Bioengineered Constructs as a Tissue Engineering-Based Therapy for Volumetric Muscle Loss

Abstract

Severe skeletal muscle injuries involving substantial tissue loss can significantly impair muscle strength and functionality, reducing the quality of life for affected individuals. Such injuries, termed volumetric muscle loss, require extensive clinical intervention, as the body’s innate healing mechanisms are insufficient to regenerate functional muscle. The current standard of care primarily involves autologous muscle tissue transfer, with some consideration of acellular synthetic constructs. However, both approaches have limited therapeutic efficacy, presenting challenges such as donor-site morbidity, infection risks, and suboptimal functional recovery. Over the past decade, skeletal muscle tissue engineering (SMTE) has emerged as a promising strategy for regenerating functional muscle through bioengineered constructs. Advanced biofabrication techniques, including bioprinting, have further enabled the development of synthetic constructs that closely mimic native muscle architecture. Given these advancements, a critical review of recent therapeutic strategies, their achievements, and limitations is necessary. This review examines the spectrum of bioengineered constructs developed from various biomaterials and evaluates their therapeutic potential. Special emphasis is placed on 3D bioprinting strategies and their role in creating physiologically relevant constructs for functional muscle restoration. In addition, the integration of machine learning in optimizing construct design, predicting cellular behavior, and enhancing tissue integration is discussed. The review indicates that despite significant progress in SMTE, key challenges remain, including replicating the complex structural organization of muscle tissue, minimizing fibrosis, and achieving vascularization and innervation to regenerate functional, strengthened muscle. Future research should address these barriers while prioritizing the development of translational, clinically relevant regenerative constructs. In addition, efforts should focus on advancing scalable, construct-based regenerative treatments that are readily available at the point of care and easily managed in surgical settings.

Impact Statement

This review highlights key advancements in skeletal muscle tissue engineering (SMTE) for treating volumetric muscle loss (VML), a condition that severely impairs muscle function and quality of life. By exploring the advancements in bioengineered constructs and the role of 3D bioprinting, this review underscores innovative strategies that closely replicate native muscle tissue, offering new prospects for functional muscle regeneration. In addition, the review discusses machine learning integration in SMTE to optimize construct design and predict cellular behavior, presenting a promising avenue for developing constructs with high therapeutic value. This review also highlights essential insights into the existing limitations in constructs and proposes future directions that should be undertaken to advance construct-based regenerative medicine toward more effective and clinically viable solutions in VML therapy.

Keywords

biomaterials bioprinting constructs machine learning skeletal muscle tissue engineering volumetric muscle loss

Introduction

As per the Global Burden of Disease 2019 data, musculoskeletal conditions affect 1.71 billion people globally, almost 25% of the world’s population.^1,2 These conditions lead to 17% of global disability-adjusted life years and cost $1 trillion annually in treatment. In the United States, musculoskeletal disabilities affect over 57 million people, making up 20% of all disability claims and resulting in an economic impact of over $213 billion annually.³

Among the leading causes of musculoskeletal disabilities are injuries and trauma to skeletal muscle, particularly volumetric muscle loss (VML)—a condition characterized by significant loss of skeletal muscle tissue, resulting in severe functional impairment and disability.⁴ VML commonly arises from high-impact trauma, including vehicular accidents, major falls, gunshot wounds, and explosive injuries, and is often associated with open fractures or compartment syndrome.⁵ While VML is most prevalent in military personnel due to battlefield injuries, it also affects civilians through gunshot wounds, severe road accidents, major falls, or tumor ablation procedures.

Skeletal muscle is a highly organized and complex tissue composed of long, cylindrical myofibers bundled into fascicles (Fig. 1A). While minor muscle injuries can heal through endogenous repair mechanisms, VML disrupts these processes, preventing functional muscle regeneration. This failure is primarily due to the loss of satellite cells (SCs) and the extracellular matrix (ECM), both essential for tissue repair and structural organization.⁶ In addition, the extensive loss of native muscle tissue eliminates critical regenerative cues needed to guide cellular responses during healing.⁷ Some studies suggest that following VML, a prolonged presence of macrophages in an intermediate (M1-to-M2) phenotype leads to excessive ECM deposition rather than muscle fiber regeneration, resulting in scar formation.^8,9 This scar/fibrotic tissue impedes SC migration and angiogenesis, compromising muscle repair. Moreover, scar tissue can obstruct the reformation of neuromuscular junctions (NMJs), which are often damaged in VML injuries^10–15 (Fig. 1B). Since NMJs are essential for muscle function and development, their disruption further exacerbates impaired regeneration. VML also results in the loss of mitochondria-rich skeletal muscle cells, leading to a decline in oxidative metabolic activity and overall metabolic imbalance. Since mitochondria function as calcium sinks, an increase in cytosolic calcium levels can overwhelm mitochondrial calcium buffering capacity, disrupting mitochondrial function and compromising cell viability; this cascade further exacerbates muscle regeneration deficits.¹⁶

FIG. 1.

Schematics showing the (A) structure of skeletal muscle and (B) neuromuscular junction.

Consequently, the pathophysiological events following VML create a highly challenging environment for endogenous muscle repair, resulting in severe loss of muscle strength and functionality. In severe cases, these deficits can lead to long-term disability or even necessitate limb amputation.^10–13

Treating VML injuries remains a significant challenge due to the complex biological processes involved in muscle regeneration. The current standard of care is autologous tissue transfer, or muscle grafting, which involves transplanting a muscle flap from a healthy donor site to the injured area.^4,10 However, this approach has several limitations, including limited donor tissue availability, risk of graft infections, and potential transplanted tissue necrosis.¹⁷ In addition, achieving full muscle restoration with adequate vascular integration and innervation remains difficult, often resulting in persistent functional deficits, reduced muscle strength, and impaired mobility.^18,19

To assist patients with mobility, advanced bracing strategies have been developed. However, these devices become a permanent part of a patient’s life and are primarily designed for lower extremity VML injuries, making them less suitable for upper extremity impairments. Meanwhile, rehabilitation approaches, although beneficial, are largely ineffective as a stand-alone treatment and are most effective when combined with physical therapy regimens following muscle grafting surgeries to enhance muscle function and strength.

To address these limitations, skeletal muscle tissue engineering (SMTE) has emerged over the past decade as a promising strategy. SMTE leverages bioengineered constructs to enhance muscle regeneration and restore function in VML injuries.²⁰ These constructs provide a three-dimensional (3D) support structure that mimics the native tissue microenvironment, promoting cell growth, differentiation, and functional integration.²¹ SMTE integrates biomaterial science, cellular and molecular biology, and bioengineering to drive the development of advanced bioengineered constructs for de novo skeletal muscle regeneration.²²

Therapeutic strategies in SMTE are categorized into in situ, in vivo, and in vitro approaches²⁰ (Fig. 2). In situ tissue engineering involves implanting acellular constructs at the injury site to stimulate endogenous regenerative mechanisms.²³ In vivo approaches use cell-laden constructs implanted directly into the injury site,²⁴ while in vitro strategies involve developing, culturing, and maturing tissue-engineered muscle grafts (TEMGs) before implantation.^20,23 In the past decade, notable developments have been made in all three strategies.^25,26 Notably, the SMTE field has seen significant advancements in biofabrication strategies, with 3D bioprinting emerging as a powerful tool for developing constructs replicating complex muscle anatomy, aiding the restoration of functional neo-muscle.

FIG. 2.

Schematic showing the strategies for skeletal muscle tissue engineering.

Given the substantial improvements in SMTE, this review provides a comprehensive overview of bioengineered constructs for VML. The discussion extends to the potential applications of artificial intelligence and machine learning (ML) in selecting biomaterials and optimizing construct manufacturing. Finally, the review highlights the current limitations, discusses the prospects within the SMTE field, and the essential advancements needed in regenerative medicine for VML injuries.

Regenerative Constructs in SMTE

Regenerative constructs for VML aim to restore damaged or lost muscle tissue by promoting regeneration and tissue growth. These constructs typically incorporate biomaterial-based scaffolds, stem cells, growth factors, and gene therapies to activate the muscle cells’ natural repair mechanisms or enhance the regeneration cascade of the transplanted cells. By providing a supportive environment that mimics native muscle architecture and composition, constructs facilitate the regeneration of muscle fibers. The ideal design of a construct suitable for functional muscle regeneration and some strategies to achieve the design are illustrated in Figure 3A. Figure 3B summarizes the biomaterials and cell types commonly used in SMTE-based constructs. The following sections discuss in detail the three main categories of constructs that have been developed to regenerate skeletal muscle.

FIG. 3.

(A) Chart showing the ideal conditions required for successful muscle regeneration and strategies to achieve these outcomes. (B) Biomaterials and cell types commonly used in skeletal muscle tissue engineering (SMTE)-based constructs.

Acellular constructs

Acellular constructs are developed using biomaterials designed to stimulate the endogenous repair mechanisms, forming new muscle fibers.²⁰ Specifically, these constructs are bioengineered with biophysical and/or biochemical cues to replicate the native ECM, creating a microenvironment that modulates host cell behavior and initiates the regeneration cascade.²⁷ One of the most significant advantages of this strategy is that the construct fabrication process is simple, eliminating the need to incorporate cells into the constructs, which leads to faster production, improved delivery, simplified storage, and reduced regulatory constraints compared with cell-laden constructs or TEMGs.²⁸

Selecting the right biomaterials is critical for developing effective regenerative acellular constructs. Biomaterials must possess specific mechanical, chemical, and biological properties that impact the strength, durability, degradation, and regenerative potential of the constructs. In SMTE, natural biomaterials such as collagen, fibrin, alginate, gelatin, chitosan, silk fibroin, hyaluronic acid (HA), laminin, agarose, and keratin are commonly used^29–32 (Fig. 3B). Among these, decellularized ECM (dECM), sourced from the tissues such as small intestinal submucosa and urinary bladder matrix is a prominent choice in SMTE. Table 1 provides an overview of studies that have used dECM as the primary material for acellular constructs in muscle regeneration. The table also includes other natural biomaterials used to make acellular constructs. Overall, natural biomaterials are appealing due to their bioactive components that mimic the native ECM, fostering cell growth, differentiation, and maturation.²⁸ Moreover, they are biodegradable, with their degradation by-products typically not exhibiting cytotoxic effects.⁴⁵ However, sourcing and preparing natural biomaterials can result in batch-to-batch inconsistencies, potentially triggering immune reactions.^46,47 In addition, natural constructs often suffer from low strength and durability, and they cannot be modified to enhance these properties.⁴⁸

Table 1.

