Engineering Large-Scale hiPSC-derived Vessel-integrated Muscle-like Lattices for Enhanced Volumetric Muscle Regeneration

Abstract

Engineering biomimetic tissue implants with human induced pluripotent stem cells (hiPSCs) holds promise for repairing volumetric tissue loss. However, these implants face challenges in regenerative capability, survival, and geometric scalability at large-scale injury sites. Here, we present scalable vessel-integrated muscle-like lattices, or VMLs, containing dense and aligned hiPSC-derived myofibers alongside passively perfusable vessel-like microchannels inside an endomysium-like supporting matrix using an embedded multi-material bioprinting technology. The contractile and millimeter-long myofibers are created in mechanically-tailored and nanofibrous extracellular matrix-based hydrogels. Incorporating vessel-like lattice enhances myofiber maturation in-vitro and guides host vessel invasion in-vivo, improving implant integration. Consequently, we demonstrate successful de novo muscle formation and muscle function restoration through a combinatorial effect between improved graft-host integration and its increased release of paracrine factors within volumetric muscle loss injury models. The proposed modular bioprinting technology enables scaling up to centimeter-sized pre-vascularized hiPSC-derived muscle tissues with custom geometries for next-generation muscle regenerative therapies.

eTOC blurb

We developed scalable vessel-integrated muscle-like lattices (VMLs) with dense, aligned hiPSC-derived myofibers and vessel-like structures using a novel bioprinting technology. These VMLs significantly improved muscle tissue regeneration and function in mice post volumetric muscle loss. These modular lattices can be adapted to large-scale muscle defects where native regeneration is impaired.

Graphical abstract

Introduction

Volumetric muscle loss is often caused by large area injuries and involves a complete loss of skeletal muscle. These injuries overwhelm the natural repair ability and typically lead to extensive fibrosis.[1] Because there are no extra cellular matrix (ECM) and satellite cells that can potentially differentiate into muscle fibers, regenerating volumetric muscle loss remains a significant challenge. Pathological environments with strong immune reactions, oxygen deficiency, and limited nutrient supply also hinder regeneration. Various therapeutic approaches for volumetric muscle regeneration include stem cell delivery, growth factor delivery, biomaterial implants, and grafting engineered muscle tissues as alternatives to autografts and allografts. However, these methods have independently proven ineffective in promoting functional muscle regeneration. Stem cells have excellent proliferation potential, differentiation ability, and regenerative functions for repairing injured tissue. However, conventional stem cell therapy (injecting stem cells into the defect area) has several limitations, including poor integration and viability at the defect site, resulting in minimal surface-level muscle regeneration and not achieving volumetric muscle tissue regeneration due to the lack of new ECM formation by stem cells alone.

Bioengineering clinically relevant muscle tissue implants that integrate in the host tissue and effectively regenerate requires reconstituting skeletal muscle architecture. This is a complex and delicate task that requires identifying and embedding skeletal muscle stem cells in a mechanically suitable scaffold, controlling cellular alignment and differentiation within the engineered scaffolds and ultimately expanding the culture to the size that matches the large defect area. Primary myogenic (progenitor) cells possess only limited proliferative capacity and lose their myogenic potential during serial in vitro passaging,[2] and face limitations in clinical translation as a treatment for volumetric muscle defects. In contrast, human induced pluripotent stem cells (hiPSCs) exhibit excellent proliferation potential [3,4] and the ability to differentiate to myogenic (progenitor) cells and regenerate skeletal muscle, similar to satellite cells.[5,6] The striking ability of hiPSC-derived muscle precursor cells (hiPSC-MPCs) (see Glossary) to self-organize by forming primary and secondary muscle fibers in a 2D in vitro culture system is similar to the muscle organization process observed in vivo.[7]

Another key factor in bioengineering clinically relevant muscle implants is to ensure they survive at the defect site, particularly in the first few hours of surgery. This is gauged by the extent to which implanted cells access nutrients and oxygen in the absence of culture medium, and how quickly the host’s blood infiltrates the implant. Volumetric tissue defects impose biologically harsh microenvironments (i.e., strong immune response and lack of oxygen, nutrient, biological reagents, and extracellular matrices (ECM)) which, in turn, impact the therapeutic ability of hiPSC-MPCs. Particularly at early stages of implantation, hiPSC-MPCs undergo cell death and dedifferentiation. This significantly decreases the therapeutic ability of hiPSC-MPCs to treat volumetric muscle defects, which recover along an extended timeline (~4 – 8 weeks) to reach restored muscle functionality and requires a robust sustainable pool of transplanted cells at the defect site throughout this period.[8–10] To fully realize the therapeutic ability of hiPSC-MPCs in volumetric muscle regeneration defects, a biomaterial must provide adequate stem cell environment, guide myogenic differentiation and maturation by providing instructive cues, and promote cell alignment.[11,12] Furthermore, blood vessel networks should be interwoven into the volumetric implants to ensure proper nutrients and oxygen delivery, and to serve as guides for the ingrowth of host endothelial cells (ECs) and hematopoietic cells, thereby achieving timely angiogenesis and preventing necrosis in the core of thick (> 1 mm) engineered tissues.[13,14] The rapid vascularization of volumetric muscle constructs based on hiPSC-MPCs is of paramount importance for the cell survival, myogenic differentiation, and maturation into functional myofibers, as these cells are metabolically active.[15–17] hiPSCs can also deliver various secreted factors including growth factors, cytokines, small RNA molecules, and extracellular vehicles (EVs) that contribute to the stimulation and drawing of host cells to the implanted area, and ultimately revive therapeutic activity at the defect site.[18–20]

A common approach for engineering 3D muscle implants involves the deposition or molding of cell-laden fibrin-based hydrogel yielding isogenic 3D hiPSC-derived artificial skeletal muscle constructs.[6,21–24] These methods can generate complex 3D constructs containing endothelium-and endomysium-like areas, albeit relying on the self-organization of ECs for vesicle formation with limited and uncharacterized perfusion capacity.[25] Indeed, these 3D constructs are fabricated by mixing and co-culturing ECs with muscle progenitors embedded in the hydrogel scaffold and have not been investigated in terms of the formation of a perfusable vascular network. Moreover, such strategies are hardly compatible with scalable integration of intricate hierarchical designs (e.g., native muscle tissue) and continue to fail in repairing volumetric muscle injuries. The constructed geometry, particularly for personalized injuries, is severely limited in this technology and would impede industrial scale up. Therefore, advanced microfabrication technique such as 3D bioprinting are being pursued.[26–32] However, many technologies still focus on developing 3D scaffolds with high printing resolution using various chemicals and reagents that sometimes induce cytotoxicity, especially in stem cells, or showing 3D constructs with cell lines that are excellently resistant to relatively harsh and vigorous external environments and reagents compared with stem cells. Therefore, to do bioprinting with stem cells, many factors should be considered and limited to select biomaterials, crosslinking reagents, crosslinking methods, addictive materials to improve printing resolution, and printing time to avoid loss of stem cell function and decrease of stem cell viability. Among numerous 3D bioprinting studies, there are few studies demonstrating therapeutic effects from 3D printed constructs with bioinks containing stem cells.[33,34] However, they must still be more advanced to achieve volumetric tissue constructs and regeneration in vivo.

Here, we propose a strategy to enhance the therapeutic effect of hiPSCs-derived implants by engineering vessel-integrated muscle-like lattices (VMLs) that emulate the intricate structural complexity and multi-dimensional hierarchy of native muscle, thus forming the basis for a new generation of muscle defect repair strategies (Fig. 1a). To fabricate scalable VMLs, we developed an unconventional embedded multi-material bioprinting (uEMB) method to enable rapid and high-resolution bioprinting of hiPSC-derived aligned myofiber and passively perfusable vessel-like lattices within a non-sacrificial, microporous, and mechanically robust but biodegradable endomysium-like supporting matrix (Fig. 1b). The endomysium-like supporting matrix provides structural support for relatively soft cell-laden or hollow microchannels in vitro and in vivo to recapitulate muscle architectural integrity.[35,36] The contractile and millimeter-long hiPSC-derived myofibers are formed in mechanically and structurally tailored ECM-based bioinks, which serve as scaffolds to ultimately guide the linear fusion of myoblasts. Incorporating a vessel-like lattice into the implants facilitates the delivery of oxygen, nutrients, and biological substances that improve the differentiation and maturation of hiPSCs-derived myofibers in large-scale constructs in vitro and host cell invasion and their growth in vivo. Finally, accelerated volumetric tissue regeneration is confirmed by in vivo volumetric muscle loss injury models by putting in the VMLs with two types of mice.

Figure 1. Large-scale skeletal muscle regeneration via enhanced regeneration inductive effect of hiPSC-derived vessel-integrated muscle-like lattices (VMLs).

(a) Schematic diagram depicts the therapeutic impact of volumetic tissue regeneration of hiPSC-derived VML implants on enhancing host cells and vessels invasion via secreted various biological factors including extracellular vesicles and cytokines. These high-frequency interactions prevent the large-scale implants core necrosis at the volumetric muscle tissue defects where lack of oxygen and biological reagents. (b) The proposed VMLs consist of millimeter-long hiPSC-MPC derived myofibers lattice around EC coated vessel-like lattice that was explored using unconventional embedded multi-material bioprinting (uEMB) to manufacture constructs that emulate the complex natural design of the native skeletal muscle, with an aim to enable the engineering of scalable and functional muscle tissues.

This study combines the following innovations. First, we create millimeter-scale iPSC-derived skeletal muscle constructs using 3D bioprinting technology for volumetric muscle loss treatment. Second, we use a multi-material bioprinting technique, directly fabricate perfusable macro vasculature and bifurcating channel network while allowing for individual optimization of the biomaterial properties of supporting bath, perfusable channels, and muscle channels (unlike molding technologies [6,21,25]). Third, we implement an efficient protocol for expansion and differentiation of iPSC-derived MPCs into skeletal muscle fibers in 3D constructs and thus eliminate the need to collect primary MPCs from human muscle biopsies [37]. Finally, we characterize the diffusion ability of the fabricated endothelialized networks and show the integration of micro-vasculature with the host’s in models of volumetric muscle loss injury.

