecular differences between embryonic myoblasts, fetal myoblasts, and satellite cells have been extensively studied [154,155,156,157][103][104][105][106], as well as summarized in narrative reviews [151,152,153][100][101][102].
The first stage of embryonic myogenesis in mice (E10.5–12.5, E = embryo age in days) produces ‘primary’ myotubes from the fusion of embryonic myoblasts [69,70,77][2][3][9]. Primary myotubes form independent of innervation, accumulate myonuclei, span tendon-to-tendon, and serve as the structural framework for the second stage of embryonic myogenesis [61,62,70,77,101,158,159,160][1][3][7][9][53][107][108][109]. The second stage in mice (E14.5–17.5) gives rise to ‘secondary’ myotubes as a result of fusion of fetal myoblasts [70,158][9][107]. Several secondary myotubes form within the basal lamina of primary myotubes and near their mid-belly [77,78][3][4]. Secondary myotube formation is dependent on innervation of primary myotubes, as well as muscle contractions [101,158,161][7][107][110].
The maturation of secondary myotubes is a critical step in embryonic myogenesis as they give rise to ~80% of the myofibers in adult muscle [101,160][7][109]. The maturation of secondary myotubes is characterized by innervation, myonuclear accretion, myofibrillogenesis, longitudinal and radial growth, and the assembly of a basal lamina [61,62,63,69,77,78,162][1][2][3][4][53][54][111]. Observations made via electron microscopy during the course of embryonic myogenesis indicate that myotubes either fuse with each other [69,70][2][9] or separate [77,78,162][3][4][111]. Prior investigators have emphasized and schematically depicted that primary and secondary myotubes can appear in a transverse section as a single cell, a cell undergoing longitudinal splitting, or two distinct cells during the course of embryonic myogenesis [69,70,160,162,163][2][9][109][111][112]. In the latter case, secondary myotubes were distinguished from primary myotubes/myofibers based on their smaller radial size [70,77,78,101,158,162][3][4][7][9][107][111]. Because secondary myotubes appear to separate from primary myotubes/myofibers throughout the course of embryonic myogenesis, investigators have noted the difficulty in quantifying myofiber number in transverse sections around the time of birth [30,78,162,163][4][24][111][112].

9. Myofiber Formation and Morphology during Trauma-Induced Muscle Regeneration

De novo myofiber formation after chemical trauma in mice is dependent on satellite cells (Pax7+ cells) [38,44,46,47,48][37][67][93][96][97]. Although several types of progenitor cells accumulate in skeletal muscle after trauma (e.g., Pax3+, fibro-adipogenic progenitors, Twist2+, mesoangioblasts, and PW1+ cells) [23[68][113][114],164,165], they are unable to fully compensate for the near absence of Pax7+ satellite cells [38,44,46,47,48][37][67][93][96][97]. This indicates that Pax7+ satellite cells are the principal, if not the sole precursor cells that gives rise to regenerating myofibers in adult mice after trauma. In contrast, de novo myofiber formation during embryonic myogenesis has been attributed to different progenitor cells, namely embryonic and fetal myoblasts [152,157][101][106].
A well-recognized phenomenon during muscle regeneration is that some myofibers appear to be either undergoing longitudinal splitting or fusion. In transverse sections, a ‘split’ or fissure within some myofibers can be observed in histologically stained sections (e.g., hematoxylin and eosin) [40[89][115][116],145,166], as well as sections treated to delineate the sarcolemma or basal lamina [167][117]. Griffin et al. [145][89] analyzed serial transverse sections and reported that 20–40% of the myofibers exhibited a ‘split’ morphology 5 days after chemical (barium chloride) trauma in mice. The incidence of ‘split’ myofibers during prolonged recovery from chemical trauma remains to be determined.
Myofibers isolated after trauma also exhibit an unusual morphology called branching [40,86,119,145,168][27][44][89][115][118]. A branched myofiber is defined as an isolated myofiber that has a membrane that is contiguous with one or more myofibers. Characteristics of a branched myofiber include a ‘split’/fissure typically near the middle of a myofiber or a bifurcation/process on their side or at their end [40,86,119,145,168][27][44][89][115][118]. Given the characteristics of myofiber branching [40[27][44][89][115][118],86,119,145,168], it is understandable how an individual branched myofiber after trauma could appear in transverse planes as a single cell, a cell undergoing longitudinal splitting or fusion, or two cells.
Myofiber branching is restricted to regenerating myofibers, as very few non-regenerating myofibers after chemical (barium chloride and cardiotoxin) trauma in mice showed signs of branching [119][44]. A moderate to high percentage of regenerating myofibers (25–60%) showed signs of branching 10–21 days after chemical trauma in mice [86,119,145,146][27][44][89][90]. The percentage of branched regenerating myofibers remains elevated (28–60%) during prolonged recovery (1 to 6 months) from chemical trauma in mice [86,119][27][44].