Strategies for Developing Acellular Constructs for Skeletal Muscle Tissue Engineering

Type of construct—biomaterials	Manufacturing techniques	Important in vitro results	Important in vivo results	Ref.
Skeletal muscle—ECM and SIS scaffold	Decellularization	Myogenic cells survived and proliferated on muscle ECM scaffolds. Muscle ECM scaffolds had growth factors, glycosaminoglycans, and structural proteins.	Defect Model: Rat abdominal wall defect model. SIS scaffold exhibited more significant scaffold degradation than muscle ECM scaffold by 14 days. By 35 days postsurgery, the remodeling outcomes induced by muscle ECM did not significantly differ from those caused by SIS scaffold.	²⁷
Acellular skeletal muscle scaffolds	Decellularization	NA	Defect Model: Extensor digitorum longus muscle in the right leg of mice. The resected muscles were replaced with implanted acellular matrices, which developed into functional new muscles. These matrices facilitated the movement and specialization of host myogenic cells and allowed for the integration of nervous fibers, vascular networks, and the homing of SC.	³³
SIS-ECM scaffold	Decellularization	Scaffold-treated VML showed more β-III-tubulin and Nestin+ cells than untreated defects.	Defect Model: Mouse quadriceps (3 × 4 mm) defect. Scaffold treatment showed myotube formation at the center and edges of the defect/scaffold, with increased MHC+ nuclei indicating muscle remodeling up to 56 days. Scaffold treatment showed enhanced Fizz1+ macrophages in the injuries.	³⁴
(i) SIS-ECM, (ii) porcine SIS-ECM, and (iii) autologous tissue graft	Decellularization	NA	Defect Model: Rat abdominal wall defect model. The newly constructed muscle generated contractile forces similar to native tissue. Autologous tissue grafts contained collagenous connective tissue and adipose tissue, with fewer skeletal muscle islands than SIS-ECM, but showed comparable fatigue resistance to native muscle.	³⁵
Rat acellular muscle matrix	Decellularization	NA	Defect Model: VML in the rat TA muscle Untreated VML injury weighed 22.7% and 19.5% less than contralateral muscles at 2 months and 4 months after injury. Untreated VML injury also exhibited a reduced peak isometric tetanic force of 28.4% and 32.5% after 2 and 4 months.	³⁶
Decellularized skeletal muscle and minced muscle autograft	Decellularization and excised from VML defect	Minced muscle addition to decellularized skeletal muscle boosted MyoD expression by 21-fold compared with decellularized skeletal muscle alone and 37-fold compared with unrepaired VML.	Defect Model: VML (8 mm diameter × 3 mm depth) in the TA muscle of the rat. Scaffold improved peak contractile force to 81 ± 3% compared with 62 ± 4% in untreated VML controls. Scaffold repair increased muscle mass restoration to 88 ± 3%, compared with 79 ± 3% in untreated VML controls. Scaffold-treated muscles showed a significant decrease in collagen-dense repair tissue formation.	³⁷
Muscle-dECM	Decellularization	Scaffold-treatment resulted in a 21.6% functional improvement. Muscle band formation indicates effective integration.	Defect Model: Rat LD muscle model with VML. Scaffold treatment resulted in a new muscle band with excellent structural integration and increased muscle force by 21.6%.	³⁸
Muscle-dECM scaffold	Decellularization	Scaffold effectively decellularized. Despite decellularization, the scaffold promoted SC proliferation.	Defect Model: VML in rat TA muscle. Scaffold-treated muscles showed reduced functional deficits and improved recovery at 2 and 4 months postinjury. Untreated muscles displayed extensive fiber damage and remodeling, but scaffold-treated muscles showed fewer injury signs.	³⁹
UBM-ECM scaffolds and autologous muscle grafts	UBM scaffolds were derived from porcine urinary bladder tissue	NA	Defect Model: A VML injury in the tibialis anterior (TA) muscle. After 2 weeks, scaffold treatment showed dense cell infiltration and a mixed M1/M2 macrophage presence with a higher M1 response, while autograft repairs had higher TGF-β+ cell presence. After 8 weeks, the scaffold was remodeled into fibrotic scar-like tissue (80%) with limited new muscle formation and reduced macrophage response.	⁴⁰
Minced muscle grafts	Autologous muscle grafts excised from TA muscle.	Minced TA muscle grafts proliferated on collagen-coated dishes for up to 14 days. Desmin+ cells, indicative of SCs and myotubes, were observed after 5 and 14 days of culture.	Defect Model: A VML injury (10 × 7 × 3 mm) in TA muscle. At 16 weeks, the untreated group showed a 12–18% reduction in TA muscle weight compared with the untreated contralateral leg and an ∼10% increase in extensor digitorum longus muscle weight. The minced graft-treated group showed comparable TA muscle weights in both legs, with TA muscles being 11–21% heavier than those in the untreated group.	⁴¹
Minced muscle graft	Autologous muscle grafts derived from ubiquitin C-green fluorescent protein (GFP) mice	NA	Defect Model: VML injury in TA muscle of nude mice. Minced grafts (≈1 mm³)-treated VML injury restored ≈34% of the lost muscle fibers after 28 days. The number of donor-derived GFP+ fibers matched the estimated number of regenerated fibers across different mouse strains. However, there was no improvement in muscle strength at 28 days postinjury.	⁴²
UBM -ECM scaffold	Laminated vacuum pressing	NA	Clinical Trial: Five VML patients treated with scaffold showed dense neo-muscle tissue formation within 6 months. Patients showed a 25% force increase and 220% task improvement. Scaffold-treated patients in the quadriceps region showed functional improvements of up to 1820%.	²⁸
polypropylene mesh + TGF-β	Polypropylene mesh was rolled into a 5-mm-diameter cylinder, soaked in PBS, and loaded with TGF-β during implantation.	NA	Defect Model: VML injury in TA muscle. At 4 weeks, the functional fibrosis treatment group showed a higher collagen content in their tissue (36.3%) compared with the untreated control group (12.2%). Functional fibrosis-treated animals showed improved in vivo muscle function compared with untreated controls at 8 weeks postinjury.	⁴³
Autologous muscle graft	Autologous muscle graft excised from TA muscle	NA	Defect Model: Partial thickness circular VML defect (8 mm diameter × 3 mm depth) in TA muscle. After 2-week treatment, muscle mass and torque recovery were similar across all treated groups, but 0^o aligned autografts showed a 2.5-fold increase in MyoD and a twofold increase in Pax7 gene expression. After 12-week treatment, aligned autografts showed better mass and torque recovery compared with 45° and 90° misaligned autografts, while misaligned autografts had more fibrosis and noncontractile tissue.	⁴⁴

ECM, extracellular matrix; dECM, decellularized ECM; VML, volumetric muscle loss; SIS, small intestinal submucosa; UBM, urinary bladder matrix; SC, satellite cell; MHC, myosin heavy chain; TGF, transforming growth factor.

On the contrary, synthetic biomaterials offer tunable strength and durability, making them well-suited for SMTE applications. These materials can be engineered to achieve specific mechanical properties, such as stiffness, which can better mimic native skeletal muscle tissue.⁴⁹ Common synthetic biomaterials include poly lactic-coglycolic acid (PLGA), polyethylene glycol (PEG), polypropylene, poly lactic acid (PLA), polyglycolic acid (PGA), and polyurethane (PU) (Fig. 3B).^50–53 Synthetic materials offer better batch consistency and greater control over mechanical properties, but they are typically bioinert, lacking the bioactive components necessary to modulate cell behavior. To address this, synthetic materials often require functionalization or modification to enhance properties such as cell adhesion, proliferation, and differentiation. In the past two decades, hybrid biomaterials combining natural and synthetic components have been developed, offering improved mechanical properties, bioactivity, and biocompatibility.

Although acellular constructs show promise, they often lead to fibrosis rather than the formation of functional muscle fibers.⁵⁴ This is partly due to a prolonged M1 macrophage response, as shown in Figure 4. Aurora et al.⁴⁰ indicated that the macrophage response transitioned to the M2 phenotype over time but did not yield substantial muscle repair. Similarly, Corona and Greising⁵⁵ highlighted that acellular scaffolds limit the migration of SCs, such as Pax7+ cells, which are essential for functional muscle regeneration. While some studies suggest that acellular scaffolds can promote neo-muscle formation, substantial evidence indicates that they are insufficient for regenerating fully functional muscle in VML injuries, mainly because they do not adequately support vascularization and NMJ formation due to excess fibrosis.^10,55 Vascularization is crucial for supplying nutrients and oxygen to regenerating muscle tissue, ensuring the viability and function of newly formed muscle fibers. Muscle regeneration is limited without proper blood vessel formation, leading to impaired healing and reduced functional outcomes.^56,57 Reinnervation is equally essential, as the formation of NMJs enables the muscle fibers to contract and function effectively. Muscle regeneration fails to integrate with the nervous system without functional innervation, impairing mobility and strength.⁵⁸ Vascularization and reinnervation are key to restoring the structural integrity of muscle tissue and its functionality, ensuring that newly formed muscle fibers can contract and contribute to movement.⁵⁹ Future research should focus on developing next-generation acellular constructs that enhance SC migration, vascularization, and NMJ formation at the injury site. In addition, constructs should biodegrade at a rate that supports muscle remodeling in the injured region.

FIG. 4.

The porcine urinary bladder matrix (UBM) scaffold induced a prolonged proinflammatory (M1) macrophage response, as shown by immunohistological staining for pan macrophages (CD68+, red), M1 macrophages (CCR7+, CD86+, green), and M2 macrophages (CD163+, CD206+, green) in the tibialis anterior (TA) muscle defect repaired with the UBM scaffold. The interface and center of the repair were analyzed at the 2-week mark, revealing the scaffold exhibiting a dominant M1 response that persisted longer than expected, thereby hindering muscle regeneration. The defect was analyzed using immunohistological staining for pan macrophages (red) (CD68+), M1 macrophages (green) (CCR7+: a–f; CD86+: g–l), and M2 macrophages (green) (CD163+: m–r; CD206+: s–x). White dashed lines indicate the approximate interface along the remaining muscle mass (*) and the porcine UBM scaffold. Scale bars = 50 μm (magnification: 400×). Quantitative analyses of the spatiotemporal macrophage response are also shown (i–iv). Two regions were included in the analysis: the interface of the injured muscle and respective repair and the center of the respective repair. Means ± SEM. n = 3 muscles/group. *p < 0.05 versus UBM interface (2 weeks); ^§p < 0.05 UBM center (2 weeks); ^#p < 0.05 versus UBM interface (4 weeks); ^£p < 0.05 versus UBM interface (8 weeks). X indicates that no measurements were made for the autograft at the 4-week time point. Figure reprinted with permission from authors.⁴⁰

In summary, acellular constructs offer key advantages, including immune compatibility, off-the-shelf availability, and faster production due to their simple fabrication process. However, they lack the cellular components necessary for immediate regeneration, leading to slower integration and functional recovery. In addition, without bioactive modifications, these constructs are prone to scar tissue formation, hindering muscle regeneration.

Cell-laden constructs

Cell-laden constructs offer a promising approach to functional muscle restoration by introducing transplanted cells in the VML region. Given that VML injuries result in the loss of SCs in the surrounding tissue, incorporating exogenous cells into bioengineered constructs can facilitate tissue regeneration and remodeling.^29,55 The transplanted cells compensate for the lost or damaged muscle cells and contribute to regeneration by modulating the microenvironment through paracrine signaling. This signaling helps recruit host cells, regulate immune responses, and promote myogenesis, thereby accelerating the formation of new muscle fibers.^60,61 Furthermore, biomaterial-based delivery systems or constructs can influence cell fate by maintaining stemness and directing differentiation, ultimately aiding in the formation of functional skeletal muscle.

Several studies have demonstrated the superior regenerative potential of cell-laden constructs compared with acellular scaffolds. For instance, Conconi et al.⁶² engineered constructs seeded with autologous myoblasts and compared their efficacy against acellular scaffolds in a rodent model. Over 30 days, the myoblast-seeded constructs exhibited substantial vascularization and retained structural and functional integrity for an additional month, whereas the acellular constructs were encapsulated by fibrotic tissue. In another study, Qiu et al.⁶³ investigated using porcine heart ECM combined with human umbilical cord mesenchymal stem cells (MSCs) for treating tibialis anterior muscle defects in rodents. Constructs seeded with MSCs exhibited superior functional recovery, as evidenced by increased isometric torque measurements at 4 and 8 weeks postimplantation compared with MSCs or ECM alone. The MSC-seeded constructs also facilitated macrophage polarization toward the M2 phenotype, activated SCs, and promoted robust myofiber formation. After 8 weeks, the ECM+MSC-treated group demonstrated complete muscle regeneration with restored muscle architecture (Fig. 5B–D), whereas treatments with only MSCs or ECM resulted in only partial mitigation of muscle atrophy. In addition, mechanical function recovery was significantly enhanced in the ECM+MSC-treated group, as shown by improved force generation at 1 month (Fig. 5E) and 2 months (Fig. 5F) postimplantation.

FIG. 5.

