38.4B: Skeletal Muscle Fibers - Biology

38.4B: Skeletal Muscle Fibers - Biology

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Skeletal muscles are composed of striated subunits called sarcomeres, which are composed of the myofilaments actin and myosin.

Learning Objectives

  • Outline the structure of a skeletal muscle fiber

Key Points

  • Muscles are composed of long bundles of myocytes or muscle fibers.
  • Myocytes contain thousands of myofibrils.
  • Each myofibril is composed of numerous sarcomeres, the functional contracile region of a striated muscle. Sarcomeres are composed of myofilaments of myosin and actin, which interact using the sliding filament model and cross-bridge cycle to contract.

Key Terms

  • sarcoplasm: The cytoplasm of a myocyte.
  • sarcoplasmic reticulum: The equivalent of the smooth endoplasmic reticulum in a myocyte.
  • sarcolemma: The cell membrane of a myocyte.
  • sarcomere: The functional contractile unit of the myofibril of a striated muscle.

Skeletal Muscle Fiber Structure

Myocytes, sometimes called muscle fibers, form the bulk of muscle tissue. They are bound together by perimysium, a sheath of connective tissue, into bundles called fascicles, which are in turn bundled together to form muscle tissue. Myocytes contain numerous specialized cellular structures which facilitate their contraction and therefore that of the muscle as a whole.

The highly specialized structure of myocytes has led to the creation of terminology which differentiates them from generic animal cells.

Generic cell > Myocyte

Cytoplasm > Sarcoplasm

Cell membrane > Sarcolemma

Smooth endoplasmic reticulum > Sarcoplasmic reticulum

Myocyte Structure

Myocytes can be incredibly large, with diameters of up to 100 micrometers and lengths of up to 30 centimeters. The sarcoplasm is rich with glycogen and myoglobin, which store the glucose and oxygen required for energy generation, and is almost completely filled with myofibrils, the long fibers composed of
myofilaments that facilitate muscle contraction.

The sarcolemma of myocytes contains numerous invaginations (pits) called transverse tubules which are usually perpendicular to the length of the myocyte. Transverse tubules play an important role in supplying the myocyte with Ca+ ions, which are key for muscle contraction.

Each myocyte contains multiple nuclei due to their derivation from multiple myoblasts, progenitor cells that give rise to myocytes. These myoblasts asre located to the periphery of the myocyte and flattened so
as not to impact myocyte contraction.

Myofibril Structure

Each myocyte can contain many thousands of myofibrils. Myofibrils run parallel to the myocyte and typically run for its entire length, attaching to the sarcolemma at either end. Each myofibril is surrounded by the sarcoplasmic reticulum, which is closely associated with the transverse tubules. The sarcoplasmic reticulum acts as a sink of Ca+ ions, which are released upon signalling from the transverse tubules.


Myofibrils are composed of long myofilaments of actin, myosin, and other associated proteins. These proteins are organized into regions termed sarcomeres, the functional contractile region of the myocyte. Within the sarcomere actin and myosin, myofilaments are interlaced with each other and slide over each other via the sliding filament model of contraction. The regular organization of these sarcomeres gives skeletal and cardiac muscle their distinctive striated appearance.

Sarcomere: The sarcomere is the functional contractile region of the myocyte, and defines the region of interaction between a set of thick and thin filaments.

Myofilaments (Thick and Thin Filaments)

Myofibrils are composed of smaller structures called myofilaments. There are two main types of myofilaments: thick filaments and thin filaments. Thick filaments are composed primarily of myosin proteins, the tails of which bind together leaving the heads exposed to the interlaced thin filaments. Thin filaments are composed of actin, tropomyosin, and troponin. The molecular model of contraction which describes the interaction between actin and myosin myofilaments is called the cross-bridge cycle.

BIO 140 - Human Biology I - Textbook

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Chapter 38

Skeletal Muscle

  • Describe the layers of connective tissues packaging skeletal muscle
  • Explain how muscles work with tendons to move the body
  • Identify areas of the skeletal muscle fibers
  • Describe excitation-contraction coupling

The best-known feature of skeletal muscle is its ability to contract and cause movement. Skeletal muscles act not only to produce movement but also to stop movement, such as resisting gravity to maintain posture. Small, constant adjustments of the skeletal muscles are needed to hold a body upright or balanced in any position. Muscles also prevent excess movement of the bones and joints, maintaining skeletal stability and preventing skeletal structure damage or deformation. Joints can become misaligned or dislocated entirely by pulling on the associated bones muscles work to keep joints stable. Skeletal muscles are located throughout the body at the openings of internal tracts to control the movement of various substances. These muscles allow functions, such as swallowing, urination, and defecation, to be under voluntary control. Skeletal muscles also protect internal organs (particularly abdominal and pelvic organs) by acting as an external barrier or shield to external trauma and by supporting the weight of the organs.

Skeletal muscles contribute to the maintenance of homeostasis in the body by generating heat. Muscle contraction requires energy, and when ATP is broken down, heat is produced. This heat is very noticeable during exercise, when sustained muscle movement causes body temperature to rise, and in cases of extreme cold, when shivering produces random skeletal muscle contractions to generate heat.

Each skeletal muscle is an organ that consists of various integrated tissues. These tissues include the skeletal muscle fibers, blood vessels, nerve fibers, and connective tissue. Each skeletal muscle has three layers of connective tissue (called &ldquomysia&rdquo) that enclose it and provide structure to the muscle as a whole, and also compartmentalize the muscle fibers within the muscle (Figure 1). Each muscle is wrapped in a sheath of dense, irregular connective tissue called the epimysium , which allows a muscle to contract and move powerfully while maintaining its structural integrity. The epimysium also separates muscle from other tissues and organs in the area, allowing the muscle to move independently.

Figure 1: Bundles of muscle fibers, called fascicles, are covered by the perimysium. Muscle fibers are covered by the endomysium.

Inside each skeletal muscle, muscle fibers are organized into individual bundles, each called a fascicle , by a middle layer of connective tissue called the perimysium . This fascicular organization is common in muscles of the limbs it allows the nervous system to trigger a specific movement of a muscle by activating a subset of muscle fibers within a bundle, or fascicle of the muscle. Inside each fascicle, each muscle fiber is encased in a thin connective tissue layer of collagen and reticular fibers called the endomysium . The endomysium contains the extracellular fluid and nutrients to support the muscle fiber. These nutrients are supplied via blood to the muscle tissue.

In skeletal muscles that work with tendons to pull on bones, the collagen in the three tissue layers (the mysia) intertwines with the collagen of a tendon. At the other end of the tendon, it fuses with the periosteum coating the bone. The tension created by contraction of the muscle fibers is then transferred though the mysia, to the tendon, and then to the periosteum to pull on the bone for movement of the skeleton. In other places, the mysia may fuse with a broad, tendon-like sheet called an aponeurosis , or to fascia, the connective tissue between skin and bones. The broad sheet of connective tissue in the lower back that the latissimus dorsi muscles (the &ldquolats&rdquo) fuse into is an example of an aponeurosis.

Every skeletal muscle is also richly supplied by blood vessels for nourishment, oxygen delivery, and waste removal. In addition, every muscle fiber in a skeletal muscle is supplied by the axon branch of a somatic motor neuron, which signals the fiber to contract. Unlike cardiac and smooth muscle, the only way to functionally contract a skeletal muscle is through signaling from the nervous system.

Skeletal Muscle Fibers

Because skeletal muscle cells are long and cylindrical, they are commonly referred to as muscle fibers. Skeletal muscle fibers can be quite large for human cells, with diameters up to 100 &mum and lengths up to 30 cm (11.8 in) in the Sartorius of the upper leg. During early development, embryonic myoblasts, each with its own nucleus, fuse with up to hundreds of other myoblasts to form the multinucleated skeletal muscle fibers. Multiple nuclei mean multiple copies of genes, permitting the production of the large amounts of proteins and enzymes needed for muscle contraction.

Some other terminology associated with muscle fibers is rooted in the Greek sarco, which means &ldquoflesh.&rdquo The plasma membrane of muscle fibers is called the sarcolemma , the cytoplasm is referred to as sarcoplasm , and the specialized smooth endoplasmic reticulum, which stores, releases, and retrieves calcium ions (Ca ++ ) is called the sarcoplasmic reticulum (SR) (Figure 2). As will soon be described, the functional unit of a skeletal muscle fiber is the sarcomere, a highly organized arrangement of the contractile myofilaments actin (thin filament) and myosin (thick filament), along with other support proteins.

Figure 2: A skeletal muscle fiber is surrounded by a plasma membrane called the sarcolemma, which contains sarcoplasm, the cytoplasm of muscle cells. A muscle fiber is composed of many fibrils, which give the cell its striated appearance.

The Sarcomere

The striated appearance of skeletal muscle fibers is due to the arrangement of the myofilaments of actin and myosin in sequential order from one end of the muscle fiber to the other. Each packet of these microfilaments and their regulatory proteins, troponin and tropomyosin (along with other proteins) is called a sarcomere .

Watch the video linked to below to learn more about macro- and microstructures of skeletal muscles. (a) What are the names of the &ldquojunction points&rdquo between sarcomeres? (b) What are the names of the &ldquosubunits&rdquo within the myofibrils that run the length of skeletal muscle fibers? (c) What is the &ldquodouble strand of pearls&rdquo described in the video? (d) What gives a skeletal muscle fiber its striated appearance?

The sarcomere is the functional unit of the muscle fiber. The sarcomere itself is bundled within the myofibril that runs the entire length of the muscle fiber and attaches to the sarcolemma at its end. As myofibrils contract, the entire muscle cell contracts. Because myofibrils are only approximately 1.2 &mum in diameter, hundreds to thousands (each with thousands of sarcomeres) can be found inside one muscle fiber. Each sarcomere is approximately 2 &mum in length with a three-dimensional cylinder-like arrangement and is bordered by structures called Z-discs (also called Z-lines, because pictures are two-dimensional), to which the actin myofilaments are anchored (Figure 3). Because the actin and its troponin-tropomyosin complex (projecting from the Z-discs toward the center of the sarcomere) form strands that are thinner than the myosin, it is called the thin filament of the sarcomere. Likewise, because the myosin strands and their multiple heads (projecting from the center of the sarcomere, toward but not all to way to, the Z-discs) have more mass and are thicker, they are called the thick filament of the sarcomere.

Figure 3: The sarcomere, the region from one Z-line to the next Z-line, is the functional unit of a skeletal muscle fiber.

The Neuromuscular Junction

Another specialization of the skeletal muscle is the site where a motor neuron&rsquos terminal meets the muscle fiber&mdashcalled the neuromuscular junction (NMJ) . This is where the muscle fiber first responds to signaling by the motor neuron. Every skeletal muscle fiber in every skeletal muscle is innervated by a motor neuron at the NMJ. Excitation signals from the neuron are the only way to functionally activate the fiber to contract.

Every skeletal muscle fiber is supplied by a motor neuron at the NMJ. Watch the video linked to below to learn more about what happens at the NMJ. (a) What is the definition of a motor unit? (b) What is the structural and functional difference between a large motor unit and a small motor unit? (c) Can you give an example of each? (d) Why is the neurotransmitter acetylcholine degraded after binding to its receptor?

Excitation-Contraction Coupling

All living cells have membrane potentials, or electrical gradients across their membranes. The inside of the membrane is usually around -60 to -90 mV, relative to the outside. This is referred to as a cell&rsquos membrane potential. Neurons and muscle cells can use their membrane potentials to generate electrical signals. They do this by controlling the movement of charged particles, called ions, across their membranes to create electrical currents. This is achieved by opening and closing specialized proteins in the membrane called ion channels. Although the currents generated by ions moving through these channel proteins are very small, they form the basis of both neural signaling and muscle contraction.

Both neurons and skeletal muscle cells are electrically excitable, meaning that they are able to generate action potentials. An action potential is a special type of electrical signal that can travel along a cell membrane as a wave. This allows a signal to be transmitted quickly and faithfully over long distances.

Although the term excitation-contraction coupling confuses or scares some students, it comes down to this: for a skeletal muscle fiber to contract, its membrane must first be &ldquoexcited&rdquo&mdashin other words, it must be stimulated to fire an action potential. The muscle fiber action potential, which sweeps along the sarcolemma as a wave, is &ldquocoupled&rdquo to the actual contraction through the release of calcium ions (Ca ++ ) from the SR. Once released, the Ca ++ interacts with the shielding proteins, forcing them to move aside so that the actin-binding sites are available for attachment by myosin heads. The myosin then pulls the actin filaments toward the center, shortening the muscle fiber.

In skeletal muscle, this sequence begins with signals from the somatic motor division of the nervous system. In other words, the &ldquoexcitation&rdquo step in skeletal muscles is always triggered by signaling from the nervous system (Figure 4).

Figure 4: At the NMJ, the axon terminal releases ACh. The motor end-plate is the location of the ACh-receptors in the muscle fiber sarcolemma. When ACh molecules are released, they diffuse across a minute space called the synaptic cleft and bind to the receptors.

The motor neurons that tell the skeletal muscle fibers to contract originate in the spinal cord, with a smaller number located in the brainstem for activation of skeletal muscles of the face, head, and neck. These neurons have long processes, called axons, which are specialized to transmit action potentials long distances&mdash in this case, all the way from the spinal cord to the muscle itself (which may be up to three feet away). The axons of multiple neurons bundle together to form nerves, like wires bundled together in a cable.

Signaling begins when a neuronal action potential travels along the axon of a motor neuron, and then along the individual branches to terminate at the NMJ. At the NMJ, the axon terminal releases a chemical messenger, or neurotransmitter , called acetylcholine (ACh) . The ACh molecules diffuse across a minute space called the synaptic cleft and bind to ACh receptors located within the motor end-plate of the sarcolemma on the other side of the synapse. Once ACh binds, a channel in the ACh receptor opens and positively charged ions can pass through into the muscle fiber, causing it to depolarize , meaning that the membrane potential of the muscle fiber becomes less negative (closer to zero.)

As the membrane depolarizes, another set of ion channels called voltage-gated sodium channels are triggered to open. Sodium ions enter the muscle fiber, and an action potential rapidly spreads (or &ldquofires&rdquo) along the entire membrane to initiate excitation-contraction coupling.

Things happen very quickly in the world of excitable membranes (just think about how quickly you can snap your fingers as soon as you decide to do it). Immediately following depolarization of the membrane, it repolarizes, re-establishing the negative membrane potential. Meanwhile, the ACh in the synaptic cleft is degraded by the enzyme acetylcholinesterase (AChE) so that the ACh cannot rebind to a receptor and reopen its channel, which would cause unwanted extended muscle excitation and contraction.

Propagation of an action potential along the sarcolemma is the excitation portion of excitation-contraction coupling. Recall that this excitation actually triggers the release of calcium ions (Ca ++ ) from its storage in the cell&rsquos SR. For the action potential to reach the membrane of the SR, there are periodic invaginations in the sarcolemma, called T-tubules (&ldquoT&rdquo stands for &ldquotransverse&rdquo). You will recall that the diameter of a muscle fiber can be up to 100 &mum, so these T-tubules ensure that the membrane can get close to the SR in the sarcoplasm. The arrangement of a T-tubule with the membranes of SR on either side is called a triad (Figure 5). The triad surrounds the cylindrical structure called a myofibril , which contains actin and myosin.

Figure 5: Narrow T-tubules permit the conduction of electrical impulses. The SR functions to regulate intracellular levels of calcium. Two terminal cisternae (where enlarged SR connects to the T-tubule) and one T-tubule comprise a triad&mdasha &ldquothreesome&rdquo of membranes, with those of SR on two sides and the T-tubule sandwiched between them.

The T-tubules carry the action potential into the interior of the cell, which triggers the opening of calcium channels in the membrane of the adjacent SR, causing Ca ++ to diffuse out of the SR and into the sarcoplasm. It is the arrival of Ca ++ in the sarcoplasm that initiates contraction of the muscle fiber by its contractile units, or sarcomeres.

Chapter Review

Skeletal muscles contain connective tissue, blood vessels, and nerves. There are three layers of connective tissue: epimysium, perimysium, and endomysium. Skeletal muscle fibers are organized into groups called fascicles. Blood vessels and nerves enter the connective tissue and branch in the cell. Muscles attach to bones directly or through tendons or aponeuroses. Skeletal muscles maintain posture, stabilize bones and joints, control internal movement, and generate heat.

Skeletal muscle fibers are long, multinucleated cells. The membrane of the cell is the sarcolemma the cytoplasm of the cell is the sarcoplasm. The sarcoplasmic reticulum (SR) is a form of endoplasmic reticulum. Muscle fibers are composed of myofibrils. The striations are created by the organization of actin and myosin resulting in the banding pattern of myofibrils.

Current Methods for Skeletal Muscle Tissue Repair and Regeneration

Skeletal muscle has the capacity of regeneration after injury. However, for large volumes of muscle loss, this regeneration needs interventional support. Consequently, muscle injury provides an ongoing reconstructive and regenerative challenge in clinical work. To promote muscle repair and regeneration, different strategies have been developed within the last century and especially during the last few decades, including surgical techniques, physical therapy, biomaterials, and muscular tissue engineering as well as cell therapy. Still, there is a great need to develop new methods and materials, which promote skeletal muscle repair and functional regeneration. In this review, we give a comprehensive overview over the epidemiology of muscle tissue loss, highlight current strategies in clinical treatment, and discuss novel methods for muscle regeneration and challenges for their future clinical translation.

1. Introduction

Skeletal muscle is one of the most abundant tissues in the human body. It accounts for 40%–45% of the total body mass and is necessary for generating forces for movement [1]. Up to a certain threshold, skeletal muscle has the capability of regenerating lost tissue upon injury [2]. Beyond this threshold, the remaining muscle tissue is unable to fully regenerate its function. This loss of skeletal muscle with lasting functional impairment is defined as “volumetric muscle loss” (VML) [3–5]. It can substantially impact the quality of life of patients by significantly reducing the functionality of the locomotion system [4].

Frequent reasons for skeletal muscle injuries are high-energy traffic accidents, blast trauma, combat injuries, surgical and orthopedic situations (e.g., after compartment syndrome or tumor resection), or contusion injury during sports that lead to an acute muscle tissue loss [6, 7]. Approximately 35–55% of sport injuries involve muscle damage at the myofiber level [8]. Those injuries that involve 20% or more of muscle loss of the respective muscle mass need reconstructive surgical procedures [9]. Progressive muscle loss can result from metabolic disorders or inherited genetic diseases such as Duchenne muscular dystrophy, Amyotrophic Lateral Sclerosis, and pediatric Charcot-Marie-Tooth disease [10–13]. Muscle atrophy can also be a consequence of peripheral nerve injuries, chronic kidney disease, diabetes, and heart failure [14, 15]. Up to 20% loss of muscle mass can be compensated by the high adaptability and regenerative potential of skeletal muscle. Beyond this threshold functional impairment is inevitable and can lead to severe disability as well as cosmetic deformities, which is why therapeutic options are in urgent demand for these patients [4, 5, 16, 17].

Muscle regeneration relies on a heterogeneous population of satellite cells, interstitial cells, and blood vessels and is mainly controlled through ECM proteins and secreted factors [18, 19]. Normally muscle mass is maintained by a balance between protein synthesis and degradation [20]. In most cases of VML, the regeneration capability of skeletal muscles is impeded, because necessary regenerative elements, mainly satellite cells, perivascular stem cells, and the basal lamina, are physically removed [21, 22]. Through denervation, protein degradation pathways (the proteasomal and the autophagic-lysosomal pathways) are activated. Therefore protein degradation rates exceed protein synthesis, which contributes to the muscle atrophy accompanied by gradual decrease of muscle wet weight and muscle fiber diameters [23, 24].

Revascularization is typically impaired. The following ischemic conditions favor fibroblast proliferation, fibrosis, and fibrotic scar tissue formation, which leads to further degeneration of the muscle [25]. The ECM composition and extent in scar tissues affect many aspects of myogenesis, muscle function, and reinnervation [26]. It can severely constrain motion and thereby aggravate the consequences of muscle tissue loss. Also in chronic muscle loss like Duchenne muscular dystrophy, fibrosis is a major problem [27]. Here, the consistent breakdown of myofibers cannot be fully compensated by satellite cell proliferation. The following inflammatory processes lead to an altered production of extracellular matrix (ECM) and consequent development of fibrosis and scar tissue formation [27–29]. This scar formation can be reduced either by injection of, for example, 5-fluorouracil and bleomycin, which antagonizes fibroblast proliferation and neoangiogenesis or by laser therapy with release of contracture and functional improvements after 6–12 months’ treatment [30, 31]. Regeneration with regression of scar tissue and functional recovery can furthermore be optimized with fat grafting [32]. However, reducing scar formation is not enough for promoting muscle tissue repair and regeneration. This reinvigorates clinical and research efforts directed at replacing or regenerating larger volumes of muscle tissue.

2. Current Methods for Treating Muscle Tissue Loss in the Clinic

Current standard of care for VML is typically based on surgical intervention with autologous muscle graft and physical therapy. Further clinically used strategies include acupuncture and application of scaffolds.

2.1. Surgical Techniques

Surgical treatment for VML includes mainly scar tissue debridement and/or muscle transposition [33]. Autologous muscle transfer is commonly performed in a clinical situation, when there are large areas of muscle loss following trauma, tumor resection, or nerve injury, which impairs the irreplaceable motor function [34, 35]. The surgeons graft healthy muscle from a donor site unaffected by the injury to restore the lost or impaired function [36]. When no adjacent muscle is available because of high-level nerve injuries or severe trauma, autologous muscle transplantation together with neurorrhaphy, in the form of free functional muscle transfer, can be applied [37, 38]. The most popular autologous muscles are latissimus dorsi muscle and gracilis muscle. Latissimus dorsi muscle transfer has been shown to be safe and efficient for restoration of elbow flexion after injuries [34]. In the case of a synovial sarcoma affecting the right gluteus medius and minimus muscles, the function of the affected hip abduction could be fully reconstructed with a free neurovascular latissimus dorsi muscle transplantation [39]. Free gracilis muscle transfer is commonly utilized to restore elbow flexion after pan-brachial plexus injury [40]. It is also applied for muscle weakness after facial palsy or pelvic floor reconstruction [41, 42]. Although functional muscle flaps can lead to at least decent functional results, they cause substantial donor site morbidity and inadequate innervation [43]. Moreover, as many as 10% of these reconstructive surgeries result in complete graft failure due to complications such as infection and necrosis [44]. Sometimes, the source of autologous muscles for grafting is a problem, if the patient is severely injured.

2.2. Physical Therapy

Exercise has the ability to prevent a decrease of skeletal muscle mass [45]. Thus, in addition to surgical techniques, physical therapy is a noninvasive/minimally invasive way to promote muscle tissue repair and regeneration. It is especially used for rehabilitation after injuries and muscle tissue transfer, or to treat chronic muscle loss.

Physical rehabilitation aims at strengthening the remaining muscles. This has been shown to accelerate muscle healing/regeneration by modulating the immune response, release of growth factors, promoting vascularization, and reducing scar formation [46–48]. Functional performance of nonrepaired VML injured muscle could be significantly improved with physical rehabilitation in the form of voluntary wheel running [49]. Interventions to enhance angiogenesis including exercise and massage are potential strategies to accelerate new muscle formation in clinically transplanted muscle grafts or other surgical situations [50]. It has been reported that physical exercise can upregulate the IGF-1 signaling pathway and decrease myostatin in muscle tissue of animals and humans, thus preventing muscle atrophy [51–53].

Physical therapy can indeed improve muscle repair and recovery however, it is unable to facilitate substantial muscle regeneration within the defect areas in VML. In addition, patients with severe diseases or injuries are frequently unable to make consistent exercise, which limits physical therapy as a treatment for VML.

2.3. Acupuncture

Acupuncture is a branch of traditional Chinese medicine, which has been widely used to treat various diseases around the world [54–56]. Electrical acupuncture treatment has been shown to suppress myostatin expression, leading to satellite cell proliferation and skeletal muscle repair [57]. Acupuncture plus low-frequency electrical stimulation (Acu-LFES) could enhance muscle regeneration and prevent muscle loss by replicating the benefits of exercise through stimulation of muscle contraction [58]. It is suitable for some patients with severe diseases, which are unable to perform exercise frequently. Acu-LFES was shown to counteract diabetes-induced skeletal muscle atrophy by increasing IGF-1 and thereby stimulating muscle regeneration [58]. Application of Acu-LFES for the treatment of diabetic myopathy and muscle loss induced by chronic kidney disease showed good functional improvement of the muscle [58, 59]. The underlying mechanism includes activation of M2 microphages and reversing mRNA expression levels of the E3 ubiquitin ligase atrogin-1.

Similar to physical exercise, acupuncture improves muscle function restoration and stimulates muscle regeneration especially in patients with muscle atrophy after chronic diseases. However, there is limited success for the regeneration of large volume muscle defects after trauma or tumor resection. Furthermore, more work needs to be done to determine the optimal timing and intensity of Acu-LFES as a standard treatment for muscle atrophy.

