It is easily observable under the electron microscope that myofibers filled with myofibrils are aligned along the longitudinal axis of the cell body. The myofibril has a diameter of approximately 1 μm and presents a periodic striped structure alternating between light and dark, in which, each I-band (light area) is divided into two parts by a Z-line and each A-band (dark area) is divided into two portions by an M-line within the H zone. The segment of myofibrils between two adjacent Z lines is commonly described as a sarcomere, which is the smallest contractile unit of the striated muscle (Figure 1). On the meta-microscale, the sarcomere primarily consists of thin and thick myofilaments that are perpendicular to the Z-line: the actin, tropomyosin, and troponin complexes (troponin T, I, and C, respectively) are organized into thin myofilaments that are anchored in Z-lines, while the myosins are assembled into thick myofilaments in the A-band that overlap with thin myofilaments (Ertbjerg and Puolanne, 2017). These structures form the basis for the “sliding filament theory” of skeletal muscle contraction: muscle contraction entails the thin myofilaments sliding over the thick myofilaments into the A-band, causing the shortening of the I-band and H zone, while the widths of the Z-line, A-band, and M-line remain unchanged (Huxley and Niedergerke, 1954; Huxley and Hanson, 1954). Specifically, myosin heads bind actin at a ∼45° angle, forming the acto-myosin crossbridges, which pull actin filaments toward the M-line through ATP hydrolysis-driven conformational changes, resulting in sarcomere shortening. Additionally, approximately 30% of crossbridges maintain passive muscle tension even during relaxation, mediated by the spring-like domains of structural titin in coordination with residual crossbridge interactions to preserve myofiber elasticity (Al-Khayat, 2013). In addition, other myofilament-associated proteins such as myosin binding protein-C (MyBP-C), nebulin, and obscurin have also been proposed to play important structural and regulatory functions during the assembly of sarcomeres.

For instance, MyBP-C is a myofilament protein binding to thin and thick myofilaments maintaining the ordered arrangement of the sarcomere and cross-bridge cycling (van Dijk et al., 2014). The interaction of MyBP-C with both myofilaments is restricted to the C zone, where actin and myosin overlap in the A-band, and MyBP-C is arranged in 7–9 transverse stripes at 43-nm intervals (Luther and Craig, 2011). MyBP-C consists of a series of immunoglobin (Ig) and fibronectin III (Fn)-like domains, and the MyBP-C N-terminal binds to actin and myosin head, while the MyBP-C C-terminal binds with meromyosin and titin (Squire et al., 2003). In addition, accumulating evidence has implicated MyBP-C in the regulation of myofilament Ca²⁺ sensitivity and enzymatic activity of myosin (van Dijk et al., 2014).

As the largest known polypeptide, titin spans the length of half of the sarcomere, anchoring its N-terminus and C-terminal at the Z-line and M-line, respectively. The C-terminals of two titin molecules overlap on both sides of the M-line, forming a continuous titin filament harboring tight binding sites for a variety of sarcomeric proteins, which functions as a scaffold, mechanical sensor, and signaling mediator. For example, within the Z-line, titin N-terminus interacts with proteins such as α-actinin and telethonin to maintain the structural integrity and resist high mechanical stress (Young et al., 1998; Mues et al., 1998). Similarly, titin C-terminal binds with several proteins at the M-line, including Muscle RING finger-1 (MURF-1), myomesin, and M-protein, functioning as scaffolding structures for A-band assembly (Witt et al., 2005). Titin is normally regarded as a “molecular spring” that generates passive tension, and the elements responsible for this elasticity primarily reside in the I-band region of titin, including tandemly arranged Ig-domains, PEVK motifs, and N2A element (Linke et al., 1998; Linke et al., 2002; Granzier and Labeit, 2006; Granzier and Labeit, 2005). Unlike the elastic I band region of titin, the A‐band region of titin is inextensible and provides a scaffold for myosin and MyBP-C, therefore determining the length and structure of thick myofilaments (Whiting et al., 1989). Moreover, as its I-band and M-line region contain Ser/Thr kinase domains, titin may be involved in signaling transduction from myofibrils to other compartments of myofiber, contributing to muscle adaptation in conditions of mechanical stress changes (Tskhovrebova and Trinick, 2008; Puchner et al., 2008).