Gross morphology and functional evaluation of TA muscle after volumetric muscle loss (VML) injury and repair. (A) Gross appearance of VML-injured muscle in the four groups: Control, extracellular matrix (ECM)-treated, mesenchymal stem cell (MSC)-treated, and ECM+MSC-treated groups. (B–D) Representative central thickness of TA muscle (B) and widths of the distal (C) and proximal (D) thirds of the TA muscle in each group. ECM-, MSC-, and ECM+MSC-treated group show partial recovery in central thickness and distal width of TA muscle. (E, F) Mechanical function evaluation by ES-induced contractions observed in each group presented at (E) 1 month and (F) 2 months. (G) Mean body weight of four groups over the 8 weeks, showing no significant differences among the groups at any time point. n = 6 per group. Data shown as mean ± SD. *p < 0.05; **p < 0.01; NS, not significant (p > 0.05). Overall, the ECM+MSC scaffold promoted functional recovery and muscle regeneration more than other treatment groups. Figures reprinted with permission from authors.⁶³

Cell-laden constructs also actively support revascularization and immunomodulation in addition to forming new muscle fibers. This can be achieved by incorporating multiple cell types in the constructs and transplanting them to the injury site. For instance, various cell types, including C2C12 muscle cells, skeletal myoblasts, adipose-derived stem cells, MSCs, endothelial cells (ECs), and regulatory T cells, have been incorporated into constructs to enhance neovascularization, promote functional integration, and improve regenerative outcomes.^64–66 However, very few studies focus on reestablishing the NMJs with cell-laden constructs, critical for improving functional recovery in the newly formed muscles. Table 2 summarizes key studies that have utilized cell-containing constructs for VML treatment.

Table 2.

Cell-Laden Construct Strategies for Volumetric Muscle Loss Treatment

Cell type	Type of construct—biomaterials used	Manufacturing techniques	Important in 'vitro results	Important in vivo results	Ref.
C2C12 myoblasts	Fibrin-based scaffold modified with agrin	Electrospinning	Myoblasts cultured on microfiber bundles with soluble agrin showed increased AChr clustering Chemically tethered agrin exhibited AChr clusters only within 10 μm from the substrate surface.	Defect Model: VML in rat TA muscle. Scaffold pretreated with soluble or tethered agrin resulted in increased NMJ formation, regeneration of myofibers, vascular infiltration, and neural infiltration compared to the untreated agrin control.	⁶⁷
	Hyaluronic acid and chondroitin sulfate hydrogel	Hydrogel Formulation and Gelation	Hydrogel supported satisfactory C2C12 myoblast proliferation and myogenic differentiation, expressing markers such as MyoD, MyoG, and MYH8.	Defect Model: Mice quadriceps (–4-mm-diameter) defect. Hydrogel scaffolds integrated well into the defects, facilitating Pax7+ SC migration, de novo myofiber formation, angiogenesis, treadmill performance, and innervation with minimal scar formation.	⁶⁸
	Microthread and fibrin gel	Microthread extrusion	Myoblasts conditioned with HGF-coated fibrin microthreads showed more Ki67+ cells after 24 h. Proliferation persisted for up to 48 h with HGF-coated EDCn microthreads but diminished by 96 h.	Defect Model: VML (critical sized) in the central region of TA muscle. The structure of the microthread scaffolds supported new myofibers to grow into the wound site, whereas fibrin gel scaffolds failed to support functional regeneration.	⁶⁹
T regulatory cells	Autologous minced muscle	Autologous minced muscle excised from TA muscle	NA	Defect Model: VML injury in TA muscle of Sprague Dawley rat. Interleukin-10 treatment improved muscle strength by 82% compared with the untreated control and by 170% compared with the PBS control. Fourteen days postinjury, interleukin-10, muscle hypertrophy, and lymphocyte signaling pathways were activated.	⁷⁰
SCs, myogenic precursors, and endothelial cells	Hyaluronic acid—photoinitiator hydrogel scaffolds	Photopolymerization	SCs and muscle progenitor cells were embedded in a hyaluronic acid—photoinitiator hydrogel.	Defect Model: Partially ablated TA muscle of Male C57BL/6J mice. Scaffold-engrafted with SCs improved muscle structure and neural and vascular networks compared with hydrogel and hydrogel with MPCs. SC-laden scaffold enhanced 20–30% functional recovery over MPC-laden scaffold.	⁷¹
Muscle progenitor cells	Keratin hydrogel and bladder acellular matrix scaffolds	The bladder acellular matrix was synthesized by decellularization of porcine bladder tissue. Keratin-based hydrogel was used to encapsulate MPCs.	NA	Defect Model: VML injuries in rat TA muscle. Twelve weeks postinjury, scaffold treatment with growth factors improved functional recovery by up to 90% compared with untreated control. Fibrosis was reduced in keratin-treated constructs notably.	⁷²
HUVECs, MSCs, NIH-3T3 fibroblasts	Collagen and methacrylate collagen hydrogel scaffold	UV photocrosslinking of hydrogel	Improved cell viability was observed with varied photoinitiator concentrations. MSCs and HUVECs were encapsulated effectively. Vascular network formation achieved.	Defect Model: VML (4 × 3 mm) quadriceps defect in mice rectus femoris and vastus intermedius muscles. The construct treatment helped in effective muscle repair and revascularization.	⁷³
ADSCs	SIS, acellular dermal matrix (ADM), and a composite of collagen–chondroitin sulfate–hyaluronic acid (Co–CS–HA).	Constructs were fabricated by mixing collagen, chondroitin sulfate, and hyaluronic acid, followed by ADSC incorporation.	ADSCs displayed better proliferation rates and maintenance of cellular viability when cultured on SIS and ADM compared with Co–CS–HA constructs.	Defect Model: 1 cm full-thickness skin excision on the dorsum of 8-week-old female mice, exposing subcutaneous muscle. Enhanced angiogenesis was observed in all ADSC-seeded ECM scaffolds. ADSC-laden scaffolds outperformed acellular scaffolds in terms of wound healing with minimal inflammation.	⁷⁴

AChR, acetylcholine receptor; SC, satellite cell; YAP, yes-associated protein; HGF, hepatocyte growth factor; HUVECs, human umbilical vein endothelial cells; MSCs, mesenchymal stem cells; ADM, acellular dermal matrix; SIS, small intestinal submucosa; ADSCs, adipose-derived stem cells; dECM, decellularized extracellular matrix; ECM, extracellular matrix; VEGF, vascular endothelial growth factor.

While cell-seeded constructs significantly enhance muscle recovery by improving cell engraftment and stimulating regeneration post-VML injury, several challenges remain. A primary limitation is maintaining long-term cell viability and function within the implanted construct. In addition, achieving a balance between multiple incorporated cell types—such as myogenic, endothelial, and neural cells—is complex yet necessary for promoting angiogenesis, myofiber maturation, and NMJ formation. Future research should focus on developing biomaterials capable of sustaining cell encapsulation while guiding cellular behavior toward the rapid restoration of vascularized and innervated muscle.

Overall, cell-laden constructs represent a transformative approach to skeletal muscle regeneration, offering active SCs that support myogenesis, angiogenesis, and faster integration with host tissue. These constructs better mimic native skeletal muscle architecture and can be customized for VML injuries. However, challenges such as limited cell viability, potential immune rejection, and the complexities of biomanufacturing and storage must be addressed to maximize their clinical potential. Future advancements should prioritize scalable, biocompatible, and functionalized constructs that ensure long-term cell survival and orchestrate a regenerative response.

Prematured tissue constructs/TEMGs

In this strategy, synthetic muscle grafts are developed in vitro and then implanted into the defects. This process begins with incorporating cells into 3D constructs, supplemented with relevant growth factors and cultured under controlled conditions to promote cellular expansion and differentiation.⁷⁵ Through a combination of biochemical and biophysical stimuli, these cells are guided to mature into myotubes or myofibers, forming TEMGs capable of integrating with host tissue postimplantation.⁷⁶ Table 3 highlights key studies that have developed TEMGs for SMTE.

Table 3.

Tissue-Engineered Muscle Graft Strategies for Volumetric Muscle Loss Treatment

Cell type	Type of construct—biomaterials used	Manufacturing techniques	Important in vitro results	Important in vivo results	Ref.
Myogenic precursors cells (MPCs) and fibroblasts	Cell coculture system	MPCs and fibroblasts were cocultured in a hydrogel-based 3D construct	MPCs with fibroblasts showed higher proliferation compared with MPCs alone. The myogenin+ cells increased with higher fibroblast numbers in the constructs	Defect Model: Quadriceps muscle injury By day 7, 25% of fibers were damaged, strongly correlating with fibroblast count. At day 30, 42% of fibers displayed embryonic /neonatal myosin, and a strong correlation with fibroblast numbers.	⁷⁷
Satellite cells (SCs)	Nuclear factor YA (NF-YA) gene construct	Lentiviral transduction	NF-YA knockout cultures exhibited an increased proportion of proliferating Pax7+/MyoD+ cells and a reduced number of quiescent Pax7+/MyoD− cells. The presence of higher EdU+/Pax7+ cells in NF-YA knockout SCs indicated stimulated proliferation.	Defect Model: VML in the rat TA muscle. NF-YA is found in the nuclei of regenerating muscles and is most abundant in SC. Mice lacking NF-YA showed severe muscle damage and persistent cell infiltration after cardiotoxin injection. These mice exhibited decreased Pax7+ cells, increased apoptosis, and SC activation.	⁷⁸
Spinal motor neurons and myocyte (MN-MYO)	Polycaprolactone (PCL) aligned nanofiber scaffolds	Electrospinning	MN-MYO coculture on nanofiber scaffolds led to neuromuscular structures aligned along the nanofibers, eventually forming NMJs. MN promoted myocyte fusion, resulting in a higher myocyte fusion index compared with only myocyte cultures.	Defect Model: VML injury (1.0 cm × 0.7 cm × 3 mm) in rat TA muscle. Preinnervated scaffolds showed a higher density of Pax7+ SC near the injury area compared with control. MN-MYO scaffold promoted revascularization, exhibited more AchR clusters near the injury area, and had higher percentages of mature NMJs than control.	¹³
Human umbilical vein endothelial cells (HUVECs) and C2C12	Bioactive glasses of silicate, borate, and aluminoborate compositions	Bioactive glass compositions were prepared by melting reagent-grade chemicals and casting into discs	The aluminoborate glass promoted faster migration and better tube formation of HUVECs compared with the other groups. Immunofluorescence imaging showed that aluminoborate glass significantly stimulated VEGF formation, supporting angiogenesis.	Defect Model: VML defect in Sprague-Dawley (S-D) rat TA muscle. All three bioactive glass compositions supported angiogenesis. C2C12 cells treated with glass extracts exhibited increased gene expression for VEGF, IGF-1, and CX43.	⁷⁹
Primary SCs	dECM combined with polycaprolactone (PCL)	Electrospinning	NA	Defect Model: VML injury in rat hindlimb muscles VEGF increased and IL-6 production decreased significantly by day 4 on the PCL scaffolds. At day 4, MyoD expression was significantly higher in aligned PCL-d-ECM scaffolds than in aligned PCL, while myogenin and α-actinin expression was lower on aligned PCL-dECM scaffolds.	⁸⁰
Murine skeletal myoblasts (C2C12) and human microvascular endothelial cells	Nanofibrillar collagen scaffolds	Facile shear-based extrusion method	Myotube length was longer in aligned scaffolds compared with randomly oriented scaffolds. Endothelialized muscle from aligned scaffolds exhibited greater contractile strength and coordinated movement than those from randomly oriented scaffolds.	Defect Model: 20% TA muscle removal in NOD SCID mice. Aligned scaffolds showed improved cell viability alignment, organization, and increased vessel and myofiber density compared with randomly oriented scaffolds.	⁸¹
C2C12 mouse myoblasts	Magnetic-controlled short nanofiber hydrogel scaffold	Advanced coaxial electrospinning	Orbicular muscle scaffolds showed circular alignment of nanofibers, which C2C12 cells adhered to, replicating the circular muscle fiber arrangement. Bipennate muscle scaffolds, created with a central PCL stick as a tendon, had nanofibers aligned in opposite diagonal directions, similar to native bipennate muscles.	Defect Model: VML injury in rat TA muscle. One week after injection, immune cell infiltration and partial hydrogel degradation were observed in the aligned nanofibers/gel and gel groups. Aligned nanofibers/gel scaffold group showed the highest myofiber count and capillary density compared with the gel and blank groups.	⁸²
Muscle SCs and bone marrow-derived stem cells	Engineered skeletal muscle units (SMUs)	SMUs are created by isolating muscle SCs, followed by culturing and differentiating.	NA	Defect Model: VML injury in rat TA muscle. The muscle constructs developed myotendinous junctions, which allowed to produce contractile force similar to that of native skeletal muscle. The SMU-treated groups were structurally similar to native muscle fibers, with laminin-rich ECMs.	⁸³
Adipose-derived stem cells (ASCs) and L6 rat skeletal myoblasts	dECM with vascular pedicles	Perfusion decellularization of adipose tissue	NA	Defect Model: VML injury in rat TA muscle. dECM+ASCs+L6 group exhibited new muscle formation with the greatest improvement in muscle contraction strength, functionality, volume, and fiber size compared with other groups.	⁸⁴
Induced skeletal myogenic progenitor cells (iMPCs)	Muscle extracellular matrix (MEM) -modified PCL scaffolds	Thermal drawing with salt-leaching	The MEM process removed 97.4% of cells, preserving essential ECM components for muscle regeneration. The myogenic differentiation of the scaffold-treated group showed the highest myogenic cell fusion and the highest gene expression levels of MyoD1, MyoG, Itgb1, Itga7 compared with the untreated control group.	Defect Model: VML model (75% muscle loss) within mouse quadriceps femoris (QF) muscles Scaffold treatment showed early signs of muscle regeneration, reduced fibrosis, and resulted in the highest AchR and α-SMA-+ arteriole expression, longest stride length, highest endurance, and restored hind limb grip strength.	⁸⁵

EdU, 5-ethynyl-2’-deoxyuridine; NMJ, neuromuscular junction; IGF-1, insulin-like growth factor 1; CX43, Connexin 43.

The scaffold or construct within TEMGs is crucial in establishing a microenvironment conducive to cell survival, differentiation, and functional tissue regeneration. Ideally, these constructs should be biodegradable, allowing progressive cellular remodeling into mature muscle fibers—a fundamental requirement for effective muscle regeneration.^86–90 Natural biomaterials, particularly dECM, are promising candidates for TEMGs due to their ability to retain native biochemical and structural cues, which support cellular attachment, differentiation, and tissue organization.^91–93 Specifically, dECM preserves essential bioactive components and ECM architecture, making it an ideal choice for engineering complex tissue constructs requiring high bioactivity.⁹⁴

Alternatively, synthetic polymers such as polycaprolactone (PCL), PLA, PGA, PLGA, and PEG are widely utilized due to their tunable mechanical properties and controlled degradation rates.⁹⁵ Composite materials that integrate natural and synthetic biomaterials are increasingly used to enhance the mechanical integrity and bioactivity of TEMGs. For example, alginate-PEG and gelatin-PCL composites have demonstrated favorable mechanical and biological properties, making them promising candidates for TEMGs.^96,97

Recent advancements in TEMGs have focused on integrating vascular and neural components to improve muscle functionality. Prevascularized and preinnervated TEMGs have demonstrated superior regenerative potential by enhancing vascular integration and NMJ. Das et al.¹³ utilized an in vitro coculture strategy by combining spinal motor neurons with skeletal myocytes on aligned nanofibrous scaffolds to develop innervated TEMGs. These preinnervated constructs successfully formed aligned neuromuscular bundles, promoting myocyte fusion and maturation. Within just 7 days of coculture, functional NMJs were observed on muscle fibers (Fig. 6A–a′), and the presence of motor neurons significantly increased the myocyte fusion index compared with constructs containing only myocytes (Fig. 6B, C). Upon implantation in a rat VML model, these preinnervated TEMGs facilitated muscle SC migration, enhanced microvascularization, and promoted NMJ formation near the injury site, leading to improved functional recovery. As innervation is a critical factor in skeletal muscle development, maturation, and functional control, TEMGs should either be preinnervated during fabrication or possess the capacity to support effective neural integration postimplantation.^58,98

FIG. 6.

Innervation of myocytes and the impact of motor neurons on myocyte maturation in vitro. (A) Rat spinal motor neurons were introduced onto a layer of myofibers differentiated on an aligned nanofiber sheet for 7 days, followed by coculture for an additional 7 days. This process resulted in the innervation of skeletal myofibers. Scale bar: 100 µm. (a′) A higher magnification view of the region marked by the white box reveals structures colabeled with the presynaptic marker synaptophysin and AchR clusters identified by Bungarotoxin staining. These observations indicate the formation of mature neuromuscular junctions (NMJs) in vitro, as highlighted by white arrows. Scale bar: 50 µm. (B) Myocytes cocultured with motor neurons (MN-MYO) displayed enhanced fusion and bundling compared with myocyte monocultures (MYO). Scale bar: 100 µm. (C) The Myocyte Fusion Index (MFI), calculated from multiple cultures (n ≥ 6), showed a significant increase in MFI in cocultures with motor neurons. Statistical analysis revealed a highly significant difference (p ≤ 0.0001****). Error bars indicate the standard error of the mean. Figures reprinted with permission from authors.¹³

Despite their high regenerative potential, TEMGs face several limitations. While they can generate contractile force, the magnitude is typically lower than that of native muscle tissue.²⁰ One of the significant barriers to TEMG efficacy is the lack of a vascular network, which restricts oxygen and nutrient diffusion, particularly in densely packed, metabolically active muscle cells. Since TEMGs often exceed the diffusion limit, developing a vascular network is essential to sustain cell viability.⁹⁸ In addition, the prolonged in vitro culture required for TEMG maturation presents challenges in clinical scalability, cost-effectiveness, off-the-shelf availability, and regulatory compliance.⁹⁹

The variability in biomaterials, cell types, culture conditions, construct dimensions, stimulation protocols, bioreactors, and preclinical models also contributes to inconsistent outcomes in TEMG-based skeletal muscle regeneration.⁵⁸ For effective VML repair, engineered constructs must incorporate both vascularization and innervation to support cell survival, metabolic activity, and functional contractility. Prevascularization strategies, such as EC coculture and the delivery of angiogenic growth factors, vascular endothelial growth factor (VEGF), fibroblast growth factor, platelet-derived growth factor, have enhanced capillary formation and vascular integration.¹⁰⁰ On the contrary, innervation strategies, including motor neuron coculture, electrical stimulation (E-Stim), topographical guidance cues, and neurotrophic factor delivery (BDNF, NGF), facilitate NMJ formation and enhance functional recovery.¹⁰¹

By integrating vascular and neural components into engineered muscle grafts, TEMGs can more effectively restore muscle function following VML. Future research should prioritize scalable, biomimetic strategies that ensure long-term cell survival, enhance functional integration, and accelerate clinical translation.

3D Bioprinted Constructs in SMTE

Despite advancements in conventional manufacturing techniques, replicating the structural and compositional complexity of the ECM remains a significant challenge. In addition, precisely arranging cells within a 3D construct to promote myofiber alignment, neo-vascularization, and innervation is complex using traditional fabrication approaches. Bioprinting, an advanced form of 3D printing that integrates living cells within biomaterial-based inks, enables the precise deposition of cells and ECM components in spatially defined regions. This technique facilitates the rapid fabrication of complex, tissue-mimetic structures, closely replicating the in vivo microenvironment of skeletal muscle.^102,103 Bioprinting methods are classified into inkjet-based, extrusion-based, and laser-assisted techniques, each offering distinct advantages and limitations.¹⁰⁴ Among these, extrusion-based bioprinting (EBB) is the most widely used in SMTE due to its ability to develop anisotropic, multicellular 3D constructs that mimic the architecture of native muscle tissue.

Hydrogels and their role in bioprinting for SMTE

Hydrogels play a critical role in bioprinting by serving as bioinks that encapsulate cells and provide a supportive microenvironment for their survival and differentiation. In SMTE, commonly used hydrogel materials include alginate, gelatin, fibrin, collagen, and dECM, selected for their biocompatibility, biodegradability, and ability to mimic the native ECM.¹⁰⁵ One of the most critical factors governing hydrogel performance in bioprinting is viscosity, which determines the structural integrity and printability of the constructs. EBB, for instance, requires higher viscosity hydrogels to ensure the stability of printed structures while maintaining optimal cell viability.^106,107 Carefully selecting hydrogel properties is crucial to achieving high-resolution constructs with appropriate mechanical support and bioactivity.

Bioprinting for cellular alignment and functional muscle regeneration

Beyond material selection, topographical and biophysical cues introduced during bioprinting significantly influence cellular behavior, alignment, and differentiation. By guiding cell orientation and fiber formation, bioprinting enables the fabrication of highly organized, anisotropic muscle constructs that closely resemble native skeletal muscle architecture.¹⁰⁸ It can also help fabricate constructs with spatial and temporal gradients of growth factors, such as VEGF, for enhanced vascularization and repair.¹⁰⁹ Table 4 overviews various cell-alignment techniques, emphasizing strategies to enhance muscle regeneration through controlled cellular organization.

Table 4.

Various Cell-Alignment Techniques for the Engineered Construct for Volumetric Muscle Loss Treatment

Cell alignment technique	Important result	Ref.
Aligned electrospun fibers and topographical cues	Provide structural guidance that promotes muscle cell elongation, fusion, and myotube formation, enhancing muscle fiber organization and regeneration.	¹¹⁰
Anchor-based gel compaction	Induces myotube alignment by applying mechanical constraints during gel compaction.	¹¹¹
Shear stress in extrusion-based bioprinting	Shear forces during extrusion align hydrogel fibers, guiding directional cell spreading and enhancing anisotropic tissue formation.	¹¹²
Electrical stimulation	Promotes myoblast orientation, enhances myogenic differentiation, and accelerates neuromuscular junction (NMJ) formation.	⁷⁶
Microgrooved substrates	Enhances myotube alignment and maturation by providing parallel microgrooved surfaces.	¹¹³
Ultrasound-induced cell patterning	Creates ordered cellular patterns by utilizing ultrasound standing waves, promoting muscle fiber orientation.	¹¹⁴
3D-printed aligned fiber scaffolds	Provides a highly organized scaffold with aligned fibers, mimicking native muscle structure.	¹¹⁵
Hydrogel Stiffness Gradient	Stiffness anisotropy directs C2C12 myotube alignment via contractility-driven collective dynamics, mimicking native skeletal muscle structure.	¹¹⁶

Several innovative bioprinting techniques have also been developed to enhance structural integrity and functional integration within engineered skeletal muscle constructs, as mentioned below.

Multimaterial printing

Figure 7A shows this approach combines sacrificial and supportive biomaterials, such as PCL pillars coprinted with gelatin-based sacrificial materials, to create spatially controlled architectures.

FIG. 7.

Extrusion-based 3D bioprinting for SMTE with enhanced structural stability. (A) Multimaterial extrusion printing with sacrificial and support structures for SMTE. To form large muscle constructs, polycaprolactone (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). (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 was demonstrated to be biocompatible (iv). Rolling of the composite sheets formed a large-scale biomimetic muscle structure (v). (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 with the cells infiltrated into a sponge scaffold or those encapsulated in bulk hydrogel through casting. Figures reprinted with permission from authors.¹⁰⁶

Layered scaffold fabrication

Figure 7B shows that a combination of 3D printing and electrospinning is used to create PCL-based scaffolds, followed by the extrusion of a cell-laden bioink, which is then rolled into a tubular construct for mechanical reinforcement and muscle fiber alignment.