2.4. Biological Scaffolds

Biological scaffolds composed of extracellular matrix (ECM) proteins are commonly used in regenerative medicine and in surgical procedures for tissue reconstruction and regeneration. The scaffolds can promote the repair of VML by providing a structural and biochemical framework [60]. For smaller amounts of muscle loss, several tissue-derived scaffolds have been tested in animal models and translated into the clinic for surgical application [6]. Xenogeneic extracellular matrix and autologous tissue have been utilized to restore functional muscle and simultaneously generate a biological niche for recovery [61]. A multilayered scaffold made of ECM derived from porcine intestinal submucosa has been applied for reconstruction of vastus medialis muscle in patients [16]. The patient showed marked gains in isokinetic performance 4 months after surgery and new muscle tissue at the implant site was demonstrated by computer tomography. Porcine small intestinal submucosa-extracellular matrix has also been utilized for the treatment of abdominal musculoskeletal wall defects, where it was sutured at the defect corners and subcuticularly closed with a vicryl-suture [61]. Also, porcine ECM from urinary bladder has been implanted in an attempt to treat VML in human beings [60]. Functional improvement with formation of muscle tissue was observed in three of the five human patients in this study.

However, allograft or xenogeneic scaffolds can still induce adverse immune response after decellularization and there might be potential risk of infectious disease transmission. Therefore, there is a clinical need to develop new strategies that can facilitate safe bigger muscle tissue repair and regeneration.

3. Developing Technologies for Muscle Tissue Engineering and Regeneration

To address remaining clinical problems and explore novel strategies for muscle tissue engineering and regeneration, new technologies have been investigated intensively. While tissue bioengineering approaches aim to construct complex muscle structures in vitro for subsequent implantation and replacement of the missing muscles, tissue regeneration approaches develop tissue-like scaffolds that can be implanted to enhance new muscle formation from remaining tissue in vivo [62]. Both approaches mainly rely on combinations of scaffolds, cells, and molecular signaling with differing focus.

3.1. Scaffold-Based Strategies

Biomaterials can provide chemical and physical cues to transplanted cells or host muscle cells to enhance their survival, promote their functional maturation, protect them from the foreign body responses, and recruit host cells and regenerate muscle tissues [63]. Biological scaffolds are used in a variety of clinical tissue engineering applications and have been studied in preclinical skeletal muscle VML injury models frequently over the last decade. They are mainly made of natural polymers, synthetic polymers, or ECM and attempt to create a microenvironment niche to favorably control the behavior of resident cells.

Natural polymers such as alginate, collagen, and fibrin have been utilized extensively in skeletal muscle engineering [64–66]. They possess intrinsic bioactive signaling cues to enhance cell behavior [67–69]. Alginate gels with a stiffness of 13–45 kPa were found to maximize myoblast proliferation and differentiation [70]. Freeze-dried collagen scaffolds facilitated the integration of aligned myotubes into a large muscle defect, which were capable of producing force upon electrical stimulation [71]. Collagen could also supply necessary growth factors to the wound site to increase muscle cell migration [72, 73]. Fibrin gels were reported to promote myoblast survival and differentiation into myofibers when integrated in tissues [74]. Fibrin scaffolds with microthread architecture were also shown to support the healing of VML in mouse models [75].

As the natural polymer only offers limited mechanical stiffness and can be easily degraded, a variety of synthetic materials have been used for skeletal muscle regeneration such as PGA, PLA, and PLGA [66, 76–78]. Myoblasts seeded onto electrospun meshes with aligned nanofiber orientation can fuse into highly aligned myotubes [78]. Furthermore, synthetic scaffolds can be easily engineered to facilitate the controlled release of growth factors for inducing muscle regeneration [75, 79]. The main disadvantages include typically poorer cell affinity compared to natural polymers and the risk of stimulation of a foreign body response by the polymer or its degradation products [79].

To improve regeneration of muscle tissues, the in vivo microenvironment of the scaffolds ideally would mimic native tissues and thereby facilitate remodeling of the neotissue [80]. An attractive approach for the repair of VML is therefore the transplantation of a myoinductive decellularized scaffold that attracts the cells required for myogenesis from the host. That is why muscle-derived ECM scaffolds are popularly investigated. These ECM scaffolds can fill the defect and restore morphology temporarily [17]. They can further be filled by bone-marrow derived mesenchymal stem cells (MSCs) after implantation. This enriched matrix gains more blood vessels and regenerates more myofibers than “conventional” extracellular matrix [17, 81]. Indeed, hydrogels derived from decellularized skeletal muscle matrix have been shown to enhance the proliferation of skeletal myoblasts when injected into an ischemic rat limb [82]. An alternative method could be to utilize minced skeletal muscle tissue that has not been decellularized, which has been reported to show better muscle regeneration than devitalized scaffolds [83]. Comparable to muscle-derived matrix, small intestinal submucosa-extracellular matrix can lead to contractile sheets of skeletal muscle with comparable contractile force [61]. For in vitro muscle tissue engineering, rat myoblasts have also been preconditioned on a porcine bladder acellular matrix in a bioreactor and then implanted in nude mice at a muscle defect to restore muscular tissue [80].

Another obstacle in muscle regeneration is the musculotendinous junction. This can be partly restored in absence of implanted cells by extracellular matrix-based platforms that have been shown to withstand half of the force of the contralateral site after complete resection in a mammalian model [80]. The newly formed muscle cells have shown better adherence to 3D polyurethane-based porous scaffolds with low stiffness and larger roughness values [84].

3.2. Cell-Based Strategies

Muscle fiber regeneration is performed by cells and consequently cell-based strategies for regeneration have been pursued [83, 85]. The cell types utilized for treating muscle loss mainly include myoblasts, satellite cells (SCs), mesoangioblasts, pericytes, and mesenchymal stem cells (MSCs) [86–88]. The most well characterized muscle stem cell is the satellite cell (SC). SCs are able to contribute extensively to the formation of new muscle fibers [86, 89]. SCs transplanted into dystrophin-deficient mdx mice yielded highly efficient regeneration of dystrophic muscle and improved muscle contractile function [90]. Unfortunately, in vitro expansion of SCs results in significant reduction of their ability to produce myofibers in vivo [91] and consequently, obtaining a sufficiently large number of fresh SCs for clinical application is impractical [92]. Myoblasts have been used for reconstructing muscle tissue defects with a variety of scaffolds [87, 93, 94]. They were shown to functionally integrate into the existing musculature of the host. Injection of a larger number of myoblasts into muscles showed promising results for the treatment of dystrophin-deficient models [95]. Also MSCs could be involved in myotube formation through heterotypic cell fusion after myogenic gene activation [88]. Mesoangioblasts and pericytes have been studied for treating muscular dystrophy, which resulted in increasing the force [96]. They have also been utilized in tissue engineered hydrogel carriers, with some reported success for promoting muscle regeneration [97].

Stem-cell-based therapies provide notable therapeutic benefits on reversing muscle atrophy and promoting muscle regeneration. Stem cell therapy (e.g., umbilical cord blood stem cell transplantation) showed positive results for treating Duchenne muscular dystrophy [98]. After application of stem cells, an increase of dystrophin positive muscular fibers was found. Biopsies of calf muscle showed growing myoblasts cells and muscular tubes and an improvement in arms and legs during physical examination was reported.

3.3. Molecular Signaling Based Strategies

Beside cues from the ECM, also a diversity of stimulatory and inhibitory growth factors such as IGF-1 and TGF-ß1 can drive endogenous skeletal muscle regeneration by activating and/or recruiting host stem cells [22]. They can be loaded on scaffolds for controlled delivery to the injured areas [72, 99]. Sustained delivery of VEGF, IGF-1, or SDF-1a was shown to enhance myogenesis and promote angiogenesis and muscle formation [73, 100–102]. Rapid release of hepatocyte growth factor (HGF) loaded on fibrin microthread scaffolds promoted remodeling of functional muscle tissue and enhanced the regeneration of skeletal muscle in mouse models [75]. Combination therapy of h-ADSCs and bFGF hydrogels resulted in functional recovery, revascularization, and reinnervation in lacerated muscles with minimal fibrosis [103]. Furthermore, PEDF peptide was reported to promote the regeneration of skeletal muscles [104].

Research into the pathogenesis of sarcopenia as one of the most frequent muscular diseases has elucidated different molecular pathways. The most promising targets include BMP and myostatin [105]. Indeed, medication with human recombinant BMP-2/7 and antimyostatin can help to reduce sarcopenic symptoms [106]. Cachexia is addressed with anamorelin, a ghrelin agonist, and selective androgen receptor modulator as well as anticytokines/myokines [107]. Another factor involved in muscle healing seems to be TGF-β. Increased TGF-β1 levels, which could be detected after the use of nonsteroidal anti-inflammatory drugs, helped to regenerate muscle tissue [108–110].

Spinal muscular atrophy arises from mutations in the survival motor neuron 1 (SMN1) gene, which often leads to the deficiency of the ubiquitous SMN protein [111]. Therefore, one of the most promising strategies is to increase the levels of full-length SMN [112]. Nusinersen is an antisense oligonucleotide drug developed for the treatment of spinal muscular atrophy (SMA), which has been approved by the US Food and Drug Administration (FDA) and European Medicines Agency (EMA) [113]. It can modulate the pre-mRNA splicing of the survival motor neuron 2 gene and showed significant improvement of muscle function after treatment. Clinical trials on infants showed significant mean improvements in developmental motor milestones including sitting, walking, and motor function [114].

3.4. Other Developing Techniques

The effect of heat stress on skeletal muscle regeneration was investigated in experimental rats [115]. Results showed that applying heat packs immediately after crush injury accelerated the degeneration process at the injured site, facilitated migration of macrophages, proliferation, and differentiation of satellite cells, and promoted muscle tissue regeneration.

Low-level laser therapy (LLLT) has also been evaluated as a therapeutic approach for stimulating muscle repair and recovery after endurance exercise training in rats [116]. Other results from the rat model suggest that it could also be an option to reduce fibrosis and myonecrosis triggered by bupivacaine and accelerate the muscle regeneration process [117]. As possible mechanisms, decreased inflammation and muscle creatine kinase levels are discussed. The combination of LLLT with platelet rich plasma (PRP) produced better results for promoting muscle regeneration after injuries compared to the isolated use of LLLT or PRP [118].

The effect of neuromuscular electrical stimulation (NMES) on skeletal muscle regeneration was assessed in healthy subjects. It increased the proliferation of myogenic precursor cells (MPCs) and their fusion with mature myofibers, which improved the regenerative capacity of skeletal muscle [119]. The effect on models with muscle injury or VML needs to be further investigated.

4. Challenges and Future Perspectives

4.1. Mechanical Properties of Biomaterials

Biomaterials for muscle tissue engineering and regeneration should persist long enough to support organized functional muscle regeneration and could be degraded gradually along with new tissue formation. The scaffolds created with natural polymers are usually associated with poor mechanical stiffness and rapid degradability, when not chemically crosslinked [120]. Synthetic polymers provide an artificial alternative with flexible mechanical properties [121, 122]. However, the use of synthetic scaffolds can be associated with side effects such as inhibition of cell migration and cell-to-cell communication [123].

A challenge for the near future will be to join the advantageous properties of natural and artificial polymers. Design of scaffolds combining favorable cell interaction with mechanical strength will facilitate implantation, give direct support to the tissue, and allow remodeling and therefore regeneration of the impaired tissue. Ideally these materials can then be used in combination with 3D-printing technology to tailor the scaffold based on the individual loss of muscle.

The mechanical and surface properties of the scaffold can be further engineered to affect the cell behavior in terms of adhesion, proliferation, migration, and differentiation [124]. If stem cells are seeded onto such scaffolds, they may therefore be guided to differentiate into different types of cells based on the scaffold properties [125, 126]. Moreover, degradation products from an ECM scaffold might contribute to the recruitment of host cells for tissue remodeling by chemoattraction [127]. Thus, better understanding of cell-scaffold interaction and development of a carrier scaffold that stimulates the niche environment for ongoing remodeling processes are further goals for future development in this area.

4.2. Vascularization in the Process of Regeneration

For engineering muscle constructs in vitro, one of the major limitations is the lack of vascularization [128]. It has been shown that myoblasts need to be within 150 μm of the supply route for oxygen and nutrients (typically vessels) to survive, proliferate, and differentiate [129]. This limits the size of constructs without a functional vascular network. Insufficient vascularization can lead to nutrient deficiencies and hypoxia deeper in the scaffolds, which results in nonuniform cell differentiation and integration, and thus decreases tissue functionality [130].

Also for in vivo muscle tissue regeneration facilitated by bioengineered muscle tissue constructs, the absence of immediate blood supply is one main reason for failure [131]. Complete revascularization of scaffolds by ingrowth of bed vessels into the graft can take up to 3 weeks, which significantly limits the capacity to obtain scar free tissue regeneration [132]. An inability of fast vascularization inevitably results in cell death and in the worst case loss of the tissue [133].

In order to solve this problem, different approaches for improved vascularization are conceivable: One way is administration of growth factors like bFGF, which can accelerate neoangiogenesis in the early stages of healing [134]. Another possibility is a coculture with endothelial cells [135]. In addition, integration of vascular networks into the bioengineered scaffold by microfluidic methods or bioprinting is expected to provide solutions in the near future [128, 136–138]. Maybe the combination of several approaches will eventually solve the current vascularization deficit of the designed tissues.

4.3. Innervation of Regenerated Muscles

A critical step for regenerating functional muscle tissue after VML injuries is achieving de novo innervation of regenerated myofibers (e.g., reestablishment of neuromuscular junctions, NMJs) otherwise, the regenerated muscle will become atrophic [139]. In all cases of autologous muscle transplantation, the force developed following direct or nerve stimulation is weaker than normal [140]. This is partially due to increased connective tissue and the failure of regeneration of some muscles. Another critical factor is the poor reinnervation at the sites of the original NMJs, which influences the force output [24]. It is unclear to what extent the innervation of the regenerated muscles can be restored. To rebuild the NHJs in newly regenerated muscle fibers, nerves need to be regenerated and new motor endplates have to be formed. The motor endplates not only confer functional control over the newly regenerated muscles, but also influence muscle fiber type, alignment, and size [141]. So far studies on the reinnervation of skeletal muscles have been limited to in vitro coculture of muscle cells and neurons [142, 143]. Those results showed better contractile force in nerve-muscle constructs and then in muscle-only constructs. However, full reestablishment of new nerves and motor endplates within new muscles has proven difficult, which needs to be further investigated.

4.4. Immune System Problems with Scaffolds and Cells

Matrix derived from both allografts and xenografts is often rejected because of host immune responses arising from antigens present in the donor tissue (e.g., Gal epitope, DNA, and damage associated molecular pattern molecules) [127, 144, 145]. They are typically processed by decellularization and/or chemical crosslinking to remove or cover antigenic molecules [146]. Specific decellularization techniques seem to alleviate some of these problems for ECM [147, 148]. However, remnant DNA within biological scaffolds after decellularization can still induce inflammatory reactions following implantation [149]. The host immune response to biological scaffolds differs among the sources of the raw materials from which the ECM is harvested, the processing steps, to the intended clinical application [127]. The cellular response to porcine SIS crosslinked with carbodiimide was shown to be predominated by a neutrophilic-type response, whereas foreign-body response associated with multinucleate giant cells was observed at the surgical site implanted with human dermis and porcine dermis. The host tissue response to porcine SIS showed organized connective tissue formation and muscle cells proliferation whereas the tissue response to human dermis was predominated by a persistent low-grade chronic inflammation with fibrous connective tissue formation, which might form an adverse environment for muscle tissue regeneration [150]. Therefore, the host immune reaction to biomaterials is a challenge that needs to be overcome by either designing materials that do not elicit such effects or modulating the adverse immune response.

Also for polymeric biomaterials, immunological compatibility remains a problem and limited biocompatibility sometimes causes local morbidity and chronic inflammation [108]. One reason could be that polymeric biomaterials attract multinucleated giant cells for disintegration [151].

Whether immune activation results in tissue regeneration or scarring is determined also by the availability of a stem or progenitor cell pool [152]. The cell source seems to be important with less immunogenicity in embryonic and adult stem cells [153]. Consequently, cells isolated from cord blood and autologous stem cells would be preferred for clinical application in such materials. Induced pluripotent stem cells (iPSCs) have a wide possible range of application as their production is relatively straight forward and they can differentiate in nearly every cell type. They might be able to overcome immunogenicity and ethical concerns. However, safety concerns for the use of iPSCs in patients currently result in very high regulatory barriers that will inhibit clinical translation for the foreseeable future [154]. The interactions between immune cells and resident cells are important in skeletal muscle regeneration. Macrophages, eosinophils, and regulatory T cells have been shown to activate satellite cells, which contribute to myofibers formation after injury [155–157]. In depth understanding of the immune reactions to both biological scaffolds and transplanted cells may provide clues to therapeutic avenues to promote muscle tissue regeneration. Study of the immunomodulation by scaffolds, materials, and cells in combination with subtle signaling might provide new strategies for enhancing muscle tissue regeneration through guided cell response.

5. Conclusion

Skeletal muscle injury or loss occurs in many clinical situations. Surgical techniques are highly developed and can provide good results for reconstructing muscle function, if all goes well. Surgery is always associated with considerable risks and high costs and even if successful, usually better function at one location is traded for impaired function at another location that is less important for the patient. Research into tissue engineering and regenerative cell therapy may overcome these problems. Tissue engineering solutions will have to combine biomimetic scaffolds which guide muscle tissue growth with growth factors, embedded supply routes, and relevant cells. These cells will have to directly improve local myogenic cell amount in injured or atrophic muscles, which can be expected to promote muscle regeneration. Such creative solutions will have to rely on a deep understanding of the regeneration process required for functional muscle regeneration (cell response to scaffolds, vascularization, myogenesis, and innervation), which will require further studies.

Conflicts of Interest

The authors declare that they have no conflicts of interest.


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Copyright © 2018 Juan Liu et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Systems biology of skeletal muscle: fiber type as an organizing principle

Skeletal muscle force generation and contraction are fundamental to countless aspects of human life. The complexity of skeletal muscle physiology is simplified by fiber type classification where differences are observed from neuromuscular transmission to release of intracellular Ca(2+) from the sarcoplasmic reticulum and the resulting recruitment and cycling of cross-bridges. This review uses fiber type classification as an organizing and simplifying principle to explore the complex interactions between the major proteins involved in muscle force generation and contraction.

Copyright © 2012 Wiley Periodicals, Inc.


Four motor unit types - slow (type S), fast-twitch fatigue resistant (type FR),…

Muscle force generation depends on…

Muscle force generation depends on muscle fiber length. Underlying the force-length relationship of…

During muscle fiber activation, Ca…

During muscle fiber activation, Ca 2+ released from the sarcoplasmic reticulum binds to…

Materials And Methods


Transgenic FVB mice carrying a rat mIGF-1 cDNA driven by skeletal muscle–specific regulatory elements from the rat MLC-1/3 locus have been described previously (Musarò et al., 2001). Expression of the MLC/mIGF-1 transgene in these mice is restricted to skeletal muscle and elevated in muscles enriched in fast fibers, such as gracilis anterior and posterior muscles. Transgene expression is reduced in slow muscles such as the soleus, where the MLC regulatory cassette is expressed at very low levels. Five three-month-old mice in each of the four categories were used: WT, MLC/mIGF-1, WT-EX, and exercised MLC/mIGF-1 (IGF-EX).


WT and transgenic mice were exercised to investigate skeletal muscle hypertrophy (n = 5 in each group) (Allen et al., 2001). Two male mice were exercised simultaneously in a S.A.M. Palace Kit™ cage fitted with an additional wheel. The distance and time spent exercising on the wheels was measured with a Sigma Sport BC 600 bicycle computer and a magnet glued to the periphery of each wheel. Exercised mice had free access to the wheels, food, and water. The weight of WT-EX mice transiently dropped 2–3 g several days after the beginning of exercise (unpublished data), presumably due to loss of fat before muscle hypertrophy. After 4 wk, exercised or sedentary age- and sex-matched controls were killed and muscles were processed for histology, immunohistochemistry, RNA expression, and protein analysis by Western blotting and immunohistochemistry.

AChE staining

The patterns of mouse gracilis muscle innervation were investigated as described previously (Schwarzacher, 1957). Gracilis anterior and gracilis posterior mouse muscles were dissected and stored in 1% paraformaldehyde solution. Muscles were washed three times in PBS and permeabilized by soaking in 0.2% Triton/PBS solution followed by three washes in PBS. Neuromuscular and myo–myonal junctions were stained by incubating in AChE staining solution for up to 1 h (Karnovsky and Roots, 1964), rinsed in PBS, and photographed with a Leica Wild M10 microscope. Longitudinal 8-μm cryosections of fixed gracilis anterior muscle were also stained for AChE and neuromuscular and myo–myonal junctions were photographed using a Nikon Optiphot-2 microscope.

Muscle fiber number and average area

Skeletal muscle hypertrophy was investigated in gracilis posterior muscle (no intrafascicularly terminating fibers), in gracilis anterior muscle (containing intrafascicularly terminating fibers), and soleus muscle from WT, IGF, WT-EX, and IGF-EX mice. Five muscles of each type were embedded in TBS tissue freezing medium and snap frozen in liquid nitrogen–cooled isopentane. Transverse 8-μm sections were cut through the middle of the muscle. Sections were stained with the Fast Green modification of the Van Gieson stain (Clark, 1981), resulting in green muscle fibers and black nuclei. The sections were photographed on a Nikon Optiphot-2 microscope and all muscle fibers counted in cross-section. The total area of the muscle cross-section was calculated by weighing a paper cut-out of the silhouette of the muscle and adjusting the area according to the known weight of 1 mm 2 paper. The average area of individual muscle fibers was then calculated by dividing the muscle area by the number of fibers.

The total number of muscle fibers with centralized nuclei was counted in these cross-sections of each muscle.

Fiber type analysis

Myosin heavy chain isoform changes were investigated in gracilis anterior and gracilis posterior muscles in WT, IGF, WT-EX, and IGF-EX mice. Transverse 8-μm sections were cut through the center of each muscle and blocked in 10% goat serum for 15 min. The sections were incubated with either anti-MyHC I antibody (American Type Culture Collection A4.951 1:10 dilution in PBS/1% BSA) or anti-MyHC IIB antibody (American Type Culture Collection BF-F3 1:10 dilution in PBS/1% BSA), washed three times in PBS, and incubated in secondary antibody (Jackson ImmunoResearch Laboratory peroxidase-conjugated goat anti–mouse IgG 115-035-164, or peroxidase-conjugated goat anti–mouse IgM 115-035-075, both 1:500 in PBS/1% BSA). The peroxidase was visualized with Sigma-Aldrich FAST 3,3′-diaminobenzidine tablets, D-4168. The fibers with each phenotype were counted and the number of MyHC IIA/IIX calculated by subtraction from the total number of muscle fibers.

Acid digestion of muscle fibers

The total number of fibers was counted in three WT and IGF-1 gracilis anterior muscles by acid digestion. Muscles were dissected, fixed for 1 h in 4% paraformaldehyde and stored in 80% glycerol for at least 1 wk. Muscles were then briefly rinsed in PBS and placed in 8 N HCl at 60°C for up to 1 h until individual fibers began to separate. The HCl was removed and the muscles placed in PBS. The total number of muscle fibers could then be counted by peeling off individual fibers under a dissecting microscope.

RNA preparation and Northern blot

Total RNA was obtained from thigh muscles of exercised and unexercised WT and MLC/mIgf-1 transgenic muscles by RNA-TRIZOL extraction (GIBCO-BRL). Total RNA (2 μg) was fractionated by electrophoresis on 1.3% agarose gels, transferred to GeneScreen Plus Membrane overnight, and hybridized with full-length cDNA probes, including Exon-1 (muscle specific localized form) of the rat IGF-1 gene. 25 ng of probe were labeled with P32 isotope using the Megaprime DNA labeling system RPN1604 from Amersham Pharmacia Biotech.

Protein analysis by Western blot

Protein was extracted from gracilis anterior and gracilis posterior muscles from WT, IGF, WT-EX, and IGF-EX muscles. Muscles were snap frozen directly in liquid nitrogen, macerated, and placed in RIPA buffer (25 mM Tris, pH 8.2, 50 mM NaCl, 0.5% sodium deoxycholate, 0.1% SDS, 0.1% sodium azide, 0.5% NP-40) with 5% protease inhibitor cocktail (Sigma-Aldrich P 2714). 30 μg of total protein was added to an equal volume of 2× reducing SDS sample buffer, boiled, and separated on a 9% SDS-PAGE gel. Proteins were transferred to Immobilon PVDF membrane using the Bio-Rad Laboratories semidry transfer system at 100 mA for 1 h. The membrane was blocked in MTTBS (TBS, 0.1% Triton, 5% nonfat milk powder) for 1 h, followed by incubation in the primary antibody (mouse monoclonal anti–GATA-2, Santa Cruz Biotechnology, Inc. CG2-96, dilution 1:500 in MTTBS). The membrane was washed three times in MTTBS and incubated in secondary antibody (Jackson ImmunoResearch Laboratory peroxidase-conjugated goat anti–mouse IgG 115-035-164, dilution 1:10,000 in MTTBS). The HRP-conjugated protein was detected using chemiluminescence according to the manufacturer's protocol (Renaissance NEN Life Science Products) and visualized on Kodak Biomax ML film.

Protein analysis

Expression patterns of GATA-2, neonatal myosin, and proliferating cell nuclear antigen (PCNA) were investigated on longitudinal 8-μm frozen sections of WT, IGF, WT-EX, and IGF-EX gracilis anterior and posterior muscles with immunohistochemistry. The procedure was identical to that described earlier. Primary antibodies used were anti-GATA-2 (Santa Cruz Biotechnology, Inc. CG2-96, dilution 1:100) in PBS/BSA (PBS with 1% bovine serum albumin), antineonatal myosin (gift from Schiaffino, dilution 1:100), and PCNA (Santa Cruz Biotechnology, Inc. PC10, dilution 1:100). The antibodies were detected with anti–mouse IgG-Cy3, Amersham Pharmacia Biotech PA43002 used at 1:1,000 dilution resulting in red fluorescence. Sections were costained with Hoechst 33258 dye (blue fluorescence) and the acetylthiocholinesterase method described earlier. For clearer presentation, the blue filter was processed into a green image using Adobe Photoshop ® 5.

Skeletal muscle fiber, nerve, and blood vessel breakdown in space-flown rats

To whom correspondence should be addressed, at: Department of Anatomy & Cellular Biology, Medical College of Wisconsin, 8701 Watertown Plank Road, Milwaukee, WI 53226, USA.Search for more papers by this author

Institute of Biomedical Problems, USSR Ministry of Health, Moscow, USSR

Department of Chemistry, San Jose State University, San Jose, California, 95192 USA

Department of Anatomy and Cellular Biology, Medical College of Wisconsin, Milwaukee, Wisconsin, 53226 USA

Department of Anatomy and Cellular Biology, Medical College of Wisconsin, Milwaukee, Wisconsin, 53226 USA

Department of Anatomy and Cellular Biology, Medical College of Wisconsin, Milwaukee, Wisconsin, 53226 USA

Department of Anatomy and Cellular Biology, Medical College of Wisconsin, Milwaukee, Wisconsin, 53226 USA

To whom correspondence should be addressed, at: Department of Anatomy & Cellular Biology, Medical College of Wisconsin, 8701 Watertown Plank Road, Milwaukee, WI 53226, USA.Search for more papers by this author

Institute of Biomedical Problems, USSR Ministry of Health, Moscow, USSR

Department of Chemistry, San Jose State University, San Jose, California, 95192 USA

Department of Anatomy and Cellular Biology, Medical College of Wisconsin, Milwaukee, Wisconsin, 53226 USA

Department of Anatomy and Cellular Biology, Medical College of Wisconsin, Milwaukee, Wisconsin, 53226 USA

Department of Anatomy and Cellular Biology, Medical College of Wisconsin, Milwaukee, Wisconsin, 53226 USA


Histochemical and ultrastructural analyses were performed postflight on hind limb skeletal muscles of rats orbited for 12.5 days aboard the unmanned Cosmos 1887 biosatellite and returned to Earth 2 days before sacrifice. The antigravity adductor longus (AL), soleus, and plantaris muscles atrophied more than the non-weight-bearing extensor digitorum longus, and slow muscle fibers were more atrophic than fast fibers. Muscle fiber segmental necrosis occurred selectively in the AL and soleus muscles primarily, macrophages and neutrophils infiltrated and phagocytosed cellular debris. Granule-rich mast cells were diminished in flight AL muscles compared with controls, indicating that mast cell secretion contributed to interstitial tissue edema. Increased ubiquitination of disrupted myofibrils implicated ubiquitin in myofilament degradation. Mitochondrial content and succinic dehydrogenase activity were normal, except for subsarcolemmal decreases. Myofibrillar ATPase activity of flight AL muscle fibers shifted toward the fast type. Absence of capillaries and extravasation of red blood cells indicated failed microcirculation. Muscle fiber regeneration from activated satellite cells was detected. About 17% of the flight AL end plates exhibited total or partial denervation. Thus, skeletal muscle weakness associated with spaceflight can result from muscle fiber atrophy and segmental necrosis, partial motor denervation, and disruption of the microcirculation.— R iley , D. A. I lyina -K akueva , E. I. E llis , S. B ain , J. L. W. S locum , G. R. and S edlak , F. R. Skeletal muscle fiber, nerve, and blood vessel breakdown in space-flown rats. FASEB J 4: 84-91 1990.

Skeletal Muscle Fibers

/>One focus of our lab is understanding the mechanisms behind how skeletal muscle cells differentiate into a specific muscle fiber type, either fast twitch or slow twitch. We are using zebrafish and mouse models to study the functions of new factors, such as Pbx homeodomain transcription factors, and how they control muscle fiber-type development.

Certain muscular dystrophies preferentially affect either fast or slow twitch muscle fibers. In human and mouse models of Duchenne Muscular Dystrophy (DMD), fast muscle fibers are more susceptible to damage than slow fibers. It is not clear how fiber-type identity confers susceptibility or resistance to muscular dystrophy. A goal of this project is to test a fundamental hypothesis: that factors that promote slow muscle differentiation will ameliorate the effects of DMD. Recent studies, including work from our lab, now create an opportunity to directly test whether fiber-type modulation is a viable therapeutic approach for muscular dystrophies. We take advantage of zebrafish models to address whether factors that regulate fiber-type differentiation enhance or suppress the zebrafish dmd muscle degeneration phenotype. Our approach is to manipulate fiber-type regulators that function early in development in dmd zebrafish embryos. We are also working to identify new epigenetic factors that regulate muscle fiber type. We are screening epigenetic chemicals for their abilities to enhance or suppress the zebrafish dmd phenotype. We expect that this project will identify genetic and epigenetic regulators of muscle fiber-type identities that confer susceptibility or resistance to muscular dystrophy. This project is aimed at an ultimate goal of manipulating skeletal muscle fiber type as a treatment for muscular dystrophy.


Fetal Data

Both M. vastus lateralis and M. vastus medialis weights showed a highly significant increase with increasing gestational age (P ≤ 0.001) and both muscles were significantly larger in the DM animals relative to NM (54% for VL, 30% for VM P ≤ 0.001 Table 1).

Days Vastus lateralis Vastus medialis
NM DM NM DM Sex ratio (M:F)
wt (g) SEM wt (g) SEM wt (g) SEM wt (g) SEM NM DM
120 2.7 0.1 3.48a a P ≤ 0.001.
0.1 0.9 0.1 1.3b b P ≤ 0.01.
0.1 1:4 2:3
160 14.5 0.5 19.8a a P ≤ 0.001.
0.6 4.8 0.4 5.9c c P ≤ 0.05.
0.4 2:3 3:2
210 47.7 1.5 66.4a a P ≤ 0.001.
2.6 17.1 1.2 23.2d d Not significant.
2.2 3:2 1:2
260 123.6 4.1 177.9a a P ≤ 0.001.
7.7 33.3 2.6 61.2a a P ≤ 0.001.
6.2 3:2 1:2
  • * Values are least-squares means, adjusted within groups for body weight and sex ratio, ± SEM (n = 3–5 per group). Data are back-transformed after log transformation for analysis.
  • a P ≤ 0.001.
  • b P ≤ 0.01.
  • c P ≤ 0.05.
  • d Not significant.

Fiber Typing and Morphological Analysis

Muscle fiber type.

Qualitative analysis of sections stained using mATPase histochemistry showed similar fiber morphology between NM and DM at all gestational ages. In both breeds, however, there was a noticeable difference between 120 and 160 days gestation in the morphology of the presumptive primary myotubes, from being large and vacuolated to more closely resembling mature myofibers. The smaller size of the type 1 fibers in the DM muscles was readily seen at 260-day gestation (Fig. 1H).

Myosin ATPase staining in M. vastus lateralis at four gestational ages: 120 days (A and B), 160 days (C and D), 210 days (E and F), and 260 days (G and H) of gestation. NM in panels on left (A, C, E, and G) DM in panels on right (B, D, F, and H). A and B stained after preincubation at pH 4.2 C-G stained after preincubation at pH 5.0 according to modified histochemical staining procedure. Dotted lines indicate primary myotubes open arrows indicate type 1 fibers. Scale bar = 50 μm.

The percentage of type 1 fibers in both DM and NM showed a biphasic pattern of change throughout development (Fig. 2), with numbers decreasing between 120 and 160 days and then increasing again between 210 and 260 days (P ≤ 0.001). There were consistently fewer type 1 muscle fibers in DM than in NM (P ≤ 0.001 Fig. 2).

Average percentages of type 1 muscle fibers in M. vastus lateralis from NM (filled circle) and DM (open circle) fetuses at four gestational ages (n = 3 per group). Values are mean ± pooled SEM. Triple asterisk, P ≤ 0.001 asterisk, P ≤ 0.05.


This study has shown a similar pattern of MHC expression in both DM and NM at 120-day gestation, with both presumptive primary and presumptive secondary fibers staining positively for embryonic MHC (Fig. 3). All primary fibers were also positive for slow MHC and all secondary fibers were also positive for fast MHC. At 160-day gestation, the pattern was similar, but there were a number of fibers in NM that were negative for embryonic MHC (Fig. 3G). At 210 days, a number of presumptive secondary fibers in the DM were negative for embryonic MHC and others were positive for slow MHC (Fig. 4). Some presumptive secondary fibers in NM and DM were positive for all MHC isoforms (Fig. 4A–F). At 260 days, all fibers were negative for embryonic MHC in NM (Fig. 4G), while immunostaining remained quite strong in smaller presumptive secondary fibers in DM (Fig. 4J).

Photomicrographs showing MHC immunohistochemistry in M. vastus lateralis at 120 (AF) and 160 days (GL) of gestation. NM in rows A–C and G–I, and DM in rows D–F and J–L. Embryonic MHC is shown in left column, slow MHC in central column, and fast MHC in right column. Dotted lines indicate type 1 fibers. Scale bar = 50 μm.

Photomicrographs showing MHC immunohistochemistry in M. vastus lateralis at 210 (AF) and 260 days (GL) of gestation. NM in rows A–C and G–I, and DM in rows D–F and J–L. Embryonic MHC is shown in left column, slow MHC in central column, and fast MHC in right column. Solid arrows indicate type 1 fibers, open arrows indicate fibers positive for all MHC isoforms. Scale bar = 50 μm.

Muscle fiber size.

The average area of type 1 fibers in both DM and NM decreased from 120 to 160 days of gestation (Fig. 5a), as their morphology changed from that of primary myotubes with a central region devoid of myofibrils to more mature muscle fibers surrounded by developing secondary fibers. By 210 days, type 1 fibers of NM muscles markedly increased in size as they continued to mature, while DM fibers had only very small increases in size, failing to regain the size they had previously been at 120 days. By main-effect analysis, type 1 fibers were significantly smaller overall in DM than in NM (P ≤ 0.001) due to the reduced size at 210 and 260 days (Fig. 5a). The average area of type 2 muscle fibers increased with age (P ≤ 0.001) and was less in DM than in NM (P ≤ 0.05). This effect was consistent across age groups from 160 to 260 days (Fig. 5b).

Average cross-sectional area of (a) type 1 and (b) type 2 muscle fibers in M. vastus lateralis NM (filled circle) and DM (open circle) fetuses at four gestational ages (n = 3 per group). Values are mean ± pooled SEM. A t-test was used for the calculation of significant differences within time periods. Triple asterisk, P ≤ 0.001 asterisk, P ≤ 0.05.

One of the most sensitive measures of overall fiber type composition is total % area of a specific fiber type, which is the product of average fiber area and average numerical percentage of each fiber type. The total % area of type 1 fibers declined between 120 and 160 days of gestation in both NM and DM, then remained relatively constant throughout the remainder of the period studied. The net effect of the changes in size and proportions of type 1 fibers in the DM animals was that the muscle overall had a significantly lower proportion of total area given over to type 1 fibers at all gestational ages (P ≤ 0.001 Fig. 6).

Total percentage of area of type 1 fibers calculated from average fiber number per fascicle multiplied by average fiber area in M. vastus lateralis from NM (filled circle) and DM (open circle) fetuses at four gestational ages (n = 3 per group). Values are least-squares mean ± SEM. Data was log-transformed for analysis. Triple asterisk, P ≤ 0.001 double asterisk, P ≤ 0.01.

Skeletal Muscle Anatomy

There are 3 types of muscle tissue in the human body: skeletal, smooth, and cardiac muscle. In this video and article, I’m going to discuss skeletal muscle tissue.

Skeletal Muscle Tissue

Skeletal muscles most commonly attach to bones, and they help you move your body. Unlike the other two types of muscle tissue, skeletal muscles contract on a voluntary basis via the somatic nervous system, allowing you to move your body at will.

Skeletal muscles also serve important functions, such as supporting your posture, protecting delicate organs, and they even produce heat during contraction, which helps the body maintain a proper temperature.

Skeletal Muscle Structure

Each skeletal muscle is considered an organ, and it’s made up of connective tissue layers, muscle fibers, blood vessels, and nerves. Skeletal muscles attach to the bones through tendons or through a direct attachment.

As you look at this muscle diagram, you’ll notice an outer layer of connective tissue called epimysium. The prefix “epi” means upon or over (epidermis is the layer upon your skin), and “mysium” comes from a Greek word that means “muscle.” Therefore, the epimysium is a layer of connective tissue that is over or upon the entire muscle organ.

Next, you’ll notice that the muscle fibers are bunched together into something called fascicles, which means “bundles.” These fascicles are surrounded by connective tissue called perimysium. “Peri” means “around,” and again, “mysium” refers to muscle. So the perimysium is around the fascicles that bundle up these muscle fibers.

Inside the fascicles, another connective tissue layer called the endomysium surrounds individual muscle cells. “Endo” means “within,” so that will help you remember that it surrounds the individual muscle fibers within the fascicle.

Muscle Fibers

Now let’s take a look at the individual muscle cells, which are called muscle fibers. These fibers are long and cylindrical, and they contain several nuclei. These muscle fibers are wrapped in a cell membrane called sarcolemma.

Inside each muscle fiber, there are tiny rods called myofibrils, which are surrounded by sarcoplasm. These myofibrils, also called fibrils, consist of repeating segments called sarcomeres, which are the tiny units responsible for skeletal muscle contraction.

Sarcomere Structure

As we take a closer look at the structure of a sarcomere, you’ll notice these zigzag sections that mark the end point of each sarcomere. These are called Z discs or Z lines, and they allow for the attachment of the thin (actin) filaments, as well as an elastic protein called titin.

Each sarcomere contains thin (actin) filaments and thick (myosin) filaments. The thin (actin) filaments, represented below in blue, anchor to the Z disc.

These thick (myosin) filaments, represented below in red, attach to an elastic, springy protein called titin, which then attaches to the Z disc. The actin and myosin filaments engage during muscle contraction, which I’ll discuss in a moment. The “M lines” or “M bands” anchor the center of the myosin filaments, holding them together while also acting as a shock absorber.

Sarcomere Bands & Zones

To help us understand the parts of the sarcomere, anatomists divide the sections into bands or zones. The arrangement of filaments within these bands accounts for the striated (striped) appearance of the skeletal muscle fibers. That’s an important characteristic about skeletal and cardiac muscle that you’ll want to remember: they both contain striations.

  • A band: First, there is an “A band” on each sarcomere, which is a section that contains the entire length of a thick myosin filament, along with overlapping portions of the thin actin filaments. This section makes up the dark part of the striation pattern.
  • I band: The “I band” is the section of the sarcomere that surrounds the Z disc and contains only thin (actin) filaments. This section makes up the lighter band in the striation pattern.
  • H zone: The H zone is the section within the A band that consists of the thick myosin filaments and its embedded M lines. There are no thin filaments in this section of the sarcomere when it is relaxed.
  • Z disc: And again, the Z disc is the zig zag portion that marks the end of each sarcomere and allows for the attachment of actin filaments and titin.

Skeletal Muscle Contraction

During muscle contraction, thick (myosin) filaments located within the sarcomere bend, and the knobby head part attaches to the thin actin filaments, sliding them toward the midline of the sarcomere. This sliding of filaments causes the sarcomere to shorten, or contract. As this takes place along all the sarcomeres within the myofibrils, the entire muscle fiber contracts, which ultimately causes the entire muscle organ to shorten or contract.

This is known as the sliding filament theory of muscle contraction.

Free Quiz and More Anatomy Videos

Take a free skeletal muscle anatomy quiz to test your knowledge, or review our skeletal muscle video. In addition, you might want to watch our anatomy and physiology lectures on YouTube, or check our anatomy and physiology notes.