Ultrastructural results with monoclonal antibodies have established that nebulin is oriented with its N-terminus at the tips of thin myofilaments in the I-band and its C-terminus at the Z-line, whose configuration both stabilizes the thin myofilaments and regulates the Z-line structure (Wright et al., 1993; Wang and Wright, 1988; Witt et al., 2006). Actually, nebulin is non-extensible because nebulin epitopes maintain a fixed distance from the Z-line, regardless of sarcomere stretching (Kruger et al., 1991). Numerous studies indicate that nebulin can bind to various proteins, including actin and myosin components, as well as proteins associated with its unique N- and C-terminal regions, especially for the central super repeats of nebulin, through which nebulin laterally interacts with the thin myofilament proteins actin, tropomyosin, and troponin complexes, functioning as the scaffold to control thin myofilament length (Kruger et al., 1991; Labeit et al., 1991). In addition, nebulin’s super-repeat region also binds to myosin, which may inhibit actomyosin ATPase activity and the sliding velocity of actin on myosin in a calcium-/calmodulin-dependent manner (Root and Wang, 1994). Differing in sequence from the central super repeats, the N-terminal sequence carries glutamate residues and can interact with tropomodulin to specify thin myofilament length, whereas the C-terminus is rich in serine residues, and the SH3 domain can directly bind to Z-line protein myopalladin and CapZ and titin to mediate the myofibrillar assembly and mechanochemical signaling (Mcelhinny et al., 2001; Wear et al., 2003; Bang et al., 2001; Kontrogianni-Konstantopoulos et al., 2009).

In contrast to MyBP-C, titin, and nebulin presenting within sarcomeres, obscurin wraps around myofilaments over the M-line and Z-line, contributing to their assembly and integration with other sarcoplasmic elements such as the SR (Kontrogianni-Konstantopoulos et al., 2003). Given its structure composing of tandem adhesion modules and signaling domains, obscurin can associate tightly and periodically with the myofilaments and the periphery of the myofibril. Specifically, the N-terminal Ig domain of obscurin binds directly to myofibrillar components such as titin, myomesin, and MyBP-C, while obscurin tightly connects with the SR component small ankyrin-1 (sAnk1) through the C-terminus to its last Ig-like domain (Kontrogianni-Konstantopoulos and Bloch, 2005; Porter et al., 2005). Other kinetic studies showed that in addition to structural support function, obscurin has a possible involvement in RhoA- and Ca²⁺-mediated signaling pathways via its Rho-GEF/PH and IQ motif, and the presence of obscurin at the NMJ is consistent with this, suggesting its direct or indirect involvement in signal transduction at the synaptic terminal (Fukuzawa et al., 2005; Young et al., 2001; Carlsson et al., 2008). Initial subcellular distribution of obscurin studies using antibodies indicated that obscurin briefly accumulates at the Z-line and primarily concentrates at the M-line during embryogenesis; however, it eventually loses Z-line distribution in later developmental stages (Young et al., 2001). Actually, multiple alternatively spliced forms of obscurin exist in distinct sarcomeric locations, such as obscurin A, obscurin B, and several additional variants that differ in the combination of C-terminal structural domain elements and localize to the M-line, A/I junction, and Z-line at the muscle resting state, respectively (Bowman et al., 2007). Notably though, when muscles are stretched, obscurin can redistribute to different locations along the sarcomere length (Bowman et al., 2007). Considering the preferential integration of these obscurin isoforms with certain regions of the sarcomere, it further implies specialized functions of these isoforms during myofibrillogenesis and at maturity. Collectively, myofibrillogenesis depends on the coordinated assembly and integration of a large number of myofilament-associated proteins into the sarcomeres. These molecules may interact with several protein ligands to function as scaffolds, regulate signaling cascades, and localize them to particular sites within or surrounding sarcomeres to coordinate sarcomeric arrangement.