Embedded bioprinting

Figure 7C shows that freeform reversible embedding of suspended hydrogels (FRESH) involves bioprinting within a gelatin microgel support bath, which preserves ECM bioink integrity during crosslinking. This method enhances fiber-rich constructs, improving cell viability, nutrient diffusion, and myogenic differentiation.¹⁰⁶

State-of-the-art bioprinted constructs

Bioprinting offers unparalleled control over the tissue microenvironment, enabling the precise integration of multiple cell types, including myogenic progenitors, fibroblasts, and ECs, within a single construct.¹⁰⁶ This multicellular approach, explored in the last few years, is crucial for regenerating functional skeletal muscle, ensuring cell–cell interactions, vascularization, and ECM remodeling. By recapitulating the biochemical and mechanical properties of native muscle tissue, bioprinted constructs significantly improve cell viability, structural organization, and functional restoration, making bioprinting an indispensable tool for SMTE.¹¹⁷ Table 5 highlights key studies using EBB to develop constructs for VML treatment.

Table 5.

Studies That Used Extrusion-Based 3D Bioprinting to Develop Therapeutics for Skeletal Muscle Tissue Engineering

Biomaterials used	Cell type	Important in vitro results	Important in vivo results	Ref.
Gelatin methacryloyl (GelMA) and fibrinogen-based scaffold	Muscle precursor (C2C12) cells	Scaffolds induced significant cell alignment, with 26.2 % of cells aligning within ±10° orientation, compared with 12.7% in the gel group. MHC staining after 9 days revealed extensive myotube formation and elongation in the aligned scaffolds.	Defect Model: VML injury (15 × 10 mm × 5 mm) in rat TA muscle. Scaffold-treated group showed lower inflammation, significant microvessel formation, and newer myofiber formation compared to the gel group. After six months, scaffold treatment showed better physical activity and myo-physiological response compared to the gel groups and untreated control.	¹³⁹
Sodium alginate and PEG-fibrinogen scaffold	C2C12 cells	Over 21 days, myotubes showed sarcomerogenesis, resulting in striated myofibers indicative of advanced muscle maturation and functionality. Immunofluorescence analysis showed that >90% of myotubes were aligned in parallel with less than a 10° deviation, and the average length of myotubes in the constructs was 346 μm.	Defect Model: Subcutaneous implantation in the back of immunocompromised severe combined immunodeficiency (SCID) mice. The scaffold treatment resulted in the formation of organized muscle tissue after 28 days. Histological analysis showed well-oriented MHC+ myofibers with denser organization compared with bulk-hydrogels.	¹⁴⁰
GelMA and hyaluronic acid methacryloyl bioinks	C2C12 cells	Ti3C2Tx MXene nanoparticles (NPs) positively affected cell growth and differentiation, with significant increases in F-actin spreading, number of nuclei, and MHC expression in cells cultured within bioinks. qRT-PCR analysis showed significant upregulation of myogenic markers (MyoD, PGC-1α, MyoG, Myh4) and downregulation of myostatin in the bioink group.	Defect Model: VML injury in TA muscle of male C57BL/6 mice. At day 7, bioprinted scaffolds resulted in significant increases in muscle mass and fiber diameter, with low inflammation. Grip strength tests showed a 64.83% improvement in restoration of motor function in the GelMA and hyaluronic acid group and 84.27% in the bioink group, compared with nontreated controls.	¹⁴¹
GelMa-based hydrogel	C2C12 cells	The precultured cell-laden GelMa bioink demonstrated full alignment of C2C12s with the printing direction, resulting in improved myotube alignment and maturation compared with the standard bioink.	NA	¹⁴²
Skeletal muscle dECM and vascular dECM bioink	Human skeletal muscle cell	The 3D cell-printed muscle constructs showed lower hypoxia and higher cell viability (91.5%) compared with dECM hydrogels (34.5%) and sponges (87.1%). Printed scaffolds showed better myotube formation in the constructs, with densely packed and aligned myotubes throughout the structure.	Defect Model: VML injury (≈ 40% ablation of the TA muscle mass) in Sprague-Dawley rat. The 3D-printed muscle constructs showed minimal fibrosis, organized de novo muscle formation, and enhanced contractile force recovery (71% of uninjured tissue function) compared with the dECM sponges and hydrogels.	¹⁴³
Gelatin and PCL	hMPC	Bioprinted constructs showed high cell viability (86.4%) on day 1 and maintained it throughout the culture period, while nonprinted constructs showed lower viability and suffered from cell death by day 5. By day 7, bioprinted constructs showed an 11.53-fold increase in MHC+ myofibers compared with nonprinted constructs, indicating successful differentiation.	Defect Model: VML injury (10 × 10 mm × 3 mm) in rat TA muscle. Bioprinted constructs integrated with MHC+ myofibers and HLA+ cells after 4 and 8 weeks postimplantation. By 8 weeks, muscle force restoration reached 82% of the normal TA muscle, and muscle weight recovery was up to 82.5% of the contralateral TA muscle.	¹⁴⁴
(i) GelMA, (ii) GelMA xanthan gum (GelXA) FIBRIN, and (iii) FIBRIN bioink	C2C12 murine myoblasts	FIBRIN and GelX hydrogels both showed high cell viability throughout the culture period. MyoD and MCK gene expressions were significantly higher in FIBRIN 3D constructs than controls at 21 and 28 days. Myogenic gene expression was upregulated at 7 and 14 days compared with 2D samples, with no significant differences at 21 days.	NA	¹⁴⁵
PEG-Fibrinogen	Murine mesoangioblasts and hMSCs	Murine mesoangioblasts in bioprinted constructs showed significant muscle fiber differentiation over 30 days. Muscle fibers in bioprinted constructs were more organized, with an average fiber length twice as long as those in bulk constructs, and displayed greater alignment, with a 3.5-fold higher coherency than bulk constructs.	Defect Model: VML injury in TA muscle of SCID/Beige mice. 3D-bioprinted constructs resulted in complete tissue recovery in the treated TA muscle with proper vascularization and innervation and the development of NMJ and Pax7-positive SCs. Human muscle-derived MSC-laden constructs showed MyHC+ fibers, lamin A/C+ human-specific nuclei, and Pax7+ SCs in the regenerated area.	¹⁴⁶
(i) (Gel-MA)/PVA (ii) dECM-MA, and (iii) dECM-MA/PVA bioinks	hMPCs	At 14 and 21 days, Gel-MA/PVA and dECM-MA/PVA constructs formed aligned multinucleated myofibers, whereas the dECM-MA construct without PVA did not show aligned myofibers.	Defect Model: VML injury (10 × 7 mm × 3 mm) in TA muscle of Rowett rat. Tetanic muscle force recovery was the highest in dECM-MA/PVA constructs. dECM-MA/PVA also resulted in organized myofiber and NMJ formation. Vascularization was initially higher in dECM-MA and dECM-MA/PVA constructs but declined at 8 weeks, while Gel-MA/PVA showed gradual vascular buildup.	¹⁴⁷

*F2.5G5 and F5G5, concentration ratio of fibrinogen and GelMA as 2.5 wt% : 5 wt% and 5 wt% : 5 wt%.

ColMA, collagen methacrylate; hASCs, human adipose Stem Cells; hMPCs, human muscle precursor cells; PEG, polyethylene glycol; MHC, myosin heavy chain; dECM, decellularized extracellular matrix.

ML in SMTE

SMTE often faces challenges in developing constructs or TEMGs that closely mimic the innate muscle anatomy. ML addresses these challenges by automating data analysis and predictive modeling to optimize construct design and tissue regeneration.¹¹⁸ ML includes supervised, unsupervised, and deep learning (DL) approaches.¹¹⁹

Supervised learning involves training models using labeled data to make predictions, while unsupervised learning identifies patterns in unlabeled data.¹²⁰ DL is a subset of ML that utilizes neural networks with multiple layers.¹²¹ DL models include artificial neural networks (ANNs) and convolutional neural networks (CNNs), commonly used for image recognition and understanding natural language tasks.¹²² In SMTE, ML has influenced the optimization of bioprinting. By analyzing mechanical testing data, ML models can recommend optimal combinations of bioinks to replicate the ECM of muscle tissue. ML also aids in optimizing culture conditions for cell differentiation and proliferation and enhances bioactive piezoelectric constructs by predicting the effects of electrical and mechanical cues.^123–125 ML can help design constructs that accurately replicate mechanical properties such as stiffness, ensuring a better microenvironment for cell maturation.¹²⁶

Furthermore, ML algorithms can be used for in-process quality control in bioprinting. These processes often rely on data from integrated cameras or sensors within bioprinters. For instance, Jin et al.¹²⁷ used a pneumatic extrusion bioprinter to create two-layer structures and used a CNN to classify prints as “good” or “error” with high accuracy (Fig. 8A).¹²⁹ In another study, Gerdes et al.¹²⁹ incorporated infrared thermocouples to measure extrusion temperatures and trained ML models to predict print flaws and parameters such as line width (Fig. 8B). Similarly, Bonatti et al.¹³⁰ used CNNs for real-time monitoring of 3D printing with varying parameters. The prints were classified as “ok,” underextruded, or overextruded, and the CNN accurately identified defects, allowing for early termination of faulty prints and optimization of printing parameters (Fig. 8B). Sun et al.¹³¹ proposed a novel system for real-time monitoring of the Taylor cone and jet formation in electrospinning. An optical microscope was integrated into the setup to capture images during the printing process. These were then classified into eight categories, including one for successful deposition and seven for different types of errors. Using a CNN with majority voting on image sequences, the system effectively identified and corrected deposition issues in real-time (Fig. 8C). In another study, Ruberu et al.¹³² collected 177 samples to reproducibly form constructs using Bayesian optimization.

FIG. 8.

Machine learning (ML) application in in-process quality control (QC) for 3D bioprinting. (A) Camera imaging of two-layer patterns in extrusion-based bioprinting (EBB) detects inhomogeneities using a convolutional neural network (CNN) algorithm. (B) Dataset for 3D scaffolds obtained via EBB, with a CNN classifying “ok,” underextrusion, and overextrusion prints, optimizing print monitoring and error detection. (C) Monitoring jet formation quality in electrohydrodynamic (EW) printing with a digital microscope and a classification algorithm for jet regime analysis. The UBM scaffold, while used for VML treatment, has limitations in regenerating muscle tissue due to its restricted ability to support cellular responses and promote muscle regeneration over time. Figures reprinted with permission from authors.¹²⁸

ML can also help design constructs replicating mechanical properties such as stiffness, ensuring a better microenvironment for cell maturation.¹²⁶ For instance, predictive modeling can recommend the best combination of alginate, gelatin, and chitosan bioinks used in bioprinting to create constructs that closely mimic muscle tissue’s natural ECM.^125,133 Moreover, ML can predict how engineered tissues respond to mechanical stimuli such as stretch or compression and help design constructs that provide appropriate mechanical cues.

In terms of cell culture and tissue development, ML can aid in optimizing culture conditions for cell differentiation and proliferation.^123–125 ML can improve muscle models for in vitro and in vivo studies. By analyzing data from in vitro experiments, such as cell viability assays and mechanical testing, ML can predict the long-term behavior of engineered constructs in vivo. For instance, Sujeeun L.Y. et al.¹³⁴ used ML to predict the performance of electrospun polymeric constructs for skin tissue engineering. They trained six supervised learning algorithms to correlate properties with cell viability and achieved the highest accuracy with random forest regression. Lim J Y et al.¹³⁵ studied skeletal muscle gauge (SMG) as an imaging marker for sarcopenia. They developed an ML algorithm to predict SMG using clinical and inflammatory markers in colorectal cancer patients. The study of 1094 patients showed low SMG in 21.6% and 20.5% of patients in training and test sets, respectively. The LP-SMG model was an independent predictor of low SMG and outperformed individual clinical variables. It can be an effective tool for detecting sarcopenia without relying on CT scans during treatment.

Figure 9 summarizes the current and potential applications of ML in SMTE. ML can help predict cellular responses to constructs, design personalized constructs, and automate constructs’ quality control during fabrication. ML can uncover patterns that improve muscle regeneration understanding by analyzing large datasets. In the next few years, ML can advance therapeutics for VML, enhancing efficiency and therapeutic efficacy while reducing development timelines.

FIG. 9.

Chart showing the existing and potential applications of ML for the construct design for VML treatment.

Research Gap and Prospects for the Future

Despite significant progress in biomaterials, biofabrication, and cell-based therapies, SMTE faces significant challenges that hinder the clinical translation of therapies and promote functional muscle regeneration. Addressing these gaps is essential for developing scalable, biofunctional, and physiologically relevant muscle constructs capable of restoring vascular, neuromuscular, and mechanical integrity in VML injuries.

Enhancing NMJ formation and functional integration

One of the most significant obstacles in SMTE is achieving functional integration between newly formed muscle and host tissue. Specifically, reinnervation, essential for restoring NMJs, remains particularly difficult, resulting in newly formed muscle with low functional recovery and limited force generation. Current strategies to promote NMJ formation include coculturing myogenic and neural progenitors, delivering neurotrophic factors (e.g., brain-derived neurotrophic factor, nerve growth factor), and applying E-Stim to guide synaptogenesis. Future research should integrate ML-driven predictive modeling to optimize scaffold architectures and biochemical cues that facilitate NMJ formation, nerve infiltration, and functional recovery. Moreover, stand-alone “smart” (responsive) constructs that can guide reinnervation in vivo without added E-Stim protocols should also be explored.

Addressing fibrosis and immune modulation in SMTE

Fibrosis is a major obstacle in VML treatment, often leading to scar tissue formation, impaired regeneration, and loss of function. To mitigate fibrosis, researchers have explored antifibrotic agent delivery, immune-responsive biomaterials that balance proinflammatory and anti-inflammatory macrophage activity, and controlled release of bioactive factors to fine-tune the inflammatory response and prevent excessive scar tissue formation. Despite promising findings, comprehensive studies evaluating the clinical translatability of these approaches are still lacking. Future research should focus on developing immunomodulatory scaffolds that actively regulate inflammation while promoting vascularization, myogenesis, and functional muscle regeneration and minimizing scar tissue formation.

Overcoming limitations in vascularization and oxygen transport

Inadequate vascularization remains a critical bottleneck in engineered muscle constructs, leading to ischemia, inadequate oxygen diffusion, and limited construct survival postimplantation. Although strategies such as prevascularization, angiogenic growth factor delivery, and coculturing ECs have improved capillary formation, sustaining long-term vascular integration remains challenging. Future research should focus on (1) developing hybrid bioprinting techniques incorporating vascularized cell-laden bioinks to create hierarchically structured capillary networks, (2) utilizing ML-driven optimization to predict optimal multicell ratios, scaffold properties, and spatial gradients of angiogenic cues for enhanced vascular integration, and (3) designing bioactive materials that release proangiogenic factors in a spatiotemporally controlled manner to promote sustained capillary formation.

Leveraging ML for SMTE optimization

ML offers immense potential in tissue engineering, but its applications in SMTE remain underdeveloped due to limited high-quality datasets. Many ML models are trained on small, nongeneralizable datasets, leading to overfitting and poor real-world applicability.^128,136 To harness ML effectively in the future, researchers must (1) develop standardized, publicly accessible large datasets to improve model reliability, (2) use feature selection techniques to identify key biomaterial–cell interactions that enhance myogenic differentiation and functional integration, and (3) integrate ML with bioprinting to optimize scaffold architectures, predict mechanical and biochemical properties, and enable real-time adaptive printing for reproducible construct fabrication.

Advancing biofabrication for complex muscle architectures

Although bioprinting and electrospinning have significantly improved the fabrication of anisotropic, hierarchical muscle structures, challenges remain in recapitulating the multiscale organization of native muscle. Future directions should include 4D bioprinting, allowing constructs to dynamically adapt to external stimuli for enhanced tissue remodeling. In this regard, electroactive biomaterials such as polyaniline, polypyrrole, polythiophene, graphene, and metallic nanoparticles can be explored to promote electromechanical stimulation, facilitating myotube formation, NMJ development, and contractile function. Moreover, hybrid biomaterials integrating conductive nanocomposites, bioactive factors, and mechanical cues can also be explored to create stimuli-responsive constructs for personalized and effective regenerative therapies.¹³⁷

Understanding cell–material interactions

A limited understanding of how cell types (myoblasts, mesoangioblasts, stem cells) interact with various biomaterials hinders construct optimization.¹³⁸ Although natural and synthetic biomaterials have been widely explored, the precise mechanisms by which scaffold physicochemical properties influence cell behavior remain unclear. Further research is needed to elucidate (1) the impact of electrical and mechanical stimulation on muscle regeneration and functional recovery and (2) optimal biomaterial compositions that support cell adhesion, proliferation, and long-term construct stability.

Bridging the gap between preclinical research and clinical translation

Although several constructs have been developed over the past two decades, almost none is available in the clinics. Translating SMTE technologies into clinically viable therapies requires addressing scalability, standardization, and regulatory approval challenges. Many engineered constructs fail to restore function due to inadequate vascularization, reinnervation, and mechanical integration. Future efforts should focus on the following: (1) Developing scalable manufacturing processes to produce off-the-shelf, patient-specific muscle grafts. (2) Strengthening collaborations among bioengineers, clinicians, regulatory officials, and insurance companies to facilitate the translation of clinically relevant constructs that align with surgical implantation techniques and postoperative rehabilitation protocols. (3) Establishing preclinical models that are similar to or close to human VML models to assess the long-term safety, efficacy, and functional restoration of engineered constructs.

By addressing these research gaps, SMTE has the potential to transform muscle regenerative medicine, offering clinically viable bioengineered solutions for VML and musculoskeletal disorders in the upcoming decade. The combined efforts in biomaterial innovation, bioprinting technology, computational modeling, and regulatory science will pave the way for personalized, scalable, and effective therapies to restore skeletal muscle function.

Authors’ Contributions

S.Y.S.: Reviewed all the studies and prepared the original draft of the article. P.S.: Contributed to editing, preparing the final draft, and overseeing project administration. Both authors reviewed and approved the final version of the article.

Footnotes

Author Disclosure Statement

The authors have nothing to disclose.

Funding Information

This work was supported by R03 Grant No. 1R03EB036096-01 and R01 Grant No. 1R01AR083865-01A1.

References

Organization WH. Musculoskeletal health. 2022. Available from: https://www.who.int/news-room/fact-sheets/detail/musculoskeletal-conditions

Liu

, Wang

, Fan

, et al. Global burden of musculoskeletal disorders and attributable factors in 204 countries and territories: A secondary analysis of the Global Burden of Disease 2019 study. BMJ Open, 2022; 12(6):e062183.

Council NS. National Safety Council Releases New Findings on Workplace Injury Prevention. 2022. Available from: https://www.prnewswire.com/news-releases/national-safety-council-releases-new-findings-on-workplace-injury-prevention-301623981.html

Grogan

, Hsu

, Consortium

STR

, Skeletal Trauma Research Consortium. Volumetric muscle loss. J Am Acad Orthop Surg, 2011; 19(Suppl 1):S35–S37.

Corona

, Rivera

, Owens

, et al. Volumetric muscle loss leads to permanent disability following extremity trauma. J Rehabil Res Develop, 2015; 52(7).

Mukund

, Subramaniam

. Skeletal muscle: A review of molecular structure and function, in health and disease. Wiley Interdiscip Rev Syst Biol Med, 2020; 12(1):e1462.

Liu

, Saul

, Böker

, et al. Current methods for skeletal muscle tissue repair and regeneration. Biomed Res Int, 2018; 2018(1):1984879.

Hurtgen

, Ward

, Garg

, et al. Severe muscle trauma triggers heightened and prolonged local musculoskeletal inflammation and impairs adjacent tibia fracture healing. J Musculoskelet Neuronal Interact, 2016; 16(2):122–134.

Hyldahl

, Nelson

, Xin

, et al. Extracellular matrix remodeling and its contribution to protective adaptation following lengthening contractions in human muscle. Faseb J, 2015; 29(7):2894–2904.

10.

Garg

, Ward

, Hurtgen

, et al. Volumetric muscle loss: Persistent functional deficits beyond frank loss of tissue. J Orthop Res, 2015; 33(1):40–46.

11.

Turner

, Badylak

. Regeneration of skeletal muscle. Cell Tissue Res, 2012; 347(3):759–774.

12.

Hoffman

, Raymond-Pope

, Pritchard

, et al. Differential evaluation of neuromuscular injuries to understand re-innervation at the neuromuscular junction. Exp Neurol, 2024; 382:114996.

13.

Das

, Browne

, Laimo

, et al. Pre-innervated tissue-engineered muscle promotes a pro-regenerative microenvironment following volumetric muscle loss. Commun Biol, 2020; 3(1):330.

14.

Sarrafian

, Bodine

, Murphy

, et al. Extracellular matrix scaffolds for treatment of large volume muscle injuries: A review. Vet Surg, 2018; 47(4):524–535.

15.

Sorensen

, Hoffman

, Corona

, et al. Secondary denervation is a chronic pathophysiologic sequela of volumetric muscle loss. J Appl Physiol (1985), 2021; 130(5):1614–1625.

16.

Downing

, Prisby

, Varanasi

, et al. Old and new biomarkers for volumetric muscle loss. Curr Opin Pharmacol, 2021; 59:61–69.

17.

Lin

C-H

, Lin

Y-T

, Yeh

J-T

, et al. Free functioning muscle transfer for lower extremity posttraumatic composite structure and functional defect. Plast Reconstr Surg, 2007; 119(7):2118–2126.

18.

Bianchi

, Copelli

, Ferrari

, et al. Free flaps: Outcomes and complications in head and neck reconstructions. J Craniomaxillofac Surg, 2009; 37(8):438–442.

19.

Lawson

, Levin

. Principles of free tissue transfer in orthopaedic practice. J Am Acad Orthop Surg, 2007; 15(5):290–299.

20.

Carnes

, Pins

. Skeletal muscle tissue engineering: Biomaterials-based strategies for the treatment of volumetric muscle loss. Bioengineering, 2020; 7(3):85.

21.

De Pieri

, Rochev

, Zeugolis

. Scaffold-free cell-based tissue engineering therapies: Advances, shortfalls and forecast. NPJ Regen Med, 2021; 6(1):18.

22.

Ostrovidov

, Salehi

, Costantini

, et al. 3D bioprinting in skeletal muscle tissue engineering. Small, 2019; 15(24):e1805530.

23.

Qazi

, Mooney

, Pumberger

, et al. Biomaterials based strategies for skeletal muscle tissue engineering: Existing technologies and future trends. Biomaterials, 2015; 53:502–521.

24.

Alarcin

, Bal-Öztürk

, Avci

, et al. Current strategies for the regeneration of skeletal muscle tissue. Int J Mol Sci, 2021; 22(11):5929.

25.

Kang

H-W

, Lee

, Ko

, et al. A 3D bioprinting system to produce human-scale tissue constructs with structural integrity. Nat Biotechnol, 2016; 34(3):312–319.

26.

Das

, Pati

, Choi

Y-J

, et al. Bioprintable, cell-laden silk fibroin–gelatin hydrogel supporting multilineage differentiation of stem cells for fabrication of three-dimensional tissue constructs. Acta Biomater, 2015; 11:233–246.

27.

Wolf

, Daly

, Reing

, et al. Biologic scaffold composed of skeletal muscle extracellular matrix. Biomaterials, 2012; 33(10):2916–2925.

28.

Sicari

, Rubin

, Dearth

, et al. An acellular biologic scaffold promotes skeletal muscle formation in mice and humans with volumetric muscle loss. Sci Transl Med, 2014; 6(234):234ra58.

29.

Badylak

, Dziki

, Sicari

, et al. Mechanisms by which acellular biologic scaffolds promote functional skeletal muscle restoration. Biomaterials, 2016; 103:128–136.

30.

Badylak

. The extracellular matrix as a biologic scaffold material. Biomaterials, 2007; 28(25):3587–3593.

31.

Badylak

, Freytes

, Gilbert

. Extracellular matrix as a biological scaffold material: Structure and function. Acta Biomater, 2009; 5(1):1–13.

32.

Garg

, Ward

, Corona

. Asynchronous inflammation and myogenic cell migration limit muscle tissue regeneration mediated by a cellular scaffolds. Inflamm Cell Signal, 2014; 1(4).

33.

Urciuolo

, Urbani

, Perin

, et al. Decellularised skeletal muscles allow functional muscle regeneration by promoting host cell migration. Sci Rep, 2018; 8(1):8398–8320.

34.

Dziki

, Sicari

, Wolf

, et al. Immunomodulation and mobilization of progenitor cells by extracellular matrix bioscaffolds for volumetric muscle loss treatment. Tissue Eng Part A, 2016; 22(19–20):1129–1139.

35.

Valentin

, Turner

, Gilbert

, et al. Functional skeletal muscle formation with a biologic scaffold. Biomaterials, 2010; 31(29):7475–7484.

36.

, Corona

, Chen

, et al. A standardized rat model of volumetric muscle loss injury for the development of tissue engineering therapies. Biores Open Access, 2012; 1(6):280–290.

37.

Kasukonis

, Kim

, Brown

, et al. Codelivery of infusion decellularized skeletal muscle with minced muscle autografts improved recovery from volumetric muscle loss injury in a rat model. Tissue Eng Part A, 2016; 22(19–20):1151–1163.

38.

Chen

, Walters

. Muscle-derived decellularised extracellular matrix improves functional recovery in a rat latissimus dorsi muscle defect model. J Plast Reconstr Aesthet Surg, 2013; 66(12):1750–1758.

39.

Corona

, Wu

, Ward

, et al. The promotion of a functional fibrosis in skeletal muscle with volumetric muscle loss injury following the transplantation of muscle-ECM. Biomaterials, 2013; 34(13):3324–3335.

40.

Aurora

, Corona

, Walters

. A porcine urinary bladder matrix does not recapitulate the spatiotemporal macrophage response of muscle regeneration after volumetric muscle loss injury. Cells Tissues Organs, 2016; 202(3–4):189–201.

41.

Corona

, Garg

, Ward

, et al. Autologous minced muscle grafts: A tissue engineering therapy for the volumetric loss of skeletal muscle. Am J Physiol Cell Physiol, 2013; 305(7):C761–C775.

42.

Corona

, Henderson

, Ward

, et al. Contribution of minced muscle graft progenitor cells to muscle fiber formation after volumetric muscle loss injury in wild‐type and immune deficient mice. Physiol Rep, 2017; 5(7):e13249.

43.

Dolan

, Motherwell

, Franco

, et al. Evaluating the potential use of functional fibrosis to facilitate improved outcomes following volumetric muscle loss injury. Acta Biomater, 2022; 140:379–388.

44.

Kim

, Kasukonis

, Roberts

, et al. Graft alignment impacts the regenerative response of skeletal muscle after volumetric muscle loss in a rat model. Acta Biomater, 2020; 105:191–202.

45.

Sadtler

, Estrellas

, Allen

, et al. Developing a pro-regenerative biomaterial scaffold microenvironment requires T helper 2 cells. Science, 2016; 352(6283):366–370.

46.

Kasravi

, Ahmadi

, Babajani

, et al. Immunogenicity of decellularized extracellular matrix scaffolds: A bottleneck in tissue engineering and regenerative medicine. Biomater Res, 2023; 27(1):10.

47.

Badylak

. Decellularized allogeneic and xenogeneic tissue as a bioscaffold for regenerative medicine: Factors that influence the host response. Ann Biomed Eng, 2014; 42(7):1517–1527.

48.

Alaribe

, Manoto

, Motaung

. Scaffolds from biomaterials: Advantages and limitations in bone and tissue engineering. Biologia, 2016; 71(4):353–366.

49.

Reddy

MSB

, Ponnamma

, Choudhary

, et al. A comparative review of natural and synthetic biopolymer composite scaffolds. Polymers (Basel), 2021; 13(7):1105.

50.

Satchanska

, Davidova

, Petrov

. Natural and synthetic polymers for biomedical and environmental applications. Polymers (Basel), 2024; 16(8):1159.

51.

Prete

, Dattilo

, Patitucci

, et al. Natural and synthetic polymeric biomaterials for application in wound management. J Funct Biomater, 2023; 14(9):455.

52.

Duman

, Şendemir Ürkmez

, Zeybek

. Electrospinning of nanofibrous polycaprolactone (PCL) and collagen-blended polycaprolactone for wound dressing and tissue engineering. Usak University Journal of Material Sciences, 2014; 3(1):121–121.

53.

, Xu

, Shen

, et al. Emulsion electrospun PLA/calcium alginate nanofibers for periodontal tissue engineering. J Biomater Appl, 2020; 34(6):763–777.

54.

Garg

, Ward

, Rathbone

, et al. Transplantation of devitalized muscle scaffolds is insufficient for appreciable de novo muscle fiber regeneration after volumetric muscle loss injury. Cell Tissue Res, 2014; 358(3):857–873.

55.

Corona

, Greising

. Challenges to acellular biological scaffold mediated skeletal muscle tissue regeneration. Biomaterials, 2016; 104:238–246.

56.

Gholobova

, Terrie

, Gerard

, et al. Vascularization of tissue-engineered skeletal muscle constructs. Biomaterials, 2020; 235:119708.

57.

Levenberg

, Rouwkema

, Macdonald

, et al. Engineering vascularized skeletal muscle tissue. Nat Biotechnol, 2005; 23(7):879–884.

58.

Gilbert‐Honick

, Grayson

. Vascularized and innervated skeletal muscle tissue engineering. Adv Healthc Mater, 2020; 9(1):e1900626.

59.

Das

, Gordián-Vélez

, Ledebur

, et al. Innervation: The missing link for biofabricated tissues and organs. NPJ Regen Med, 2020; 5(1):11.

60.

Boldrin

, Elvassore

, Malerba

, et al. Satellite cells delivered by micro-patterned scaffolds: A new strategy for cell transplantation in muscle diseases. Tissue Eng, 2007; 13(2):253–262.

61.

Sicari

, Londono

, Badylak

. Strategies for skeletal muscle tissue engineering: Seed vs. soil. J Mater Chem B, 2015; 3(40):7881–7895.

62.

Conconi

, De Coppi

, Bellini

, et al. Homologous muscle acellular matrix seeded with autologous myoblasts as a tissue-engineering approach to abdominal wall-defect repair. Biomaterials, 2005; 26(15):2567–2574.

63.

Qiu

, Liu

, Zhang

, et al. Mesenchymal stem cells and extracellular matrix scaffold promote muscle regeneration by synergistically regulating macrophage polarization toward the M2 phenotype. Stem Cell Res Ther, 2018; 9(1):88–15.

64.

Jodat

, Zhang

, Al Tanoury

, et al. hiPSC-derived 3D bioprinted skeletal muscle tissue implants regenerate skeletal muscle following volumetric muscle loss. 2021.

65.

Hwangbo

, Lee

, Jin

E-J

, et al. Bio-printing of aligned GelMa-based cell-laden structure for muscle tissue regeneration. Bioact Mater, 2022; 8:57–70.

66.

, Lei

, Li

, et al. Living cell‐laden hydrogels: Unleashing the future of responsive biohybrid systems. Responsive Materials, 2023; 1(1):e20230009.

67.

Gilbert-Honick

, Iyer

, Somers

, et al. Engineering 3D skeletal muscle primed for neuromuscular regeneration following volumetric muscle loss. Biomaterials, 2020; 255:120154.

68.

Narayanan

, Jia

, Kim

, et al. Biomimetic glycosaminoglycan-based scaffolds improve skeletal muscle regeneration in a Murine volumetric muscle loss model. Bioact Mater, 2021; 6(4):1201–1213.

69.

Grasman

, Do

, Page

, et al. Rapid release of growth factors regenerates force output in volumetric muscle loss injuries. Biomaterials, 2015; 72:49–60.

70.

Huynh

, Reed

, Blackwell

, et al. Local IL-10 delivery modulates the immune response and enhances repair of volumetric muscle loss muscle injury. Sci Rep, 2023; 13(1):1983.

71.

Rossi

, Flaibani

, Blaauw

, et al. In vivo tissue engineering of functional skeletal muscle by freshly isolated satellite cells embedded in a photopolymerizable hydrogel. Faseb J, 2011; 25(7):2296–2304.

72.

Passipieri

, Baker

, Siriwardane

, et al. Keratin hydrogel enhances in vivo skeletal muscle function in a rat model of volumetric muscle loss. Tissue Eng Part A, 2017; 23(11–12):556–571.

73.

Chen

C-L

, Wei

S-Y

, Chen

W-L

, et al. Reconstructing vascular networks promotes the repair of skeletal muscle following volumetric muscle loss by pre-vascularized tissue constructs. J Tissue Eng, 2023; 14:20417314231201231.

74.

Liu

, Zhang

, et al. Synergistic angiogenesis promoting effects of extracellular matrix scaffolds and adipose-derived stem cells during wound repair. Tissue Eng Part A, 2011; 17(5–6):725–739.

75.

Madden

, Juhas

, Kraus

, et al. Bioengineered human myobundles mimic clinical responses of skeletal muscle to drugs. Elife, 2015; 4:e04885.

76.

Ahadian

, Ostrovidov

, Hosseini

, et al. Electrical stimulation as a biomimicry tool for regulating muscle cell behavior. Organogenesis, 2013; 9(2):87–92.

77.

Mackey

, Magnan

, Chazaud

, et al. Human skeletal muscle fibroblasts stimulate in vitro myogenesis and in vivo muscle regeneration. J Physiol, 2017; 595(15):5115–5127.

78.

Rigillo

, Basile

, Belluti

, et al. The transcription factor NF-Y participates to stem cell fate decision and regeneration in adult skeletal muscle. Nat Commun, 2021; 12(1):6013.

79.

Jia

, Hu

, Li

, et al. Glass-activated regeneration of volumetric muscle loss. Acta Biomater, 2020; 103:306–317.

80.

Patel

, Dunn

, Talovic

, et al. Aligned nanofibers of decellularized muscle ECM support myogenic activity in primary satellite cells in vitro. Biomed Mater, 2019; 14(3):035010.

81.

Nakayama

, Quarta

, Paine

, et al. Treatment of volumetric muscle loss in mice using nanofibrillar scaffolds enhances vascular organization and integration. Commun Biol, 2019; 2(1):170.

82.

Wang

, Li

, Wang

, et al. Injectable remote magnetic nanofiber/hydrogel multi-scale scaffold for functional anisotropic skeletal muscle regeneration. Biomaterials, 2022; 285:121537.

83.

Nutter

, VanDusen

, Florida

, et al. The effects of engineered skeletal muscle on volumetric muscle loss in the tibialis anterior of rat after 3 months in vivo. Regen Eng Transl Med, 2020; 6(4):365–372.

84.

Liang

, Han

, Li

, et al. Perfusable adipose decellularized extracellular matrix biological scaffold co-recellularized with adipose-derived stem cells and L6 promotes functional skeletal muscle regeneration following volumetric muscle loss. Biomaterials, 2024; 307:122529.

85.

Jin

, Shahriari

, Jeon

, et al. Functional skeletal muscle regeneration with thermally drawn porous fibers and reprogrammed muscle progenitors for volumetric muscle injury. Advanced Materials, 2021; 33(14):2007946.

86.

Ulery

, Nair

, Laurencin

. Biomedical applications of biodegradable polymers. J Polym Sci B Polym Phys, 2011; 49(12):832–864.

87.

Farshidfar

, Iravani

, Varma

. Alginate-based biomaterials in tissue engineering and regenerative medicine. Mar Drugs, 2023; 21(3):189.

88.

Haider

, Khan

, Iqbal

, et al. Chitosan as a tool for tissue engineering and rehabilitation: Recent developments and future perspectives–A review. Int J Biol Macromol, 2024; 278(Pt 1):134172.

89.

Rao

. Hyaluronic acid–based hydrogels for tissue engineering. In: Biomaterials for Organ and Tissue Regeneration. Elsevier: 2020; pp. 551–565.

90.

Sanz-Horta

, Matesanz

, Gallardo

, et al. Technological advances in fibrin for tissue engineering. J Tissue Eng, 2023; 14:20417314231190288.

91.

Parenteau-Bareil

, Gauvin

, Berthod

. Collagen-based biomaterials for tissue engineering applications. Materials, 2010; 3(3):1863–1887.

92.

Mohanto

, Narayana

, Merai

, et al. Advancements in gelatin-based hydrogel systems for biomedical applications: A state-of-the-art review. Int J Biol Macromol, 2023; 253(Pt 5):127143.

93.

Sonaye

, Ertugral

, Kothapalli

, et al. Extrusion 3D (bio) printing of alginate-gelatin-based composite scaffolds for skeletal muscle tissue engineering. Materials, 2022; 15(22):7945.

94.

Crapo

, Gilbert

, Badylak

. An overview of tissue and whole organ decellularization processes. Biomaterials, 2011; 32(12):3233–3243.

95.

Middleton

, Tipton

. Synthetic biodegradable polymers as orthopedic devices. Biomaterials, 2000; 21(23):2335–2346.

96.

Utech

, Boccaccini

. A review of hydrogel-based composites for biomedical applications: Enhancement of hydrogel properties by addition of rigid inorganic fillers. J Mater Sci, 2016; 51(1):271–310.

97.

Jang

T-S

, Jung

H-D

, Pan

, et al. 3D printing of hydrogel composite systems: Recent advances in technology for tissue engineering. Int J Bioprint, 2018; 4(1):126.

98.

Novosel

, Kleinhans

, Kluger

. Vascularization is the key challenge in tissue engineering. Adv Drug Deliv Rev, 2011; 63(4–5):300–311.

99.

Martin

, Passey

, Player

, et al. Factors affecting the structure and maturation of human tissue engineered skeletal muscle. Biomaterials, 2013; 34(23):5759–5765.

100.

Webber

, Khan

, Sydlik

, et al. A perspective on the clinical translation of scaffolds for tissue engineering. Ann Biomed Eng, 2015; 43(3):641–656.

101.

Afshar Bakooshli

, Lippmann

, Mulcahy

, et al. A 3D culture model of innervated human skeletal muscle enables studies of the adult neuromuscular junction. Elife, 2019; 8:e44530.

102.

Leberfinger

, Dinda

, Wu

, et al. Bioprinting functional tissues. Acta Biomater, 2019; 95:32–49.

103.

Xiang

, Miller

, Guan

, et al. 3D bioprinting of complex tissues in vitro: State-of-the-art and future perspectives. Arch Toxicol, 2022; 96(3):691–710.

104.

, Chen

, Fan

, et al. Recent advances in bioprinting techniques: Approaches, applications and future prospects. J Transl Med, 2016; 14:271–215.

105.

Unagolla

, Jayasuriya

. Hydrogel-based 3D bioprinting: A comprehensive review on cell-laden hydrogels, bioink formulations, and future perspectives. Appl Mater Today, 2020; 18:100479.

106.

Samandari

, Quint

, Rodríguez‐delaRosa

, et al. Bioinks and bioprinting strategies for skeletal muscle tissue engineering. Advanced Materials, 2022; 34(12):2105883.

107.

Mobaraki

, Ghaffari

, Yazdanpanah

, et al. Bioinks and bioprinting: A focused review. Bioprinting, 2020; 18:e00080.

108.

Donderwinkel

, Van Hest

JCM

, Cameron

. Bio-inks for 3D bioprinting: Recent advances and future prospects. Polym Chem, 2017; 8(31):4451–4471.

109.

Ashammakhi

, Ahadian

, Xu

, et al. Bioinks and bioprinting technologies to make heterogeneous and biomimetic tissue constructs. Mater Today Bio, 2019; 1:100008.

110.

Ferraris

, Spriano

, Scalia

, et al. Topographical and biomechanical guidance of electrospun fibers for biomedical applications. Polymers (Basel), 2020; 12(12):2896.

111.

Chen

, Vigliotti

, Bacca

, et al. Role of boundary conditions in determining cell alignment in response to stretch. Proc Natl Acad Sci U S A, 2018; 115(5):986–991.

112.

Prendergast

, Davidson

, Burdick

. A biofabrication method to align cells within bioprinted photocrosslinkable and cell-degradable hydrogel constructs via embedded fibers. Biofabrication, 2021; 13(4):044108.

113.

Gao

, Xiao

, Wei

, et al. Regulation of myogenic differentiation by topologically microgrooved surfaces for skeletal muscle tissue engineering. ACS Omega, 2021; 6(32):20931–20940.

114.

Chansoria

, Narayanan

, Schuchard

, et al. Ultrasound-assisted biofabrication and bioprinting of preferentially aligned three-dimensional cellular constructs. Biofabrication, 2019; 11(3):035015.

115.

Choi

, Lee

, Kim

, et al. Fibre-infused gel scaffolds guide cardiomyocyte alignment in 3D-printed ventricles. Nat Mater, 2023; 22(8):1039–1046.

116.

Skillin

, Kirkpatrick

, Herbert

, et al. Stiffness anisotropy coordinates supracellular contractility driving long-range myotube-ECM alignment. Sci Adv, 2024; 10(22):eadn0235.

117.

Decante

, Costa

, Silva-Correia

, et al. Engineering bioinks for 3D bioprinting. Biofabrication, 2021; 13(3):032001.

118.

, Jiang

. A perspective on using machine learning in 3D bioprinting. Int J Bioprint, 2020; 6(1):253.

119.

Chahal

, Gulia

, Department of Computer Science and Applications, Maharishi Dayanand University, Rohtak, India. Machine learning and deep learning. IJITEE, 2019; 8(12):4910–4914.

120.

Mobarak

, Mimona

, Islam

, et al. Scope of machine learning in materials research—A review. Applied Surface Science Advances, 2023; 18:100523.

121.

Mishra

, Reddy

, Pathak

. The understanding of deep learning: A comprehensive review. Mathematical Problems in Engineering, 2021; 2021(1):1–15.

122.

Sun

, Yao

, An

, et al. Machine learning and 3D bioprinting. Int J Bioprint, 2023; 9(4):717.

123.

Zhu

, Huang

, Wu

, et al. Deep learning-based predictive identification of neural stem cell differentiation. Nat Commun, 2021; 12(1):2614.

124.

Jovel

, Greiner

. An introduction to machine learning approaches for biomedical research. Front Med (Lausanne), 2021; 8:771607.

125.

Rafieyan

, Ansari

, Vasheghani-Farahani

. A practical machine learning approach for predicting the quality of 3D (bio) printed scaffolds. Biofabrication, 2024; 16(4):045014.

126.

Sun

, Yao

, Huang

, et al. Machine learning applications in scaffold based bioprinting. Materials Today: Proceedings, 2022; 70:17–23.

127.

Jin

, Zhang

, Shao

, et al. Monitoring anomalies in 3D bioprinting with deep neural networks. ACS Biomater Sci Eng, 2023; 9(7):3945–3952.

128.

Bonatti

, Vozzi

, De Maria

. Enhancing quality control in bioprinting through machine learning. Biofabrication, 2024; 16(2):022001.

129.

Gerdes

, Gaikwad

, Ramesh

, et al. Monitoring and control of biological additive manufacturing using machine learning. J Intell Manuf, 2024; 35(3):1055–1077.

130.

Bonatti

, Vozzi

, Chua

, et al. A deep learning quality control loop of the extrusion-based bioprinting process. Int J Bioprint, 2022; 8(4):620.

131.

Sun

, Jing

, Fan

, et al. Electrohydrodynamic printing process monitoring by microscopic image identification. Int J Bioprint, 2019; 5(1):164.

132.

Ruberu

, Senadeera

, Rana

, et al. Coupling machine learning with 3D bioprinting to fast track optimisation of extrusion printing. Applied Materials Today, 2021; 22:100914.

133.

Al-Kharusi

, Dunne

, Little

, et al. The role of machine learning and design of experiments in the advancement of biomaterial and tissue engineering research. Bioengineering, 2022; 9(10):561.

134.

Sujeeun

, Goonoo

, Ramphul

, et al. Correlating in vitro performance with physico-chemical characteristics of nanofibrous scaffolds for skin tissue engineering using supervised machine learning algorithms. R Soc Open Sci, 2020; 7(12):201293.

135.

Lim

, Kim

, Lee

, et al. Skeletal muscle gauge prediction by a machine learning model in patients with colorectal cancer. Nutrition, 2023; 115:112146.

136.

, Shirzad

, Kim

, et al. Rheology-informed hierarchical machine learning model for the prediction of printing resolution in extrusion-based bioprinting. Int J Bioprinting, 2023; 9(6):1280–1324.

137.

Gahlawat

, Oruc

, Paul

, et al. Tissue engineered 3D constructs for volumetric muscle loss. Ann Biomed Eng, 2024; 52(9):2325–2347.

138.

Judson

, Rossi

. Towards stem cell therapies for skeletal muscle repair. NPJ Regen Med, 2020; 5(1):10.

139.

, Hou

, Wang

, et al. Bioprinted anisotropic scaffolds with fast stress relaxation bioink for engineering 3D skeletal muscle and repairing volumetric muscle loss. Acta Biomater, 2023; 156:21–36.

140.

Costantini

, Testa

, Mozetic

, et al. Microfluidic-enhanced 3D bioprinting of aligned myoblast-laden hydrogels leads to functionally organized myofibers in vitro and in vivo. Biomaterials, 2017; 131:98–110.

141.

, Kang

, Heo

, et al. Skeletal muscle regeneration with 3D bioprinted hyaluronate/gelatin hydrogels incorporating MXene nanoparticles. Int J Biol Macromol, 2024; 265(Pt 1):130696.

142.

Kim

, Kim

. 3D bioprinting of functional cell-laden bioinks and its application for cell-alignment and maturation. Applied Materials Today, 2020; 19:100588.

143.

Choi

Y-J

, Jun

Y-J

, Kim

, et al. A 3D cell printed muscle construct with tissue-derived bioink for the treatment of volumetric muscle loss. Biomaterials, 2019; 206:160–169.

144.

Kim

, Seol

Y-J

, Ko

, et al. 3D bioprinted human skeletal muscle constructs for muscle function restoration. Sci Rep, 2018; 8(1):12307.

145.

Ronzoni

, Aliberti

, Scocozza

, et al. Myoblast 3D bioprinting to burst in vitro skeletal muscle differentiation. J Tissue Eng Regen Med, 2022; 16(5):484–495.

146.

Fornetti

, De Paolis

, Fuoco

, et al. A novel extrusion-based 3D bioprinting system for skeletal muscle tissue engineering. Biofabrication, 2023; 15(2):025009.

147.

Lee

, Kim

, Lee

, et al. Self-aligned myofibers in 3D bioprinted extracellular matrix-based construct accelerate skeletal muscle function restoration. Appl Phys Rev, 2021; 8(2):021405.