Results

VMLs contains dense and aligned hiPSC-derived myofiber and vessel-like lattices

The uEMB strategy consists of directly printing a hiPSC-MCPs-laden bioink within a pregelled GelMA supporting matrix, unlike conventional embedded printing methods in which inks are typically printed within microgels and slurries.[35,36] To form millimeter-long myofibers, hiPSC-MPCs must be anchored via abundant Arginyl-glycyl-aspartic acid (RGD) binding sites on the engineered ECM matrix, which ultimately organize the linear fusion of myoblasts, similar to the native ECM.[38,39] The bioink for muscle fibers consists of gelatin and GelMA, both of which contain denatured collagen, a major ECM component of skeletal muscle tissue with abundant RGD binding sites. By adjusting the concentration of GelMA and the crosslinking density, we can easily tune the porosity and degradation of the hydrogels[40] in order to achieve adequate mechanical support and degradation behaviors for the engineering of myofibers with hiPSC-MPCs. The role of the GelMA supporting matrix is comparable to the hierarchical ECM structure of the native endomysium, which is mainly composed of type I and type III collagens.[41] This surrounding connective tissue physically supports densely bound aligned myofiber bundles and perfusable vessels,[42] preventing structural collapse during muscle contractions and movements. In addition, the incorporation of GelMA pre-polymers in both the supporting matrix and the bioink prevents the delamination of the bioprinted layers upon photo-crosslinking.

The thermally reversible gelation properties of both gelatin and GelMA enable a straightforward temperature-controlled printing process.[43,44] Complex multi-layered patterns can be fabricated by harnessing the reversible thermal-healing properties of the GelMA supporting matrix just below the gelation temperature (between ~8 °C and ~20 °C, sol-gel state) (Figure S1 and S2). To maintain shape fidelity, the bioink stiffness could be increased by increasing polymer concentration (Figure S2a). However, higher mechanical stiffness is a tradeoff that reduces biocompatibility with soft tissues and laden cells. Alternatively, gelatin can be added to the GelMA bioink to temporarily increase its stiffness and viscosity for bioprinting, followed by a post-crosslinking removal step at ~37 °C,[45] creating non-merged parallel channels (Figure S2b). By adjusting ink flow rates and nozzle translational speed, the diameter of the bioprinted channels could be tuned (160 ± 8 to 220 ± 16 μm) (Figure S2c and d). Changing the center-to-center inter-channel distance did not significantly affect the shape fidelity and the distance could be reduced to 320–350 μm before the channels started to merge (Figure S2e). To obtain aligned channels resembling muscle architecture, we printed vertically aligned microfibers within the supporting matrix with this minimal spacing, thus increasing the density of the muscle tissue. We set out to use a photo-crosslinking strategy to transform the embedding bath, including the printed patterns, into a solid hydrogel scaffold that was robust enough to enable in vitro culture, in vivo implantation, and ultimately, muscle tissue repair. To this end, various concentrations of GelMA and photo-crosslinking conditions (i.e., UV exposure time) were assessed to obtain a solid construct with mechanical properties optimized for muscle stem cell differentiation and maturation. This screening was also based on our previous studies that focused on identifying the necessary mechanical properties for a hydrogel to mimic the native skeletal tissue ECM (i.e., 10–50 kPa and >30% elastic strain).[46] Using a 5% (w/v) GelMA matrix, we investigated stiffness as a function of UV exposure duration (Figure S3a). The optimum UV condition (55 s, 5.5 J/cm²), with a matrix stiffness of ~8 kPa after 21 days of in vitro culture (Figure S3b), resulted in a stable construct that prevented the collapse of the printed patterned geometries during long-term tissue culture. It was observed that HUVECs embedded in 5% (w/v) GelMA started to proliferate after three days of in vitro culture, while HUVECs within 7% (w/v) GelMA did not (Fig. S3c).

The uEMB of the recurring micropatterns was demonstrated by rapidly printing various longitudinally aligned patterns with high printing density within a relatively large-sized supporting bath (~1 cm³), thus abstractly representing the vessels and muscle fibers in a native muscle fiber bundle (Figure 2a, b). Longitudinal patterns could also be combined with the printing of transversal patterns (Figure 2c). Complex 3D shapes such as fractal-like branched networks containing aligned microfibers could also be printed in this supporting gel to recapitulate the architecture of four different types of natural vascularized skeletal muscle flaps (Figure S4a, < 5 min print time per construct). The proposed uEMB technology enables the creation of large vascularized tissue implants by adjusting the size and geometry of the supporting matrix. Alternatively, centimeter-scale building blocks can be assembled into larger constructs using 3% (w/v) GelMA as light-curable glue. For instance, combining nine uniquely patterned blocks forms large, complex, and perfusable 3D constructs with volumes up to 10 cm³ (Figure S4b).

Figure 2. Fabrication of VMLs contain hiPSC-derived millimeter-long myofiber and vessel-like lattices using the uEMB.

(a-c) Various designs of 3D printed VMLs composed of aligned and patterned microfibers via optimized bioinks formulations and printing parameters, as visualized using inks spiked with red and green polystyrene. (d) Transversal and longitudinal cross sectional scanning electron microscopy (SEM) imaging to evalaute hollow microchannel lattcie inside the microporous supporting matrix. (e) Oxygen concentration in the microporous supporting matrix by the inter-channel distance between the hollow microchannel lattice (‘**’: p<0.001; Kruskal Wallis test; n=3). (f, g) Seeding HUVECs into the hollow microchannels resulted in the formation of endothelialized microchannels throughout the construct as confirmed by confocal fluorescence imaging of positive CD31 staining after three weeks of culture. (h) Ve-cadherin (green) and CD31 (red) expression indicated endothelial cell junctions of the HUVECs coated on wall of the hollow microchannels in 3D printed iPSC-MPCs-laden constructs after three weeks of culture. (i) Optimal formulation of hiPSC-MPCs (> 20 × 10⁶ cell/mL)-laden 2% (w/v) gelatin/7.5 (w/v) GelMA hydrogel supported the long-term 3D in vitro culture and differentiation of hiPSC-MPCs indicated by positive (j) Actn2, Pax7, MyoG, and (k) Titin and MyHC stainings. (l) Calcium transition analysis of spontaneous contractile activity of hiPSC-derived myofibers in the optimizied gelatin/GelMA hydrogels after 15 days differentiation. (m) The optimized hiPSC-MPCs laden hydrogel formulation also formed a bioink that could be printed within a GelMA support matrix to engineer patterned (i.e., aligned) muscle fibers spanning the entire construct as confirmed by confocal fluorescence imaging of viable actin-stained cells. (n) MyHC and ACTN2 expression and millimeter-long myofiber in the printed hiPSC-MPCs laden gelatin/GelMA hydrogel channels were observed on day 20. (o) SEM showed aligned nanofibrous structures that were uniquely present in the printed gelatin/GelMA hydrogel channels, but not in the GelMA support matrix.

An efficient way to create open channels within thick, soft biomaterials is to use sacrificial materials like agarose, carbohydrate glass, or gelatin as templates.[47–50] To create perfusable vascular-like networks, we formulated a thermo-reversible gelatin ink, which could be removed by warming up the photocured GelMA supporting matrix to 37 °C in PBS or cell culture media. The method readily enabled the fabrication of a bifurcating channel network within a ~1 cm³ supporting bath and enabled the controlled screening of bioengineering parameters such as vessel diameter, number, and density. Although the fabricated channels (diameter = 148.2 ± 4.3 μm) were larger than the native blood vessels in skeletal muscle, we anticipated that the addition of such perfusable vessels inside a large implant could accelerate the host’s blood invasion post-implantation, facilitate the formation of micro-capillaries, and ultimately, promote implant integration. Given the average diameter sizes (184.4 ± 22.3 μm for the muscle channel and 148.2 ± 4.3 μm for the perfusable channel), we were able to achieve a perimeter to perimeter distance of ~183.7 μm between muscle and perfusable channels which is comparable to the diffusion limit of the native skeletal muscle environment and thus helps with the engineering of large viable tissues.[51–53] Scanning electron microscopy (SEM) imaging of freeze-dried constructs confirmed printed channel patency (Figure 2d), which was corroborated by successfully perfusing the channels with fluorophores that also represented the perfusion and diffusion of nutrients, growth factors, and waste products in these constructs (Figure S5a and Movie S1). Time-lapse confocal fluorescence imaging revealed a uniform diffusion gradient of the fluorescent-labeled dextran (~20 kDa) within the supporting matrix, which was characterized by a relatively high diffusional permeability (P_d = 7.25 ± 0.675 × 10⁻⁵ cm s⁻¹), thus favoring the exchange of cellular nutrients and waste products and corresponding to previously reported values (Figure S5b).[54][55] We also observed excellent oxygen permeability confirmed by a significantly higher oxygen level in the supporting matrix containing vessel-like lattices that printed with 350 μm distance (Fig. 2e). Furthermore, HUVECs grew along the channel wall, thereby forming an endothelialized vessel-like lattice that covered the entire channel’s interior throughout the construct as confirmed by CD31 and VE-cadherin expression (Fig. 2f–h).

Next, we set out to explore the optimal bioink conditions for hiPSC-MPCs to mature and differentiate inside perfusable bioprinted constructs. Among the printable gel mixtures inside a 5% GelMA supporting matrix 7.5% (w/v) GelMA bioink (stiffness = 40±8 kPa, further reduced to 22±2 kPa after 3 weeks of incubation in standard culture medium) provided a 3D microenvironment that was mechanically ideal for the myogenic induction of hiPSC-MPCs (i.e., 10–50 kPa and >30% elastic strain) (Fig. S6a–c).[46,56] To this end, 3- to 4- week old hiPSC-derived progenitor cells (NCRM-1) were obtained (Fig. S6d)[5,57] and encapsulated in 7.5% (w/v) GelMA and cultured the constructs for 4 weeks with the goal to investigate cellular behavior modulated by UV crosslinking and cell density (Fig. S6e and f).[58–60] A higher cell density (> 20 × 10⁶ cell/mL) was most compatible for the development of striated myofibers with well-organized sarcomeres, while also containing a pool of satellite cells, indicated by positive Actn2, Pax7, MyoG, Titin, and MyHC staining (Fig. 2i–k and S6g, h). Additionally, the spontaneous contractile activity of in situ formed hiPSC-derived myofibers was emulated in the optimized muscle bioink (Fig. 2l and Movie S2). Also, the incorporation of another hiPSC cell line (GM23338; TTN-GFP) in the muscle bioink showed striated myofibers with well-organized sarcomeres with no statistically significant difference in the expression of key markers, compared with that of NCRM-1 cell line constructs (Fig. S7).

Using the uEMB with high cell concentration-laden bioinks that are typically difficult to print with conventional bioprinting techniques, we successfully obtained vertically-aligned hiPSC-MPC-laden microfibers at a density of ~7 fibers/mm² with high cell viability (>85%) (Fig. 2m and S8). After 20 days of differentiation, the printed hiPSC-MPCs-laden microfibers showed striated, millimeter-long, tertiary muscle fiber structures (Fig. 2n), which might be induced by aligned and nanofibrous morphology of the bioinks formed by the shear stress of the printing nozzle when the muscle bioink extruded (Fig. 2o).[61] Implanted constructs undergo incessant physical stress and friction at the implant-host tissue interface during body movements. However, constructs with printed fibers and channels remained intact for 30 days of in vitro culture, showing no significant differences in stiffness and elongation at break compared to the bulk supporting matrix without printed channels and fibers (Fig. S9). Furthermore, the printed constructs showed persistent elastic behavior during cyclic mechanical loading up to 40% strain and did not demonstrate any fatigue, a behavior similar to native skeletal muscle that tolerates cyclic contraction-relaxation stress with excellent elasticity during muscle activity.[62,63] These results indicate that the mechanical properties of the printed constructs could aid in preventing the collapse of perfusable vessels and could physically support the aligned myofibers against cyclic stresses during the regeneration process after in vivo implantation. The maximum distance of hollow channels for maintaining high cell viability was ~350 μm (Fig. S10) and ~1:4 hollow channel-to-fiber ratio rescued the engineered myogenic phenotype. The printed hiPSC-MPCs within the supporting matrix with and without vessel-like lattice demonstrated high cell growth, alignment, and spatial confinement confirmed by F-actin staining (Fig. 3a and 3b). However, the printed hiPSC-MPCs in vessel-like lattice integrated VMLs showed higher Pax7, MyHC, and MyoG expression, and improved myofiber alignment that was confirmed by a single peak orientation (Fig. S11) and ~5–10-fold higher fusion index (Fig. 3c). In the endothelialized vessel-like lattice integrated VMLs, the printed hiPSC-MPCs showed significant higher levels of adult muscle protein (MYOG, MYH1, MYH2, MYH3, MYH7 and MYH8) indicating both accelerated proliferation and maturation of myofibers compared to the VMLs without a vessel-like lattice at day 20 (Fig. 3d). Therefore, efficient endothelialization, which can transfer biological factors and oxygen in the large-sized VMLs, enhanced myogenic differentiation, leading to MyHC and ACTN2 expression (Fig. S12).

Figure 3. Effect of vessel-like lattice on the maturation of hiPSC-derived myofibers at VMLs and its degeneration in harsh microenvironments.

(a) After up to three weeks of in vitro culture, 3D printed muscle fibers were still viable and functional as confirmed by positive Actn2, Pax7, MyoG, and MyHC immunostaining. Bioprinted hiPSC-laden constructs with vessel-like lattice exhibited enhanced expression of myogenic stem cell and muscle maturation markers over the group without lattice. (b) Alignment index of printed hiPSCs in constructs with and without the vessel-like lattice was measured using fast Fourier transform analysis. A sample of F-actin-stained images of the constructs, with and without the vessel-like lattice, processed and used for this analysis, is provided in Figure S15. Accordingly, both groups revealed highly aligned cellular fibers with no statistically significant difference (‘ns’: p=0.667; Mann-Whitney unpaired t test; n=4). (c) Fusion index of matured muscle fibers expressing MyHC in 3D printed constructs after 10 and 20 days of culture. Quantifying the expression percentage of these markers (i.e., % specific marker positive cells among the total number of DAPI-positive cells in a specific region of interest) revealed significant differences between the non-perfusable versus perfusable constructs (‘*’: p<0.01; ‘**’: p<0.0019; ‘***’: p<0.0004; ‘****’: p<0.0001; Two-way ANOVA Sidak’s multiple comparisons test; n=3). (d) RT-PCR gene expression analysis of 3D printed constructs with and without HUVEC-coated vessel-like lattice recorded on day 10 and 20 of culture. Y-axis represents expression level of various genes involved in the differentiation of hiPSC-MPCs to skeletal muscle relative to the undifferentiated hiPSC-MPC control, normalized to the housekeeping genes (‘**’: p<0.001; ‘*’: p<0.05; Kruskal Wallis test; n=3). (e) Schematic diagram depicting how differentiation and cell viability of hiPSC-MPCs is impacted by a subset of environmental conditions (normoxia/hypoxia, with or without differentiation reagents supplements (w and w/o Diff reagents, respectively). (f, h) Comparison of cell viability in hiPSC-MPCs under anoxic and normoxic conditions with or without differentiation reagent enrichment (‘****’: p<0.0001, ‘***’: p<0.0005, ‘**’: p<0.002, ‘*’: p<0.05; Two-way ANOVA Tukey’s multiple comparisons test; n=5). (g) Comparison of MyHC expression of hiPSC-MPCs under anoxic and normoxic conditions with or without differentiation reagent enrichment. (i) Comparison of % of fusion index upon myogenic differentiation of hiPSC-MPCs under anoxic and normoxic conditions with or without differentiation reagent enrichment (‘****’: p<0.0001, ‘***’: p<0.0005, ‘**’: p<0.002, ‘*’: p<0.05; Owo-way ANOVA Tukey’s multiple comparisons test; n=3).

Maturation of hiPSC-derived myofibers in VMLs and their degeneration in harsh microenvironments

To understand the behavior of hiPSC-MPCs-laden implants at the volumetric muscle defect area, we cultured the hiPSC-MPC-laden hydrogel after one week of differentiation in the oxygen and biological reagents depletion conditions (Fig. 3e). Although these conditions do not entirely mimic muscle defect area, we expect a rapid decrease in oxygen concentration due to cellular oxygen consumption and constrained delivery of oxygen and nutrients into the large scale cell-laden hydrogels from host skeletal muscle and vessels (~5%).[64] The severe hypoxic/diff. reagent-starved group underwent cell death or de-differentiation within the first 7 days as observed by mononucleated morphology (Fig. 3f–i). The severe hypoxic/diff. reagent-rich group survived an additional week but eventually underwent the same process. This could be attributed to the ischemic necrosis caused by inefficient transfer of both biological factors and oxygen in these constructs. On the contrary, the normoxic/diff. reagent-supplied group demonstrated >90% cell viability across the 14 days of the study as well as ~77% higher muscle fusion by day 14 as compared with the severe hypoxic/diff. reagent-starved group. Following these results, we concluded that further measures such as aided-endothelialization through the incorporation of vessel-like components is needed to accelerate host’s blood invasion in implants for oxygen and nutrients supply in 7 days.

Enhanced vascularization of endothelialized vessel-like lattice integrated constructs in vivo

To quantitatively investigate the impact of vessel-like lattice on rapid vascularization, we explored the subcutaneous in vivo grafting of the printed constructs (~1 cm³) with four different designs (G1-G4) (Fig. 4 and S13). One-week post-implantation, endothelialized vessel-like lattice with (G1) and without (G2) human umbilical vein endothelial cells (HUVECs) encapsulated in the supporting matrix were infiltrated by red blood cells indicating anastomosis with the host (Fig. 4b and 4d). After three weeks, G1 and G2 demonstrated full infiltration with many host capillaries into the HUVEC coated channels (Fig. 4b and 4d, (ii)), which in turn increased by 1.5 to 2-fold over time (Fig. 4e and 4f). Moreover, encapsulated HUVECs in the supporting matrix (G2) proliferated (Fig. S3c) and promoted sprouting near the edges of the printed HUVEC-coated channels in vitro (Fig. 4c). This behavior was observed in vivo, whereby the migration and outgrowth of transplanted HUVECs connected to the neighbor channels (Fig. 4d (ii)). In G1 and G2 specifically, after one week, several vessels consisting of concentrically patterned rat and human ECs were observed in the surrounding tissue area (Fig. 4b and 4d, (iii)), corroborating with the excellent graft-host integration, similar to previous studies.[65,66] In the tissue surrounding G2, a prevalent presence of migrated HUVECs from the pre-vascularized channels to the host-tissue interface was observed in comparison with G1, possibly due to larger HUVEC density and the formation of sprouting networks at a higher rate (Fig. 4b and 4d, (iv)). Moreover, the presence of endothelialized channels significantly increased the survival of cells in the transplanted constructs compared with the control, HUVEC-laden bulk gel (G4) (Fig. 4g) and induced relatively fast degradation compared with acellular hollow channel in the supporting bath (G3) and G4 (Fig. 4h). Presumably due to their endothelialized vessel-like lattice, the VMLs allowed for accelerated biomaterial degradation through the accumulation of host cytokines including TNF-α, which would a trigger foreign body response and an increased expression of metalloproteinase1/3 (MMP1/3), leading to GelMA degradation.[67]

Figure 4. Rapid vascularization of endothelialized vessel-like lattice integrated constructs in vivo.