The skeletal muscle contains a specialized endoplasmic reticulum network called SR, which is responsible for protein and calcium homeostasis regulation. Highly precise views under electron microscopes (EMs) showed that the SR consists of tubules and cisternae around each myofibril, in which the thin tubules are named longitudinal SR (l-SR) and are positioned around the A- and I-band, and its enriched Ca²⁺ ATPase (SERCA) is responsible for removing Ca²⁺ from the cytoplasm to the lumen; the tubule edges merge into the terminal cisternae at the boundaries of the A- and I-band and collectively form the triad structure, with the transverse tubules (TT) originating from the sarcolemma, maintaining excitation–contraction coupling (ECC) in the skeletal muscle (Rossi et al., 2022a) (Figure 2; Table 1). Within the muscle field, ECC is defined as the process linking the sarcolemma depolarization induced by motor–neuron stimulation and Ca²⁺ release from the SR. In particular, there are two Ca²⁺ channel proteins dihydropyridine receptor (DHPR) and ryanodine receptor 1 (RyR1) located on the TT and terminal cisternae region facing the TT terminal junctional SR (j-SR), respectively. Following sarcolemma depolarization, the DHPR undergoes conformational changes and directly transmits signals to open the RyR1 to trigger Ca²⁺ release into the myoplasm, thereby initiating muscle contraction (Barone et al., 2015). In fact, DHPR and RyR1 tightly wound together to form the core of the macromolecular complex known as the Ca²⁺ release unit, which includes numerous interacting proteins, and each of them influences the entire ECC process (Dulhunty, 2006). These proteins include TT biogenesis proteins amphiphysin 2/bridging integrator-1 (BIN1), caveolin 3 (CAV3), and dysferlin (DYSF); triad formation proteins mitsugumins (MG), junctophilin (JPH), myotubularin (MTM1), triadin, junctin, and the junctional sarcoplasmic reticulum protein 1 (JP-45); and Ca²⁺-binding proteins calsequestrin (CASQ), sarcalumenin (SAR), and HRC (Figure 2). For instance, BIN1 is highly expressed in the skeletal muscle and capable of driving membrane tubulation during TT maturation, in which BIN1 works separately or in cooperation with a GTPase dynamin 2 (DNM2) (Lee et al., 2002; Cowling et al., 2017). BIN1 knockout mouse myofibers present abnormal TTs and deficient ECC, and DNM2 downregulation improved its motor and histopathological phenotypes, suggesting that the balance between BIN1 and DNM2 is necessary for TT maturation, which is mediated by the interaction of the BIN1 SH3 domain with the proline-rich domain of DNM2 (Silva-Rojas et al., 2022; Fujise et al., 2021; Takei et al., 1999). As a member of the integral membrane protein family, CAV3 is not only implicated in caveolae membrane formation at the sarcolemma but also localizes at the developing TTs, and its deletion leads to TT membrane disorder with lateral loss. However, CAV3 is undetectable in the mature TTs, suggesting that CAV3 may be involved in the early development of the TT system in the skeletal muscle (Tang et al., 1996; Parton et al., 1997; Galbiati et al., 2001). DYSF mainly localizes to the sarcolemma for membrane fusion and repair, and DYSF-deficient muscle shows subsarcolemmal vacuoles contiguous with the TT system (Selcen et al., 2001). Similarly to mice deficient in CAV3, DYSF mouse mutants display dilated and longitudinally oriented TTs, and CAV3 and DYSF show partial colocalization and interaction in developing TT system, suggesting a specific role of DYSF in TT biogenesis (Hernandez-Deviez et al., 2006; Matsuda et al., 2001). Although the precise mechanism is not yet clear, there is a hypothesis that DYSF may facilitate the fusion of vesicles containing CAV3 with TTs (Klinge et al., 2010). These findings suggest that TT biogenesis requires advanced mechanisms of membrane fusion, and additional factors involved in TT formation are needed.

An abundance of membrane proteins are involved in triad formation, including the maturation of SR terminal cisternae and the connection between TT and SR. A notable example is the mitsugumin 29 (MG29), a member of the synaptophysin family that appears preferentially at the SR membrane and is then transferred to the triad proteins during the early stages of skeletal myogenesis (Takeshima et al., 1998; Komazaki et al., 1999). Analysis of MG29-deficient mice exhibited an accumulation of transport vesicles and incomplete formation of triad and SR networks, suggesting that MG29 may facilitate the docking and fusion of transport vesicles, processes that lead to the creation of the SR network and TT to form the triad (Nishi et al., 1999; Komazaki et al., 2001). Moreover, MG29-deficient mice showed impaired store-operated Ca²⁺ entry (SOCE) in the skeletal muscle, in which MG29 directly interacts with canonical-type transient receptor potential cation channel type 3 (TRPC3) on TT membranes to regulate Ca²⁺ transients during skeletal muscle contraction (Woo et al., 2015). Another synaptophysin, MG53 (also termed TRIM72), contributes to intracellular vesicle translocation and accumulation, and it has been recognized as an essential component of the membrane repair machinery in the striated muscle (Cai et al., 2009a; Whitson et al., 2021). It has been suggested that MG53 can interact with CAV3 and DYSF to facilitate membrane repair at injury sites by forming a molecular complex (Cai et al., 2009b).

In the skeletal muscle, JPHs (JPH1 and JPH2) provide a structural basis for physiological coupling between the plasma membrane and SR: they bridge the TT membrane through their cytosolic N-terminal domain with the j-SR via their C-terminal transmembrane domain (Takeshima et al., 2000). Apart from their structural functions, JPHs can also interact with various proteins such as DHPR, RyR, CAV3, triadin, and CASQ, serving as a scaffold for assembling Ca²⁺-release complexes to ensure effective ECC (Perni and Beam, 2022; Golini et al., 2011).

MTM1 is a phosphoinositide phosphatase localized at triads, and in phosphoinositide metabolism, MTM1 can specifically dephosphorylate phosphatidyl-inositol-3-phosphate (PtdIns3P) and phosphatidyl-inositol-3,5-bisphosphate (PtdIns(3,5) P2), which mediate endocytosis and membrane trafficking (Hnia et al., 2012). Moreover, MTM1 knockout in animal models leads to disruption of triad morphology and Ca²⁺ homeostasis, while its overexpression causes membrane stack accumulation in the subsarcolemmal region, indicating the delicate balanced expression of MTM1 in triad formation (Al-Qusairi et al., 2009; Dowling et al., 2009; Buj-Bello et al., 2008).

Triadin is an integral span membrane protein family localized at the triad, and all isoforms share common N-terminal transmembrane domains but have different C-terminal segments (Marty et al., 2009; Oddoux et al., 2009). The classic isoform of approximately 95 kDa (Trisk 95) is located at the terminal cisternae of the skeletal muscle, and its knockout mice model presented abnormal triads and Ca²⁺ homeostasis, but no significant changes were observed in ECC (Shen et al., 2007). It has been demonstrated that Trisk 95 is able to interact with RyR1, JPH1, and Ca²⁺-binding protein CASQ1 to maintain the triad organization and intracellular Ca²⁺ homeostasis regulation (Shen et al., 2007; Rossi et al., 2022b; Lee et al., 2001). Similarly, a membrane-spanning protein structurally homologous to triadin, named junctin, is an alternative splicing product of the aspartate β-hydroxylase gene (Dulhunty et al., 2009). Junctin and Trisk 95 both interact with RyR1 and CASQ1, anchoring CASQ1 near RyR1; however, their binding sites on RyR1 and CASQ1 and the detailed functional effects are extremely different. For example, in contrast to one same binding site (KEKE motifs) on Trisk 95 for RyR1 and CASQ1, there are multiple binding sites on junctin for RyR1 and CASQ1. Due to the presence of multiple RyR1/junctin and junctin/CASQ1 binding sites, compared to Trisk 95, junctin could assure transmittance of signals from CASQ1 to RyR1 (Beard et al., 2008; Dulhunty et al., 2017). Moreover, another integral membrane protein JP-45 is highly enriched in j-SR, which interacts with DHPR via its N-terminal domain and with CASQ1 through its C-terminal segment (Anderson et al., 2003). A substantial body of research indicates that both overexpression and ablation of JP-45 leads to a reduction in voltage-dependent Ca²⁺-release, which may be closely correlated with changes in the collaboration between JP-45 and DHPR/CASQ1 (Anderson et al., 2006; Gouadon et al., 2006; Yasuda et al., 2013).

It is well-known that Ca²⁺ is preserved in the SR and mainly buffered by the low-affinity and high-capacity Ca²⁺ binding protein CASQ to polymerize into elongated linear polymers under physiological conditions (Wang et al., 1998; Perni et al., 2013). Actually, CASQ is localized exclusively in the j-SR by binding to RyR, triadin, and junctin, as well as creating a complex three-dimensional cross-linked network. In mammals, the CASQ1 isoform is exclusively expressed in the fast-twitch skeletal muscle, while the CASQ2 isoform is found in the slow-twitch skeletal muscle and heart; especially, CASQ1 is capable of storing 80% of the Ca²⁺ in the SR (Beard et al., 2004; Murphy et al., 2009). It has become increasingly clearer that CASQ1 interacts with RyR, triadin, and junctin in a Ca²⁺-dependent manner, which is attributed to changes in the Ca²⁺ concentration that induces a conformational change in CASQ1: CASQ1 maximally inhibits the RyR1 when the free Ca²⁺ concentration reaches 1 mM, and the inhibition gradually diminishes as the Ca²⁺ concentration is altered, while CASQ1 dissociates from the RyR1 complex at Ca²⁺≤ 100 μM or≥ 5 mM (Wang and Michalak, 2020; Royer and Rios, 2009). Recently, new functions of CASQ1 have been proposed, including regulation of SOCE by binding to STIM1/OraiI and ER stress responses interacting with IRE1α in the skeletal muscle (Michelucci et al., 2020; Wang et al., 2019). Similar to the j-SR, the l-SR also contains Ca²⁺ binding proteins, and the most plentiful one is SAR. It has been demonstrated that SAR colocalizes and directly interacts with SERCA, enhancing SERCA complex stability to pump Ca²⁺ into the lumen of the SR to initiate muscle relaxation (Yoshida et al., 2005; Conte et al., 2023). Another secondary SR Ca²⁺-binding protein is HRC, which is far less abundant than CASQs but structurally similar to CASQs, functioning as Ca²⁺ buffer. Actually, the local and rapid changes of Ca²⁺ levels within the SR may result in different multimer forms of HRC and affect its interactions with other SR components (Suk et al., 1999). It has been suggested that HRC could interact with either the RyR or SERCA complex to manage Ca²⁺ release and uptake, respectively. As the SR Ca²⁺ increases, the HRC/SERCA interaction gradually diminishes, causing the detachment of SERCA from HRC. The released HRC then regulates Ca²⁺ release through the RyR complex, while locally reduced SR Ca²⁺ concentration facilitates the reconnection of HRC/SERCA, resulting in the reuptake of cytoplasmic Ca²⁺ (Arvanitis et al., 2011). However, current research studies on HRC mainly focus on the cardiac muscle, and the HRC/triadin and HRC/SERCA interactions and their functional implications on Ca²⁺-uptake and release in skeletal muscle remain unclear.

As a highly metabolic tissue, the skeletal muscle relies on mitochondria to convert substrates into ATP via oxidative phosphorylation. This ATP powers critical processes such as myofibrillar contraction (through myosin ATPase activity), ion transport (e.g., Ca²⁺ reuptake by the SR), and cellular homeostasis. Beyond energy production, the mitochondria are central to metabolite biogenesis, cell-cycle regulation, and apoptosis/autophagy signaling (Mcbride et al., 2006; Dominy and Puigserver, 2013; Tilokani et al., 2018; Palmer et al., 2011; Ploumi et al., 2017). Actually, mitochondrial function is dynamically regulated by cycles of fission (division), fusion (merging), and motility—collectively termed mitochondrial dynamics (Ni et al., 2015). These processes depend on coordinated nuclear gene expression, mtDNA transcription, and protein import (Ploumi et al., 2017). Dysregulation of fission/fusion disrupts mitochondrial membrane integrity, exacerbating apoptosis and autophagy through pathways such as Bax/Bcl-2-imbalance and mPTP opening (Jeong and Seol, 2008; Xian and Liou, 2021).

The mitochondria in myofibers exhibit distinct morphological and functional specialization based on their subcellular localization: the sub-sarcolemmal mitochondria (SSM) reside beneath the sarcolemma and are usually spherical in form with few branches, supplying ATP to sarcolemmal pumps (e.g., Na⁺/K⁺-ATPase); the perinuclear mitochondria (PNM) are in proximity to nuclei/capillaries and have structural similarities with SSM, supplying ATP to the nuclear pores, and both SSM and PNM are termed peripheral mitochondria (PM); the intra-myofibrillar mitochondria (IFM) are located between myofibrils and have a close contact with the SR displaying reticular network structure, which specialize in ATP production for contractile machinery (myofibrils) and SR Ca²⁺-ATPase (SERCA) (Vincent et al., 2019; Swalsingh et al., 2022). IFM exhibit higher oxidative enzyme activity, respiratory chain complex density, and nucleotide metabolism proteins compared to SSM, aligning with their role in sustaining contraction (Ferreira et al., 2010).

Emerging studies demonstrate that mitochondrial subpopulations diverge in stress responses and signaling. For instance, some studies demonstrated that IFM possess higher levels of cytochrome c/apoptosis-inducing factor (AIF) release and lower membrane potential, ROS production rate, and Bax-to-Bcl-2 ratio compared with SSM (Adhihetty et al., 2005). Considering its greater oxidative enzyme activities and respiration rates, IFM may represent the larger pro-apoptotic protein pool in the skeletal muscle. A recent spatial distribution study of mitochondrial enzymes showed that complexes responsible for membrane potential (complex IV) and ATP (complex V) production are preferentially located in PM and IFM, respectively, and it has been postulated that membrane potential generated in PM can be directly distributed to IFM through mitochondrial connections, promoting ATP production (Willingham et al., 2021). It has also been reported that IFM is structurally and functionally associated with SR, and their contact sites are referred to mitochondria-associated membranes (MAMs) for exchanging Ca²⁺. In the mitochondria, Ca²⁺ influx increases ATP production by enhancing the activity of mitochondrial oxidases, while the exceeding physiological needs of Ca²⁺ can promote the opening of the mitochondrial permeability transition pore (mPTP), thereby triggering apoptosis or necrotic cell death (Rossi et al., 2019; Matuz-Mares et al., 2022). Therefore, mitochondrial subcellular specialization is tightly linked with skeletal muscle quality and function.

In the skeletal muscle, the cytoskeleton serves as a cellular scaffold determining the morphology of myofibers and transmitting mechanical force between myofibrils and extracellular skeleton. In addition to myofibrils themselves, the cytoskeleton component also includes costameres, IF, and microtubule. Costameres are positioned underneath the sarcolemma and vertical to the longitudinal axis of myofibers, connecting the sarcomere to sarcolemma via the Z-line and M-line. Along with assembling and stabilizing the sarcomere, costameres also regulate the interaction between the cytoskeleton and the extracellular matrix (ECM), all of which are based on its intricate protein composition. Current evidence supports the existence of major costameric proteins including the dystrophin–glycoprotein complex (DGC) and the vinculin–talin–integrin system (Figure 3). The DGC connects the cytoskeletal actin and the laminin via dystrophin and dystrophin-associated glycoprotein (DAG) interactions, respectively, in which the cytoskeletal protein dystrophin binds to cytoplasmic actin at its N-terminus, whereas its C-terminus is associated with DAGs that can be classified as cytoplasmic, supramembrane, and extracellular subcomplex (Blake et al., 2002; Sweeney and Barton, 2000). The cytoplasmic subcomplex comprises the α-dystrobrevin (α-DB) and syntrophin (syn), which are directly linked to dystrophin C-terminus. The supramembrane subcomplex includes glycosylated transmembrane protein sarcoglycan (SG) and sarcospan (SP), which need to be correctly assembled into a complex with dystrophin for maintaining sarcolemma integrity. For the extracellular subcomplex, the primary component α-dystroglycan (α-DG) strengthens the physical linkage between the ECM and the basement membrane components via binding to laminin-2 in the ECM and β-dystroglycan (β-DG) in the sarcolemma. The β-DG can also interact with the dystrophin, thus establishing the connection between the actin-based cytoskeleton and the ECM. Hence, it is clear that dystrophin serves a pivotal role in DGC system, whose absence or dysfunction leads to increased sarcolemma fragility, thereby weakening its functions to protect against mechanical stress (Jaka et al., 2015). Similarly, the vinculin–talin–integrin system is also fundamental for facilitating actin cytoskeleton interactions with the surrounding matrix, especially in skeletal muscle cell adhesion and signaling (Mukund and Subramaniam, 2020). Here, the dimeric protein talin links vinculin with integrin: on the one hand, talin binds to the cytoplasmic tails of integrins to activate ECM adhesion; on the other hand, the interaction between vinculin and talin provides the connection of actin filaments to the sarcolemma, reinforcing the adhesion site. Moreover, the complex can also participate in signal transduction pathways, which influence various cellular responses, such as changes in gene expression and cell behavior.

Another cytoskeletal element, IF, has been demonstrated to tether the contractile myofibrillar apparatus to the sarcolemma and other organelles such as the myonucleus, mitochondria, SR, and lysosome, which forms the transverse fixation system of the muscle together with the costameres (Figure 3). Based on similarities among their sequence and structure, IF proteins are grouped into six types, namely, types I and II (keratin), type III (desmin, vimentin, glial fibrillary acidic protein (GFAP), and peripherin), type IV (neurofilament-like protein), type V (lamin A/C), and type VI (large-molecular weight proteins such as nestin, synemin, paranemin, and syncoilin or a special group for CP49/phakinin and filensin/CP115), and the list is still growing (Hnia et al., 2015; Costa et al., 2004; Eldirany et al., 2021). During myogenesis, myoblasts initially express vimentin and nestin, while desmin is induced early in myogenic differentiation and then integrated into the pre-existing network of vimentin, and the mature skeletal muscle mainly contains desmin and keratin (Schweitzer et al., 2001). These filamentous structures form extended IF networks around the Z-line of the myofibrils, thereby linking adjacent myofibrils together, especially desmin and keratin (K8 and K19), which are linked to sarcolemma through costameric components. In addition, desmin knockout in mice causes disruption in the nuclear position, myofibril alignment, mitochondrial function, and even NMJ and MTJ integrity, indicating a more prominent role of desmin in maintaining cytoskeletal stability (Capetanaki et al., 1997; Castanon et al., 2013). Actually, molecular studies demonstrate that desmin can bind or copolymerize with IF proteins vimentin, nestin, synemin, paranemin, and syncoilin and widely interact with other functional proteins. In early myogenesis, nestin co-polymerizes with vimentin and later on with desmin to form filaments, whereas vimentin completely disappears and nestin is selectively expressed only in the NMJ and MTJ in mature skeletal myofibers (Romero et al., 2013; Hol and Capetanaki, 2017; Cizkova et al., 2009). Synemin is another desmin-associated molecule and is considered to link the heteropolymeric IFs to the sarcolemma by directly interacting with desmin/vimentin, α-actinin, α-dystrobrevin, and vinculin (a microfilament-associated protein) (Sun et al., 2008; Etienne-Manneville, 2018). For an IF component of developing skeletal muscle, paranemin has been shown to be required for desmin filaments organization, but not nestin or synemin (Schweitzer et al., 2001). Unlike the majority of IF proteins that form filaments by homopolymerization or heteropolymerization, syncoilin is a non-filament-forming member that binds directly to desmin and α-dystrobrevin, thus linking desmin to the sarcolemma and contributing to the lateral force transmission in the skeletal muscle. Except for being located near the sarcolemma, syncoilin is enriched at the NMJ and MTJ and around the nuclei in mature skeletal myofibers (Moorwood, 2008; Blake and Martin-Rendon, 2002). In the skeletal muscle, IFs are also located in the nuclei as lamins, mainly represented by lamin A/C, to form a nucleoskeleton beneath the inner nuclear membrane and hold mechanosignals. LINC complex (SUN1/2-nesprin3) on the nuclear membrane binds to lamin A/C at the inner nuclear membrane (through SUN1/2) and to desmin-binding protein plectin 1 at the outer nuclear membrane (through nesprin3), thus allowing the association of the nucleoskeleton with the cytoskeleton (Etienne-Manneville, 2018). Furthermore, via the nuclear pores, cytoplasmic IFs may directly connect lamins, but it remains to be determined in the skeletal muscle (Lockard and Bloom, 1993; Georgatos et al., 1987; Capetanaki et al., 2007). Desmin filaments have also been suggested to connect other cell components such as lysosome (via the myospryn), SR (via the MTM1), microtubule (via the dynein), and M-line (via the MyoM1) (Hnia et al., 2015), (Kielbasa et al., 2011). Especially, the desmin-binding protein plectins are responsible for linking desmin IF networks with nuclei (via the plectin 1), mitochondria (via the plectin 1b), Z-line (via the plectin 1 day), and costamere (via the plectin 1f) (Castanon et al., 2013), (Carlsson et al., 2000; Konieczny et al., 2008). These desmin interactions are likely to coordinate the activity of specific kinases and phosphatases, thus regulating signal transduction.

In addition to the costamere and IF, myofibers also contain microtubules to transmit mechanical force, which form a grid containing longitudinal and transverse components across the fiber. During myogenesis, myoblast fusion, nuclear localization to the cell periphery, and assembly of the sarcomere require microtubules to ensure proper myofiber structure and function (Lucas and Cooper, 2023). It is well known that microtubule assembly nucleates at the microtubule-organizing center (MTOC), and MTOC transferring from the centrosomal to non-centrosomal location is a marker of differentiation (Sanchez and Feldman, 2017). During skeletal muscle differentiation, mononuclear myoblasts cease proliferation and elongation to form myoblasts with fusogenic capability, attributed to the MTOC switching from centrosomes to nuclear membranes and bi-polar extension of microtubules toward the cell (Sanchez and Feldman, 2017; Clark et al., 2002). Myoblast fusion occurs at the tips of elongated myoblasts through close contacting and merging of the cell membranes; hence, the failure of myoblast elongation can lead to a lack of fusion (Kim et al., 2015). As fusion proceeds, the nucleus of a newly fused myoblast rapidly migrates to the central myotube nuclei, which is mediated by dynein motor–Par6 complex that connects the nuclear envelope with microtubules (Bone and Starr, 2016; Starr, 2017). Then, these nuclei spread apart and are evenly spaced throughout the myotube, and the process is currently thought to be regulated by a variety of mechanisms: the kinesin-1 motor-nesprin1/2 complex links the nuclei to the microtubules and transports the nuclei along the microtubules; the kinesin-1 motor-microtubule-associated protein 7 (MAP7) complex drives the sliding of anti-parallel microtubules that extend outward from adjacent nuclei, thereby pushing neighboring nuclei apart; Nesprin-α recruits centrosomal protein Akap450 to the nuclear envelope independently of kinesin to nucleate microtubules (Starr, 2017; Gimpel et al., 2017). Eventually, the nuclei move from the center of the myofiber to the sarcolemma in a desmin-dependent process; except for some nuclei that migrate to the NMJ and anchor there, the remaining nuclei are evenly spaced around the myofiber periphery (Bone and Starr, 2016). Moreover, microtubules bound by myosin-binding protein Muscle RING finger-2 (MURF-2) provide a scaffold for the association of sarcomeric myosin with the A-band portion of titin, thereby generating the mature A band during myofibril assembly (Pizon et al., 2002; Pizon et al., 2005). In a similar manner, another microtubule-binding protein dystrophin serves as the scaffold for microtubule growth, resulting in microtubules aligning with dystrophin in both longitudinal and transverse directions of the sarcomere (Prins et al., 2009). Currently, research on microtubule organization within myofibers is continually deepening, such as its involvement in fiber type-dependent, Golgi apparatus relevance, mitochondrial trafficking, and primary cilia signaling (Ng et al., 2021; Oddoux et al., 2013; Giovarelli et al., 2020).