Similar to traditional 2D printing, which ejects a tiny amount of ink onto paper to obtain an edited document, inkjet-based 3D bioprinting deposits nano- or micro-liter droplets of cell-laden bioinks toward a substrate in a drop-on-demand manner to stack-up a three-dimensional objective [34]. In general, the bioink containing living cells is loaded in a cartridge connected to the printing heads of an inkjet printer (Figure 2A). During the printing process, printing heads are usually deformed by a piezoelectric actuator or by thermal variation to squeeze the bioink, thus creating droplets. The piezoelectric heads include an element composed of special materials (e.g., crystals and ceramics) that are sensitive to electrical charges. As a pulsed voltage is provided, the elements bend backward, forcing an accurate amount of bioink from the cartridge onto the substrates [35]. A thermal inkjet printer imposes a rapid increase in temperature near the emission tip to generate a vapor bubble that outputs a droplet of bioink [36]. Recently, electrostatic and electrohydrodynamic-based modules have been adopted for 3D inkjet bioprinting techniques, which can achieve an identical effect by intermittently applying pressure to the bioink cartridge [37]. Several important parameters of inkjet-based 3D bioprinting must be considered to control and stabilize the droplets. The size of the droplets depends on the viscosity and surface tension of the bioink. The vibrational frequency of electrical signals that activate the actuators mainly dominates the speed of droplet deposition; the pulse and amplitude determine the consistency of the droplet number and size during each ejection.

Because of its ability to print small volumes of bioink, inkjet-based bioprinting has a high resolution, which is as fine as 2 μm, i.e., the dimension of a single cell [38]. Moreover, the rapid and tunable printing speed (1–250,000 droplets per second) is another merit of this technology [39]. However, the working mechanism requires careful selection of bioink with low viscosity (<0.1 Pa·s) to prevent nozzle clotting [40], which drastically limits the cell density upper range and narrows the range of available biomaterials.

Comparable with inkjet printing, laser-assisted printing also generates cell-laden droplets using a laser beam and deposits them on a substrate to build a 3D construct. A typical laser-assisted bioprinting system consists of three elements: a light source that focuses a pulsed laser beam; a ribbon composed of a laser-absorbing layer sandwiched between a transparent donor slide and a layer of cell-laden biomaterial; and a receiving reservoir or substrate that facilitates droplet deposition and cell adhesion [41]. During the printing process, as the laser beam is focused on the ribbon, the materials in the absorbance layer (e.g., Au, Ti) produce vapor pockets that induce the formation of droplets and propel them onto the substrate (Figure 2B). The key parameters to manipulate a laser-assisted bioprinter include laser effluence (e.g., laser wavelength, pulse energy), elimination frequency, the surface tension of the substrate, the distance between the ribbon and substrate, and the thickness and viscosity of the cell-laden layer.

Laser-assisted bioprinting has the advantages of ultrafine resolution (20–30 μm) and rapid printing speed (5 kHz) [42]. In addition, because of the nozzle-less working mechanism, this technology is not subjective to the cell density and viscosity of biomaterials. Despite these advantages, owing to the sophisticated laser system and composite ribbon, the costs of establishing a laser-assisted bioprinting system are inevitably high, which limits the widespread adoption of this technique.

Driven by a pneumatic or mechanical system, microextrusion-based bioprinting continuously forces cell-laden bioinks through a nozzle, forming a fine filament. By controlling the movement of the nozzles, the extruded filaments are stacked to create 3D constructs (Figure 2C). A pneumatic system applies stable air pressure provided by a clean compressed air source to the bioinks residing in a cartridge. Because the pressure values can be flexibly manipulated, a wide range of biomaterials with various viscosities (30 mPa·s to over 6 × 10⁷ mPa·s) can be fabricated for bioprinting tissue/organ equivalents [43]. In comparison, the mechanical system directly imposes forces on the bioinks via a piston or screw module. While the steady movement of the piston module is beneficial for tuning the flow rate of the bioink, the screw module offers better spatial control on the cartridge, thereby extruding the bioinks with higher viscosity [44]. However, the severe mechanical force might have detrimental effects on the cells encapsulated in the bioink owing to the potential damage to the cell membrane, leading to cell death.

Although microextrusion-based bioprinting has a broader pool of applicable biomaterials, the printing resolution, in comparison with the other two aforementioned technologies, is inevitably lower (>100 μm) owing to the shear stress [45]. During the extrusion process, the shear stress can exceed a threshold that secures the viability of the loaded cells when the nozzle size is small and the viscosity of the bioink is high. A study has reported that highly sensitive cells retain high cell viability (≥90%) after printing as long as a low shear stress (~100 Pa) is ensured [46]. Therefore, to fabricate a highly viable tissue construct, it is important to increase the nozzle size and reduce the viscosity of the bioink, which may result in a decreased printing resolution. However, the relatively larger nozzle used in microextrusion-based bioprinting systems can lead to a high printing speed, enabling rapid fabrication of large-volume constructs. Moreover, the rough resolution allows for direct bioprinting of pre-processed cell spheroids/aggregates, which could rapidly create functional tissues via cell self-assembly.

In summary, the important considerations and performances of three different 3D bioprinting systems are compared in Table 1. In general, each bioprinting technique has its advantages and limitations. Despite the relatively low resolution, microextrusion-based bioprinting offers a solution for the rapid fabrication of volumetric constructs using viscose bioinks with high cell density. Owing to these advantages, among bioprinting technologies, the microextrusion-based approach is the most promising technology for building tissue/organ substitutes thus far. Several pioneering reports have successfully created human-sized ears, muscles, tendons, and hearts.

Decellularization is a process that removes cellular and genetic molecules and preserves the acellular ECM components to maintain the mechanical and biochemical properties of tissue-specific microenvironments for artificial tissue engineering. The process can be classified into four main treatment categories: physical, chemical, enzymatic, and combinatorial [53]. Physical decellularization, including thermal, mechanical, and electrical treatments, has been reported in some studies. The main purpose of these physical approaches is to destroy the cellular and plasma membranes and remove biomolecules from the tissue. Thermal shock methods, including rapid freezing and snap freezing, are frequently employed. The growth of ice crystals weakens the cellular membrane and facilitates the release of cellular lysates. Direct abrasion with mechanical force through sonication and agitation accelerates cell-ECM dissociation [57]. However, physical methods should be followed by post-processing, coupled with chemical and enzymatic treatments to remove the membranous and intracellular lysates by promoting the diffusion of decellularizing agents [58].

Chemicals accelerate decellularization by decomposing chemical junctions, lysing cellular membranes, and removing cytoplasmic components. Diverse chemical decellularization reagents, including acids and alkalis, hypo/hypertonic solutions, ionic/non-ionic/bipolar detergents, and enzymes, have been employed [59]. Acids and alkalis, such as peracetic acid, hydrochloric acid, sodium hydroxide, and ammonium hydroxide, hydrolyze cytoplasmic biomolecules, including nucleic acids, and sterilize materials by neutralizing microbiomes. However, these pH-shifting solutions affect the ECM composition and lead to structural/functional changes ^[60][61][60,61]. Treatments with ionic/non-ionic/bipolar detergents induce ionic disturbance, resulting in dissociation of the cellular membrane and cytoplasmic proteins. Ionic detergents, such as SDS, have either a cationic or negatively charged head and are usually used to solubilize cellular and cytoplasmic membranes. However, due to their high destructivity, essential ECM components, including glycosaminoglycans (GAGs), collagen, and growth factors, can be eliminated [62]. Although nonionic detergents, including Triton X-100, are effective for delipidation, they have relatively weak detrimental effects that require long-term treatment, which may be an effective decellularization method to preserve biochemical and mechanical properties [63]. Bipolar detergents such as CHAPS, SB-10, and SB-16 have a neutral net charge and possess moderate disruptive capacity between that of ionic and nonionic detergents, which are appropriate characteristics for removing ECM proteins and DNA [64]. Hypotonic (e.g., Tris-HCl) and hypertonic (e.g., NaCl) solutions break down the matrix molecules and dissociate the junction between DNA and protein via an osmotic effect. Chelating agents, including EGTA and EDTA, bind to divalent cations, including Ca²⁺ and Mg²⁺, which mediate cell-ECM adhesion and disturb the protein–protein interactions, coupled with other enzymes and detergents [59].

Enzymes, including proteases, esterases, and nucleases, cleave intercellular and extracellular junctions as well as intracellular interactions. Proteolytic enzymes, including trypsin and dispase, cleave peptide bonds with high specificity. In particular, trypsin cuts the peptide bond between lysine and arginine and disrupts the ultrastructure of the native tissue in a time-dependent manner, preserving GAGs and breaking collagen and elastin, which changes their mechanical properties. Dispase cleaves basement membrane proteins, including collagen type IV and fibronectin. α-Galactosidase dissects alpha-galactosyl moieties of glycoproteins and glycolipids and prevents adverse xenoimmune responses. Nucleases, including endonucleases and exonucleases, cleave nucleic acids, including ribonucleotides and deoxynucleotides [65]. However, long-term enzymatic treatment drastically changes the mechanical and biochemical properties of the ECM. Furthermore, after enzymatic treatment, the lysate, including undesired ECM residues, needs to be removed through further processes to prevent an immune response.

As the terminal step of decellularization, sterilization is a critical process which removes the remaining pathogenic compounds and prevents an adverse immune response prior to in vitro analysis or implantation. Several sterilization methods can be employed, including dry heat, pressurized steaming, electron beam irradiation, gamma ray irradiation, and chemical treatments using ethylene oxide and peracetic acid. Because the biochemical and physical properties of dECM can be affected by disinfection, selecting the proper sterilization method is very important [66]. In addition to sterility, the effects on the deformation of growth factors and cytokines and the preservability of structural stability and biocompatibility should be satisfied.

The content of the prepared dECM should be quantified and qualified. The goals of decellularization are maximal clearance of cellular and genetic molecules and minimal loss of ECM components. Hematoxylin and eosin staining and DNA quantification are commonly conducted to verify the absence of cells and cell debris in the dECM. Using hematoxylin and eosin staining, the efficiency of decellularization can be determined by visualizing the presence of nuclei in the cells. Because double-stranded DNA (dsDNA) induces an immune response, the content of dsDNA should be quantified and should be lower than 50 ng dsDNA per mg dECM. Additionally, xenogenic factors such as the Gal epitope should be eliminated to prevent adverse inflammatory reactions in the recipient [67].

The preservation of ECM components, such as collagen, GAGs, and elastin, should also be estimated. Decellularization changes the composition of the extracellular matrix. In particular, the loss of components related to cellular functionality in the ECM influences tissue-specific cellular phenotypes due to the lack of attachment sites, growth factors, and other molecules. For instance, in the case of bone tissues, the content of collagen and hydroxyapatite should be precisely quantified because the components provide flexibility and strength for load-bearing functions. Recently, proteomic analysis has been highlighted because of its complementary strategy, providing both compositional and quantitative data. It also allows an in-depth study of enriched proteins such as growth factors and cytokines in dECM [68]. In addition, as well as the development of analysis tools, the process of decellularization should be also optimized in order to qualify the dECM components.

Conventionally, dECM-based materials have been used in the form of whole organs or cell sheets for in vitro modeling. However, several limitations for tissue engineering, such as dimensional differences and a lack of tailored microgeometries, have been raised. To overcome these drawbacks, the soluble form of dECM can be utilized along with the development of 3D bioprinting technology [56]. A gel-like substance with the desired concentration is essential for constructing tissue analogs. Preserving tissue-specific biochemical properties, including essential growth factors and peptides, is important because each tissue has different cells that secrete different functional factors. Furthermore, the rheological properties need to be fulfilled for use as a bioink for 3D printing. This material is regarded as a new class of biomimetic hydrogels.

Solubilization is usually accomplished using enzymatic acidic digestion of the dECM to be printed. Briefly, small pieces of lyophilized dECM are digested using pepsin, an endopeptidase that catalyzes protein hydrolysis at pH 2.0–3.5. Together with physical stirring, a gel-like substance with the desired concentration can be obtained [69]. After digestion, the hydrogel is neutralized at a proper temperature to recover and spontaneously reform intramolecular bonds of the solubilized dECM protein [58]. Through this process, various dECM bioinks have been developed; several bioprinting companies, such as T&R Biofab and Innoregen, have recently successfully commercialized dECM bioinks including Bone deCelluid™, Cartilage deCelluid™, Skin deCelluid™, and Gel4Tissue^{® [30][70][71][72]}[30,70,71,72]. The dECM bioink is then ready to be equipped for 3D bioprinting.

Excellent printability refers to the capacity of a bioink to support precise 3D bioprinting of a living-cell-laden construct with pre-designed structures [73]. Achieving satisfactory printability is highly dependent on the properties of applied bioinks, bioprinter configurations, and biofabrication strategies ^[74][75][74,75]. The dECM bioink should possess an appropriate viscosity that enables tissue fabrication using 3D bioprinting technology, as analyzed in Section 2. Viscosity is a natural characteristic of a fluidic material, referring to the internal frictional force between adjacent layers of fluid that are in relative motion. As in other fluids, the viscosity of dECM bioink is determined by the nature of the dECM materials, such as molecular weight, molecular structure, and interactions between chains. Because different tissues contain various ECM components, the resultant tissue-specific dECM bioinks inevitably show disparate viscosities. For instance, the reported dECM bioink obtained from the porcine brain has much lower viscosity (2.1 ± 0.3 Pa, 10 mg/mL) [76] than that harvested from porcine skin (300–500 Pa, 10 mg/mL) [30], which might be due to the lack of elastic proteins (e.g., collagen and elastin) in brain tissues. Although viscosity is the nature of a material, it can be tuned by controlling temperature; in general, viscosity is positively correlated with temperature [77]. However, due to the thermal sensitivity of collagen components as well as considering cell viability, the range over which temperature can be tuned during 3D bioprinting is usually confined between 4 °C and 37 °C. In addition to manipulating their temperature, increasing the concentration of dECM bioinks could also effectively elevate their viscosities. However, dECM materials have limited water solubility. Hence, it is difficult to control viscosity freely by changing their concentration. Intractable viscosity is a common drawback of bioinks originating from natural polymers. Supplementing thick materials as additives may be an effective way to solve this problem.

In addition, a rapid and irreversible sol–gel transition behavior is another key criterion for good printability. Ideally, the liquid bioink should be immediately polymerized after being deposited on demand, forming stable solid hydrogels that retain the printed structures. Most of the currently available dECM bioinks rely on the thermal gelation of collagen components, a process of fibril formation induced by peptide bonds between collagen molecules [78]. However, this process usually requires over 10 min to accomplish [79], and thus leads to the poor printability of dECM bioinks. To overcome this limitation, various strategies have been applied to improve the performance of tissue-specific bioinks, such as the incorporation of additives/crosslinkers and the chemical modification of dECM bioinks, which can help enable other rapid polymerizations (e.g., ionic and photo-gelation) ^[80][81][80,81].

Achieving high cell viability in cell-laden constructs is the first objective of 3D bioprinting. To ensure that the cells remain viable during the microextrusion-based printing process, the dECM bioink must exhibit a shear-thinning behavior to reduce the generated shear stress when flowing through a narrow nozzle. Reportedly, shear-induced cell damage can be minimized when the shear stress is less than 5 kPa [82]. Thus far, all types of tissue-specific bioinks have exhibited non-Newtonian behavior and shear-relaxation properties. However, because the shear stress is determined by both the viscosity of the bioink and the size of the printing nozzles, it is important to consider the detrimental effects of shear stress before selecting the printing devices and formulating dECM bioinks. The range of suitable concentrations of currently developed bioinks that can guarantee cell viability and function is 0.6–3% [53].

Nutrient/oxygen supply and metabolic byproduct removal are essential for the survival of cells. To secure cell viability, one of the easiest ways is to physically regulate the microstructure of the biological construct, so that the necessary regulating processes can be realized in vitro. Adjusting the porosity of bioink hydrogels can efficiently control the permeability of the cell-residing microenvironment, and thus the metabolic transportations can be effectively mediated [83]. The recently developed advanced 3D bioprinting strategies have achieved high fabrication resolution, dozens of micro-meters, that can be used to reconstruct microchannels and micro-vasculatures. For example, heart and liver dECM bioinks were successfully applied to construct biomimetic tissues with microchannels and interconnected vascular networks (30 μm to 180 μm wide) using a scanningless and continuous 3D bioprinting method [84]. Such defined ultrafine structures can directly deliver nutrients and oxygens to living cells.

In addition, the polymerization process that converts bioink into a stable hydrogel is another stage that might affect cell viability. The three main methods of polymerizing bioinks are thermal crosslinking, photo-crosslinking, and ionic crosslinking. First, as previously mentioned, the gelation of the dECM bioink depends on the polymerization of collagen components. Although thermal gelation via simple incubation does not cause detrimental effects on cell viability, different concentrations of dECM bioinks might result in altered gelation kinetics, which subsequently requires a prolonged or reduced incubation period to avoid insufficient gelation or dehydration-induced cell damage. Second, for photo-crosslinking, photo-initiators are necessary to initiate free radical reactions under ultraviolet (UV) light irradiation. Although dECM materials do not support photopolymerization, it is possible to achieve this by blending photo-crosslinkable materials with dECM bioinks. For instance, one study applied riboflavin to reinforce the cardiac dECM bioink after photopolymerization under UV light [85]. However, photo-initiators are generally toxic to cells, and UV light can damage cell DNA, thereby reducing cell viability. Therefore, to ensure high cell viability, it is important to carefully consider the concentration of the photo-initiator and control UV irradiation to provide a cell-friendly polymerization process. Lastly, ionic gelation relies on the presence of ion crosslinkers in the bioink. Alginate is a commonly used bioink that can be ionically crosslinked. When this polysaccharide polymer encounters divalent cations (e.g., Ca²⁺, Ba²⁺, Mg²⁺), instant crosslinking of the carboxylate groups of the guluronate groups occurs on the polymer backbone, forming hydrogels. A hybrid bioink containing vascular dECM and alginate has been formulated to facilitate coaxial bioprinting of blood vessel constructs involving human endothelial cells [86]. The concentration of ions should be sufficiently high to fully polymerize the alginate components. However, excessively high ion concentrations may adversely cause cell death due to osmotic pressure.

As the printed bioink is polymerized to form hydrogels, its role changes from a cell carrier to a scaffold that mechanically supports the printed tissue analogs and provides biological cues to promote the activities of embedded cells until they synthesize and assemble sufficient ECM instead. Therefore, appropriate mechanical strength and stable compositional disintegration are critical requirements for dECM bioinks.

First, as a scaffold, the bioink hydrogel should be sufficiently strong to resist gravity-caused structural collapse to sustain the complex 3D constructs. Moreover, because the hydrogel is composed of tightly bonded polymer chains, the 3D network absorbs additional water content from the culture medium during the incipient incubation period, leading to structural swelling. A hybrid bioink containing 3 wt% vascular dECM and 2 wt% alginate was reported to have swelled by 150.61 ± 6.37% before reaching equilibrium (10 h) [87]. Structural dilation may cause loss of resolution and a reduction in porosity. Furthermore, as the encapsulated cells attach, proliferate, and migrate, the generated force can induce severe shrinkage of the hydrogel, resulting in the dilapidation of the fabricated constructs. Hence, attention should be paid to minimizing the swelling and contraction of the hydrogel by modulating the mechanical strength of the dECM bioink. Various strategies have been proposed for 3D bioprinting of robust hydrogels. For example, a bioink consisting of alginate and poly (ethylene glycol) diacrylate was developed as an interpenetrating network via ionic–covalent bonding entanglement [88]. The printed structures could resist mechanical stress without significant plastic deformation while maintaining high cell viability (75.5 ± 11.6%) over 7 days.

In addition, upon 3D bioprinting of cell-laden constructs, dynamic ECM remodeling is an important process for successful tissue morphogenesis. In particular, the bioink hydrogel should guide the residing cells to proliferate, differentiate, aggregate, and reorganize; meanwhile, it needs to be gradually degraded and replaced by the de novo ECM produced by cells. The dECM bioinks can provide a superior microenvironment for cell-cell and cell-ECM interactions. However, achieving a tunable degradation rate of bioink hydrogels remains challenging. Rapid degradation might cause compositional instability, weakening the advantage of the dECM bioink, while retarded disintegration of the hydrogel could impede cell activities. Although it is possible to add degradation-active/inert ingredients (e.g., alginate and agarose) to manipulate the degradation rates of the bioink [89], the options remain limited considering the viability and activities of cells. Thus, for 3D bioprinting of functional tissues and organs, the degradation rate of bioink hydrogels must be prudently designed.

The mechanism of neovascularization can be roughly divided into angiogenesis and vasculogenesis. Angiogenesis is the growth of new capillaries from pre-existing blood vessels, while vasculogenesis refers to the formation of a primitive vascular network from endothelial primary or progenitor cells. Accordingly, two prevailing strategies have been established for 3D bioprinting of vascularized tissue constructs: the incorporation of angiogenic factors and pre-vascularization.

The first strategy attempts to spatially involve growth factors (e.g., vascular platelet-derived growth factor, vascular endothelial growth factor (VEGF), fibroblast growth factors) that can induce the ingrowth of host blood vessels into the fabricated construct, realizing in vivo angiogenesis. Thus, numerous studies have successfully achieved vascularized bone and muscle tissue. For instance, one study utilized 3D bioprinting technology to fabricate a biomimetic vascularized bone construct with regional immobilization of BMP-2 and VEGF. The gradient of growth factors loaded in the engineered constructs exhibited a superior ability to enhance microcapillary formation [140]. However, biomolecules usually have a short half-life and are rapidly eliminated in vivo, which might lead to insufficient dosage for the generation of viable vascular networks that support implanted grafts. Although encapsulating growth factors into polymeric particles and microspheres can achieve prolonged release kinetics, maintaining the gradient angiogenic signals remains challenging. The application of dECM bioink might be an alternative to such an elaborate design. Gao et al. developed a vascular dECM bioink from porcine aorta tissue for treating ischemic diseases. In particular, a tubular construct containing human endothelial progenitor cells and microspheres loaded with atorvastatin were successfully fabricated using coaxial 3D bioprinting technology [141]. After implantation into the ischemic limb of mice for 28 days, such a cell/drug co-laden vessel showed remarkable neo-vessel formation and limb salvage, demonstrating the potential of VdECM to promote neovascularization. Despite these remarkable achievements, angiogenesis-based vascularization is highly dependent on complex in vivo microenvironments, cellular status, construct design, and release profile of growth factors, which inevitably make the strategy difficult to control.

In addition, coaxial 3D bioprinting of the VdECM bioink could also contribute to the prevascularization of tissue constructs. Using this technique, Choi et al. combined the advantages of MdECM and VdECM to directly bioprint a core/shell filament that spatially carries human umbilical vein endothelial cells and human skeletal muscle cells that emulate the hierarchical architecture of vascularized muscle fibers [115]. Subsequently, multiple prevascularized muscles were fabricated together to scale-up a large-volume muscular construct for treating VML created in rat models, which extensively improved muscle fiber formation, neovascularization, and recovery of muscle contractile force. Apart from the direct printing of vascular constructs and positioning endothelial cells, 3D bioprinting can also realize the prevascularization of tissue constructs by creating hollow channels in bulk cell-laden hydrogels. Kolesky et al. applied cell-laden GelMA to embed a template of vasculature 3D printed using Pluronic F-127 [142]. Upon polymerization of GelMA, Pluronic F-127 was liquefied to facilitate the evacuation of the template. After seeding by endothelial cells, the resultant interconnected microchannels in the matrix resembled the native vasculature. Although engineering hollow channels can be controlled to effectively configure the pattern and design of vasculatures, the shrinkage of the hosting matrix caused by the activity of embedded cells might induce structural deformation, leading to destructive dilapidation and channel blockage. Therefore, careful design of the bioink formulation and evaluation of mechanical and rheological properties are important when this strategy is considered.

Compared with vascularization, research regarding tissue innervation began relatively late, and its progress is still in its infancy. Denervated skeletal muscles lose their contractility and suffer from muscle atrophy. Tissue-engineered skeletal muscle constructs consisting of living muscle cells are similar to denervated native counterparts, which require rapid integration with the host nervous system. Otherwise, muscle atrophy and functional recovery failure occur. The primary methods to encourage innervation of regenerating muscle after VML treatment are neurotization and rehabilitative exercise. However, because both methods result in only moderate success, advanced constructs with mature neural tissues must be developed to improve the therapeutic effects for VML or to better understand the development and maturation of NMJs.

The 3D bioprinting technique is advantageous in precisely integrating multiple cells and biomolecules within a complex entity and thus could promise effective incorporation of nerves with muscle constructs. Kim et al. investigated the feasibility of establishing a neural cell-integrated human skeletal muscle equivalent using a 3D bioprinting technique [143]. This construct was fabricated by 3D bioprinting of a hybrid bioink (HA/fibrinogen/gelatin/glycerol) containing HMPCs and/or human neural stem cells (HNSCs) onto a PCL anchor fabricated previously. To avoid hypoxia-induced cell death, fugitive Pluronic F-127 was utilized to fabricate the embedded channels. The involvement of neural cells in the skeletal muscle construct promoted myofiber generation, long-term survival, and NMJ formation in vitro. Moreover, the integration of HNSCs facilitated rapid innervation, restoration, maturation, and functionalization of muscle tissue in a rodent model of muscle defect injury, suggesting that the neural-skeletal muscle construct can be rapidly integrated with the host neural network, leading to accelerated muscle regeneration. In addition, the coaxial 3D bioprinting technique has also been utilized to co-print primary myoblasts and NSC-34 cells (a motor neuron cell line) [144]. Such a core/shell structure mimics the muscle/neuron physiological system by bringing differentiating motor neuron-like cells in proximity with skeletal myoblasts. Although the formation of mature NMJs has not been reported, the improved differentiation of NSC-34 cells in proximity to skeletal myofibers in the coaxial scaffolds is in accordance with the presence of skeletal myoblasts, which are known to enhance the development of neuromuscular junctions.