Myofiber ‘splitting’ and/or branching also occurs in hypertrophying [169,170][119][120] and diseased muscle (e.g., in mdx mice) [95[6][44][121][122],119,171,172], as well as a result of aging [119][44]. Although each condition in which ‘split’ and/or branched myofibers are observed is associated with muscle regeneration to some degree, the environment and the phenotype of regenerating, hypertrophying, and diseased muscle is distinctly different. This has resulted in wide speculation on the cause and physiological significance of ‘split’ and/or branched myofibers. A comprehensive overview of the abnormal morphology of some myofibers in the context of muscle hypertrophy [169,170][119][120] and disease [172][122] can be found elsewhere.
In trauma models of muscle regeneration, ‘split’/branched myofibers are thought to represent active/incomplete fusion of myotubes/myofibers to each other [86,119,168,173][27][44][118][123]. This premise is supported by qualitative observations of myotube/myofiber fusion after muscle trauma [40,83,84,85][26][35][115][124]. Furthermore, molecules expressed by myogenic cells that facilitate or augment their fusion in vitro have also been reported to influence myofiber branching after chemical (barium chloride) trauma in mice [86,145,146][27][89][90]. The sustained presence of branched myofibers after trauma is believed to reflect a protracted or stalled fusion, as well as to confound interpretations of an increased number (hyperplasia) of myofibers after chemical trauma [40,122,166][47][115][116]. Specifically, some believe that myofiber branching produces a pseudo hyperplasia when myofiber number is quantified in transverse sections.
When considering the morphology of myofibers during embryonic myogenesis, another possibility exists for the formation of ‘split’/branched myofibers during muscle regeneration. That is, the unusual myofiber morphology could reflect myotubes/regenerating myofibers that are separating from each other during the course of regeneration. To be clear, this scenario is distinctly different from the premise that extreme myofiber hypertrophy causes longitudinal splitting of the affected myofiber [169,170][119][120]. Prior investigators have reported that myotubes form within and outside of the basal lamina of necrotic myofibers and that they assemble their own basal lamina after chemical (barium chloride) and physical (e.g., cold exposure and crush) trauma [37,39,40,41,167][66][115][117][125][126]. Schmalbruch et al. [40][115] noted that “usually, one basal lamina tube contained several myotubes” after physical trauma. The assembly of a basal lamina by myotubes/regenerating myofibers after trauma is thought to ultimately replace the basal lamina of necrotic myofibers [37,39,40,41,167][66][115][117][125][126]. The extent to which myotubes/regenerating myofibers assemble their own basal lamina could influence the degree to which they fuse or separate from each other, as the basal lamina would likely serve as a physical barrier for their fusion. Nevertheless, closely aligned myotubes/regenerating myofibers could appear as ‘split’/branched myofibers if they are separating from each other during the course of regeneration. In this scenario, the sustained presence of ‘split’/branched myofibers during prolonged recovery could reflect a protracted or stalled assembly of a basal lamina for each myofiber.
The possibility that myofiber ‘splitting’ and branching during regeneration reflects the separation of myotubes/regenerating myofibers is intriguing for two reasons. One, the morphology of ‘split’/branched myofibers during regeneration resembles the morphology of primary and secondary myotubes/myofibers separating from each other during the course of embryonic myogenesis. However, only the ‘split’ morphology has been noted during muscle development [69,70,101,162[2][7][9][111][112],163], as no study has examined isolated myotubes/myofibers for signs of branching during embryonic myogenesis. Thus, the extent to which myotubes/myofibers that are separating from each other would appear as a branched myofiber after being liberated from embryonic muscle is unknown. Furthermore, myotubes/myofibers formed during embryonic myogenesis assemble their own basal lamina [77[3][127][128],174,175], whereas their formation during trauma-induced muscle regeneration primarily occurs within existing basal lamina (i.e., the basal lamina of necrotic myofibers) [37,39,40,41,167][66][115][117][125][126]. This difference further complicates the extrapolation for findings from embryonic myogenesis to trauma-induced muscle regeneration. The premise that myotube/regenerating myofiber separation explains myofiber ‘splitting’/branching during regeneration is also intriguing because researchers have noticed that the total number of regenerating myofibers in transverse sections increases markedly from 7 to 28 days after chemical (barium chloride) trauma in mice. This increase could be attributable to a pseudo hyperplasia resulting from incomplete myotube/myofiber fusion, myofibers separating from each other during the course of regeneration, or sustained de novo myotube/regenerating myofiber formation. The latter possibility seems unlikely as satellite cell/myoblast proliferation and markers of de novo myofiber formation (e.g., embryonic myosin heavy chain expression) are substantially reduced or non-existent after 7 days of recovery from chemical trauma in mice [36,37,38,96,97,98][38][39][41][43][66][67]. Nevertheless, the underlying processes that cause myofiber ‘splitting’ or branching after trauma-induced muscle regeneration remain to be determined.

References

  1. Harris, A.J.; Duxson, M.J.; Fitzsimons, R.B.; Rieger, F. Myonuclear birthdates distinguish the origins of primary and secondary myotubes in embryonic mammalian skeletal muscles. Development 1989, 107, 771–784.
  2. Fidzianska, A. Human ontogenesis. I. Ultrastructural characteristics of developing human muscle. J. Neuropathol. Exp. Neurol. 1980, 39, 476–486.
  3. Ontell, M.; Kozeka, K. The organogenesis of murine striated muscle: A cytoarchitectural study. Am. J. Anat. 1984, 171, 133–148.
  4. Ontell, M.; Kozeka, K. Organogenesis of the mouse extensor digitorum logus muscle: A quantitative study. Am. J. Anat. 1984, 171, 149–161.
  5. Romero, N.B.; Mezmezian, M.; Fidzianska, A. Main steps of skeletal muscle development in the human: Morphological analysis and ultrastructural characteristics of developing human muscle. Handb. Clin. Neurol. 2013, 113, 1299–1310.
  6. Merrick, D.; Stadler, L.K.; Larner, D.; Smith, J. Muscular dystrophy begins early in embryonic development deriving from stem cell loss and disrupted skeletal muscle formation. Dis. Model. Mech. 2009, 2, 374–388.
  7. Harris, A.J. Embryonic growth and innervation of rat skeletal muscles. I. Neural regulation of muscle fibre numbers. Philos. Trans. R. Soc. Lond. B Biol. Sci. 1981, 293, 257–277.
  8. Harris, A.J.; Fitzsimons, R.B.; McEwan, J.C. Neural control of the sequence of expression of myosin heavy chain isoforms in foetal mammalian muscles. Development 1989, 107, 751–769.
  9. Kelly, A.M.; Zacks, S.I. The histogenesis of rat intercostal muscle. J. Cell Biol. 1969, 42, 135–153.
  10. Goldspink, G. Changes in striated muscle fibres during contraction and growth with particular reference to myofibril splitting. J. Cell Sci. 1971, 9, 123–137.
  11. Goldspink, G. The proliferation of myofibrils during muscle fibre growth. J. Cell Sci. 1970, 6, 593–603.
  12. Roman, W.; Martins, J.P.; Carvalho, F.A.; Voituriez, R.; Abella, J.V.G.; Santos, N.C.; Cadot, B.; Way, M.; Gomes, E.R. Myofibril contraction and crosslinking drive nuclear movement to the periphery of skeletal muscle. Nat. Cell Biol. 2017, 19, 1189–1201.
  13. Cramer, A.A.W.; Prasad, V.; Eftestol, E.; Song, T.; Hansson, K.A.; Dugdale, H.F.; Sadayappan, S.; Ochala, J.; Gundersen, K.; Millay, D.P. Nuclear numbers in syncytial muscle fibers promote size but limit the development of larger myonuclear domains. Nat. Commun. 2020, 11, 6287.
  14. Williams, P.E.; Goldspink, G. Longitudinal growth of striated muscle fibres. J. Cell Sci. 1971, 9, 751–767.
  15. Wada, K.I.; Katsuta, S.; Soya, H. Natural occurrence of myofiber cytoplasmic enlargement accompanied by decrease in myonuclear number. Jpn. J. Physiol. 2003, 53, 145–150.
  16. Gattazzo, F.; Laurent, B.; Relaix, F.; Rouard, H.; Didier, N. Distinct Phases of Postnatal Skeletal Muscle Growth Govern the Progressive Establishment of Muscle Stem Cell Quiescence. Stem Cell Rep. 2020, 15, 597–611.
  17. Bachman, J.F.; Klose, A.; Liu, W.; Paris, N.D.; Blanc, R.S.; Schmalz, M.; Knapp, E.; Chakkalakal, J.V. Prepubertal skeletal muscle growth requires Pax7-expressing satellite cell-derived myonuclear contribution. Development 2018, 145, dev167197.
  18. Huh, M.S.; Young, K.G.; Yan, K.; Price-O’Dea, T.; Picketts, D.J. Recovery from impaired muscle growth arises from prolonged postnatal accretion of myonuclei in Atrx mutant mice. PLoS ONE 2017, 12, e0186989.
  19. White, R.B.; Bierinx, A.S.; Gnocchi, V.F.; Zammit, P.S. Dynamics of muscle fibre growth during postnatal mouse development. BMC Dev. Biol. 2010, 10, 21.
  20. Cardasis, C.A.; Cooper, G.W. An analysis of nuclear numbers in individual muscle fibers during differentiation and growth: A satellite cell-muscle fiber growth unit. J. Exp. Zool. 1975, 191, 347–358.
  21. Schultz, E. Satellite cell proliferative compartments in growing skeletal muscles. Dev. Biol. 1996, 175, 84–94.
  22. Enesco, M.; Puddy, D. Increase in the Number of Nuclei and Weight in Skeletal Muscle of Rats of Various Ages. Am. J. Anat. 1964, 114, 235–244.
  23. Montgomery, R.D. Growth of human striated muscle. Nature 1962, 195, 194–195.
  24. Rowe, R.W.; Goldspink, G. Muscle fibre growth in five different muscles in both sexes of mice. J. Anat. 1969, 104, 519–530.
  25. Li, M.; Zhou, X.; Chen, Y.; Nie, Y.; Huang, H.; Chen, H.; Mo, D. Not all the number of skeletal muscle fibers is determined prenatally. BMC Dev. Biol. 2015, 15, 42.
  26. Allbrook, D. An electron microscopic study of regenerating skeletal muscle. J. Anat. 1962, 96, 137–152.
  27. Martin, R.A.; Buckley, K.H.; Mankowski, D.C.; Riley, B.M.; Sidwell, A.N.; Douglas, S.L.; Worth, R.G.; Pizza, F.X. Myogenic Cell Expression of Intercellular Adhesion Molecule-1 Contributes to Muscle Regeneration after Injury. Am. J. Pathol. 2020, 190, 2039–2055.
  28. Clark, W.E. An experimental study of the regeneration of mammalian striped muscle. J. Anat. 1946, 80, 24–36.
  29. Liu, J.; Huang, Z.-P.; Nie, M.; Wang, G.; Silva, W.J.; Yang, Q.; Freire, P.P.; Hu, X.; Chen, H.; Deng, Z. Regulation of myonuclear positioning and muscle function by the skeletal muscle-specific CIP protein. Proc. Natl. Acad. Sci. USA 2020, 117, 19254–19265.
  30. Wada, K.; Katsuta, S.; Soya, H. Formation process and fate of the nuclear chain after injury in regenerated myofiber. Anat. Rec. 2008, 291, 122–128.
  31. Buckley, K.H.; Nestor-Kalinoski, A.L.; Pizza, F.X. Positional Context of Myonuclear Transcription During Injury-Induced Muscle Regeneration. Front. Physiol. 2022, 13, 845504.
  32. Newlands, S.; Levitt, L.K.; Robinson, C.S.; Karpf, A.B.; Hodgson, V.R.; Wade, R.P.; Hardeman, E.C. Transcription occurs in pulses in muscle fibers. Genes. Dev. 1998, 12, 2748–2758.
  33. Lash, J.W.; Holtzer, H.; Swift, H. Regeneration of mature skeletal muscle. Anat. Rec. 1957, 128, 679–697.
  34. Buckley, K.H.; Nestor-Kalinoski, A.L.; Pizza, F.X. Intercellular Adhesion Molecule-1 Enhances Myonuclear Transcription during Injury-Induced Muscle Regeneration. Int. J. Mol. Sci. 2022, 23, 7028.
  35. Jirmanova, I.; Thesleff, S. Ultrastructural study of experimental muscle degeneration and regeneration in the adult rat. Z. Zellforsch. Mikrosk. Anat. 1972, 131, 77–97.
  36. Capetanaki, Y.; Bloch, R.J.; Kouloumenta, A.; Mavroidis, M.; Psarras, S. Muscle intermediate filaments and their links to membranes and membranous organelles. Exp. Cell Res. 2007, 313, 2063–2076.
  37. Lepper, C.; Partridge, T.A.; Fan, C.M. An absolute requirement for Pax7-positive satellite cells in acute injury-induced skeletal muscle regeneration. Development 2011, 138, 3639–3646.
  38. Whalen, R.G.; Harris, J.B.; Butler-Browne, G.S.; Sesodia, S. Expression of myosin isoforms during notexin-induced regeneration of rat soleus muscles. Dev. Biol. 1990, 141, 24–40.
  39. d’Albis, A.; Couteaux, R.; Janmot, C.; Roulet, A.; Mira, J.C. Regeneration after cardiotoxin injury of innervated and denervated slow and fast muscles of mammals. Myosin isoform analysis. Eur. J. Biochem. 1988, 174, 103–110.
  40. Waisman, A.; Norris, A.M.; Costa, M.E.; Kopinke, D. Automatic and unbiased segmentation and quantification of myofibers in skeletal muscle. Sci. Rep. 2021, 11, 11793.
  41. Sartore, S.; Gorza, L.; Schiaffino, S. Fetal myosin heavy chains in regenerating muscle. Nature 1982, 298, 294–296.
  42. Rosenblatt, J.D.; Woods, R.I. Hypertrophy of rat extensor digitorum longus muscle injected with bupivacaine. A sequential histochemical, immunohistochemical, histological and morphometric study. J. Anat. 1992, 181, 11–27.
  43. Hardy, D.; Besnard, A.; Latil, M.; Jouvion, G.; Briand, D.; Thepenier, C.; Pascal, Q.; Guguin, A.; Gayraud-Morel, B.; Cavaillon, J.M.; et al. Comparative Study of Injury Models for Studying Muscle Regeneration in Mice. PLoS ONE 2016, 11, e0147198.
  44. Pichavant, C.; Pavlath, G.K. Incidence and severity of myofiber branching with regeneration and aging. Skelet. Muscle 2014, 4, 9.
  45. Grubb, B.D.; Harris, J.B.; Schofield, I.S. Neuromuscular transmission at newly formed neuromuscular junctions in the regenerating soleus muscle of the rat. J. Physiol. 1991, 441, 405–421.
  46. Louboutin, J.P.; Fichter-Gagnepain, V.; Noireaud, J. Comparison of contractile properties between developing and regenerating soleus muscle: Influence of external calcium concentration upon the contractility. Muscle Nerve 1995, 18, 1292–1299.
  47. Lee, A.S.; Anderson, J.E.; Joya, J.E.; Head, S.I.; Pather, N.; Kee, A.J.; Gunning, P.W.; Hardeman, E.C. Aged skeletal muscle retains the ability to fully regenerate functional architecture. Bioarchitecture 2013, 3, 25–37.
  48. Rosenblatt, J.D. A time course study of the isometric contractile properties of rat extensor digitorum longus muscle injected with bupivacaine. Comp. Biochem. Physiol. Comp. Physiol. 1992, 101, 361–367.
  49. Duguez, S.; Feasson, L.; Denis, C.; Freyssenet, D. Mitochondrial biogenesis during skeletal muscle regeneration. Am. J. Physiol. Endocrinol. Metab. 2002, 282, E802–E809.
  50. Wagner, K.R.; Max, S.R.; Grollman, E.M.; Koski, C.L. Glycolysis in skeletal muscle regeneration. Exp. Neurol. 1976, 52, 40–48.
  51. McArdle, A.; Edwards, R.H.; Jackson, M.J. Release of creatine kinase and prostaglandin E2 from regenerating skeletal muscle fibers. J. Appl. Physiol. 1994, 76, 1274–1278.
  52. Gregorevic, P.; Plant, D.R.; Stupka, N.; Lynch, G.S. Changes in contractile activation characteristics of rat fast and slow skeletal muscle fibres during regeneration. J. Physiol. 2004, 558, 549–560.
  53. Zhang, M.; McLennan, I.S. During secondary myotube formation, primary myotubes preferentially absorb new nuclei at their ends. Dev. Dyn. 1995, 204, 168–177.
  54. Evans, D.; Baillie, H.; Caswell, A.; Wigmore, P. During fetal muscle development, clones of cells contribute to both primary and secondary fibers. Dev. Biol. 1994, 162, 348–353.
  55. de Lima, J.E.; Bonnin, M.A.; Bourgeois, A.; Parisi, A.; Le Grand, F.; Duprez, D. Specific pattern of cell cycle during limb fetal myogenesis. Dev. Biol. 2014, 392, 308–323.
  56. Stickland, N.C. Muscle development in the human fetus as exemplified by m. sartorius: A quantitative study. J. Anat. 1981, 132, 557–579.
  57. Mantilla, C.B.; Sill, R.V.; Aravamudan, B.; Zhan, W.Z.; Sieck, G.C. Developmental effects on myonuclear domain size of rat diaphragm fibers. J. Appl. Physiol. 2008, 104, 787–794.
  58. Pawlikowski, B.; Pulliam, C.; Betta, N.D.; Kardon, G.; Olwin, B.B. Pervasive satellite cell contribution to uninjured adult muscle fibers. Skelet. Muscle 2015, 5, 42.
  59. Gokhin, D.S.; Ward, S.R.; Bremner, S.N.; Lieber, R.L. Quantitative analysis of neonatal skeletal muscle functional improvement in the mouse. J. Exp. Biol. 2008, 211, 837–843.
  60. Bachman, J.F.; Chakkalakal, J.V. Insights into muscle stem cell dynamics during postnatal development. FEBS J. 2022, 289, 2710–2722.
  61. Keefe, A.C.; Lawson, J.A.; Flygare, S.D.; Fox, Z.D.; Colasanto, M.P.; Mathew, S.J.; Yandell, M.; Kardon, G. Muscle stem cells contribute to myofibres in sedentary adult mice. Nat. Commun. 2015, 6, 7087.
  62. Bruusgaard, J.C.; Liestol, K.; Gundersen, K. Distribution of myonuclei and microtubules in live muscle fibers of young, middle-aged, and old mice. J. Appl. Physiol. 2006, 100, 2024–2030.
  63. Beaucage, K.L.; Pollmann, S.I.; Sims, S.M.; Dixon, S.J.; Holdsworth, D.W. Quantitative in vivo micro-computed tomography for assessment of age-dependent changes in murine whole-body composition. Bone Rep. 2016, 5, 70–80.
  64. Somerville, J.M.; Aspden, R.M.; Armour, K.E.; Armour, K.J.; Reid, D.M. Growth of C57BL/6 mice and the material and mechanical properties of cortical bone from the tibia. Calcif. Tissue Int. 2004, 74, 469–475.
  65. Burkholder, T.J.; Fingado, B.; Baron, S.; Lieber, R.L. Relationship between muscle fiber types and sizes and muscle architectural properties in the mouse hindlimb. J. Morphol. 1994, 221, 177–190.
  66. Webster, M.T.; Manor, U.; Lippincott-Schwartz, J.; Fan, C.M. Intravital Imaging Reveals Ghost Fibers as Architectural Units Guiding Myogenic Progenitors during Regeneration. Cell Stem Cell 2016, 18, 243–252.
  67. Murphy, M.M.; Lawson, J.A.; Mathew, S.J.; Hutcheson, D.A.; Kardon, G. Satellite cells, connective tissue fibroblasts and their interactions are crucial for muscle regeneration. Development 2011, 138, 3625–3637.
  68. Wosczyna, M.N.; Rando, T.A. A Muscle Stem Cell Support Group: Coordinated Cellular Responses in Muscle Regeneration. Dev. Cell 2018, 46, 135–143.
  69. Panci, G.; Chazaud, B. Inflammation during post-injury skeletal muscle regeneration. Semin. Cell Dev. Biol. 2021, 119, 32–38.
  70. Dumont, N.A.; Bentzinger, C.F.; Sincennes, M.C.; Rudnicki, M.A. Satellite Cells and Skeletal Muscle Regeneration. Compr. Physiol. 2015, 5, 1027–1059.
  71. Chazaud, B. Inflammation and Skeletal Muscle Regeneration: Leave It to the Macrophages! Trends Immunol. 2020, 41, 481–492.
  72. Taylor, L.; Wankell, M.; Saxena, P.; McFarlane, C.; Hebbard, L. Cell adhesion an important determinant of myogenesis and satellite cell activity. Biochim. Biophys. Acta Mol. Cell Res. 2022, 1869, 119170.
  73. Krauss, R.S. Regulation of promyogenic signal transduction by cell-cell contact and adhesion. Exp. Cell Res. 2010, 316, 3042–3049.
  74. Krauss, R.S.; Joseph, G.A.; Goel, A.J. Keep Your Friends Close: Cell-Cell Contact and Skeletal Myogenesis. Cold Spring Harb. Perspect. Biol. 2017, 9, a029298.
  75. Millay, D.P. Regulation of the myoblast fusion reaction for muscle development, regeneration, and adaptations. Exp. Cell Res. 2022, 415, 113134.
  76. Petrany, M.J.; Millay, D.P. Cell Fusion: Merging Membranes and Making Muscle. Trends Cell Biol. 2019, 29, 964–973.
  77. Lehka, L.; Redowicz, M.J. Mechanisms regulating myoblast fusion: A multilevel interplay. Semin. Cell Dev. Biol. 2020, 104, 81–92.
  78. Moss, F.P.; Leblond, C.P. Satellite cells as the source of nuclei in muscles of growing rats. Anat. Rec. 1971, 170, 421–435.
  79. Lepper, C.; Conway, S.J.; Fan, C.M. Adult satellite cells and embryonic muscle progenitors have distinct genetic requirements. Nature 2009, 460, 627–631.
  80. Arnold, L.; Henry, A.; Poron, F.; Baba-Amer, Y.; van Rooijen, N.; Plonquet, A.; Gherardi, R.K.; Chazaud, B. Inflammatory monocytes recruited after skeletal muscle injury switch into antiinflammatory macrophages to support myogenesis. J. Exp. Med. 2007, 204, 1057–1069.
  81. Bates, P.C.; Millward, D.J. Characteristics of skeletal muscle growth and protein turnover in a fast-growing rat strain. Br. J. Nutr. 1981, 46, 7–13.
  82. Millward, D.J.; Garlick, P.J.; Stewart, R.J.; Nnanyelugo, D.O.; Waterlow, J.C. Skeletal-muscle growth and protein turnover. Biochem. J. 1975, 150, 235–243.
  83. Petrany, M.J.; Swoboda, C.O.; Sun, C.; Chetal, K.; Chen, X.; Weirauch, M.T.; Salomonis, N.; Millay, D.P. Single-nucleus RNA-seq identifies transcriptional heterogeneity in multinucleated skeletal myofibers. Nat. Commun. 2020, 11, 6374.
  84. Proud, C.G. Regulation of mammalian translation factors by nutrients. Eur. J. Biochem. 2002, 269, 5338–5349.
  85. von Walden, F. Ribosome biogenesis in skeletal muscle: Coordination of transcription and translation. J. Appl. Physiol. 2019, 127, 591–598.
  86. Kim, M.; Franke, V.; Brandt, B.; Lowenstein, E.D.; Schowel, V.; Spuler, S.; Akalin, A.; Birchmeier, C. Single-nucleus transcriptomics reveals functional compartmentalization in syncytial skeletal muscle cells. Nat. Commun. 2020, 11, 6375.
  87. Plant, D.R.; Colarossi, F.E.; Lynch, G.S. Notexin causes greater myotoxic damage and slower functional repair in mouse skeletal muscles than bupivacaine. Muscle Nerve 2006, 34, 577–585.
  88. Wang, X.; Jia, Y.; Zhao, J.; Lesner, N.P.; Menezes, C.J.; Shelton, S.D.; Venigalla, S.S.K.; Xu, J.; Cai, C.; Mishra, P. A mitofusin 2/HIF1alpha axis sets a maturation checkpoint in regenerating skeletal muscle. J. Clin. Investig. 2022, 132, e161638.
  89. Griffin, C.A.; Kafadar, K.A.; Pavlath, G.K. MOR23 promotes muscle regeneration and regulates cell adhesion and migration. Dev. Cell 2009, 17, 649–661.
  90. Simionescu-Bankston, A.; Leoni, G.; Wang, Y.; Pham, P.P.; Ramalingam, A.; DuHadaway, J.B.; Faundez, V.; Nusrat, A.; Prendergast, G.C.; Pavlath, G.K. The N-BAR domain protein, Bin3, regulates Rac1- and Cdc42-dependent processes in myogenesis. Dev. Biol. 2013, 382, 160–171.
  91. Goh, Q.; Song, T.; Petrany, M.J.; Cramer, A.A.; Sun, C.; Sadayappan, S.; Lee, S.J.; Millay, D.P. Myonuclear accretion is a determinant of exercise-induced remodeling in skeletal muscle. eLife 2019, 8, e44876.
  92. Englund, D.A.; Figueiredo, V.C.; Dungan, C.M.; Murach, K.A.; Peck, B.D.; Petrosino, J.M.; Brightwell, C.R.; Dupont, A.M.; Neal, A.C.; Fry, C.S.; et al. Satellite Cell Depletion Disrupts Transcriptional Coordination and Muscle Adaptation to Exercise. Function 2021, 2, zqaa033.
  93. McCarthy, J.J.; Mula, J.; Miyazaki, M.; Erfani, R.; Garrison, K.; Farooqui, A.B.; Srikuea, R.; Lawson, B.A.; Grimes, B.; Keller, C.; et al. Effective fiber hypertrophy in satellite cell-depleted skeletal muscle. Development 2011, 138, 3657–3666.
  94. Egner, I.M.; Bruusgaard, J.C.; Gundersen, K. Satellite cell depletion prevents fiber hypertrophy in skeletal muscle. Development 2016, 143, 2898–2906.
  95. Goh, Q.; Millay, D.P. Requirement of myomaker-mediated stem cell fusion for skeletal muscle hypertrophy. eLife 2017, 6, e20007.
  96. Fry, C.S.; Lee, J.D.; Mula, J.; Kirby, T.J.; Jackson, J.R.; Liu, F.; Yang, L.; Mendias, C.L.; Dupont-Versteegden, E.E.; McCarthy, J.J.; et al. Inducible depletion of satellite cells in adult, sedentary mice impairs muscle regenerative capacity without affecting sarcopenia. Nat. Med. 2015, 21, 76–80.
  97. Sambasivan, R.; Yao, R.; Kissenpfennig, A.; Van Wittenberghe, L.; Paldi, A.; Gayraud-Morel, B.; Guenou, H.; Malissen, B.; Tajbakhsh, S.; Galy, A. Pax7-expressing satellite cells are indispensable for adult skeletal muscle regeneration. Development 2011, 138, 3647–3656.
  98. Millay, D.P.; Sutherland, L.B.; Bassel-Duby, R.; Olson, E.N. Myomaker is essential for muscle regeneration. Genes Dev. 2014, 28, 1641–1646.
  99. Bi, P.; McAnally, J.R.; Shelton, J.M.; Sanchez-Ortiz, E.; Bassel-Duby, R.; Olson, E.N. Fusogenic micropeptide Myomixer is essential for satellite cell fusion and muscle regeneration. Proc. Natl. Acad. Sci. USA 2018, 115, 3864–3869.
  100. Biressi, S.; Molinaro, M.; Cossu, G. Cellular heterogeneity during vertebrate skeletal muscle development. Dev. Biol. 2007, 308, 281–293.
  101. Stockdale, F.E. Myogenic cell lineages. Dev. Biol. 1992, 154, 284–298.
  102. Chal, J.; Pourquie, O. Making muscle: Skeletal myogenesis in vivo and in vitro. Development 2017, 144, 2104–2122.
  103. Biressi, S.; Tagliafico, E.; Lamorte, G.; Monteverde, S.; Tenedini, E.; Roncaglia, E.; Ferrari, S.; Ferrari, S.; Cusella-De Angelis, M.G.; Tajbakhsh, S.; et al. Intrinsic phenotypic diversity of embryonic and fetal myoblasts is revealed by genome-wide gene expression analysis on purified cells. Dev. Biol. 2007, 304, 633–651.
  104. Hutcheson, D.A.; Zhao, J.; Merrell, A.; Haldar, M.; Kardon, G. Embryonic and fetal limb myogenic cells are derived from developmentally distinct progenitors and have different requirements for beta-catenin. Genes. Dev. 2009, 23, 997–1013.
  105. Lepper, C.; Fan, C.M. Inducible lineage tracing of Pax7-descendant cells reveals embryonic origin of adult satellite cells. Genesis 2010, 48, 424–436.
  106. Relaix, F.; Rocancourt, D.; Mansouri, A.; Buckingham, M. A Pax3/Pax7-dependent population of skeletal muscle progenitor cells. Nature 2005, 435, 948–953.
  107. Duxson, M.J.; Usson, Y.; Harris, A.J. The origin of secondary myotubes in mammalian skeletal muscles: Ultrastructural studies. Development 1989, 107, 743–750.
  108. Dennis, M.J.; Ziskind-Conhaim, L.; Harris, A.J. Development of neuromuscular junctions in rat embryos. Dev. Biol. 1981, 81, 266–279.
  109. Harris, A.J. Embryonic growth and innervation of rat skeletal muscles. III. Neural regulation of junctional and extra-junctional acetylcholine receptor clusters. Philos. Trans. R. Soc. Lond. B Biol. Sci. 1981, 293, 287–314.
  110. Ross, J.J.; Duxson, M.J.; Harris, A.J. Neural determination of muscle fibre numbers in embryonic rat lumbrical muscles. Development 1987, 100, 395–409.
  111. Ontell, M.; Dunn, R.F. Neonatal muscle growth: A quantitative study. Am. J. Anat. 1978, 152, 539–555.
  112. Ontell, M. Neonatal muscle: An electron microscopic study. Anat. Rec. 1977, 189, 669–690.
  113. Liu, N.; Garry, G.A.; Li, S.; Bezprozvannaya, S.; Sanchez-Ortiz, E.; Chen, B.; Shelton, J.M.; Jaichander, P.; Bassel-Duby, R.; Olson, E.N. A Twist2-dependent progenitor cell contributes to adult skeletal muscle. Nat. Cell Biol. 2017, 19, 202–213.
  114. Kuang, S.; Charge, S.B.; Seale, P.; Huh, M.; Rudnicki, M.A. Distinct roles for Pax7 and Pax3 in adult regenerative myogenesis. J. Cell Biol. 2006, 172, 103–113.
  115. Schmalbruch, H. The morphology of regeneration of skeletal muscles in the rat. Tissue Cell 1976, 8, 673–692.
  116. Davis, C.E.; Harris, J.B.; Nicholson, L.V. Myosin isoform transitions and physiological properties of regenerated and re-innervated soleus muscles of the rat. Neuromuscul. Disord. 1991, 1, 411–421.
  117. Bassaglia, Y.; Gautron, J. Fast and slow rat muscles degenerate and regenerate differently after whole crush injury. J. Muscle Res. Cell Motil. 1995, 16, 420–429.
  118. Tamaki, T.; Akatsuka, A. Appearance of complex branched fibers following repetitive muscle trauma in normal rat skeletal muscle. Anat. Rec. 1994, 240, 217–224.
  119. Murach, K.A.; Dungan, C.M.; Peterson, C.A.; McCarthy, J.J. Muscle Fiber Splitting Is a Physiological Response to Extreme Loading in Animals. Exerc. Sport. Sci. Rev. 2019, 47, 108–115.
  120. Jorgenson, K.W.; Phillips, S.M.; Hornberger, T.A. Identifying the Structural Adaptations that Drive the Mechanical Load-Induced Growth of Skeletal Muscle: A Scoping Review. Cells 2020, 9, 1658.
  121. Lovering, R.M.; Michaelson, L.; Ward, C.W. Malformed mdx myofibers have normal cytoskeletal architecture yet altered EC coupling and stress-induced Ca2+ signaling. Am. J. Physiol. Cell Physiol. 2009, 297, C571–C580.
  122. Chan, S.; Head, S.I. The role of branched fibres in the pathogenesis of Duchenne muscular dystrophy. Exp. Physiol. 2011, 96, 564–571.
  123. Hurme, T.; Kalimo, H.; Lehto, M.; Jarvinen, M. Healing of skeletal muscle injury: An ultrastructural and immunohistochemical study. Med. Sci. Sports Exerc. 1991, 23, 801–810.
  124. Robertson, T.A.; Papadimitriou, J.M.; Grounds, M.D. Fusion of myogenic cells to the newly sealed region of damaged myofibres in skeletal muscle regeneration. Neuropathol. Appl. Neurobiol. 1993, 19, 350–358.
  125. Vracko, R.; Benditt, E.P. Basal lamina: The scaffold for orderly cell replacement. Observations on regeneration of injured skeletal muscle fibers and capillaries. J. Cell Biol. 1972, 55, 406–419.
  126. Caldwell, C.J.; Mattey, D.L.; Weller, R.O. Role of the basement membrane in the regeneration of skeletal muscle. Neuropathol. Appl. Neurobiol. 1990, 16, 225–238.
  127. Jenniskens, G.J.; Hafmans, T.; Veerkamp, J.H.; van Kuppevelt, T.H. Spatiotemporal distribution of heparan sulfate epitopes during myogenesis and synaptogenesis: A study in developing mouse intercostal muscle. Dev. Dyn. 2002, 225, 70–79.
  128. Patton, B.L.; Miner, J.H.; Chiu, A.Y.; Sanes, J.R. Distribution and function of laminins in the neuromuscular system of developing, adult, and mutant mice. J. Cell Biol. 1997, 139, 1507–1521.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/)
ScholarVision Creations
Feedback