(a) Endothelialized channels in the supporting bath (G1). Seeding HUVECs into the hollow lattice resulted in the formation of endothelialized vessel-like lattices throughout the construct as confirmed by confocal fluorescence imaging of positive CD31 staining after three weeks of culture. (b) H&E stained and immunostained images of G1 harvested on week 1 and week 3. (c) Endothelialized channels in the HUVEC-laden supporting matrix (G2). The embedding HUVECs in the GelMA (i.e., in addition to seeding HUVECs in the channels) promoted sprouting from the endothelialized channels into the bulk material. (d) H&E stained and immunostained images of G2 harvested on week 1 and week 3. Red blood cells are indicated with yellow arrows in H&E images. Human-specific CD31 is colored in red and rat-specific CD31 is colored in green. Human-specific CD31 is colored in red and rat-specific CD31 is colored in green. (e) Number of total human and rat vessels per high-power field (HPF) found in the supporting bath and inside the channels after 1 and 3 weeks of in vivo culture (‘ns’: p=0.57; Two-way ANOVA Sidak’s multiple comparisons test; n=3). The formation of human and rat vessels could have been triggered much faster in presence of the ECs in the G2 supporting matrix (f) Number of vessels found in the ratpositive and human-positive CD31 immunostained samples after harvest on week 1 and week 3. (‘ns’: p=0.8; Two-way ANOVA Sidak’s multiple comparisons test; n=3). In the surrounding tissue, rat vessels were initially found more prevalently than human vessels. However, as the HUVECs grew out into the host tissue, the ratio of rat to human vessels decreased over time.. (g) Comparing the percentage of apoptotic cells in groups G2 and G4 using the results of TUNEL staining (‘**’: p<0.0062; unpaired t-test, n=4). (h) Remaining hydrogel area after harvesting the implanted samples on week 1. (‘**’: p<0.0027; ‘***’: p<0.0002; ‘****’: p<0.0001; One-way ANOVA Tukey’s multiple comparisons test; n=5).

hiPSC-derived VMLs enhance muscle regeneration and functional recovery

For evaluating therapeutic ability, hiPSC-derived VMLs were implanted into a full thickness muscle defect model with a cylindrical injury (~16π mm³) in the posterior muscle compartment in the lower leg involving the gastrocnemius and soleus in the immunodeficient mice (NOD-scid IL2Rgamma^null strain) (Fig. 5a).[68,69] To prevent the sample dislocation and to facilitate integration with the host, we integrated the implants with suturable nanofibrous scaffolds made of a mixture of polyglycerol sebacate and polycaprolactone (PGS/PCL). Previously, we demonstrated the biocompatibility and biodegradability of these electrospun scaffolds for in vivo wound repair and stimuli-responsive drug delivery.[70] Here, we used a photo-crosslinkable GelMA glue to attach the 3D bioprinted constructs to the scaffolds using UV irradiation (Figure S14). Successful adhesion was revealed by SEM imaging of the hydrogel-nanofibrous scaffold interface (Figure S14a). The mechanical characterization of the sutured nanofibrous electrospun scaffolds revealed an elastic modulus of 16.4 ± 5.8 MPa and an ultimate tensile strength (UTS) of 1.5 ± 0.7 MPa, which were comparable to reported values (Figure 5b,c and S14b,c).[71] Stretching the composite scaffold did not result in fracture at the suture point, suggesting that the suture could tolerate high mechanical stress and large dynamic movement post-surgery. After 4-weeks post-surgery, we observed ~< 3.6 % area fibrosis in the V3 group with a significant reduced fibrotic area compared to that observed in the untreated muscle defect (V1) and the acellular VML (V2) (Fig. 5d and 5e). Moreover, the V3 group samples at the 8-week harvest timepoint exhibited a significantly smaller remaining hydrogel (Fig. 5d). We identified newly generated myofibers with MyHC expression and centrally located nuclei (shown with arrows) at a higher quantity in the V3 group compared to the control conditions (Fig. 5f and 5g).

Figure 5. hiPSC-derived VMLs enhance muscle regeneration and functional recovery using full thickness muscle defect model in NOD-scid IL2Rgamma^null mice.

(a) The hiPSC-derived VMLs can be directly sutured into the volumetric muscle loss injury site via a suturable PGS/PCL graft. Harvested quadriceps after 8 weeks implantation of the PGS/PCL graft showed that printed constructs integrated into the volumetric muscle loss injury site. (b) Tensile stress-strain curve of the two sutured PGS/PCL grafts (n=3). (c) The interface between the GelMA supporting matrix and the PGS/PCL graft that were strongly attached to each other. (d) Quantification of the fibrotic tissue and remaining hydrogel area in all conditions 4 weeks and 8 weeks post-surgery (‘****’: p<0.0001, ‘***’: p<0.001, ‘*’: p<0.05; Two-way ANOVA Sidak’s multiple comparisons test; n=3). (e) Masson trichrome staining images of 3D VML and control groups 8 weeks post-surgery. The implanted hydrogel site is marked with arrows. (f) Immunostaining the samples against MyHC (green) and DAPI (blue) showed the presence of muscle with centrally located nuclei as denoted by the white arrows. (g) The number of myofibers identified with centrally located nuclei per high power field 8 weeks post-surgery in harvested samples of all conditions (‘****’: p<0.0001; Two-way ANOVA Sidak’s multiple comparisons test; n=15). (h) Human CD31 (red)/mouse CD31 (green)/DAPI immunostaining images of harvest samples near the implanted samples in V3 and V2 groups as well as the untreated injury site in V1, 8 weeks post-surgery. (i, j) Quantification of the (i) number (‘****’: p<0.0001; Two-way ANOVA Sidak’s multiple comparisons test; n=15) and (j) diameter (‘ns’: p=0.48; Two-way ANOVA Sidak’s multiple comparisons test; n=15) of the vessels found in the harvested samples 8 weeks in post-surgery. (k) Co-staining of βIII-tubulin (red)/AchR (green)/DAPI (blue) in the three volumetric muscle loss groups (V1 = untreated injury, V2 = 3D acellular gel, V3 = 3D vascularized muscle implants, VML) 8 weeks post-surgery. (l) Quantification of AchR-positive muscle fibers per high power field in all conditions (n=10), including V0 = sham. (m-o) Muscle function recovery was measured in ankle plantar flexion torque response and in situ twitch and tetanus test. All tests performed using One-way ANOVA Tukey’s multiple comparisons test: (m) Torque (‘****’: p<0.0001; ‘***’: p<0.0008; ‘**’: p<0.032; ‘ns’: p=0.2615; n=6), (n) Twitch strength (‘**’: p<0.0021; ‘*’: p<0.0330; n=6), (o) Tetanus Strength (‘***’: p<0.0007; ‘*’: p<0.0492; n=6). The forces in n and o were normalized to animal’s body weight.

A significantly large number of vessels expressing human and mouse CD31-positive cells were prevalently found among mouse vasculature near the implant interface and the surrounding host tissue in the V3 compared to the control groups (Fig. 5h and 5i), in line with the subcutaneous study (Fig. 4d). Although the average diameter of vessels was not significantly different in all groups (Fig. 5j), we observed larger vessels (>100 μm in diameter) expressed with mouse and human CD31-positive cells in V3 (Fig. 5h, Fig. S15a). Host neuronal invasion and formation of neuromuscular junctions was relatively high in the V3 group (co-expression of acetylcholine receptor (AChR) and βIII-tubulin (TUJ1) at the host-scaffold interface) compared to V2 and V1 groups (Fig. 5k, 5l and Fig. S15b).[72] This result was harmonious with the improved muscle function recovery observed in the V3 group (Fig. 5m–o), suggesting that new neuromuscular junctions could have contributed to improved functional muscle recovery.[37] The V3 group exhibited a significant increase in ankle plantar flexion torque response compared to the V1 group and gradual average improvement from weeks 4 to 8, as well as values closer to the average sham torque than other groups (Fig. 5m). Moreover, the presence of well-integrated contraction units and muscular junctions in the V3 group suggested higher muscle recovery over other groups.[73] This observation was corroborated from the excitation of the isolated gastrocnemius (GA) muscle during in situ twitch and tetanus test which revealed a significant increase in maximal twitch force and tetanic muscle contraction in V3 group compared to V1 and V2 groups (Fig. 5n and 5o).

We investigated the donor origin of the regenerated muscle 8 weeks post-implantation to assess whether the newly formed myofibers were triggered by the implanted hiPSC-MPCs. We observed a regeneration area that showed a similar morphology of staining as to previous studies.[24,57,68,74–76] Especially, we observed that distinct expression of human Lamin A/C (hLamin A/C) in the V3 group overlapped with nuclei similar to the positive control result, and a behavior not observed in the V2 group (Fig. S16a). Clear co-expression of the MYH/MyH3 and human Leukocyte antigen (HLA-A) at the interface of implanted V3 group with the host muscle tissue suggested that the newly generated myofibers were of human origin (although constituting a small cell number) (Fig. S16b and 16c). These results were similar to the HLA-A/MHC staining result shown in research by Kim and colleagues.[77] However, there remained inconclusive evidence of the transplanted hiPSCs-MPCs within the hydrogel area or the muscle itself due to the non-specific expression of hLamin A/C.

Cross-validation of muscle regeneration with hiPSC-derived VMLs in humanized mice

To further validate behavior of the implanted hiPSC-MPCs, hiPSC-derived VMLs were implanted at a cylindrical defect (~16π mm³) in the posterior muscle compartment in the lower leg involving the gastrocnemius and soleus in a humanized mice (RAG2−/−γc−/− strain) that has the stable absence of T and B cells and has been shown to confirm xenotransplantation of human cells (Fig. 6a).[78,79] Immunostaining harvested samples after 1- and 4-weeks post-surgery revealed a clear distinction among groups that showed significantly increased expression of hLamin A/C markers overlapped with MyHC marker in the V3 group, particularly at the areas adjacent to the implanted constructs (Fig. 6b–d). In 1-week harvested samples, hLamin A/C was found sparsely (and only) inside printed channels while this expression expanded to the rest of the implant site at the 4-week timepoint due to degradation of the supporting matrix (Fig. 6e). The expression of hLamin A/C overlapping with MyHC was however predominantly observed in the innermost areas of the implant compared to the peripheral sections of the hydrogel or inside native host muscle area (Fig. 6f), suggesting partial invasion of the implanted human cells to native host muscle tissue. Specifically, in V3 samples, co-staining of hLamin A/C, hSpectrin and hDystrophin (i.e., cytoplasmic markers of human muscle fiber), verified the presence of human cells (Fig. 6g) which was not observed in V2 samples (Fig. S17). Clear co-expression of Pax7 and hLamin A/C in the hiPSC-laden group confirmed that the implanted constructs were able to maintain and migrate hiPSCs into the host’s tissue (Fig. 6h, i). In the RAG2−/−γc−/− strain, the number of human specific vessels formed in the V3 group was statistically significant compared to the control groups (Fig. S18). Interestingly, human CD31⁺ vessels were more observed in RAG2−/−γc−/− strain confirmed by compare between Fig. 5i and S18. These results suggested that vessel invasion was accelerated in V3 as in agreement with the higher levels of VEGF secretion observed in hiPSC-derived VMLs over time (Fig. 6j).

Figure 6. Cross-validation of muscle regeneration with hiPSC-derived VMLs in RAG2−/−γc−/− mice.

(a) Schematic image depicting host invasion and migration of hiPSC-derived myofibers and HUVECs outwards into the hydrogel and surrounding area 1 week and 4 weeks post implantation. (b) Co-staining of human lamin A and lamin C (hLaminA/C, green) with myosin heavy chain (MyHC, red) in V1, V2 and V3 groups 1 week and 4 weeks post-surgery. Yellow arrows on the magnified images indicate areas with overlapping Lamin A/C, DAPI and MyHC staining, an indication of cells with human origin. (c) Quantification of hLaminA/C-positive cells per high power field (HPF) in the hydrogel area for V3 and V2 (‘****’: p<0.0001; Two-way ANOVA Sidak’s multiple comparisons test; n=10). (d) Quantification of hLaminA/C signals overlapped with MyHC positive cells per high power field in the hydrogel area for V3 and V2 (‘****’: p<0.0001; Two-way ANOVA Sidak’s multiple comparisons test; n=10. (e) V3 samples at 1 and 4 weeks after implantation immunstained with hLamin A/C (green) and MyHC (red) demonstrating presence of these markers first around the channel area (week 1) and later all across the hydrogel implant area with (f) a representative magnified image of these samples indicating bulk of hLamin A/C expression within the boundaries of the implant area after 4 weeks. (g) Co-staining of hLaminA/C (green) with human Spectrin (top row, green) and human dystrophin (bottom row, green) 4 weeks post-surgery. (h) Co-staining of hLaminAC (green) and Pax7 (red) 4 weeks post-surgery. Yellow arrows indicate nuclei with overlapping staining, indicating the specificity toward cells with human origin. (i) Quantification of hLaminA/C signals overlapped with Pax7 positive cells per high power field in the hydrogel area for V3 and V2 (‘****’: p<0.0001; Two-way ANOVA Sidak’s multiple comparisons test; n=10. (j) Secreted VEGF from the 3D printed muscle tissue with aligned hiPSC-MPCs microfibers (n=3). DAPI staining is indicated by blue across all images.

Discussion

Endowing the large-scale constructs with organized and functional vessel networks is of vital importance for sustaining the therapeutic function of hiPSC-derived skeletal muscle cells, while also promoting implant-host integration and facilitating the delivery of biological factors in volumetric muscle loss injuries. In previous studies, 3D hiPSC-laden fibrin hydrogels [6,21–24], 3D bioprinted vasculature with decellularized ECM [80], and the complex 3D architecting of soft hydrogels using in-situ multi-photon-based printing [81] have shown improved muscle regeneration. However, these strategies are hardly compatible with the scalable integration of intricate hierarchical designs of the native muscle into engineered centimeter-scale muscle tissue using hiPSCs.

The current study is the combinatory work to directly fabricate seamlessly integrated and scalable VMLs containing hiPSC-derived myofiber and endothelialized vessels through modulating the geometry of the endomysium-like supporting matrix while allowing for individual optimization of the biomaterials properties for achieving excellent hiPSC-MPCs maturation and differentiation. This endomysium-like supporting matrix physically supported densely bound aligned myofiber- and vessel-like lattices preventing structural collapse against constant physical stress and friction at the implant-host tissue interface during body movements, resulting in resembling the function of native ECMs. An efficient protocol to expand and differentiate hiPSC-MPCs that were anchored via abundant RGD binding sites on the aligned nanofibrous 3D matrix, which ultimately organize the linear fusion of myoblasts, to form millimeter-long and contractile muscle fibers, was implemented.[38,39] This eliminates the need for primary MPCs collection from human muscle biopsies as formerly pursued in other studies.[37] Also, these results highlight the capacity of hiPSC-laden printed fibers to manufacture mesoscale functional muscle tissue in vitro.

Pre-endothelialized vessel-like lattice and supporting matrix showed a significant impact on host vessel infiltration to aid sustained oxygen and biological reagent supply at the time of implantation to the hiPSC-derived VMLs. This finding was previously observed in organized human EC constructs patterned into cords which acted as templates for guided vascularization in vivo.[82] Therefore, the VMLs well incorporated in host tissue and showed significantly decreased fibrotic tissue formation and higher muscle regeneration along with large-sized vessels and functional recovery in defect sizes larger (~40% muscle removed) than in previously reported mice defect models (Table 1). We verified the human origin in the newly formed myofibers, induced by the implanted hiPSC-MPCs, through cross-validation with two different mice strain (Immune-deficient mice (NOD-scid IL2Rgamma^null) and Humanized mice (RAG2−/−γc−/−)). Significantly higher expression of hLamin A/C markers overlaps with MyHC marker in the VMLs. Also, the behavior of implanted human cells would be continual and directed in the absence of an immune response, with the conspicuous expressions of human specific markers such as hLaminc A/C, hSpectrin and hDystrophin. Observing the definitive presence of human markers in the implantation area confirmed that the hiPSC-MPCs at the transplantation site play a significant role for improving recovery after volumetric muscle loss by contributing to significantly greater functional muscle recovery.

Table 1.

Comparison of type of implants, fabrication technique, volumetric muscle loss defect size, and animal type in several studies and current our study.

Implant (sample)	Fabrication technique	Size of the implant	Defect size	Animal	Ref.
hiPSC-MPCs laden GelMA hydrogel	uEMB	4πx4 mm3	4π x 4 mm3 (50.24 mm3, 26.9 ± 3.1mg)	Mice-RAG2−/−γc−/−-NOD-scid IL2Rgammanull	Our study
Muscle stem cells (MuSCs) and muscle resident cells (MRCs) laden ECM-based scaffold	Blending prepolymer solution in bioreactor	2×7×2 mm3	2×7×2 mm3 (28 mm3, 15 ± 1.5 mg)	Mice-NOD/SCID-NSG	[109]
Laminin enriched fibrin hydrogel	Punching after blending prepolymer solution	6 mm punching construct (not mentioned size)	3mm biopsy punch (18.56 ± 0.41 mg)	Mice-C57BL6 mice	[110]
Minced muscle	After mincing muscle, implantation	1 mm3	2mm biopsy punch (8.1 ± 0.3 mg)	Mice-C57BL/6J mice-Foxn1nu mice	[111]
Fibrin microthreads loaded with HGF	Filing hydrogel in injury site	4×2×2 mm3	4×2×2 mm3 (16 mm3, ~30mg)	Mice-SCID mice (SHO)	[112]
hiPSCs with fibrinogen and thrombin solution	Filing hydrogel in injury site	3×3×6 mm3	3×3×6 mm3 (54 mm3)	Mice-NSG mice	[24]
Minced muscle	After mincing muscle, implantation	3×3×1.5 cm3	3×3×1.5 cm3 (~5g)	Pig- Yorkshire-cross Pig	[98]
skeletal muscle progenitor cells	Delaminating monolayer of myogenic differentiated skeletal muscle progenitor cells	0.25π x 14 ± 1 cm3	5.26 ± 0.76g	Sheep -Polypay sheep	[99]

Previous studies have introduced that the therapeutic mechanism of implanted or injected iPSC-laden hydrogels or iPSCs in muscle defect or disease models that show improved muscle regeneration based on successful engraftment,[6,24,83–87] the secretion of factors (i.e., EVs),[88] or immune modulation[89] (Table S1). Recently, Hicks and colleagues reported that host satellite cells are one of the significant factors to inhibit the engraftment of hiPSCs due to competition; in their study, the population of hiPSCs significantly decreased in a mouse defect model.[90] Although we observed human markers at 4 and 8 weeks that might be considered partial engraftment of the implanted hiPSC-MPCs, we expect that the dominant population at the implant area might be host cells, not implanted human cells, over longer time. Therefore, the positive impacts of hiPSC-derived VMLs on inducing muscle recovery could be attributed to the Paracrine effect of hiPSC-MPCs observed in other transplanted tissues, particularly constructs of human donor origin.[91,92] Also, we measured VEGF secretion in a medium supernatant of in vitro cultured hiPSC-derived VMLs (no HUVEC) (Fig. 6j) and observed a 3-fold increase in VEGF secretion over the course of the second to third week after the induction of hiPSC-MPC differentiation, which contributed to axonal growth, angiogenesis and later muscle regeneration over the course of the in vivo study.[93–97] As a result, we could suggest that the hiPSC-derived VMLs have contributed to the functional recovery and regenerative capacity of the volumetric muscle loss tissue by increasing the secretion of factors and cytokines.

Together, this study provides a volumetric muscle loss repair strategy based on pre-vascularized large-scale hiPSC-derived skeletal muscle implants. To further optimize cell engraftment in the host’s tissue at a clinically ready stage and achieve full muscle reconstitution post- volumetric muscle loss, we recommend expanding testing to larger animals that better mimic human skeletal muscle cell proliferation and injury recovery. Also, the proposed uEMB technology, designed to print stem cell-laden bioinks, is able to produce centimeter-scale stem cell-laden structures (~1 cm³) by modulating the size and geometry of the supporting matrix. Furthermore, centimeter-scale building blocks could be assembled into even larger constructs using a low concentration of GelMA pre-polymer solution as light-curable glue. Gluing together nine uniquely patterned building blocks readily resulted in the formation of large, complex, and perfusable 3D constructs with volumes of almost 10 cm³ (Figure S4b). In the case of large animals such as pigs, a volumetric muscle loss model is created by removing approximately 5g of muscle tissue (approximately 3 × 3 × 1.5 cm defect size), as proposed in several studies.[98,99] Therefore, by enlarging the endomysium-like supporting matrix or assembling centimeter-scale printed building blocks, the proposed uEMB technology could help create implants tailored to defect areas in large animal models. However, in fabricating excessively large implants, the overall printing time using a single or dual printing nozzle takes a long time (> a few tens mins), mainly affecting stem cell viability and other behaviors. Despite demonstrating proof of concept in Figure S4, it is necessary to enhance current uEMB technology. Previously developed multimaterial multi-nozzle 3D printheads[100] or Lego-inspired assemble concepts[101] are good candidates for rapidly obtaining volumetric multi-material constructs and expanding the size of implants using printed building blocks. However, these fascinating technologies need to be re-designed or modified to use stem cell-laden ECM-based bioinks. Combining these modified technologies into the uEMB system makes it possible to create large-sized printing constructs in a rapid manner. A sophisticated assembly process in a favorable environment for stem cells also needs to be developed. These advanced stem cell bioprinting technologies can move one step further in biomanufacturing to obtain clinically relevant large-sized tissue implants with stem cells. Later, biomanufactured tissue constructs are expected to make significant contributions to biomedical engineering fields such as regenerative medicine and drug test models.

Concluding remarks

Regenerating muscle and retrieving its function after a volumetric muscle loss incident remains a significant clinical challenge despite advancements in therapies. To solve this problem, this work introduces a combination strategy of innovative biofabrication and stem cell technologies to create large and scalable pre-vascularized muscle fiber bundles. Specifically, we use an unconventional embedded multi-material bioprinting method with hiPSC-laden photo-crosslinkable biomaterials within a non-sacrificial supporting hydrogel that enables the fabrication of complex geometric features. The optimized muscle fiber bioink supports the 3D culture of hiPSC-MPCs, differentiating and maturing these cells into skeletal myofiber bundles. The inclusion of endothelialized, perfusable microchannels like vessels sustains the viability and function of muscle stem cells both in vitro and in vivo, promoting better graft-host integration. The 3D pre-vascularized tissue constructs with hiPSC-MPCs were implanted in a volumetric muscle loss-injured animal model and demonstrated improved human cell engraftment in pre-vascularized bioprinted constructs. The advanced bioprinting technology can print centimeter-scale stem cell-laden building blocks (~1 cm³) by adjusting the size and shape of the supporting matrix. These building blocks can be assembled with a biomaterial glue to form significantly larger implant blocks clinically relevant to human defect sizes. This advanced biofabrication approach could enable the high-throughput production of high-fidelity biomimetic tissue constructs (see Outstanding Questions). This technology promises to enhance volumetric muscle regeneration therapies, offering a practical and scalable solution to a critical clinical challenge.

Outstanding Questions.

• Proposed technology demonstrated improved integration of muscle myofibers and endothelial cells in implanted in vivo models. How much more improvement is needed before it can be applied in translational clinical applications? How can this design be integrated with chimeric antigen receptor-T cell (CAR-T) therapy to enhance the healing process during the first few hours of surgery?

• To address large-scale muscle defects, we proposed a scalable fabrication and modular assembly method. Creating such patient-derived VML modules depends on obtaining or expanding a large cell pool from the patient. Is this clinically feasible within the post-injury healing timeframe?

• Our tests showed VML constructs can endure cyclic stress up to 40% strain, similar to native muscle. Would large defect implants (made using assembled VML modules) maintain this elastic behavior?

Technology readiness

To treat volumetric muscle loss in practice, implants should contain enough cell density to compensate for cell death during the implantation procedure and in early hours of host integration. To this end, we have optimized construct design to achieve highly dense muscle-laden channels with millimeter-long and densely aligned human induced pluripotent stem cell (hiPSC)-derived myofibers. This cell density has proven effective in achieving functional regeneration in the mice tested. However, more studies are needed in larger animals to quantify the number of cells needed for large defects and this study may result in construct pattern redesign. The hiPSC-derived VMLs demonstrate successful de novo muscle formation and muscle function restoration through a combinatorial effect between improved graft-host integration and increased release of paracrine factors. As such, we believe that we have achieved a TRL 3 level technology readiness in proving that the concept works in vitro and in a relevant environment, e.g. in vivo. Next, the modular building blocks should be assembled and tested in larger defects to validate continuous elastic behavior, large-scale muscle regeneration capacity of the construct and implant-wide cellular viability. Moreover, the assembly should be implanted in larger animals, for longer durations of culture to validate long term muscle functionality to achieve TRL 4. Ultimately, upon successful completion of the mentioned tests, the constructs should be implanted in human defects during clinical trials to inform regeneration efficiency and muscle function restoration.

STAR★ Methods

RESOURCE AVAILABILITY

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Su Ryon Shin (sshin4@bwh.harvard.edu)

Materials availability

This study did not generate new unique reagents.

Data and code availability

• Authors can confirm that all relevant data are included in the paper and/ or its supplementary information files.

• This paper does not report original code.

• Any additional information required to reanalyze the data reported in this work paper is available from the lead contact upon request.

EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

Cell lines

Human Umbilical Vein Endothelial Cells (HUVECs) with prescreened angiogenesis were purchased from Lonza. Cells were cultured in EBM^™-2 Basal medium supplemented with EGM^™-2 pre and post printing. Mouse C2C12 cells were purchased from ATCC (CRL-1772) and were cultured and maintained following the vendor protocol. To differentiate C2C12s into skeletal muscle cells, the application of proliferation medium (Dulbecco’s modified Eagle medium (DMEM), 10% FBS, 1% P/S) was stopped and DMEM supplemented with 2% horse serum (Gibco) and 1% P/S was used to induce differentiation and continue culture by refreshing the medium every other day. Two hiPSC lines, NCRM-1 (RRID:CVCL_1E71) derived from umbilical cord blood and GM23338 as fluorescently labeled with titin, TTN-GFP (RRID:CVCL_F182) derived from arm skin, were used and validated for the 3D cultures in this study. The hiPSCs were cultured according to previous procotols.[39] Briefly, Matrigel-coated plates (BD) were used to seed and culture the cells using mTesR1 medium (Stem Cell Technologies). Aggregate and single-cell passaging could both be used. The cells were verified free of mycoplasma. hiPSC-derived myogenic progenitors were generated following a serum-free differentiation method.[5,57] Three- to four-week-old wild-type hiPSC-derived primary myogenic populations were detached to replate (cell density = 3.5–4 × 10⁴ cm²) myogenic progenitors on Matrigel-coated plates (Corning) and the cells were then cultured in skeletal muscle growth medium (SKGM-2, Lonza) supplemented with 10 μM ROCK inhibitor for 24 hours. Afterwards, the ROCK inhibitor was removed and the culture was continued with SKGM-2 for ~2 days until they were dissociated using Accutase (Stemcell Technologies) and were frozen or prepared for bioprinting.

Mice

Animal procedures of subcutaneous test (hydrogel implantation) using nude rat were approved by the Institutional Animal Use and Care Committee of Harvard Medical School (Protocol number: 2017N000114). 12-week-old nude rats (NIH-Foxn1^rnu Rat, Charles River) were acclimatized for a week prior to surgery in the vivarium of Brigham and Women’s Hospital. Animal procedures of volumetric muscle loss model (hydrogel implantation) were approved by the Institutional Animal Use and Care Committee of Harvard Medical School (Protocol number: 2016N000375). 10-week-old male mice (Jackson Laboratories; Bar Harbor, ME, USA) were acclimatized for a week prior to surgery in the vivarium of Brigham and Women’s Hospital. Twelve RAG2−/−γc−/− and thirty-five NOD-scid IL2Rgammanull mice were used in this study for the iPSC-laden and acellular hydrogel implants, respectively.

METHOD DETAILS

Embedded multi-material (EMB) bioprinting of 3D constructs

Following previous GelMA synthesis protocols, we synthesized medium degree GelMA[102] by administering 5% (v/v) MA to the gelatin mix at a 0.5 ml/min mixing rate. To prepare the supporting matrix, molds were fabricated with polydimethylsiloxane (PDMS) and sterilized using 70% ethanol, overnight UV exposure in the biosafety cabinet, and sterile PBS washing. Next, 0.25 wt% PI and 5 wt% freeze-dried GelMA were mixed in 40% HBSS and 60% hiPSC-MPC proliferation medium and the solution was incubated for 1 hour at 37 °C (GelMA pre-polymer solution). Next, the solution was poured into the mold, and was allowed to gelate (sol-gel state) at 4 °C for ~5–8 minutes.

The muscle bioink was prepared with 7.5% GelMA, 2 wt% gelatin and 0.25% PI dissolved in HBSS and hiPSC-MPC proliferation medium (2:3) and was followed by a 1-hour incubation step at 37 °C. At the time of bioprinting, a pellet of hiPSC-MPCs was uniformly dissolved in 600 μl bioink prepolymer solution to make the bioink, which was then transferred to a 1ml syringe with a printhead needle (30G, blunt, BD). 3D bioprinting was performed using a Cellink Inkredible bioprinter that was calibrated prior to each run with respect to the positional accuracy of the needle and a reference mold in XYZ directions. In the case of dual ink printing, the second nozzle (perfusable channel ink) was also calibrated in reference to the muscle ink nozzle. The muscle ink was extruded at a 4 μl/min rate using an ad hoc syringe pump (New Era Pump Systems). To create perfusable channels, a soluble 5% gelatin ink was prepared and printed inside the bath in the form of highly aligned channels. To ensure the printability of the ink, gelatin was thermally gelated and printed at 19–22 °C. Next, the GelMA construct with bioprinted hiPSC and gelatin channels were exposed to a UV light source (~100 mW cm⁻², OmniCure S2000) to be chemically crosslinked. Constructs were then taken out from the molds, and were washed (PBS) and collected onto a 24-well plate pre-filled with warm hiPSC-MPC proliferation medium, followed by incubation at 37 °C and 5% CO₂. The media was changed after 1 hour to remove the dissolved gelatin from the constructs. The medium was refreshed on day 1 after printing and regularly every 2 days afterwards, until the differentiation medium was changed, or until fixation or implantation of the constructs. By submerging the constructs in medium and incubating the construct at 37°C, the gelatin inside the printed channels gradually dissolved into the solution, leaving a fully hollow channel inside the crosslinked GelMA bath.

For bioprinting the muscle flap geometries, a CAD drawing of each geometry was used to lasercut and create customized PDMS molds. Next, a 3D gcode was created using the Cellink Heartware software. An appropriate angle of printing was chosen such that the ratio of Z travel to X travel (and Z travel to Y travel) remained larger than 1. The mold was cut open and the nozzle traveled in the Z direction to bioprint the structure. The construct was then bioprinted and crosslinked under UV light using the parameters mentioned above.

To extrude multi-material coaxial filaments, a coaxial nozzle was fabricated by adhering concentric 25G inner and 18G outer needles using hot glue. To bioprint the sheath, 5% GelMA and 2% alginate was fed into the outer needle at 16 μl/min. To bioprint the core, 5% GelMA and 2% gelatin supplemented with 0.25% PI and 1% CaCl₂ was fed to the inner needle at 8 μl/min. The core-sheath crosslinking was achieved through the Ca²⁺ crosslinking of alginate as well as GelMA covalent gelation upon UV irradiation.

The approximate time for bioprinting constructs with the density that was used for in vivo volumetric muscle loss experiments is ~20 minutes/construct. This time depends on the printing density (e.g., the number of muscle channels and perfusable channels per mm area), geometry of the construct (e.g., cylindrical and symmetrical geometries vs asymmetrical structures such as those in Figure 2l–o). To decrease this time, it is possible to create smaller modular units which can then be assembled into a centimeter sized construct using 3% (w/v) GelMA UV-curable glue to allow for the fabrication of larger 3D assemblies for larger muscle defect sizes. Given the modularity of this fabrication method, constructs can be printed in parallel or asynchronously, therefore mitigating the risks of cell viability reduction and material compromise during printing.

Characterization of mechanical and rheological properties

To perform compression mechanical tests on the constructs (cylindrical samples, diameter = 0.9 cm, height = 0.5 cm), a parallel plate platform was used to compress the samples at room temperature until rupture (ADMET, MTESTQuattro). The linear portion of the resulted stress-strain curve (i.e., <20% strain) was used to calculate the Young’s modulus.

Cyclic compression tests were performed and recorded on an Instron 5966 Universal Testing System with 500 N load cell. To ensure symmetry, the constructs were 3D bioprinted in the same cylindrical molds as mentioned above and were cut in half (perpendicular to the bioprinted channels). 2 mm sample cuts showed nearly identical results. Five cycles (0–10% strain) were performed sequentially to ensure good contact and the absence of hysteresis. Then, incremental strain ranges were used to perform sequential cyclic tests (starting at 0% strain, going up to 10%, 20%, 30%, and 40%), returning to 0% strain in between the increments. To ensure reproducibility, the strain incremental measurement was repeated for one sample and an excellent agreement was obtained.

To measure the rheological properties of the bioinks (Anton Paar MCR 302 rheometer), a 10-millimeter diameter concentric cylinder Couette DIN bob with a 10.84 mm diameter cup (~1 mL sample volume) was used for all measurements, giving a geometry gap between the inner and outer cylinders of 0.42 mm. This geometry was selected to minimize the impact of drying and to increase the surface area to obtain a better signal than possible via a parallel plate or cone and plate geometry. A C-PTD200 module (bottom), chiller (Julabo circulating chiller) and solvent trap attachments (Anton Paar) were used to maintain temperature control. Experimental data recording and analysis were performed with Anton Paar RheoCompass. Temperature sweeps were done between 4 and 37 °C; all samples were allowed to equilibrate for at least 10 minutes at 4 °C prior to measurement. For oscillatory temperature ramps, the ramp rate was 1 °C/min; the strain amplitude and frequency were γ0 = 0.3% and ω=10 rad/s, respectively, chosen to be in the linear viscoelastic regime (LVE) at 4 °C determined by amplitude sweeps. The sweep was then reversed to examine rheological hysteresis. Before each sample measurement, calibrations were done to the measuring system inertia of the upper plate, as well as the motor that compensated for residual friction. Next, after gap recalibration, the constructs were introduced to the machine and were allowed to reach an equilibrium before running the test (10 minutes).

Characterization of microporous structures of hydrogels

To perform SEM imaging, the constructs were first incubated in PBS overnight at 37 °C. The constructs were then collected and freeze-dried inside 1 ml Eppendorf tubes that were punctured on the cap for air release. Next, the constructs were cut at different horizontal and vertical cross-sections parallel or perpendicular to the bioprinted channels. SEM imaging was performed at the Harvard center for nanoscale systems.

Characterization of perfusable microchannels

To characterize the perfusability of the hollow channels, a perfusion test was performed using FITC-Dextran dye. Briefly, 5% FITC-Dextran powder was dissolved in PBS and covered by aluminum foil. The constructs were first placed horizontally (channels were oriented parallel to the ground) on a petri dish on a confocal microscope and a stacked image (image 0) was captured from the channels using the brightfield channel and the green fluorescent laser. Next, the constructs were gently flipped vertically (channels were now oriented vertically) and 10 μl of the FITC solution was added to the channels from the top. The constructs were immediately flipped over to the initial orientation, and the diffusion was recorded using the time series function of the confocal microscope in a 20-minute span. Stacked images of the green fluorescent laser and brightfield were recorded every 10 minutes. The images were then analyzed using ImageJ to calculate fluorescentablece intensity near the channels. To calculate the diffusional permeability, the following formula was used following a previous protocol[103]:

$$P_d=\frac{1}{l_1-l_b}\left(\frac{l_2-l_1}{t}\right)\frac{d}{4}$$

Where P_d represented the diffusional permeability, l₁ was the average intensity at image 0, l₂ was the average intensity in the later timepoints (e.g. t ~10 min), l_b was the background intensity before FITC addition, and d was the diameter of the printer channel (~345 μm). The portion of the intensity curve on the left side of the peak was used to calculate the fluorescent intensity.

Fabrication of cell-laden bulk hydrogels

HUVEC-laden GelMA constructs:

To fabricate bulk hydrogels containing embedded HUVECs, 5% GelMA bath pre-polymer solution was prepared as explained in the previous sections. Next, HUVECs were trypsinized and the pellet was resuspended in the pre-polymer solution to reach a final cell density of 7.5 × 10 ⁶ cells/ml. Next, the pre-polymer solution was poured into PDMS molds as explained above and incubated in 4 °C for 5–8 minutes. The constructs were then crosslinked under UV light (100 mW/cm²) for 55 seconds, and were extracted from the molds, washed with PBS and placed in 24-well plates supplemented with HUVEC culture medium. The medium was refreshed 1 hour after crosslinking, the day after, and after that regularly every 2 days.

iPSC-MPC-laden GelMA:

To fabricate bulk hydrogels containing embedded hiPSC-MPCs, 7.5% GelMA bath pre-polymer solution containing 0.25% PI was prepared in a solution containing 40% HBSS and 60% SKGM-2+Ri medium. hiPSC-MPCs were trypsinized and embedded in this pre-polymer solution at a density of 7.5 million cells/ml. Next, the pre-polymer solution containing cells was poured into PDMS molds and incubated at 4 °C for 5–8 minutes. The constructs were next crosslinked under UV light (100 mW/cm²) for 55 seconds, and were extracted from the molds, washed with PBS and placed in 24-well plates supplemented with SKGM-2+Ri culture medium. The medium was refreshed 1 hour after crosslinking, the day after, and then regularly every 2 days.

Fabrication of 3D vascularized (HUVEC-seeded) constructs

5% soluble gelatin ink was first prepared according to the soluble ink preparation instructions. Next, cells were trypsinized and centrifuged for 5 minutes at 1000 rpm. Afterwards, HUVECs were added to the ink at a density of 4 × 10 ⁷ cells/ml. Meanwhile, the media of the wells containing constructs was aspirated by pipette. Constructs were placed vertically using a small spatula in 48-well plates. 10 μL of HUVEC-laden bioink (4 × 10⁵ cells) were injected two times on top of each construct through the cross section of the hollow channels using a pipette. Constructs were immediately flipped on the side. After adding cells to all constructs, the well plate was returned to the incubator for 45 minutes to allow cells to loosely attach to the wall. At this time, no media was added to the constructs. After this incubation period, the constructs were placed vertically again and another two rounds of 10 μL HUVEC-laden bioink were injected inside all channels in each construct. Constructs were again flipped horizontally and incubated for 45 minutes at 37°C. After this time, constructs were checked under the microscope to confirm the presence of cells inside the channels. HUVEC media was gently added to each construct and the constructs were incubated overnight. The media was changed the next day to avoid possible contaminations. The constructs were replated in new 48-well plates after 2–3 days to avoid consumption of media by the cells that might have migrated to the bottom of the well plate.

Preparation of hiPSC-MPCs for 3D bioprinting

2 days prior to 3D bioprinting, frozen vials of hiPSC-MPCs were thawed and seeded onto Matrigel-coated 6-well plates (Matrigel hESC-qualified, Corning). Specifically, the cells were thawed, resuspended in SKGM-2+Ri medium (SKGM-2+Ri (10 μM) + 0.2% Pen/Strep) and were seeded at a density of 8*10⁴ cells /cm². The medium was replaced with SKGM-2+0.2% P/S after 24 hours and until bioprinting.

Differentiation of hiPSC-MPCs

To differentiate hiPSC-MPCs into skeletal muscle cells, a medium cocktail of the following components was used (iPSC-Diff media) following a previously established hiPSC-MPC differentiation protocol[5]: Briefly, after 24–48 hours, cells were induced for myogenic differentiation in DMEM/F-12, GlutaMAX^™ (Gibco) supplemented with 1 μM Chiron (Tocris), 2% knock-out serum replacement (KSR, Invitrogen), 0.2% Pen/Strep (Life Technologies), Insulin-Transferrin-Selenium (ITS, Life Technologies), 10 μM prednisolone (Sigma-Aldrich), and 10 μM TGF-β inhibitor SB431542 (Tocris,). After differentiation induction, medium was refreshed every day until day 2 and afterwards, every other day.

Characterization of in vitro cultured samples

Fixation of cell-laden constructs was done at different timepoints (e.g., day 10, day 20, day 30). Briefly, the constructs were treated with 4% paraformalehyde solution (Thermo Fisher scientific) for 20 minutes. Next, the constructs were treated with 0.1% Triton-X 100 for 10 min, and then washed with DPBS three times. The constructs were next blocked with 10% goat serum in DPBS for 1 hour at 25°C. To stain the cytoskeleton, F-actin (1:40) was added to the constructs. Primary antibodies that were used in this study included mouse anti-myoG (1:200), mouse anti-Pax7 (1:100), mouse anti-beta-III tubulin monoclonal antibody (1:500) and mouse anti fast myosin heavy chain (1:100). To stain the HUVECs, CD31 antibody (1:100) and VE-cadherin antibody (1:100) were used. Conjugated secondary antibodies (goat anti-mouse (rabbit, rat) IgG, Invitrogen Alexa Fluor 594, 488, and 647 (1:200) were added and the constructs were incubated at 4 °C for 6 hours. DAPI (1:1000) was added 30 minutes prior to imaging and was used for visualization. For imaging the constructs, a fluorescence (Zeiss Axio Observer D1) and a confocal microscope (SP7, Leica) were employed. To acquire Z-stack, tiled and time-lapse 3D images (e.g. the endothelialized constructs and the dye diffusion tests), a Zeiss LSM 880 airyscan confocal microscope was used. Image processing and data analysis of the stacked data was performed using Zen Blue and Fiji.

Fast Fourier transform (FFT) analysis

To quantify the extent of muscle fiber alignment, FFT analysis was performed using a previously described method.[104] Briefly, the images were stored and analyzed as uncompressed. TIFF files and were converted to grayscale 8-bit images. ImageJ was used to conduct 2D FFT analysis on four images per print condition. The obtained image was then analyzed using a MATLAB program we developed to plot the FFT alignment intensity versus angle or orientation in degrees. From each orientation histogram, the angle at which most fibers were aligned was determined. Alignment index was calculated as the fraction of cells aligned within 20° of the peak angle.[105] The larger the alignment index was, the higher was the fraction of fibers aligned near a peak angle. A completely randomly aligned matrix was indicated by an alignment index of 0.

In vivo subcutaneous study

HUVEC-laden constructs cultured for 3–4 weeks were selected for implantation. Four groups of constructs containing HUVECs were fabricated as below:

G1 - Endothelialized perfusable channels in the supporting bath: perfusable channels were fabricated in a 5% GelMA supporting bath as described in the previous sections. The channels were later seeded with HUVECs and cultured for 2 weeks before surgery.

G2 - Endothelialized perfusable channels in the HUVEC-laden supporting bath: HUVEC-laden bulk GelMA pre-polymer solution was prepared as previously described. This bath was used to 3D bioprint perfusable channels according to the protocol described previously.

G3 - Perfusable channels in the supporting bath. This group did not contain any cells and was bioprinted as a control group for G1. Briefly, perfusable channels were bioprinted in a 5% GelMA bath as described previously. The constructs were cultured for 2 weeks before surgery.

G4 - HUVEC-laden supporting bath. This group contained a bulk cell-laden hydrogel with no 3D bioprinting involved. The constructs were prepared according to the protocol described above for fabricating bulk hydrogels. The constructs were cultured for 2 weeks before surgery. This group served as a control group for G2, investigating the effect of incorporating bioprinted perfusable channels in the constructs.

Subcutaneous embedding test was performed on nude rats anesthetized with Isoflurane anesthetic. Briefly, subcutaneous space was accessed through dorsal incision with the length of 2 cm. Subsequent subcutaneous pockets were created by blunt separation. Next, three hydrogel constructs were introduced into the subcutaneous pockets on bilateral sides of the cut. Total of 6 constructs were implanted for each group. To alleviate post-surgery pain of rats, carprofen (5 mg/kg BW/day s.c. as a single injection) was postoperatively administered for 72 hours and the rats were monitored hourly for 6 hours post-surgery and twice per day for 5 days. Samples of each group were harvested at two timepoints (weeks 1 and 4). The implanted gels and surrounding tissue were removed for further staining and studying angiogenic properties. Next, the harvested cell-laden printed constructs were washed with PBS and fixed in 4% paraformaldehyde for 24 hours before staining with Haemotoxylin and Eosin (H&E) and human-specific and rat-specific CD31 antibodies (iHisto.io).

Fabrication and characterization of suturable PGS/PCL scaffolds

PGS Synthesis:

Poly(glycerol sebacate) (PGS) was synthesized using an established protocol.[106] Equimolar quantities of sebacic acid (Sigma-Aldrich) and glycerol (Fisher Scientific) were mixed using a magnetic stirrer. The mixture was reacted under microwave radiation in 1 minute intervals for a total of 7 minutes at 500 W (Hamilton Beach, VA). After every minute interval, the microwave door was opened for 10 seconds to allow vapors to be removed. The synthesized PGS was allowed to cool to room temperature.

PGS/PCL Scaffold fabrication:

Polycaprolactone (PCL) Mn 80,000 (Sigma-Aldrich) was dissolved at 20% (w/v) in hexafluoroisopropanol (HFIP) (CovaChem, IL) using a magnetic stirrer in a covered container to prevent evaporation. In a separate covered container, the synthesized PGS was dissolved in HFIP at 20% (w/v). Equal volumes of the PGS and PCL solutions were added together and mixed on a magnetic stirring for 3 hours at room temperature. The solution was transferred to a syringe and electrospun at 18 kV using a 22 G blunt-tipped needle. The fibers were collected on a rotating drum coated with aluminum foil. The drum was rotated at a speed of 50 rpm and set at a distance of 22 cm from the tip of the needle. The electrospun mats and aluminum foil were removed from the drum and desiccated for 24 hours.

Preparation and sterilization of scaffolds for surgery:

On the day before surgery, PGC/PCL scaffolds were sterilized with 70% ethanol and were left inside the biosafety cabinet overnight. On the day of the surgery, the scaffolds were detached from the foil gently using two forceps inside the biosafety cabinet. The bioprinted constructs were attached to the scaffolds by adding a drop (~15μL) of GelMA prepolymer solution (7.5% GelMA+ 0.5% PI) to the mesh and placing the constructs on top of the pre-polymer solution. The constructs were crosslinked under UV light (100 mW/cm²) for 5 seconds. The adhesion of construct-mesh makeup was tested by applying tensile forces to the construct with forceps. Next, the constructs attached to the mesh were placed in 48-well plates, supplied with SKGM-2 medium and incubated at 37 °C until surgery.

Mechanical characterization of PGS/PCL scaffolds:

Using a surgical suture, 20mm x 6mm x 0.1mm rectangular shaped specimens of the PGS/PCL scaffold were sutured. The tensile strength of both the sutured and non-sutured scaffolds were studied using a uniaxial material testing system (Instron 3342). Rectangular-shaped specimens (40mm x 6mm x 0.1mm) were stretched at a strain rate of 7mm per minute, in compliance with the ASTM D822 standard. Five samples were tested for each group. The Young’s modulus ‘E’ of the fibers was calculated from the linear region of the stress-strain curve corresponding to 0%−30% of the tensile strain.

Morphological characterization of PGS/PCL scaffolds:

Electrospun PGS/PCL sheets were imaged using scanning electron microscopy. First, desiccated sheets were coated in platinum and palladium for 10 seconds using a Sputter Coater 108 Auto (Cressington, UK). The coated sheets were imaged on a high vacuum at 2 kV using a FEI Quanta 200 ESEM (FEI Company, Hillsboro, OR). To image scaffold-hydrogel interfaces, a 1cm² piece of the PGS/PCL scaffold was photo-crosslinked on a polymerized GelMA hydrogel cube. The scaffold was pre-wet with liquid GelMA before putting on the hydrogel cube and crosslinked with a UV light for 20 seconds operating at 850 mW/cm² intensity. The PGS/PCL-GelMA construct was freeze-dried for 3 days. The samples were sputter-coated with a 5nm thin layer of Pt/Pd alloy (80:20) for increased conductivity and imaged using JEOL JSM-IT100 InTouchScope^™ scanning electron microscope operated at an accelerated voltage of 5–10 kV. Fiber diameters of the scaffold and pore sizes of the GelMA hydrogel were measured using Image J software. Three samples were analyzed, and 10 images taken from different areas of the cross-section.

In vivo volumetric muscle loss study

Mice were anaesthetized using a 2–4% isoflurane vaporizer induction chamber and the surgical site was disinfected with chlorhexidine and 70% isopropanol (Contec, Spartanburg, SC). Then the mice were randomly assigned to one of the following four groups: volumetric muscle loss treated with 3D vascularized muscle tissue (V3, n=11 for the NOD-scid IL2Rgamma^null strain and n=4 bilateral implantation for the RAG2 strain), volumetric muscle loss treated with acellular bioprinted constructs (V2, n=9 for the NOD-scid IL2Rgamma^null strain and n=4 bilateral implantation for the RAG2 strain) and untreated volumetric muscle loss (V1, n=8 for the NODscid IL2Rgamma^null strain and n=4 for the RAG2 strain) and sham (V0, n=7 for the NOD-scid IL2Rgamma^null strain and n=4 for the RAG2 strain, with similar procedures on the skin and fascia but no injury to the muscle). For the rectangular defect model, bilaterally, a longitudinal incision at quadriceps muscle was created followed by a sharp resection of the quadriceps muscle in parallel to the femur length and in all study groups, approximately 34.7 ± 0.7 mg of the muscle was cut, leaving only a base of the skeletal muscle.[107] In the final and optimized volumetric muscle loss model, blunt dissection of the incised skin, fascia, and hamstring muscles was performed to expose the gastrocnemius (GA) muscle. A 4-mm diameter biopsy punch was used to create a cylindrical defect size of 4πx4 mm³(DxL, ~20% of muscle mass was removed).[108] The injury was resulted in a full thickness muscle defect to the level of the tibia, and removed 26.9 ± 3.1mg of skeletal muscle (approximately 20% of muscle mass).

In the groups receiving treatment (V3 and V2), the construct (acellular bioprinted or vascularized muscle tissue) was attached to a PGS/PCL scaffold and placed with the scaffold side up into the volumetric muscle loss defects. The implanted 3D constructs (which underwent some degradation during in vitro culture to ~4×4 mm (DxL) for cylindrical defects and 2×3×6 mm³ for rectangular defects) filled in the defect cavity. The muscle defects in the untreated volumetric muscle loss group (V1) were left without any fillings. At the end of the procedures, the fascia and skin incisions were closed with simple sutures (4–0 Silk, Ethicon, