Bioinks and bioprinting strategies for skeletal muscle tissue engineering

Abstract

Skeletal muscles play important roles in critical body functions and their injury or diseases can lead to limitation of mobility and loss of independence. Current treatment modalities result in variable functional recovery, while reconstructive surgery, as the gold standard approach, is limited due to donor shortage, donor site morbidity and limited functional recovery. Skeletal muscle tissue engineering (SMTE) has generated enthusiasm as an alternative solution for treatment of diseased or lost tissue and serve as functional disease models. In recent years, bioprinting has emerged as a promising tool for recapitulating the complex and highly organized architecture of skeletal muscles at clinically relevant sizes. Here, we discuss skeletal muscle physiology, muscle regeneration following injury, current treatments following muscle loss, and then critically review bioprinting strategies implemented for SMTE. Subsequently, we discuss recent advancements that have led to improvement of bioprinting strategies to construct large muscle structures, boost myogenesis in vitro and in vivo, and enhance tissue integration. Bioinks for muscle bioprinting, as an essential part of any bioprinting strategy, are discussed and their benefits, limitations, and areas to be improved are highlighted. Finally, the directions that field should expand to make bioprinting strategies more translational and overcome the clinical unmet needs are discussed.

1. Introduction

Skeletal muscles make up about 45 percent of the mass of the human body ^[1]. These tissues generate contractile force to direct the dynamic movement of the body parts. In addition, they act as an external barrier to support internal organs while they also enable vital functions, such as chewing and maintaining homeostasis. Skeletal muscles have high regenerative capacity following small injuries, while severe muscle defects can overwhelm their innate regenerative ability and cause chronic functional deficits ^[2]. These injuries not only result in significant burdens on the patients, but also cost healthcare system and the governments billions of dollars annually ^{[2b, 3]}. The outcome of current surgical interventions is limited by inadequate innervation, muscle regeneration, and functional recovery ^[4]. Skeletal muscle tissue engineering (SMTE) and regenerative medicine are being explored as a powerful alternative solution to existing clinical strategies ^[5]. Traditional tissue engineering approaches such as cell delivery or implantation of scaffolds fabricated through molding, electrospinning etc. fail to recapitulate the complex structure of skeletal muscle in clinically relevant size scales. 3D bioprinting, however, enables maintaining a precise control over the deposition of cells and biomaterials ^[6]. SMTE through 3D bioprinting also offers advances in disease and drug modeling, food production, and biorobotics ^[6–7].

In the following sections, the physiology of skeletal muscles and common myopathies will first be discussed. Then, fundamentals of SMTE and traditional strategies will be highlighted. Following that, recent advances in bioprinting methods and incorporated bioinks for SMTE will be critically reviewed, the limitations will be listed, and potential developments to overcome those limitations will be discussed. Furthermore, recent developments in application of 3D bioprinting for in vivo muscle regeneration are highlighted. The application of 3D bioprinting for integration of vascular and neural systems, as well as interfaced tendon tissues, as indivisible components of musculoskeletal systems is discussed later. At the end, the potential directions that field should move for successful translation of research outputs to clinical application are provided.

2. Skeletal Muscle Physiology, Injuries, and Therapies

2.1. Skeletal muscle anatomy and physiology

Skeletal muscles are a major component of the musculoskeletal system, and they connect to the bones through tendons, which are collagenous connective tissues ^[8]. To effectively generate and transmit contractile force, skeletal muscles have a hierarchically striated structure (Figure 1A) ^[9]. Muscles are covered in an epimysium, a smooth extracellular connective tissue that protects muscles and enables their gliding motion with respect to other tissues. The epimysium, along with perimysium and endomysium compose the muscle connective tissue, which is mainly made of extracellular matrix and fibroblasts ^[10]. Each muscle is constructed from highly packed fascicles which are protected by the perimysium. The fascicles are composed of individual multinucleated muscle cells called muscle fibers that are surrounded by endomysium. Each muscle fiber is composed of packed fibrous structures referred to as myofibrils. Myofibrils are long parallel protein fibers that run through the length of muscle fibers. A membrane-bound structure called sarcoplasmic reticulum surrounds each myofibril and acts as a calcium ion bank for muscle contraction ^[11]. Finally, each myofibril has lengthwise units of intercalated myosin and actin filaments called sarcomeres, the functional contractile unit of skeletal muscle. Each muscle fiber possesses hundreds of thousands of sarcomeres in each cell which are bound together at the z-discs, also called z-lines. The z-discs are the location where neighboring actin filaments of adjoining sarcomeres are connected through α-actinin crosslinking. The main components of sarcomeres are the thick filaments, composed of bundled myosin molecules, and thin filaments, composed of wound tropomyosin and actin. The sliding of the thick over thin filaments powered by the myosin heads results in the macroscopic contraction of the muscle fiber.

Figure 1.

Physiology of skeletal muscle, its regeneration mechanism following injury and main current treatment approaches. (A) Anatomy of hierarchical tissue architecture of skeletal muscle. Muscles are made of organized multiscale fibrous structures namely fascicles, muscle fibers, myofibrils, and intrinsic myosin and actin filaments, respectively from millimeter to nanometer scales, packed with extracellular matrix, capillaries and nerves. (B) The natural regenerative mechanism of skeletal muscle following an injury through a close interplay between myogenic immune cells. Upon injury, immune cells infiltrate to the defect cite, stablish a pro-inflammatory environment, helping the activation and proliferation of satellite cells (SCs), and clean the defect cite from cellular debris. SCs which were initially reside in sarcolemma in a quiescence state become active (myoblasts) and proliferate to provide cellular support for repairing the muscle, followed by their differentiation to myocytes, capable of fusing each other and forming multinucleated cells. Finally, multinucleated cells form muscle fibers, fuse the remnant tissue and mature to functional fibers. Immune cells also contribute to the early differentiation and maturation of muscle fibers by direct generation of myogenic factors, or indirectly through regulating the behavior of other cells, including fibroblasts, for deposition of ECM and secretion of myogenic factors.

Muscles have a high metabolic rate and as a result are highly vascularized to facilitate the transport of oxygen, nutrients, and waste. The intracellular space between the cell membrane of the muscle fiber, the sarcolemma, and the endomysium is filled with extracellular nutrients to support muscle fiber function. Satellite cells (SCs), the multipotent precursors to skeletal muscle cells, which are the main contributors in muscle regeneration are located under the basal lamina surrounding myofibers ^[12]. Each fiber is also innervated by a somatic motor neuron, which can generate action potentials that enables synchronized muscle contraction ^[1b]. The terminal of the motor axon forms a synapse with the muscle fiber at the neuromuscular junction (NMJ). Muscle fiber contraction is initiated by a motor nerve impulse ^[1b]. The nerve impulse reaches the NMJ and causes vesicles containing acetylcholine (ACh) to be released from the axon terminal, and bind to ACh receptors (AChRs) on the myofiber membrane generating an action potential. The action potential causes the release of a large number of calcium ions into the myofibers ^[13]. The calcium ions interact with the shielding proteins on the actin and myosin filaments causing them to interact and pull past each other resulting in contraction ^{[1b, 6]}.

The above-mentioned brief explanation of muscle anatomy and physiology demonstrated the importance of hierarchical fibrous muscle structure and its integration with other tissues, including vascular and neural system, in proper tissue function. This structure needs to be maintained and recapitulated following an injury through the natural repair of skeletal muscle or its directed regeneration with various treatment approaches.

2.2. Natural regeneration mechanism of the skeletal muscle

Adult skeletal muscles can perform efficient regeneration thanks to a population of stem cells called satellite cells ^[14]. During development, axial and limb skeletal muscles develop from muscle progenitors originating from the dermomyotome, the dorsal compartment of somites, which are periodic epithelial structures generated from paraxial mesoderm after gastrulation ^[15]. Alternatively, muscles from the head derive from the pre-chordal and cranial mesoderm ^[16]. Paraxial muscles develop through two main consecutive phases known as embryonic (or primary), and fetal (or secondary) myogenesis ^{[15, 17]}. During this process, mesoderm precursor cells commit to the myogenic lineage, becoming proliferating myoblast, which differentiate into postmitotic myocytes that later fuse to form multinucleated muscle fibers ^[18]. In late stages of fetal myogenesis, a subset of muscle progenitors expressing paired box protein Pax7 is set aside to become SCs ensuring postnatal growth and regeneration of the skeletal muscle ^[19]. Moreover, during this stage, muscle fibers grow first by fusion of progenitors and then by hypertrophy and become innervated by motoneurons. Later stages of muscle fiber maturation involve further hypertrophy, a switch from embryonic to adult myosin heavy chain (MHC) isoforms, and establishment of the slow or fast fiber phenotypes ^[15].

In adulthood, skeletal muscle has low turnover and SCs remain mostly quiescent, but upon injury, SCs can become activated and contribute to regenerate the muscle ^[20]. Minor injuries such as local plasma membrane damage caused by muscle contractions, can be repaired without the contribution of the SCs ^[12]. However, in the event of major injury, skeletal muscle undergoes regeneration through an orchestrated cascade of physiological events regulated by SCs ^[21] (Figure 1B). The SCs go through a process reminiscent of myogenesis to regenerate the tissue ^[21c]. However, SCs alone are not able to regenerate the injured tissue and a close interplay between myogenic cells and immune system is required for successful healing of muscle injuries ^[22].

Initially, the damaged muscle fibers undergo necrosis through restriction of blood flow and disruption of the muscle fibers membrane, concomitant with increased calcium-dependent proteolysis and inflammatory responses that drives their degeneration ^{[12, 21a, 23]}. Similar to other injuries, the immune cells are rapidly recruited at the site of damage ^[24]. Neutrophils are the first to be recruited within couple of hours to help degrade the cellular debris produced by releasing proteases and prepare inflammatory conditions for subsequent immune cells. Following neutrophil invasion, macrophages are recruited and enter the injury site enriched with pro-inflammatory factors including interferon‑γ (IFNγ) and tumor necrosis factor (TNF), which induce the activation of macrophages to a pro-inflammatory state called M1. The number of M1 macrophage increases around 24–48h post-injury and they contribute to phagocyte the cellular debris and secrete further pro-inflammatory cytokines. The pro-inflammatory cytokines released by neutrophils and M1 macrophages including IFNγ and TNF not only affect the inflammation state, but also contribute to myogenesis through regulation of SCs activation, proliferation and early differentiation ^{[12, 21c, 25]} (Figure 1B). It has been reported that the increase in IFNγ expression at the injury site coincides with an elevated number of activated satellite cells (myoblasts; PAX7⁺/MYF5⁺/MYOD⁺) ^[26]. Furthermore, IFNγ induces proliferation of myoblasts by inhibiting their differentiation ^[27]. TNF, on the other hand, suppresses the proliferation of myoblasts and promotes their early differentiation to postmitotic myocytes expressing myogenin ^[28]. Interestingly, immune cells, particularly M1 macrophages, release a significant amount of insulin-like growth factor 1 (IGF1), which is known to affect both myogenesis and inflammation state ^[29]. Although skeletal muscle is a rich source of IGF1, it has been argued that autologous IGF1 may not be sufficient for regeneration of severe muscle injuries. Therefore, macrophages release IGF1 to promote myoblast proliferation, expand and support the tissue repair. M1 macrophage-derived IGF1 further induce the phenotype transformation of macrophages from a pro-inflammatory state to an anti-inflammatory type called M2, which is crucial for late stages of regeneration.

The differentiated myocytes then fuse to each other to form muscle fibers and continue to differentiate ^[21a]. Myocytes further undergo secondary fusion as well as fusion into existing fibers to increase their volume and contractility. Immune cells also play key roles in this myogenesis stage. Regulatory T cells secrete the amphiregulin protein, which contributes to expression of myogenic regulatory factors for later differentiation of muscle fibers ^[21c]. Within this stage, revascularization and reinnervation begins ^{[21a, 30]}. The final stage of muscle regeneration is muscle remodeling ^[21a]. The myofibers continue to mature and the extracellular matrix (ECM) is remodeled to match the remnant tissue ^[21a]. TGFβ released by M2 macrophages in this stage can promote proliferation of fibro-adipogenic progenitors (FAPs), which reside in the tissue and differentiate them toward fibroblasts ^[21c]. The fibroblasts synthesize ECM and release additional IGF required for maturation of muscle fibers ^[21c].

2.3. Major skeletal muscle defects

Skeletal muscle defects can result from diseases such as muscular dystrophies, or traumatic incidents caused by varied etiologies including accidents, surgical resection or reconstruction, combat injuries and extensive exercising ^[31]. While disease-induced myopathies typically cause widespread muscle dysfunction and need to be treated through more systemic approaches such as gene therapies ^[32], traumatic skeletal muscle injuries are usually restricted to a discrete area, in which the healing can be promoted through administration of local treatments. Traumatic skeletal muscle injuries have high prevalence, involving around 77% of the battlefield casualties and 35%−55% of all sport injuries ^{[2b, 33]}. Over 4.5 million reconstructive surgeries are performed each year to address traumas, tumor resection, and other major losses of muscle ^[31a]. In many cases, such traumatic injuries cause more than 20% loss of the muscle mass, termed as volumetric muscle loss (VML), which is above the natural regenerative threshold ^[34]. The depletion of this large amount of skeletal muscle tissue deprives the necessary cellular, structural, and chemical components needed to regenerate functional tissue ^{[23a, 35]}. For example, in post-VML injury, several important myogenic factors, including IGF1, that are important for maintenance and regeneration of skeletal muscle tissue are downregulated ^[23a]. The catastrophic injury creates an environment that prevents the normal apoptosis of fibroblasts with a prolonged state of inflammation that generates significant fibrous scar tissue ^[36]. The profibrotic nature of VML injuries is due to the over expression of several growth factors, including TGFβ, leading to aggressive fibroblast proliferation ^[37]. The lack of satellite cells in the wound bed, the disruption in the native skeletal muscle ECM, the inflamed biochemical environment, and the fibrous scar tissue formation impedes and prevents the generation of new functional skeletal muscle. Therefore, when VML occurs in an extremity, which is the most frequent location, there is an irreversible loss of muscle mass and function secondary to fibrosis and a loss of muscle mass ^[23a].

2.4. Current clinical treatments for VML

While there are some established approaches for treatment of small muscle injuries through helping the natural regeneration mechanism with physical therapy or a combination of resting, icing, compressing and elevating (RICE), large-scale muscle damage that happens in VML generally requires surgical intervention ^[31a]. Surgical intervention involves scar debridement, followed by muscle transplantation, transposition, or even amputation and prosthetic implantation ^{[2a, 34, 38]}. Because the muscle transposition is not always an available option and amputation is not preferred, autologous transplantation is the current standard of care, in which a muscle flap is removed from a healthy donor tissue of the same person and implanted into the muscle defect. While some muscle function can be regained through the use of autologous muscle flap transfer, they often result in donor site morbidity, variable integration with remnant tissue, and incomplete strength reconstitution ^{[2b, 39]}. Therefore, autologous grafting is not an ideal method as it results in substantial donor injury with limited functional benefit to the recipient tissue. Therefore, treatments relying on the strengthening of surrounding musculature, although the loss of function is dramatic and permanent, are warranted. Physical therapy alone or after surgical grafting is frequently applied to promote tissue regeneration and/or integration after implantation ^[34]. However, physical therapy is impossible for some VML patients due to their extreme disability, and its regenerative effects and final functional recovery are limited ^[34]. Tissue engineering therefore attracted increasing attention owing to its potential in developing donor-free strategies for partial to full recovery of lost muscle function.

2.5. Skeletal muscle tissue engineering (SMTE)

Tissue engineering and regenerative medicine have been used to open new avenues not only for resolving the above-mentioned challenges associated with current clinical strategies in skeletal muscle regeneration, but also for disease modeling, food production, and biorobotics ^[6–7]. In the context of skeletal muscle, “regenerative medicine” focuses on the engineering of biomaterials, cells, biological factors or their combinations for delivery into the damaged or diseased muscle to promote the innate healing mechanism ^[40], whereas “tissue engineering” is a broader terminology which can include the application of engineered muscle for in vitro disease modeling, meat biomanufacturing, and biorobotics. While the main focus of this review is on the SMTE for healthcare applications, readers are referred to recent reviews on the implementation of SMTE for in vitro muscle models ^[41], meat biomanufacturing ^[42], and biorobotics ^[43].

Generally, SMTE methods can be classified into two main groups: acellular and cellular approaches. Acellular approaches rely on the engineering of biomaterials and biological factors and their application for muscle regeneration, which makes them simpler, cheaper and more translatable to clinical practice due to less stringent regulatory requirements ^[44]. As a result, clinical trials on the treatment of VML have mainly been reported with acellular scaffolds ^[45]. However, cellular methods which incorporate cells in the engineered construct generally result in higher regeneration rate and function restoration compared to their acellular counterparts ^[46]. The cell source in these strategies are myoblast cell lines, muscle progenitors derived from small biopsies, or stem cells directly harvested from other tissues or prepared through induction of pluripotency ^[47]. In cellular strategies for regeneration of skeletal muscles, cells can be administrated alone, in the forms of cell suspension ^[48], cell sheets ^[49], and minced muscle tissues ^[50], or in combination with scaffolding structures and biological factors. Scaffolding materials have been utilized to provide mechanical strength, binding moieties, 3D migration potential, and ECM signaling in SMTE ^[51]. Such engineered muscle grafts can be constructed by the encapsulation of cells and bioactive factors inside the scaffolding material precursor, followed by biofabrication of the muscle construct through solidification of the biocomposite, or by cellularization of a pre-fabricated solid scaffold. Alternatively, the biocomposite can be directly injected into the muscle defect for its regeneration ^{[44, 52]}. Nevertheless, the artificial muscle grafts need to recapitulate the structural and biological characteristics of skeletal muscle for proper regeneration and functional recovery of defected muscle, as discussed before. Therefore, directionality has always been an important factor for engineering skeletal muscle grafts ^[53]. Three main strategies have been traditionally used to engineer muscle constructs with organized directionality: (i) cellularization of pre-fabricated substrates/scaffolds with intrinsic cues for induction of cellular alignment, (ii) micropatterning of the muscle progenitors in linear architectures using external fields during biofabrication of muscle construct, and (iii) alignment of the cells using internal or external stimuli post-biofabrication. Each approach will be discussed briefly here.

Different approaches have been developed to fabricate substrates/scaffolds with chemical or topological cues for guiding muscle cell alignment ^[54]. Electrospinning is the most widely used approach for the fabrication of scaffolds with a directional organization for muscle tissue engineering. Multiple researchers reported the development of aligned nanofibrous scaffolds, followed by their cellularization with muscle progenitors for SMTE ^[53]. Notably, it has been shown that a fibrin/alginate electrospun scaffold with partial nanofiber alignment seeded with C2C12 myoblasts could induce cell alignment and differentiation in vitro, and muscle regeneration as well as functional recovery in a VML model ^[46a]. However, micro-patterned substrates and electrospun scaffolds are usually suitable for the generation of small, 2D constructs and limited for fabrication of realistic large, 3D muscle grafts with irregular morphology for clinical applications. Therefore, researchers endeavored to incorporate morphological cues in 3D scaffolds for SMTE ^[53].

One method to form aligned architecture in 3D scaffolds for SMTE is directional freeze-drying of hydrogels ^[55]. Kroehne et al. ^[55a] applied this method to form collagen-based grafts with aligned differentiated C2C12 cells and recovered muscle function in a mouse VML model post implantation of the graft. Another method is to use unidirectionally aligned removable/sacrificial filaments in the scaffold ^[56]. Interestingly, this method has been used for in vivo formation of endogenous 3D scaffolds with aligned topographical cues; this was accomplished through the implantation of a sacrificial PCL fibrous scaffold, in vivo cellularization from the surrounding tissue, scaffold remodeling, explanation, and decellularization ^[56a]. Mechanical strain is also reported to form unidirectional fibrous architecture for SMTE in specific hydrogels such as collagen ^[57]. However, the small inherent porosity of the scaffolds fabricated using this method did not allow for easy cellularization after biofabrication. Therefore, Jin et al. encapsulated cells in the hydrogel precursor, partially crosslinked the scaffold, and exposed the solution to mechanical strain to induce aligned fibrillar architecture in the collagen and decellularized muscle ECM ^[57b]. While both of these scaffolds demonstrated a high ability to induce directionality of encapsulated muscle progenitors, the decellularized scaffold caused a higher level of myogenesis both in vitro and in a VML mouse model ^[57b]. 3D scaffolds with directionality for SMTE were also biofabricated by first generating individual fibers and then assembling these fibers into larger constructs using various textile strategies ^[58]. This strategy allows for the formation of multicellular structures and enables directionality both in cellular organization and scaffold architecture. Although the biofabrication methods mentioned above can address the requirement for 3D muscle grafts in regenerative medicine, they suffer from multiple cumbersome processing steps and are unable to create complex and multimaterial structures required for vascularized and innervated grafts that are necessary for the engineering of large skeletal muscle constructs.

Cells could also be patterned using external fields into linear structures during biofabrication to form biomimetic muscle tissues. Acoustic ^[59], magnetic ^[60], and electrical fields ^[61] have been implemented for SMTE. However, these strategies require complex facilities, while they are limited in the construction of large complex architectures required for the clinical treatment of VML. Alternatively, external fields were applied during the cell incubation time post-biofabrication using various bioreactors to align cells unidirectionally for SMTE ^[62] (see Section 3.3 for a more detailed discussion on the application of bioreactors for improvement and maturation of biofabricated skeletal muscle tissues). Mainly, static or dynamic electrical and mechanical stimuli have been applied for induction of directionality into cultured muscle progenitors, while cell-induced internal mechanical stimulation is the most widely traditional approach used for SMTE ^{[52, 62–63]}. In this method, a scaffolding material encapsulating cells is molded and constrained at different points (anchoring sites), followed by its incubation in culture conditions. In contractile muscle cells, a cytoskeleton-mediated internal tension will be induced along the line passing the anchoring sites, stimulating the cells for their alignment and myogenic differentiation ^[64]. Multiple researchers have been utilizing this method recently for fabrication and maturation of miniaturized muscles for studying muscle physiology and disease in vitro ^[65], or even their implantation into small animals ^{[64b, 66]}. However, given the complexity of forming vascularized tissues required for in vitro culture of thick (>1mm) cellular constructs, such miniaturized tissues are hard to scale up for the realistic clinical application, where a large construct with irregular shape is required for the treatment of muscle defects.

While the traditional methods offer some specific advantages such as simplicity and lower costs, they suffer from setup complexity, the negative impact of the external fields on cells, limited construct size, multistep fabrication processes, insufficient vascularization, and incapability of recapitulating the complex, hierarchical, and organized structure of skeletal muscle tissues ^[6]. Alternatively, 3D bioprinting can be implemented to accurately control the spatial distribution of the biomaterials, biological factors, and encapsulated cells. Recently, the potential benefit of 3D bioprinting has been explored for SMTE. Because the main focus of this review is on the application of bioprinting for SMTE, the readers are referred to the recent reviews on other tissue engineering strategies for engineering of skeletal muscle tissues ^[67].

3D bioprinting has evolved over the past few decades, and currently, a wide variety of 3D bioprinting technologies have been introduced for tissue engineering. In this review, we will cover the 3D bioprinting approaches utilized for SMTE, discussing the advantage and shortcomings of each method, and how they can be improved to address the main requirements for SMTE. Furthermore, the bioinks implemented for the bioprinting of muscle constructs are discussed. Finally, future directions of bioprinting for SMTE are detailed.

3. 3D Bioprinting for SMTE

Bioprinting is an additive manufacturing strategy with controlled deposition of bioinks for fabrication of tissue-like constructs ^[68]. In most cases, the bioink is made of scaffolding materials carrying live cells, which may be supplemented with drugs, and biological factors. Depending on the additive process used in the fabrication of these tissue-like constructs, bioprinting strategies can be classified into extrusion bioprinting, inkjet bioprinting, stereolithography bioprinting, laser-assisted bioprinting, and non-conventional bioprinting approaches. An ideal bioprinting strategy (i) supports the viability and functionality of the encapsulated cells; (ii) enables a precise control over the spatial distribution of different scaffolding materials, biological factors, and cells over clinically and biologically relevant dimensions; (iii) offers sufficient resolution to recapitulate the architecture of the native tissue; and (iv) is compatible with the printing of diverse bioinks with different viscosities and crosslinking methods ^[68]. In bioprinting of muscle constructs, the goal is to create a construct which directs the formation of highly organized hierarchical fibrous architecture of the native tissue with sufficient vascularization and innervation. Here, we describe different bioprinting strategies implemented for SMTE and emphasize the benefits and challenges of each approach for biofabrication of muscle constructs.

3.1. Bioprinting strategies for SMTE

The selection of the bioprinting approach is usually based on the target tissue, which dictates the desired architecture, feature dimensions, bioink properties, and mechanical properties of the final scaffold required for tissue maturation and function. Generally, extrusion-based bioprinting results in a fibrous structure with the capability to fabricate large and complex constructs. Inkjet and laser-assisted bioprinting create constructs through the deposition and coalescing of small droplets to form thin constructs with high speed and cellular viability. Stereolithography-based bioprinting allows biofabrication of constructs with complex architectures mostly from a single photo-crosslinkable bioink. As a result, all bioprinting approaches are not necessarily applicable for the engineering of a specific tissue. Table 1 summarizes the main strategies specifically used in SMTE and introduces their advantages and disadvantages for the biofabrication of muscle constructs. It should be noted that some specifications mentioned in the table (such as resolution) are not only dependent on the bioprinting strategy, but also on the bioink properties, surface properties, cell type, printing parameters, and even crosslinking strategy. As a result, only comparative insights between the approaches are provided here, and readers are referred to excellent reviews for a general overview on quantitative values ^[68–69].

Table 1.

Main strategies implemented for bioprinting of skeletal muscle tissues and their promise for SMTE.

	Extrusion bioprinting	Inkjet Bioprinting	Stereolithography
Method
Advantages	• Biomimetic fibrous structure for SMTE • Simple and low-cost facilities • Compatible with complex and large structures • Compatibility with various crosslinking methods • Compatible with multimaterial and gradient printing • Decent resolution	• Simple and low-cost facilities • Compatible with multimaterial and gradient printing • High resolution • High speed	• High cell viability due to shear-free printing • High resolution • Compatible with bioinks wide range of viscosities • Compatible with complex and large structures • High structural fidelity
Disadvantages	• Limited structural fidelity with low viscosity bioinks • Increased shear stress on the cells with increasing resolution, viscosity, and cell density	• Discontinuity in the printed structure • Limited structural similarity with skeletal muscle tissue • Limited shape fidelity of the final construct • Incompatible with viscous bioinks • Low cell density	• Low throughput • Limited to photocrosslinkable bioinks • Limited capability of multimaterial and gradient printing

Extrusion bioprinting is the most widely used method implemented for engineering skeletal muscle tissues and will be discussed in more detail (Section 3.1.1). Other bioprinting strategies used for SMTE, including inkjet and stereolithography bioprinting methods, will also be explored in Section 3.1.2.

3.1.1. Extrusion-based bioprinting for SMTE

Extrusion-based bioprinting is the approach most widely implemented for SMTE. This method is associated with the controlled extrusion of a bioink through a nozzle with defined tip size using mechanical or pneumatic forces, deposition of fibrous structures, and bioink gelation to form complex scaffolds out of overlaid fibers ^[70]. The movement of the bioprinting head and the printing collector is automatically adjusted using a computer to fabricate complex structures. Therefore, the structure should be first designed using a computer-aided design (CAD) software, and then be translated to the codes readable by the machine through computer-aided manufacturing (CAM) strategies ^[71]. The bioink can be solidified upon extrusion using a wide range of gelation approaches including thermal, ionic, enzymatic, or photo-crosslinking ^{[68, 72]}. Using multiple printing heads, recent extrusion bioprinters are capable of fabricating constructs out of different bioinks which could contain various biomaterials, cells, or biological factors ^[73].

The inherent fibrous architecture of the biofabricated scaffolds using extrusion bioprinting makes this approach an ideal method for engineering muscle tissues ^[74]. Additionally, it has been reported that the shear stress induced during extrusion can direct muscle cell alignment through (i) organizing the polymeric chains and forming nanotopographical cues in the deposited fiber ^[75], (ii) mechanically stimulating the cells ^[76], and (iii) aligning high aspect ratio micro/nano features mixed with the bioink ^[77]. Multiple researchers implemented extrusion bioprinting to fabricate artificial muscles as in vitro models to elucidate the development and repair mechanisms of skeletal muscles ^{[41, 58b, 73, 75–76, 78]}, for the regeneration of muscle defects in vivo ^{[77, 79]}, and even as biological actuators for bio-robotics and microelectromechanical systems (MEMS) ^[80]. Despite all of the achievements in extrusion-based bioprinting, this strategy still suffers from two main disadvantages for engineering of muscle tissues: (i) limited shape fidelity when bioprinting of large constructs, and (ii) limited resolution to direct proper cellular alignment and differentiation. In the following sections, we will describe the achievements to resolve these challenges.

3.1.1.1. Strategies for extrusion-based bioprinting of large skeletal muscle tissues

To fabricate skeletal muscles having a clinically relevant size, the bioprinting strategy needs to offer a stable printing of 3D constructs with high shape fidelity and structural integrity. Increasing the viscosity of the bioink and the concentration of crosslinking agents may address this requirement, though it changes the scaffold stiffness and could significantly reduce cells viability, proliferation, and differentiation ^[6]. To resolve the challenge of structural integrity for bioprinting large musculoskeletal scaffolds while preserving the functionality of the encapsulated cells, researchers integrated supportive structures in the bioprinted scaffolds (Figure 2). The integration of supportive elements and bioprinted constructs is a hybrid manufacturing strategy that facilitates the formation of tissue-like structures with clinically relevant dimensions ^[81]. Three different mechanisms have been introduced for integration of supportive elements for enhanced structural stability of bioprinted muscle constructs including: (i) multimaterial printing for incorporation of supporting structures during bioprinting (Figure 2A) ^{[73, 78b, 79b]}, (ii) bioprinting on previously fabricated supportive structures (Figure 2B) ^{[78a, 78h]}, and (iii) bioprinting inside a support bath (Figure 2C) ^{[78i, 79a]}.

Figure 2.

Strategies for extrusion bioprinting of large muscle tissues with enhanced structural stability. (A) Multimaterial extrusion printing with sacrificial and support structures for SMTE. To form large muscle constructs, PCL was printed as supporting pillars, while the main bioink and sacrificial material were printed with a linear architecture entangled to the support pillars (i). After bioprinting and crosslinking of the printed structure, the sacrificial material was removed by incubation of the construct at 37 °C, leaving behind the fibrous structure (ii-iv). The printing strategy resulted in high cellular viability, alignment, and maturation in vitro (v, vi). Reproduced with permission from Nature Publishing Group ^[79b]. (B) Bioprinting on prefabricated support structures. Bioprinting on an electrohydrodynamically printed fibrous support structure followed by rolling for construction of biomimetic large-scale muscle tissues (i). C2C12-laden bioink was printed on PCL fibrous structures (ii). Electrohydrodynamic printing of PCL inside an ethanol bath formed a nanofibrous structure within the main PCL fibers, followed by stretching of the structures to generate an aligned nanofibrous architecture (iii). The biofabrication strategy demonstrated to be biocompatible (iv). Rolling of the composite sheets formed a large-scale biomimetic muscle structure (v). Reproduced with permission from American Chemical Society ^[78h]. (C) Bioprinting inside a support bath (embedded bioprinting). Cell-laden mdECM was printed inside a temporary gelled bath, followed by thermal crosslinking of the ECM-based bioink, and release of the crosslinked scaffold from the support bath (i). Inset shows an actual image of the large-scale scaffold biofabricated through embedded bioprinting. The embedded bioprinting approach supported cellular viability (ii) and myogenesis (iii), compared to the cells infiltrated into a sponge scaffold or those encapsulated in balk hydrogel through casting. Reproduced with permission from Elsevier ^[79a].

Atala and his collaborators developed a multimaterial 3D bioprinting strategy capable of printing multiple cell-laden fibers integrated with sacrificial and supporting structures for biofabrication of human-scale tissue constructs (Figure 2A–i) ^{[73, 79b]}. The presence of sacrificial and supporting constructs ensures the structural stability of the printed scaffold while allowing the application of more cell-friendly bioinks with low viscosities ^[73] (see Section 3.2). A 3D structure with aligned fibers was printed using human muscle progenitor cell (hMPC)-laden bioink, supported by with PCL pillars and sacrificial gelatin-based material (Figure 2A–ii) ^[79b]. The implemented cell containing bioink was a mixture of gelatin, fibrinogen, hyaluronic acid, and glycerol, in which the fibrinogen was crosslinked using a thrombin solution post bioprinting (Figure 2A–iii). Following the formation of a stable construct upon crosslinking, the sacrificial gelatin was washed, leaving behind the hMPC-laden fibrin fibers supported by PCL pillars (Figure 2A–iv). Despite the non-printed scaffolds, the fibrous architecture of the printed scaffolds with embedded microchannels formed upon dissociation of sacrificial material enabled high cellular viability and maturation into aligned myotubes after 7 days of differentiation in vitro (Figure 2A–v, vi). The biofabricated muscle was then implanted in a rat model for treatment of VML.

To enhance the structural stability for the bioprinting of large muscle constructs, the bioink can also be printed on prefabricated support scaffolds (Figure 2B). Yeo et al. ^{[78a, 78h]} developed an approach by combining 3D printing, electrospinning, bioprinting, and rolling for the biofabrication of large-scale biomimetic muscles. PCL mats first were 3D printed, followed by electrospinning of PCL nanofibers on top of the mats, bioprinting cell-laden alginate-based bioink on the nanofibrous structure, and finally rolling the layered sandwich sheets to form a large muscle-like construct ^[78a]. The presence of PCL nanofibers in the support scaffold could act as morphological cues to guide cellular alignment, which is an important consideration in SMTE. To fabricate hierarchical fibrous PCL support scaffolds, the authors further used an alternative approach through electrohydrodynamic PCL printing in an ethanol bath ^[78h]. This approach caused the generation of nanofibrous mats, followed by their stretching to align the internal fibers. Subsequently, a C2C12-laden bioink was printed on the fibrous mats using extrusion bioprinting, followed by rolling of the construct to form a biomimetic 3D muscle. In this hierarchical fibrous architecture, the fibrous PCL acts both as a support structure and morphological cues for cellular alignment, while a cell-laden collagen-based hydrogel, printed on the supporting structure, enables cellular adhesion and myogenic differentiation.

An alternative approach that enables stable bioprinting of large muscle constructs is extrusion bioprinting inside a temporary support bath, sometimes called freeform reversible embedding of suspended hydrogel (FRESH) or embedded bioprinting ^[82]. In this approach, a biocompatible solid-like material is utilized as a suspension bath to support the extruded filaments in their printed position, maintaining the printed structure stable until complete crosslinking. Therefore, embedded bioprinting supports the use of a broader range of bioinks and slow crosslinking chemistries. The printed structure is then released from the suspension bath through dissociation of the supporting biomaterial. Embedded bioprinting has been implemented for SMTE to address the need for regeneration of muscle tissues with clinically relevant sizes ^{[78i, 79a]}. This method was utilized for the generation of artificial muscle flaps with primary human skeletal muscle cells (hSkMCs) encapsulated in decellularized muscle-derived ECM (mdECM) (Figure 2C) ^[79a]. The support bath was prepared by mixing gelatin microgels with a 10% polyvinyl alcohol (PVA) solution. Subsequently, the extrusion bioprinting was performed within the bath. The printed structures were then placed inside a cell culture incubator at 37°C to crosslink the mdECM, melt the gelatin, and release the main bioprinted construct (Figure 2C–i). High cellular viability as well as myogenic differentiation of the muscle progenitors along the printed fibers was observed and reported to be correlated with the fibrous architecture of the released construct, with large space between the fibers, offering a rapid transport of nutrients and waste through the scaffold (Figure 2C–ii, iii).

Although the application of the supporting elements can improve the structural stability of bioprinted constructs, the incorporation of additional materials such as PCL can affect the efficacy of the scaffolds post implantation. On the other hand, the application of a support bath in embedded bioprinting increases the complexity of the fabrication, prolongs the processing time, and reduces the bioprinting resolution. The development of new bioinks with optimized printability and cell permissibility can help to better resolve this challenge (see Section 3.2 for detailed discussion).

3.1.1.2. Strategies for directing muscle cells organization with extrusion-based bioprinting

The second challenge of the extrusion bioprinting in the context of SMTE is its limited resolution for directing cellular alignment ^[58b]. It has been demonstrated that the response of encapsulated cells to morphological features reduces with increasing the size scales, with features larger than 100 μm being unrecognizable to cells ^[83]. However, the fiber diameters compatible with current extrusion bioprinting strategies are larger than this value ^[78m]. The size of deposited filaments in extrusion bioprinting is dictated by several parameters including nozzle tip diameter, physical and rheological properties of the bioink, and more importantly, the viability of encapsulated cells affected by flow-induced shear stress ^[84]. As a result, conventional extrusion bioprinting methods fail to properly direct cellular alignment required for maturation and functionality of the final muscle tissue ^[58b]. To overcome this challenge, researchers developed strategies to form intrinsic microtopographies within the printed fibers (Figure 3). An internal microstructure, with aligned microfilaments in the order of cell dimensions, can direct the cellular organization and therefore their function for engineering of muscle. This strategy could reduce the feature resolution to sub-filament sizes. Various approaches have been developed to create microfilamentous structures before (Figure 3A), during (Figure 3B), or after (Figure 3C) bioprinting.

Figure 3.

Strategies for the incorporation of internal microtopographies in extrusion bioprinting to direct cellular alignment for SMTE. (A) Development of a microstrand-based bioink for an improved cellular organization in bioprinted muscle tissues. A cell-laden pre-crosslinked hydrogel block was deconstructed into microstrands by sizing through a micro-mesh (i). The microstrand-based bioink was then extruded to form aligned internal microfilaments inside the deposited fibers by induced shear stresses (ii). The sizing process was demonstrated to support cellular viability (iii), while the filamentous nature of the construct induces cellular alignment to recapitulate the native structure of the muscle (iv). Reproduced with permission from Wiley ^[78f]. (B) Formation of internal microfilaments during bioprinting using a microfluidic strategy. A static mixer with helical Kenics™ elements was utilized to co-extrude two different bioinks and divide their streams into multiple sub-streams, forming a multicompartmental structure of intercalated striations (i). The number and size of downstream striations were dependent on the number of applied Kenics™ elements inside the nozzle (i, ii). Despite the cells encapsulated in printed fibers from homogeneously premixed bioink, the myoblasts demonstrated a highly aligned organization from the first date of culture inside multicompartmental hydrogel fibers (MCHFs) printed using the static mixer (iii, iv). Reproduced with permission from AIP publishing ^[58b]. (C) Formation of internal microfilaments after bioprinting through leaching of sacrificial materials. A bioink consisted of an mdECM-based material mixed with sacrificial PVA was used for bioprinting. Upon extrusion of the bioink, the polymeric chains were aligned with printed fiber, followed by dissociation of the PVA, which created internal topographical cues to direct cellular alignment (i). Muscle progenitor cells demonstrated a high level of alignment on day 14 post-culture in the fibers fabricated from mdECM/PVA bioink, in contrast to those cultured mdECM bioink alone (ii). Reproduced with permission from Elsevier ^[78j].

Intrinsic topographies within filaments can be formed by shear-induced alignment of a previously prepared microstrand-based bioinks during extrusion (Figure 3A). Kessel et al. ^[78f] used micro-grids to deconstruct a pre-crosslinked cell-laden bulk hydrogel into spaghetti-like microstrands with a size ranging from 40 to 100 micrometers (Figure 3A–i). They demonstrated that while microstrands had a random orientation after sizing through the micro-grids, the shear stress during the extrusion induced an aligned organization of the microstrands within the printed structures (Figure 3A–ii). The researchers reported that the sizing process did not affect the viability of the encapsulated cells (Figure 3A–iii). Additionally, microstrand-based bioink demonstrated a good printability as a result of its shear-thinning behavior. Using a gelatin-gelatin methacrylol (GelMA) hydrogel composite encapsulating C2C12 myoblasts, formation of aligned multinucleated myotubes inside the microfilaments was further reported (Figure 3A–iv). However, this strategy suffers from two main limitations. First, the scaffolding hydrogel in this approach requires a relatively high mechanical stiffness to allow deconstruction through micro-grids. Such stiff hydrogels can hinder cellular activity and differentiation. Second, the two-step preparation of the bioink with encapsulated cells in pre-crosslinked hydrogels significantly complicates the bioprinting strategy.

An alternative strategy for creating intrinsic topographies is forming internal striations within the bioink flow inside the nozzle ^{[58b, 78m, 78n, 85]}. Using an innovative approach, Madero et al. ^[78m] implemented a Kenics™ static mixer containing multiple helical elements to divide the streams of myoblast-laden and acellular alginate-based bioinks into sub-streams that were then crosslinked to form printed fibers with cellular patterning. In that system, each helical element divided the upstream flow into two sub-streams, therefore, enabling facile control of the number and size of the internal microfilaments by adjusting the number of elements. While a biomimetic muscle structure with patterned C2C12s inside the fibers could be formed, the proposed strategy failed to support cellular spreading and alignment due to the limited cell permissibility of the alginate-based bioink and lack of morphological cues in the structure. To resolve these challenges, a multicompartmental bioprinting strategy has been developed based on the co-extrusion of two different bioinks through a static mixer integrated into a coaxial microfluidic device (Figure 3B) ^[58b]. Acellular alginate and myoblast-laden GelMA bioinks were implemented in the multicompartmental bioprinting to form intercalated microfilaments of alginate and GelMA inside the fibers with different mechanical and biological properties (Figure 3B–i, ii). In the engineered microstructure, GelMA microfilaments provided a cell permissive environment, while alginate compartments offered morphological and mechanical cues for cellular alignment. Despite the cells encapsulated inside the printed fibers fabricated out of homogeneously mixed alginate and GelMA, those cultured in multicompartmental hydrogel fibers (MCHFs) demonstrated high metabolic activity, rapid cellular spreading, highly aligned cellular organization from the first date of culture (Figure 3B–iii, iv), and rapid myogenic differentiation toward formation of large fascicle-like fibers with densely packed myofibers (Figure 3B–v). However, it is preferred to keep the diameter of the filaments in sub-millimeter scales to allow proper diffusion of the nutrients across the fibers thickness.

In addition to the formation of microfilaments before or during printing, they can be created post printing by incorporation of sacrificial polymers with the main bioink ^{[78e, 78j]}. In this strategy, the sacrificial polymers will be aligned with extrusion-induced shear stress and leached out after bioprinting, leaving behind a fibrous internal structure. This strategy was implemented by Kim et al. ^[78j] through bioprinting a cell-laden mixture of methacrylated mdECM (mdECM-MA) and PVA solutions (Figure 3C). Before bioprinting, mdECM-MA and PVA polymeric chains had a random orientation, while the shear stress during the extrusion of the bioink aligned the polymeric chains in the printing direction (Figure 3C–i). Upon leaching the PVA chains, an aligned fibrous structure of mdECM-MA was formed, providing morphological cues to direct cellular alignment. This approach was demonstrated to support cellular viability and induce high cellular organization within the scaffold. The researchers demonstrated that the cells bioprinted in mdECM-MA alone did not show an aligned morphology, in contrast to those printed with mdECM-MA/PVA mixture (Figure 3C–ii). Furthermore, compared to mdECM-MA/PVA, GelMA/PVA demonstrated lower myogenesis potential, indicating that both chemical and morphological cues are required for SMTE. The formation of biomimetic fibrous structures through leaching of sacrificial material after 3D printing was also utilized to fabricate acellular scaffolds for SMTE ^[86].

3.1.2. Other bioprinting strategies for SMTE

While extrusion bioprinting has been implemented as the main bioprinting approach in SMTE, researchers explored other approaches for potential enhancement of resolution, cell viability, and bioink compatibility. The other bioprinting strategies utilized for engineering skeletal muscle tissues include inkjet and stereolithography bioprinting methods.

Inkjet bioprinting employs various droplet formation mechanisms, including thermal, piezoelectric, or electrostatic actuation, to precisely generate droplets and deposit them in computer-controlled positions for additive construction of a designed scaffold ^[87]. Inkjet bioprinting benefits from promising resolution, high throughput printing capability, and low cost tools ^[68]. However, the inherent discontinuity of the inkjet printing process reduces the compatibility of the approach with SMTE due to the limited mechanical robustness of the final construct and, more importantly, the lack of structural similarity with fibrous skeletal muscle tissues. Therefore, while few studies reported inkjet bioprinting of myoblasts-laden bioink to study the interaction of these cells with other cells in the musculoskeletal system ^[88], droplet-based bioprinting methods are mainly used for engineering of thin smooth muscle constructs ^[89].

Stereolithography bioprinting is another approach implemented for bioprinting of muscle constructs. In stereolithography, a photo-polymerization approach is applied to selectively crosslink designated regions of a photocrosslinkable bioink ^[90]. The selective light exposure can be performed by the application of photomasks, using computer-controlled lasers, or through dynamic light processing with micromirror arrays focused on a thin layer ^[91]. This approach allows high-resolution shear-free printing of cell-laden constructs, without any restriction in bioink rheological properties. Using selective UV exposure through a photomask, Aubin et al. ^[83] formed 50 μm strips of myoblast-laden GelMA and reported a high level of alignment both in the cytoskeleton and nuclei of the encapsulated cells after 5 days of culture. A similar approach was applied by Agrawal et al. ^[92] to form fascicle-like structures within a microfluidic chip. Stereolithography bioprinting by laser ^[93] and micromirror array ^[94] was also utilized for high-resolution biofabrication of myoblast-laden scaffolds. However, stereolithography bioprinting suffers from low throughput in biofabrication of large structures, specifically for regeneration of large skeletal muscle defects, as well as the requirement of the photocrosslinkable bioinks. Additionally, biocompatibility of the method needs multiple optimizations due to the potential cytotoxicity of UV exposure, photo-initiators and free radicals generated during the photocrosslinking. Moreover, the presence of a bioink bath in this bioprinting strategy makes the changing of the bioink and, therefore, multimaterial printing challenging.

In addition to the bioprinting strategies mentioned in Table 1, other additive biomanufacturing approaches such as cell electrospinning and 3D cell patterning have been developed for engineering of muscle constructs. Cell electrospinning was recently implemented for SMTE as a modified version of extrusion bioprinting ^[95]. This strategy is based on the establishment of an optimized electrical field between the printing nozzle and the collector, generally consisted of a pair of electrodes opposed to each other, to stretch a charged bioink and deposit it between the electrodes, forming aligned nanofibrous struts. The electrostatic force in this process dominates the surface tension of the liquid, significantly reduces the fiber size, and resolves the challenges of fiber resolution in normal extrusion bioprinting. However, the electric field could negatively affect the biocompatibility of the approach. Moreover, the fabrication of complex and multimaterial constructs is challenging through cell electrospinning. On the other hand, an organized cellular architecture can be formed through 3D patterning of encapsulated cells within the hydrogel precursor using an external field, followed by crosslinking of the hydrogel. For example, acoustic fields have been implemented to generate standing fields in 3D environments and pattern muscle cells in linear architecture with desired size scales for muscle tissue engineering ^[59]. A combined acoustic cell patterning with extrusion bioprinting has also been reported for SMTE through the application of acoustic nozzles ^[96]. While the application of external fields for biomanufacturing of skeletal muscle constructs is promising, such methods suffer from a lack of flexibility in the printing architecture, time-consuming preparation steps, small size scales, and affected cellular behavior ^{[58b, 97]}. Because these non-conventional biomanufacturing are not categorized as bioprinting, they are out of the scope of this review and the readers are referred to recent publication for a more detailed discussion ^[98].

3.2. Bioink formulations for SMTE

The bioink to be printed needs to be selected based on the bioprinting approach, the target tissue, and the encapsulated cells ^[99]. An ideal bioink offers good 3D printability, proper cell permissibility, and similar mechanical and structural properties of the bioprinted scaffold to that of native tissue.

The printability of a bioink is determined by its rheological properties and gelation kinetics, with respect to the selected bioprinting approach. For example, a viscous (>30 mPa.s) shear-thinning bioink with rapid crosslinking is preferred for extrusion bioprinting, inkjet bioprinting favors low viscosity bioinks (3.5–12 mPa.s) with low surface tension, and SLA bioprinting requires a photocrosslinkable bioink ^{[68, 95a, 100]}. It is worth noting that the rheological properties of the bioink can also affect the distribution of the cells inside a scaffold; this is of particular interest when printing large muscle constructs for VML treatment, because they usually take more time and increase the opportunity of cell sedimentation within the bioink before printing and crosslinking. In many cases, the bioinks have been improved with various strategies including covalent/non-covalent functionalization, nanomaterials incorporation, and thermal treatment for better printability ^[101]. After crosslinking, the bioink needs to form an environment similar to the native ECM in vivo, offering high cellular viability, cell-binding sites, biodegradability, and proper physiochemical cues to enable cell spreading and migration, scaffold remodeling, and tissue maturation through cellular differentiation ^[99]. The similarity between the mechanical and structural properties of the bioprinted scaffold to the native tissue enables efficient integration and function of the engineered tissue in vivo. Specifically, it has been demonstrated that stem cell lineage specification toward a myogenic fate and an improved quality of myogenic differentiation occurs on substrates with mechanical properties closely resembling native skeletal muscle, having a passive Young’s modulus of around 12 kPa ^[102]. While this stiffness has been used as a benchmark in some studies, most neglect the properties found in native skeletal muscle. Most commonly, the mechanical properties of acellular printed and crosslinked biomaterials are tested for compressive and tensile loading ^[78m]. The resultant stress-strain curves have been used for further material investigation such as compressive moduli, tensile moduli, and tensile strength. Less frequently, the constructs are investigated with loaded cells or after culturing ^[78g]. Almost no focus has been placed on understanding the mechanical properties of these scaffolds after experiencing remodeling. Finally, the method of implantation, such as adhesion directly to injured tissues, requires suitable adhesion characteristics that can be elucidated through ex vivo normal and shear adhesion tests to skeletal muscle ^[79d].

To meet the above-mentioned requirements, researchers have designed and optimized several bioink formulations to balance cell permissibility of the bioink with its printability and final mechanical properties of the scaffold. Synthetic and natural biopolymers, or a mixture of them, have been tested for bioprinting of various tissues ^{[99, 103]}. Natural biopolymers have been more widely used for muscle tissue engineering due to their similarities with native muscle in term of physiochemical cues required for cellular development toward functional muscle ^[64a]. Table 2 summarizes the main bioinks used for SMTE. To achieve proper cell permissibility as well as reasonable printability and mechanical stability for bioprinting of skeletal muscle constructs, researchers explored different bioinks, mostly composed of multiple biomaterials. As mentioned before, the application of support structures (Figure 2) is an approach to improve the stability in the bioprinting of low-viscosity cell-permeable bioinks. Alternatively, novel composite materials can be prepared to address printability, mechanical stability, and cell permissibility, without the need for complex supporting systems. Alginate-, gelatin-, and fibrin-based hydrogels are the main biomaterials used as the primary component of the bioinks for bioprinting of myoblast laden constructs (Table 2).

Table 2:

Bioinks implemented for bioprinting in SMTE.

Primary Bioink Material	Additive Bioink Components (Purpose)	Bioprinting method	Cells	Achievement	Ref.
Alginate	PEO (enhanced printability and scaffolds degradation rate)	Extrusion	C2C12	Improved cell alignment and differentiation in vitro	[78a]
Alginate	Pluronic (enhanced printability)	Extrusion	C2C12	Improved cell alignment in vitro	[78e]
Alginate	GelMA (enhanced cell permissibility)	Extrusion	C2C12	Formation of fibrillar hydrogel fibers	[78m]
Alginate	GelMA (enhanced cell permissibility)	Extrusion	C2C12	Formation of fibrillar hydrogel fibers. Demonstrated the expression of sarcomeric α-actinin.	[78n]
Alginate (oxidized)	Gelatin (enhanced cell permissibility)	Extrusion	C2C12	Improved cell alignment and differentiation in vitro	[75a]
Alginate (oxidized, methacrylated, and RGD functionalized)	PEGMA (enhanced SLA printability)	SLA	C2C12, HN, and ASC	Printing complex multicellular structures for investigation of cellular interaction	[93]
GelMA	Alginate (enhanced printability)	Extrusion	C2C12	Improved cell alignment in vitro	[78b]
GelMA	PEDOT:PSS (enhanced printability and conductivity)	Extrusion (embedded)	C2C12	Development of a biocompatible conductive hydrogel for SMTE	[78i]
GelMA	Gellan gum (enhanced printability)	Extrusion	C2C12	Development of a bioink with improved printability	[104]
GelMA	CMCMA, AlgMA, or PEGDA (enhanced printability and fidelity)	Extrusion	C2C12	Enhanced printability and fidelity of the construct as well as cell alignment and differentiation	[78d]
GelMA	NA	Extrusion (in situ)	C2C12	Improved cell alignment and differentiation in vitro and enhanced tissue integration in vivo (VML model)	[79e]
GelMA or ColMA	NA	Extrusion	C2C12 or hMPC	Improved cell alignment and maturation in vitro. Demonstrated the expression of sarcomeric α-actinin.	[76]
[GelMA-gelatin] microstrands	NA	Extrusion	C2C12	Improved cell alignment and differentiation in vitro	[78f]
Multicompartmental [GelMA-alginate]	NA	Extrusion	C2C12	Controled cell alignment and maturation in vitro	[58b]
GelMA	PEGDMA (enhanced printability)	Inkjet	hSkMC and rat tenocyte	In vitro muscle and tendon-muscle models. Demonstrated sarcomeric patterns and calcium signaling.	[88a]
GelMA	NA	SLA	C2C12	Improved cell alignment in vitro with cell confinement	[83]
GelMA	NA	SLA	C2C12	Printing complex multicellular muscle structure	[94]
GelMA	NA	SLA	C2C12	Forming maturated skeletal muscle model in vitro	[92]
GelMA	NA	SLA (in vivo)	ASC	Treatment of VML in mice using a non-invasive in vivo bioprinting	[105]
HCC-gelatin	NA	SLA (in vivo)	MuSC and fibroblast	De novo formation of aligned myofibers in mice using a non-invasive internal in vivo bioprinting	[106]
Gelatin	Alginate (enhanced printability)	Extrusion	C2C12 and HUVEC	Formation of vascularized muscle tissue in vivo (subcutaneous)	[78l]
Fibrin	HA and Gelatin (enhance printability)	Extrusion	C2C12	Bioprinting of muscle-tendon interface In vitro	[78c]
Fibrin	HA, Gelatin, and Glycerol (enhance printability)	Extrusion	C2C12	Enhanced myogenesis and muscle maturation in vitro and in vivo (subcutaneous rat model). Demonstrated neural and vascular integration in vivo, electrically induced compound muscle action potential (3.7 mV) using electromyography.	[73]
Fibrin	HA, Gelatin, and Glycerol (enhance printability)	Extrusion	hMPC and hNSC	Enhanced myogenesis and muscle maturation in vitro and in vivo (VML model) through neural cell integration. Demonstrated around 100% isometric torque functional restoration in a VML model.	[79c]
Fibrin	HA, Gelatin, and Glycerol (enhance printability)	Extrusion	hMPC	Enhanced myogenesis and muscle maturation in vitro and in vivo (VML model). Demonstrated around 82% isometric torque functional restoration in a VML model.	[79b]
Fibrin	HA, Gelatin (enhance printability)	Extrusion	C2C12	Improved cell alignment and differentiation in vitro for biological actuators. Demonstrated calcium transients, electrically induced functional contraction (max ~50μN/mm2), positive force-frequency relationship	[80]
Collagen	PEO (enhanced printability and scaffolds degradation rate)	Extrusion	C2C12	Improved cell alignment and differentiation in vitro	[78h]
Collagen	Gold Nanowires (form aligned topographical cues)	Extrusion	C2C12	Improved cell alignment for enhanced myogenesis and muscle maturation in vitro and in vivo (VML model). Demonstrated the expression of sarcomeric α-actinin, electrically induced functional contraction, calcium transients in vitro.	[77]
Collagen	NA	Extrusion	C2C12	Improved cell alignment and differentiation in vitro. Demonstrated the expression of sarcomeric α-actinin, electrically induced functional contraction, calcium transients in vitro	[75b]
Muscle dECM (methacrylated)	PVA (form fibrous topographical cues upon dissociation)	Extrusion	C2C12	Improved cell alignment and maturation in vitro	[78j]
Muscle dECM	NA	Extrusion	C2C12	Improved cell alignment and differentiation in vitro. Demonstrated sarcomeric patterns, formation of AChR clusters, and electrically induced functional contraction.	[78g]
Muscle dECM	NA	Extrusion (embedded)	hSkMC and HUVEC	Enhanced myogenesis and muscle maturation in vitro and in vivo (VML model) through endothelial cell integration. Demonstrated electrically induced functional contraction in vitro (max ~0.8 mN) and 80% isometric torque functional recovery in a VML model.	[79a]
PEG-Fibrinogen	Alginate (enhanced printability)	Extrusion	C2C12	Improved cell alignment for enhanced myogenesis and muscle maturation in vitro and in vivo (subcutaneous)	[78k]
Gellan gum	Surfactant (reduce surface tension for enhanced inkjet printability and increased biocompatibility)	Inkjet	C2C12 and PC12	Neural-muscle cell integration	[88b]
PBS	NA	Inkjet	C2C12	Improved myogenesis for biological actuators. Demonstrated electrically induced functional contraction, positive force-frequency relationship	[107]

Alginate is one of the most widely used biomaterials applied in tissue engineering, owning to its simple and rapid ionic gelation, biocompatibility, and tunability of rheological and mechanical properties ^[108]. The rapid gelation and tunable viscosity of the alginate results in promising printability of this material, specifically for extrusion bioprinting ^[109]. Alginate have been used as the primary bioink material for bioprinting of myoblast laden constructs (Table 2). However, a major drawback of the alginate as a bioink in SMTE is the low cell permissibility of the hydrogel. Alginate lacks RGD binding sites and matrix metallopeptidase (MMP) biodegradable sequences ^[108a], which limits cellular activity inside the scaffold. To resolve this challenge, sacrificial materials have been mixed with alginate to create space for myoblast spreading upon dissociation and enhance the degradation rate of the scaffold ^{[78a, 78e]}. However, these scaffolds still suffer from the lack of binding sites, which reduces the cell permissibility of the bioprinted muscle. Alternatively, cell permissive gelatin-based materials have been incorporated into the bioink formulation to provide cell binding sites, though, the presence of alginate in the scaffold with limited biodegradability impairs cellular activity and maturation of muscle cells in such scaffolds ^{[78m, 78n]}.

Gelatin-based materials have been widely used for bioprinting of muscle constructs (Table 2). Gelatin is a biocompatible material with intrinsic RGD and matrix metallopeptidase (MMP) sequences, offering cell-binding sites and biodegradability ^[110]. Furthermore, gelatin benefits from low antigenity, facile processing and low cost. For rapid biocompatible crosslinking of gelatin, and therefore enhancing its printability and mechanical stability of the scaffold, gelatin have been methacrylated, rendering this material as photocrosslinkable GelMA. Given the high cell permissibility, multiple groups implemented GelMA as the primary bioink material for bioprinting skeletal muscle tissues (Table 2). However, the low viscosity of pure GelMA, when utilizing low concentrations, limits its application for extrusion bioprinting of cell-laden constructs ^[111]. While higher concentrations of the precursor solution can offer higher viscosity and thus printability, cell spreading, proliferation, and migration are hindered in the scaffolds formed with high GelMA concentration ^{[6, 111]}. A strategy to resolve this challenge is to exploit thermal gelation property of the gelatin and finely control the bioprinting temperature to print GelMA in a partially gelled state ^[79e]. Due to challenges associated with fine temperature control of the bioprinting process, various components have been alternatively added to GelMA to enhance its printability for SMTE (Table 2). Gellan gum was added to GelMA for enhancing its viscosity from 10 mPa.s (7.5% wt GelMA at 25°C) to more than 300 mPa.s (7.5% w/v GelMA with 0.2% w/v gellan gum at 25°C) while maintaining myoblast viability and spreading potential ^[104]. Similarly, GelMA was mixed with different photocrosslinkable materials to improve its printability ^[78d]. Two different polysaccharides that were functionalized with methacrylic anhydride, alginate-methacrylate (AlgMA) and carboxymethyl cellulose-methacrylate (CMCMA), and synthetic polyethylene glycol diacrylate (PEGDA) were separately mixed with GelMA to enhance its printability while maintaining the functionality of myoblasts encapsulated within the bioink. While all of the additional components enhanced the printability, GelMA+AlgMA and GelMA+CMCMA better supported the viability of the C2C12s and improved their alignment and differentiation.

Although the addition of printability enhancers to the main scaffolding material can facilitate its biofabrication, it comes with the cost of reduced cellular functionality in most of the reports ^{[78m, 78n]}. Recently, Samandari et al. ^[58b] developed a new strategy based on compartmentalization of the bioink composites to resolve the challenge of balancing printability with cell permissibility (Figure 3B). They demonstrated that the integration of alginate (2% w/v) with low-methacrylated GelMA (10% w/v) using the multicompartmental bioprinting strategy does not limit the cell permissibility but also enhances cellular alignment and maturation. In this strategy, instead of homogeneously mixing two hydrogel precursors, an optimized level of mixing is used to create distinct compartments of two hydrogels in the form of an intercalated structure consisted of multiple microfilaments of the hydrogels. As a result, the multicompartmental bioink is easily printable by rapid gelation of the alginate compartments, forming a matrix with embedded GelMA precursors. Subsequent GelMA photocrosslinking generates a favorable environment for cell functionality, while the presence of alginate in this microfilamentous structure induces cellular alignment.

An alternative strategy for balancing the printability and cell permissibility is the application of temporary polymeric networks mixed with the main bioink for entrapping the uncrosslinked chains of the primary biomaterial to allow for its secondary crosslinking followed by the removal of the temporary network ^[112]. This strategy was implemented for SMTE, where a mixture of photocrosslinkable semi-synthetic polyethylene glycol (PEG)-fibrinogen (0.8% w/v) and sodium alginate (4% w/v) was used as the bioink for the bioprinting of myoblast-laden scaffolds through a coaxial microfluidic extrusion device ^[78k]. In the bioink formulation, alginate acted as a temporary template material to enable rapid crosslinking upon exposure to CaCl₂ solution, which was extruded simultaneously with the bioink through the shell of the coaxial nozzle. Photopolymerization of PEG-fibrinogen after crosslinking, followed by alginate leaching with ethylenediaminetetraacetic acid (EDTA) calcium chelator, left a stable cell-laden scaffold, supporting cellular alignment in vitro and in vivo. However, two-step crosslinking followed by removal of the sacrificial polymers using additional chemicals makes the bioprinting complicated and less biocompatible.

Fibrin, a hydrogel network formed through polymerization of fibrinogen, is a natural material extensively used in tissue engineering ^[113]. The abundance of cell binding and degradation motifs, capability of binding cell-derived growth factors, possibility to be derived from patient’s own blood, and proper elasticity of the crosslinked network makes fibrin a promising biomaterial for muscle tissue engineering ^{[64a, 114]}. Myoblasts can spread homogeneously in the fibrin scaffolds, remodel the network, and differentiate to mature muscle fibers. However, due to low viscosity and slow crosslinking kinetics, usually through thrombin exposure, fibrin is mixed with other components to enhance its printability for SMTE (Table 2). A 2% (w/v) fibrinogen solution in high glucose DMEM supplemented with sacrificial gelatin (3.5% w/v), HA (0.3% w/v), and glycerol (10% v/v) demonstrated a good printability, followed by excellent cellular activity upon fibrinogen crosslinking and dissociation of supplemented materials ^{[73, 79b, 79c]}. However, low mechanical stability of the fibrin scaffold in such concentrations necessitated the application of supporting structures, which complicated the bioprinting procedure.

The only biomaterial that can fully recapitulate the specific physical, mechanical and biochemical properties of a target tissue for cellular activity is the native tissue ECM ^{[79d, 115]}. Therefore, researchers implemented tissue-specific decellularized ECM in bioinks for bioprinting of corresponding tissues ^[116]. A muscle-derived ECM provides mechanical and biochemical cues required during the myoblast development toward mature myotubes ^{[78g, 78j]}. Choi et al. ^[78g] prepared mdECM by harvesting porcine skeletal muscle tissue followed by its decellularization and lyophilization. The mdECM was dissolved at 1% w/v concentration and reported to have thermoresponsive property, which could form a gel at physiological temperatures. The prepared solution was used as a bioink and demonstrated shear-thinning properties, ideal for extrusion bioprinting. The bioprinted constructs using this bioink provided a myogenic environment to induce cellular alignment, differentiation, and maturation toward contractile myotubes. Interestingly, a remarkable number of AChR clusters that are involved in the development of NMJ could be detected in the bioprinted constructs using the mdECM, in contrast to those printed with collagen ^[78g]. However, the slow thermal gelation of the developed mdECM bioink (as reported ~30 min) reduces the printability of such materials.

To make the mdECM more compatible with the bioprinting of skeletal muscle constructs, it has been functionalized with methacrylic anhydride (MA) to form photocrosslinkable mdECM-MA ^[78j]. It has been reported that the synthesis of mdECM-MA supported the preservation of high levels of collagen, elastin, and glycosaminoglycans, as well as many growth factors and cytokines beneficial for muscle regeneration. On the other hand, the synthesis could eliminate cellular and DNA components to minimize the immune response of the host species. As expected, a high level of myogenesis was observed upon myoblast encapsulation and their subsequent bioprinting using the mdECM-MA bioink ^[78j]. While the dECM-based bioinks are highly cell-permissive, they suffer from low availability, considering the challenges associated with approval of xeno-sourced biomaterials for clinical applications.

Although natural polymers have been preferred as the main component of bioinks for SMTE, they suffer from batch-to-batch variation, which is a major challenge for their clinical application and for the reproduction of research findings ^[117]. Standardization of the biomaterial synthesis and bioink preparation, as well as quality control measures post-preparation, are highly required for a successful future of bioprinting in SMTE.

The printability and cell permissibility are not the only requirements of a good bioink for SMTE. Given the crucial role of directionality in muscle development and functionality, additional components may be incorporated to the bioinks to incorporate morphological, physiochemical, and mechanical cues for enhanced cell alignment and differentiation. For example, sacrificial materials can be mixed with the main bioink and printed using extrusion bioprinting to form elongated micro-porosities upon leaching. Gelatin ^[73], PVA ^[78j], and Pluronic F127 ^[78e] are the main sacrificial materials used in the bioprinting of skeletal muscle tissues because of their ability to leach out of the main printed structure via melting at physiological temperatures, hydrolysis, and melting at low temperatures, respectively. Additionally, high aspect ratio particles such as micro-/nano-rods may be added to the bioink and aligned via extrusion or external fields to form morphological cues for cell alignment ^[77].

3.2.1. Emerging bioinks as potential candidates for SMTE

Another strategy to control cellular behavior with adjustment of bioink formulation is the incorporation of self-assembly peptides for controlling scaffold architecture in sub-micrometer scales ^[118]. During self-assembly, specifically selected peptides with random organization autonomously form structurally well-defined and stable constructs through noncovalent interactions between the components ^[119]. In this process, the interaction of the peptides is spontaneous and reversible. The self-assembly peptides can form hydrogels with nanofibrous architectures to induce morphological cues for the cells, offer an ECM-like biophysical environment, and control the release kinetics of loaded therapeutics ^[119–120]. Therefore, biomaterials based on self-assembly peptides have been widely used for biomedical applications ^[119–120]. Recently, self-assembly peptides have been implemented as bioinks in bioprinting applications ^{[118, 121]}. Considering the reversible non-covalent binding of these peptides, they can be interesting candidates for extrusion bioprinting by designing them as shear-thinning hydrogels ^{[118, 122]}. Self-assembly peptides have also been investigated as bioinks for SMTE ^[123]. The fibrous architecture of assembled peptides could resemble the native collagenous ECM microenvironment and support the myogenic differentiation of C2C12 myoblasts ^[123]. Interestingly, it has been shown that the architecture of nanofibers can be controlled and aligned with the extruded filaments, which result in the alignment of encapsulated cells ^[124]. This strategy has been used for the transplantation of satellite cells into injured muscles ^[125]. The results demonstrated that the injected bioink in vivo caused co-alignment of peptide nanofibrous architecture with native myofibers, support the survival, proliferation, and differentiation of muscle progenitors ^[125]. Self-assembly peptide-based hydrogels were also used as IGF-1 carriers for their sustained delivery to enhance myogenesis in cardiac muscle tissues ^[126]. While highly interesting for SMTE, the self-assembly peptides suffer from low shape fidelity after bioprinting. To resolve this, these biomaterials can be designed to be stiffer, which in turn can increase shear stress on the cells during bioprinting ^[118].

One more important factor which needs to be considered for the development of a proper bioink in SMTE is cellular interaction in the bioprinted constructs. The bioink needs to support sufficient cell-cell communication since it is well known that proper cell-cell communication can promote the myogenesis of the muscle progenitors ^[127]. One approach is to develop conductive biomaterials which facilitate cellular interaction through ion exchange ^[128]. The addition of conductive components in bioinks has been reported to enhance cell-cell signaling and the final functionality of the bioprinted muscle construct ^[78i]. However, the direct exposure of most conductive materials to the cells embedded within the bioink typically reduces the biocompatibility of the bioink. Furthermore, intimate contact of the myogenic cells is usually required for their communication through gap junctions and transport of extracellular vesicles, growth factors, and cytokines ^{[127b, 129]}. To address this need, one strategy is to increase the cellular density for closer contact ^[130]. It has been reported that the increase of muscle progenitor density in the bioprinted construct can enhance cellular alignment, differentiation, and muscle volume post in vivo implantation ^[79b]. However, even with very high cellular densities, cellular communication is impaired due to the lack of endogenous ECM between the myogenic cells. Endogenous ECM secreted by the muscle progenitors themselves offers a native-like tissue environment, mimicking the muscle cell niche, and significantly facilitates cellular communication and local availability of various signals in the microenvironment ^[131]. 3D cell aggregates are therefore great candidates to be replaced by the individually dispersed cells in the bioink. Cell aggregates enhance cellular activity, differentiation, and protein secretion ^[132]. 3D cell aggregated can either be mixed with scaffolding biomaterials to prepare a bioink ^[133] or be printed directly as a scaffold-free bioink ^[134]. Interestingly, it has been reported that myoblast spheroids can reorganize and form aligned multinucleated myotubes in response to morphological cues after deposition using a cell electrospinning approach ^[135]. Cell aggregates can be generated in the form of cell spheroids ^[136] or strands ^{[131c, 137]}, and bioprinted into constructs with high cell densities. While cellular communication is a crucial factor in myogenesis, there is no report on the bioprinting of skeletal muscle cell aggregates to this date. Considering the potential of cell aggregates to reorganize and form aligned myotubes, outperforming single cells in myogenesis ^[135], cell aggregate-based bioinks may have a promising future in SMTE.

3.3. In vitro maturation of 3D bioprinted constructs

As mentioned briefly in Section 2.5, internal or external stimulation can be utilized post-biofabrication to improve cellular organization, myogenesis, and final maturation of the constructs. A realistic recapitulation of muscle structure requires an approximate myofibers diameter of 50–100 μm with the capability to generate contractile forces in the order of 200 mN/mm^{2 [63]}. However, there are only a few investigations that reported the measurement of contractility of the bioprinted skeletal muscles ^{[79a, 80, 138]}, where the contractile forces generated by the printed muscle grafts were a couple of orders of magnitude less than the physiological levels. Several other in vitro functional measures for quantifying the performance of bioprinted muscle have also been performed: tensile stress testing of cell-laden constructs ^[78g], passive tension after exposure to cardiotoxin ^[92], calcium signaling ^{[80, 88a]}, and contraction displacement ^[80]. Although the bioprinted constructs can be integrated into the host tissue and matured post-implantation, different strategies have been developed for in vitro enhancement of skeletal muscle maturation and functionality through various stimulations inducing exercise-mimetic and/or nutrition-mimetic conditions ^[139]. Mechanical and electrical stimulations are the main approaches for in vitro improvement of engineered skeletal muscle constructs.

Mechanical loading during both embryonic development and adulthood is known to be an important factor in skeletal muscle maturation and hypertrophy ^[63]. Therefore, mechanical stimulation has been used for in vitro maturation of skeletal muscle constructs. Particularly, cell-induced hydrogel compaction and consequently passive mechanical tension have been widely utilized for the improvement of 3D muscle constructs in vitro ^[5]. In this method, as mentioned in Section 2.5, hydrogel embedding muscle progenitors is molded and constrained using stiff anchors. The contractile nature of the muscle cells induces tension along the line passing the anchors ^{[64a, 140]}. This passive tension is shown to align the cells, improve their myogenesis, and their final maturation and functionality. This approach has been applied for maturation of bioprinted skeletal muscle constructs by simultaneous printing of anchoring elements and cell-laden filaments ^{[73, 79b, 79c]}, the printing of cell-laden bioink around prefabricated anchors ^{[80, 88a]}, or assembly of bioprinted muscle construct on anchoring elements ^[138].

Mechanical stimulation can also be applied by external static or dynamic strains using various bioreactors ^[62]. Notably, a 6-day static mechanical strain with a magnitude fluctuating between 3–10% demonstrated significant improvement in alignment and myogenesis of myoblasts encapsulated in a fibrin hydrogel ^[141]. As another example, a combination of static (growing 0–10% for 4 days, static 10% for 3 days) and dynamic (5% for 2 days, 10% for 2 days, 15% for 4 days) strains of human skeletal muscle cells in a Matrigel/collagen scaffold, respectively mimicking the embryonic development and muscle hypertrophy, demonstrated 200–300% increase in elasticity, 12% fiber diameter, and 40% myofibers area of engineered muscle construct ^[142]. However, the authors could not find a report on the application of external mechanical stimulation for the improvement of bioprinted skeletal muscle constructs.

Electrical stimulation has also been implemented for enhanced cellular organization, myogenesis, and functionality of engineered skeletal muscles. Typically, electrical stimulation is applied to mimic the in vivo neural activity when generating electrochemical signals from motor neurons ^[63]. Using 2D and 3D engineered skeletal muscle constructs, it has been demonstrated that electrical stimulation can induce alignment of muscle progenitors, enhance their differentiation, increase their contraction potential, and even change their phenotype (slow or fast-twitch) ^{[63, 139]}. Interestingly, recent studies reported that electrical stimulation with intermittent stimulation regimes at 1 Hz and 10 Hz (bipolar 70 mA amplitude and 2 ms duration pulses) could be used as an exercise-mimetic condition to increases hypertrophy and reduce muscle wasting in engineered skeletal muscles ^{[65b, 143]}. While these fascinating results are observed in biofabricated muscle constructs with traditional tissue engineering approaches, studies on the application of electrical stimuli on the improvement of bioprinted muscles are very limited and not promising ^[80], which is due to the earlier stage of muscle bioprinting compared to traditional SMTE. Considering the clear advantages of electrical stimulations on the engineered muscles, more investigation on the development and optimization of strategies for the maturation of bioprinted muscles using electrical stimulations needs to be performed.

Other examples of stimulation for improvement of engineered muscle in vitro are chemical (for example with IGF-1 growth factor ^[144] or synaptogenic agrin ^[145]) and optogenetic ^[146] stimulations, which can be found comprehensively in recent reviews ^{[62, 139]}.

The above-mentioned discussion demonstrates the benefit of various stimulations for SMTE, which can be applied to bioprinted muscles for their enhanced maturation and functionality. For this purpose, two challenges need to be addressed: (i) studies on the application of bioreactors and functional characterization of bioprinted skeletal muscle tissues are very rate and new investigations need to be performed development and standardization of such approaches; (ii) there exists a huge difference between the structure and functionality of in vivo healthy muscle and tissue-engineered ones, even post-incubation in various bioreactors, which may imply the need for implementation of the body as the bioreactor post-implantation (See Section 3.4).

3.4. Bioprinting for regeneration of muscle defects in vivo

The main goal of bioprinting for SMTE is the regeneration of muscle defects. As mentioned before, VML imposes significant financial and social burdens on the patients and healthcare system. However, as indicated in Table 2, bioprinting investigations on SMTE were mostly focused on evaluating cellular behavior in vitro. While these studies provide valuable information, several factors limit their translation in vivo ^[147]; first, most of the bioprinted muscle tissues in vitro are smaller than the clinical-relevant size scales required for regeneration of human muscle defects. Biofabrication of larger construct using many of the presented strategies, if possible, is challenging not only because of its structural stability, but also due to the lack of vascular networks for the transportation of nutrition, waste, and oxygen through the scaffold. Second, many in vivo conditions are not, or even cannot yet be recapitulated in vitro. Several physiological mechanisms including immune responses to the scaffolding material and implanted cells are challenging to be captured by in vitro experiments. Finally, the integration of biofabricated muscle with the host tissue and capability of functional muscle recovery cannot be evaluated in vitro. Therefore, in vivo experiments are required to evaluate the efficacy of the bioprinted muscles. For example, previous in vivo studies have quantified bioprinted SMTE constructs using histological analysis (muscle fiber regeneration area, fibrosis area, hypertrophy from cross-sectional area, formation of NMJs, and vascularization), muscle wound closure, and functional assessment (muscle weight, treadmill running, compound muscle action potential, and isometric and twitch force/torque generation) ^{[73, 77, 79]}. Two main strategies are available for the regeneration of muscle defects in vivo through bioprinting: (i) implantation of in vitro printed and potentially matured muscle construct and (ii) direct in vivo printing inside the muscle defect. In the following section, we will introduce each strategy and discuss its advantages and limitations.

3.4.1. Implantation of in vitro-printed muscle constructs

The main approach for regeneration of defected muscle through bioprinting is the in vitro fabrication of muscle construct followed by its implantation into the injury site. This can be accomplished through (i) scanning the defect morphology and reconstructing a CAD model, (ii) 3D bioprinting using a robotic system controlled by a CAM software, (iii) in vitro culture of the cell-laden construct, and (iv) in vivo implantation of the muscle.

Upon biofabrication, the biocompatibility and integration capability of the bioprinted muscle need to be assessed in vivo. An approach for evaluating in vivo biocompatibility is subcutaneous implantation of bioprinted muscles ^{[73, 78i, 78k, 78l]}. In a prominent study, Costantini et al. ^[78k] showed that after 7 days of in vitro culture, the myoblast-laden PEG-fibrinogen scaffold that was implanted subcutaneously on the back of immunocompromised mice was biocompatible and biodegradable. In addition, the cells encapsulated in the bioprinted scaffolds demonstrated improved maturation into aligned long myotubes after 28 days compared to those encapsulated in the bulk hydrogel. However, the scaffolds did not seem to be integrated with host vascular or neural networks. To evaluate the ability of tissue integration, the insertion of the common peroneal nerve into the biofabricated construct has been reported ^[73]. C2C12-laden bioink, composed of a hyaluronic acid (HA), gelatin, fibrinogen, and glycerol solution mixture, was printed and differentiated for 7 days in vitro followed by subcutaneous implantation into nude rats. After two weeks of implantation, a high level of cellular maturation was observed, and more importantly, implanted scaffolds demonstrated tissue integration indicated by vascularization and innervation of immunostained harvested tissues ^[73]. The functionality of the implanted construct was evaluated via electromyography, and the construct resulted in a compound muscle action potential of 3.7 mV, which was less than the 10.7 mV of the control gastrocnemius muscle but greater than the 0 mV of subcutaneous tissue. The action potential of the construct corresponded with immature but developing skeletal muscle ^[73].

A better understanding of the functionality of the bioprinted muscle in a clinical context can be obtained by its implantation into VML models ^{[77, 79a–c]}. It has been reported that the fibrous architecture of biofabricated structures using extrusion bioprinting can improve the cellular organization and enhance muscle function recovery after VML ^[79b]. Using the multimaterial extrusion bioprinting strategy shown in Figure 2A, a 10×7×3 mm³ artificial muscle flap was biofabricated encapsulating human primary muscle progenitor cells (hMPCs). Following one week of culture in vitro, the bioprinted muscle was implanted into the tibialis anterior (TA) muscle defect of nude rats to evaluate its in vivo functionality. The results demonstrated a significant improvement in muscle weight and capability of force generation, measured through peroneal nerve stimulation compared to the non-treated, gel-only treated, and cell-laden bulk hydrogel treated defects. An 82% functional recovery of the TA muscle force was reported in the injured muscles treated with bioprinted scaffolds after 8 weeks of implantation. Furthermore, immunostaining results showed that the bioprinted muscle integrated well with the hosts’ vascular and neural networks, and newly formed myofibers of the donor were identified in regenerated tissue ^[79b]. Similarly, the other efforts reporting the implantation of bioprinted muscle constructs in VML injury models resulted in isometric torque functional recovery of 80% of uninjured controls^[79a] and near 100%^[79c] (not statistically significant from sham controls). This again implies that the body can be utilized as a natural excellent bioreactor for bioprinted muscle constructs when sufficient biophysical cues are provided. Unfortunately, most of the other bioprinted SMTE constructs listed in Table 2 have not been validated in vivo and lack histological and functional characterization.

The importance of physiochemical cues for muscle tissue recovery upon injury was further confirmed by the incorporation of gold (Au) nanowires in the bioink ^[77]. A myoblast-laden collagen bioink supplemented with gold nanowires was printed using extrusion bioprinting, followed by the application of an electric field for two days in vitro to align the gold nanowires and consequently provide morphological cues for enhancing muscle cell alignment. After 8 weeks of the implantation into temporalis muscle defects of Sprague–Dawley rats, the bioprinted muscle with aligned gold nanowires demonstrated significantly reduced fibrosis and enhanced muscle tissue recovery compared to those without aligned nanowires.

The above-mentioned examples demonstrate the benefit of 3D bioprinting for the regeneration of skeletal muscle defects. However, several challenges still remain to be addressed when treating actual human muscle defects including the complexity of the procedure required for the fabrication of the scaffold that matches the irregular-shape of muscle defects, delayed treatment by CAD/CAM process and in vitro culture period, and limited implant-tissue adhesion, which reduces the opportunity of integration. In addition, the investigation into the effect of the immune system on the efficacy of muscle regeneration must be taken into consideration, which was not covered by any of the aforementioned studies because of their use of immune-deficient animals.

3.4.2. In vivo printing for SMTE

As mentioned before, conventional bioprinting strategies usually suffer from major limitations including (i) delayed treatment; (ii) limited adhesion of the implant to the tissue and therefore its poor integration; (iii) complications for matching the implant to the irregular shaped defects due to the dynamic nature of the tissues, deformation of printed construct during the in vitro culture, and limited capability of the current bioprinters for fabrication of complex structures with curved surfaces; and (iv) lack of proper in vivo physiochemical signals for tissue development before the implantation. Therefore, in vivo, also called in situ or interoperative, bioprinting strategies were introduced to resolve these limitations by offering conformal bioprinting of scaffolds that accurately match the geometry of the defect, delivering the treatment immediately, adhering the scaffold to the remnant tissue through in situ crosslinking, and implementing the body as a natural bioreactor. In vivo printing further allows real-time decision making by the surgeon to account for unexpected incidents and discrepancies in tissue margins from imaging modalities ^[148].

Recently, in vivo printing has been investigated for SMTE (Figure 4). Tamayol and his group developed multiple versions of handheld extrusion-based (bio)printers ^{[79d, 79e, 149]} for the treatment of musculoskeletal defects (Figure 4A and B). Handheld bioprinters can minimize the complexity of robotic printing approaches through the elimination of scanning modalities and the requirement of CAD/CAM processing. Using handheld bioprinters, a surgeon can inspect the muscle defect and directly print the bioink, even on curved surfaces with irregular defect shapes. The first version of their handheld bioprinter was used for in situ deposition and photocrosslinking of C2C12-laden GelMA bioinks (Figure 4A–i) ^[79e]. The results demonstrated that the in situ crosslinking of the GelMA hydrogel induced a strong adhesion between the deposited scaffold and native muscle tissue, which can resolve the challenges associated with implant fixation and implant-tissue integration. The in vitro assessment of the cellular behavior indicated that the in situ printing approach supports cellular viability, while the shear-induced stress experienced during the extrusion bioprinting enables myoblast alignment and differentiation (Figure 4A–ii). Furthermore, the translational feasibility of the approach was demonstrated through in vivo printing of acellular GelMA for the treatment of a murine VML injury model (Figure 4A–iii, iv). They further assessed the effect of the sustained release of vascular endothelial growth factor (VEGF), controlled by binding growth factors to Laponite® nanoparticles, for the improved treatment of VML using a new version of in situ handheld bioprinter (Figure 4B–i) ^[79d]. A high level of muscle recovery was revealed by functional and histological tests in the treatment group that received in vivo printed VEGF-eluting scaffolds (Figure 4B–ii, iii). However, the in vivo bioprinting with cell-laden bioinks using a handheld printer has yet to be achieved.

Figure 4.

In vivo bioprinting for treatment of muscle defects. (A) A handheld extrusion-based bioprinter developed for direct printing of the bioinks inside the muscle defects. The handheld bioprinter with integrated UV light (i) was implemented to deposit and in situ crosslink C2C12-laden GelMA hydrogels. A high level of alignment and myogenic differentiation was observed in the printed constructs (ii). The translational feasibility of the method was further evaluated by in vivo printing of acellular GelMA in a murine VML injury model. Reproduced with permission from American Chemical Society ^[79e]. (B) The application of a handheld bioprinter for treatment of murine VML models by printing growth factor-eluting bioinks. GelMA supplemented with Laponite nanoparticles loaded with VEGF used as a bioink for in vivo printing of biomimetic fibrous structures inside muscle defects (i). The functional (ii) and histological (iii) evaluations demonstrated a significant improvement in the treatment of VML by in vivo printing of VEGF-eluting bioinks (VEGF+) compared to untreated and those treated without the application of VEGF in the bioink (VEGF-). Reproduced with permission from Wiley ^[79d]. (C) Intravital non-invasive bioprinting for de novo formation of muscle. The intravital bioprinting is accomplished by injection of the bioink inside the defect site followed by selective photocrosslinking of the bioink through the skin using near-infrared exposure. Using a photocrosslinkable gelatin encapsulating muscle-derived stem cells (MuSCs) and fibroblasts, elongated structures were bioprinted inside mice muscle (ii). Despite the cells only injected into the defect site (No i3D), those bioprinted in elongated structures (i3D) demonstrated a high level of alignment and tissue integration 7 days post bioprinting (iii). Reproduced with permission from Nature Publishing Group ^[106].

The in vivo bioprinting for SMTE has been evolved even further recently by the development of non-invasive intravital bioprinting ^[105–106]. Using a stereolithography-based bioprinting approach and application of photo-initiators sensitive to near-infrared (NIR) light, researchers enabled internal bioprinting without open surgeries. The bioink is first injected into the defect site, followed by its selective NIR-photocrosslinking through the skin using two-photon excitation ^[106] (Figure 4C–i) or digital micromirror device (DMD) projection^[105]. The intravital non-invasive strategy has been reported to successfully fabricate adipose-derived stem cells-laden GelMA scaffold matching muscle defect and effectively treat the VML ^[105]. Furthermore, it has been demonstrated that this strategy can create biomimetic elongated structures for SMTE ^[106] (Figure 4C–ii). Through bioprinting of a photocrosslinkable gelatin encapsulating muscle-derived stem cells (MuSCs) and fibroblasts, the elongated structure could induce a high level of cellular alignment, myogenesis, and vascular network integration (Figure 4C–iii). Due to the embedded nature of this bioprinting strategy, it does not need high viscosity bioinks to generate scaffolds with high fidelity. However, SLA-based in vivo bioprinting is limited to photocrosslinkable bioinks.

3.5. Bioprinting for Engineering Vascularized and Innervated Muscles

Muscle injury is multifaceted and causes disruptions in the local ECM, vasculature, and neural networks. Ensuring proper vascularization and innervation is of significant interest to SMTE for both regeneration in myopathies and for in vitro muscle modeling. Lack of proper innervation prevents the maturation of skeletal muscle and coordinated action potential signaling. Furthermore, vasculature is needed for the engineering of large muscle tissue to provide adequate nutrients, oxygen, and waste transport. Fully functional muscles cannot be conceived of without these two vital components, and implanted constructs without cell populations to develop these systems are dependent on in vivo innervation and vascularization that can fail to occur or occur without proper organization ^[67a].

Two main strategies have been introduced for vascularization and innervation of the bioprinted muscles: (i) in vivo vascularization/innervation and (ii) pre-vascularization/innervation. In the first strategy, the bioprinted muscle tissue without vasculature or a neural network is implanted in vivo and the host body induces vascular and neural network formation into the artificial muscle flap ^[67a]. Although this approach is simple, in many cases, the host’s vasculature and nerves fail to properly infiltrate into the scaffold due to the improper physiochemical properties of scaffolding material, which cause limited functional recovery of the muscle ^[67a]. A possible resolution for this challenge is the formation of micro/macro porosities or capillaries inside the bioprinted muscle, for example through the implementation of sacrificial materials in bioprinting strategy to enhance the opportunity of remnant muscle to induce vascularization and innervation into the artificial flap ^{[73, 79b]}. Alternatively, in the second strategy, pre-vascularization/-innervation approach, researchers attempt to include endothelial ^{[78l, 79a, 94]} and neural ^{[79c, 88b, 93]} cells in the bioprinting of skeletal muscles and culture in vitro before implantation. Several investigations have reported bioprinting of muscle progenitors with neural and endothelial cells in vitro using extrusion bioprinting ^[78l], inkjet bioprinting ^[88b], stereolithography ^[93–94], and cell electrospinning ^[95c]. However, to evaluate the clinical relevance of these approaches, in vivo investigations are required for the evaluation of host integration of the vascular and neural systems (Figure 5).

Figure 5.

Vascularization and innervation in bioprinting of skeletal muscle tissues. (A) Bioprinting of pre-vascularized scaffold in a designed core-shell structure and its efficacy in vitro and in vivo. (A) An embedded bioprinting strategy was used for bioprinting of three types of artificial muscle flaps (i): printing: human skeletal muscle cells (hSkMCs) in muscle-derived ECM (mdECM) bioprinted with a fibrillar architecture; printing mixed: hSkMCs and human umbilical vein endothelial cells (HUVECs) in a homogeneously mixed mdECM and vascular-derived ECM (vdECM) bioprinted with a fibrillar architecture; coaxial printing: hSkMCs in mdECM as the core of the fibers and HUVECs in vdECM as the shell of the fibers bioprinted with a fibrillar architecture (ii). The in vitro investigation demonstrated an organized structure of muscle and vascular cells in the coaxial printing group (iii). Immunostaining of the harvested muscle tissue after 4 weeks of implantation in rat VML model demonstrated a high level of integration and anastomosis of the host vasculature into the prevascularized scaffolds fabricated using coaxial printing (iv). The high level of integration was further confirmed by the formation of neuromuscular junctions (NMJs) and host neural network integration in coaxially printed scaffolds (v). Reproduced with permission from Elsevier ^[79a]. (B) Bioprinting of pre-innervated biomimetic muscles for in vitro and in vivo myogenesis. A multimaterial bioprinting strategy was used to fabricate large fibrous scaffolds with embedded microchannels. The main bioink encapsulated human muscle progenitor and human neural stem cells (hMPCs and hNSCs) (i). In vitro evaluations demonstrated that the integration of neural cells can enhance the viability (ii) and maturation (iii) of the muscle progenitors. Furthermore, the in vitro formation of neuromuscular junctions (NMJs) was confirmed by AChR immunostaining results (iv). The immunostaining of the harvested tissue after 4 and 8 weeks of implantation in the rat VML model demonstrated the integration of host neural network (NF⁺ signal) and formation of NMJs (AChR⁺ signal) with the implanted muscle fibers (MHC⁺ signal) (v). On the other hand, the hNSCs in the implanted tissue (βIIIT⁺ signal) were differentiated and integrated into the host neural network (HLA⁺ signal). Reproduced with permission from Nature Publishing Group ^[79c].

The in vivo efficacy of pre-vascularized bioprinted muscle tissues was explored by Choi et al. ^[79a] using embedded extrusion bioprinting of cell-laden decellularized ECM (Figure 5A–i). Three different bioprinting approaches were considered to evaluate the effect of presence and distribution of endothelial cells in the bioprinted muscle flap (Figure 5A–ii): (i) printing of hSkMCs in mdECM, (ii) printing of a homogeneously mixed bioink consisted of hSkMCs and human umbilical vein endothelial cells (HUVECs) encapsulated in decellularized muscle and vascular ECMs (mdECM and vdECM), and (iii) coaxial extrusion bioprinting of hSkMCs in mdECM and HUVECs in vdECM, through the core and shell of the coaxial nozzle, respectively, to mimic the architecture of the muscle fibers surrounded by blood capillaries. Interestingly, the in vitro results demonstrated that the printing of mixed muscle and endothelial cells downregulate the differentiation and maturation of both cells, as indicated by the result of immunostaining (Figure 5A–iii). However, the coaxial printing resulted in the greatest vessel formation, muscle maturation, and functionality of the bioprinted muscle measured through ex vivo twitch and tetanus forces measurement. Following the promising in vitro outcomes, they implanted pre-vascularized muscle flaps into the TA defects of Sprague-Dawley rats with approximately 40% muscle loss. The pre-vascularized bioprinted muscles resulted in the highest muscle weight recovery and lowest fibrosis, compared to bioprinted muscles without HUVECs, bioprinted muscles with mixed bioink, or untreated animals. A 76.8% muscle weight recovery was reported for the animals treated with pre-vascularized bioprinted muscle flap with organized core-shell structure after 4 weeks of implantation. Immunostaining further revealed a higher level of HUVECs-derived and host capillaries in co-axially printed pre-vascularized muscle tissues, with co-localization of capillaries, indicating vasculature anastomosis (Figure 5A–iv). As a result of proper implant integration, an 85% functional recovery was observed in the capability of muscle force generation. This high level of functional recovery was further confirmed by the formation of NMJs and host neural network integration (Figure 5A–v).

In addition to pre-vascularization, the effect of innervation of the bioprinted muscle on the restoration of muscle function was recently studied ^[79c] (Figure 5B). Using the multimaterial strategy shown in Figure 2A, human neural stem cells (hNSCs) were mixed with hMPCs, encapsulated in the bioink, and printed to form fibrous structures with macro-porosities between the fibers (Figure 5B–i). Interestingly, in vitro results demonstrated that neural cell integration enhances the viability and maturation of the muscle cells (Figure 5B–ii, iii). Additionally, the formation of NMJs indicated by AChR immunostaining was reported in vitro (Figure 5B–iv). Subsequently, the pre-innervated muscle flaps were implanted into immunodeficient RNU rats with a 40% TA muscle defect. After 4 weeks of implantation, a significantly higher level of muscle weight recovery and lower level of fibrosis was observed in the animals treated with pre-innervated muscle flaps when compared to non-treated and hMPCs-only treated groups. Furthermore, immunostaining results demonstrated NMJ formation between host nerve (NF⁺) and myofibers (AChR⁺ on MHC⁺) formed in the bioprinted muscle (Figure 5B–v), as well as hNSCs differentiation and integration with host neural network (Figure 5B–vi).

Overall, the in vitro and in vivo results demonstrated that vascularization and innervation of skeletal muscle constructs are necessary, both in vitro and in vivo, for the long-term survival of the engineered tissue, accelerated myogenesis and maturation, and the functional recovery of muscle after injury. The ability to print and spatially organize endothelial and neural cells and muscle cells allow for the recreation of more physiologically relevant muscle fibers that are independent of host infiltration ^[67a].

3.6. Bioprinting for Interfaced Muscle-Tendon Engineering

Unfortunately, many musculoskeletal defects involve damage to the musculoskeletal interfaces. However, few attempts have been made to engineer musculoskeletal interfaces. In addition to regenerative medicine, a good understanding of the behavior of the skeletal muscle tissues in vitro is associated with investigating the musculoskeletal system as a unit. Given the flexibility of the bioprinting strategies in multimaterial/multicellular biofabrication, they can be adapted for engineering musculoskeletal interfaces. While bioprinting bone-tendon units are out of the scope of this study, readers are referred to recent review papers on this topic ^[8]. Here we discuss the attempts that have been made to integrate skeletal muscle and tendon tissues.

Native muscle-tendon units consist of an elastic muscle domain with densely packed organized myotubes and a stiff tendon domain with spare fibroblasts in collagen-based ECM connected at the myotendinous junction ^[78c]. To mimic this architecture, the multimaterial bioprinting strategy shown in Figure 2A was implemented for recapitulating the native structure of the myotendinous junction ^[78c]. C2C12 myoblasts and NIH/3T3 fibroblasts were mixed separately with a bioink that consisted of gelatin, HA, and fibrinogen and were loaded into two separate printing heads of an extrusion-based bioprinter. Also, for each section, a support material was selected to construct a stable structure with different mechanical properties. Polyurethane (PU), an elastomeric material, was implemented for the muscle domain and stiff PCL with its high young modulus and plastic characteristics was used for the tendon side. The bioprinting strategy demonstrated a high level of cell viability and differentiation in both muscle and tendon units as well as good cellular integration at the interface. Interestingly, gene expression analysis demonstrated the development of muscle-tendon-like interfaces by upregulated expression of paxillin, talin, vinculin, integrin β1, laminin α1, and laminin α2, compared to only-muscle bioprinted tissues. In another study, primary human skeletal-muscle-derived cells (SkMDCs) and primary rat tail tenocytes were co-printed using inkjet bioprinting to form muscle-tendon units ^[88a]. Tenocytes were encapsulated in a mixture of GelMA and polyethylenglycol dimethacrylate (PEGDMA) bioink and were printed around two vertical posts, followed by the printing myoblasts-laden GelMA/PEGDMA between the tendon domains. The results demonstrated the differentiation of the unit toward contractile tissues. These preliminary studies demonstrate the importance of studying muscle-tendon interaction in SMTE. However, more in vitro and in vivo investigations are required to better understand synergistic effects of muscle and tendon cells on their differentiation and development, as well as regeneration of muscle-tendon units.

4. Future Outlook and Conclusions

Skeletal muscles occupy a major mass of human body and play key functions essential to our living. Skeletal muscle injury and diseases are frequent and major injuries caused by trauma or tumor removal can have debilitating outcomes. 3D bioprinting for SMTE have shown a great promise to engineer functional tissues that can be implanted to recover the lost functions. In addition, tissue engineered muscles can serve as reliable tools for drug discovery and diseases modeling. Despite the vast progress that has been made in bioprinting in SMTE, several aspects of this field have yet to be addressed or fully developed for the clinical benefits to be realized ^[6].

One of the most important challenges challenge towards the clinical use of SMTE products is the limited knowledge on their interaction with the host immune system. As discussed in Section 2.2, immune system is a major contributor in the innate regeneration of skeletal muscle. However, when implementing an exogenous engineered muscle graft, the immune system is not only a contributor in muscle regeneration, but also a protective system against any unknown source. The main challenge against understanding the interaction of immune system and bioprinted muscle grafts is complexity of the signaling cascades among different cells types in the microenvironment, which makes removing any species from the system challenging or even impossible. As a result, immune deficient animals cannot properly serve as a model for the in vivo studies. Specifically in muscle regeneration, most of the immune cells including T cells play major roles in repair mechanism ^[21c]. On the other hand, the immune system of wild type animals generally doesn’t allow application of human-derived materials. Therefore, the tissue engineering grafts designed for human may not be applicable to animals and clinical trials are necessary to fully elucidate the effects of the designed tissue engineering strategy.

Another option is the use of immune compatible materials along with patients (animal)-derived biomaterials and cells, which can substantially minimize the risk of an adverse immune response towards the engineered tissue. Fibrin, one of the best candidates for primary bioink material can be harvested from the patients’ blood with a minimally invasive manner, while stem cells can be harvested directly with small biopsies and propagated in vitro, or indirectly through induction of pluripotency. However, to this date still there is no agreement towards the ideal cell source for muscle tissue regeneration.

While skeletal muscle satellite cells and induced pluripotent stem cell (iPSC)-derived muscle progenitor cells are the front runners, they suffer from main limitations. The application of patient-derived satellite cells is typically limited due to the loss of their regenerative potential during their long-term expansion in vitro necessary for generating enough cells for regeneration of large defects ^[150]. Furthermore, regulatory processes and therefore clinical implementation of iPSCs-included therapies are very cumbersome, due to the genetic manipulation of cells to be administrated in human body. Therefore, only a few number of clinical trials could be registered to date for clinical application of iPSCs ^[151], despite numerus publications on this field. Furthermore, in cases that the diseases are caused by genetic disorders, the use of patient own cells could carry a similar risk of the disease occurrence. In these cases, the use of gene editing tools to repair any adverse genetic abnormalities that may be the cause of specific disorders is necessary. Furthermore, the compatibility of these cells with various bioprinting modalities and their ability in forming hierarchical muscle mimicking tissues are not well studied yet. Despite the above-mentioned challenges, it is expected that the application of iPSCs for SMTE expands due to their unique advantages offering a patient-specific cell source with good expansion capability, as well as allowing gene editing for treatment of genetic myopathies. For more information on the application of iPSC for SMTE, readers are referred to recently published excellent reviews ^[152].

In addition to muscle progenitors, other cell types such as endothelial and neural cells need to be considered for integration into bioprinted muscle constructs, as discussed in Section 3.5. Few publications reported the bioprinting strategies for development of vascularized or innervated muscle constructs and their integration into host vascular and nervous systems. Due to the importance of these systems on the proper muscle function (See section 2.1), it is expected that effort to identify proper cell sources and strategies that allow the proper vascularization and innervation of bioprinted scaffolds to be the subject of various studies.

As the complexity of cell types and lineages incorporated into SMTE constructs increases, improved focus on the biomaterial properties, both pre and post printing, needs to be considered. While there has been a significant number of investigations into the properties of 2D substrates on myogenic and other cellular responses, understanding cellular responses and behavior in 3D environments require further study ^[153]. Further, most post crosslinking investigation has placed significant focus on time-independent metrics such as stiffness and elasticity of the scaffold, but a further understanding of the effects of time-dependent viscoelasticity, which resembles native tissue, is needed ^[154]. Finally, the effect of microenvironmental remodeling on the bulk biomaterial and any added components on cellular response, development, and disease progress also need clarification ^[155].

One more improvement that can revolutionize the field of bioprinting for SMTE is the identification and incorporation of biological factors, which are needed to modulate the environment post-implantation and prevent excessive inflammation and fibrosis at the tissue implant interface, into the bioink. However, some challenges limit the application of biological factors in the bioprinting investigations for SMTE. Growth factors are typically expensive and successful tissue regeneration requires a customized cocktail of factors based on the patients’ needs ^[156]. Patient-specific blood products have generated a great promise as a suitable source of biological factors in bioinks ^[156]. While the application of blood-derived components for bioprinting of muscle constructs has not yet investigated, several studies demonstrated the benefit of incorporation of blood-derived products into the bioink for improved cellular functionality post bioprinting of other tissue constructs ^[157]. However, the biological factors derived from blood-derived products can be denatured easily and can significantly lower their long-term benefit. The recent success of COVID-19 vaccines has redrawn significant attention towards mRNA-based therapies that utilizes endogenous cells as factories for the long-term expression of the target molecules. However, the longevity and suitability of these therapies for treatment of chronic disease and traumatic injuries has remained to be investigated.

Another aspect which needs specific attention for realization of 3D bioprinting for SMTE is the development of intraoperative bioprinting strategies, as discussed in Section 3.4.2. While the development of efficient and ergonomic in vivo bioprinters has still a long way to go and need more investigations, a special attention needs to be paid toward the bioinks and their preservation. In some cases, the bioinks need to be immediately available for application upon unexpected injuries. Usually, the preparation of muscle progenitor cell population for incorporation in intraoperative bioprinting requires a long time, which needs to be considered beforehand. The development of approaches for proper preservation of bioinks components (biomaterials, cell bank, etc) or the whole bioink composite is of particular interest, specifically for muscle injuries in the battle field or space missions ^[158].

A key factor that is needed for the effective translation of bioprinted scaffolds to clinical use is their proper standardization. So far, there are limited efforts to develop standards and quality control measures in the area of bioprinting, specifically for SMTE. This limitation stems from the fact that the field is young, interdisciplinary, and underdeveloped. Regardless, an interdisciplinary effort led by governments is needed to develop standard test protocols to characterize various mechanical, biological, and functional properties of bioprinted scaffolds at different stages of culture. Therefore, in parallel with the technological advancements, regulatory hurdles, quality control, and manufacturing standards must be achieved and established, respectively ^[159]. With the progression toward clinical viability, SMTE and skeletal muscle bioprinting will open a new paradigm toward treatment of muscle diseases and injuries.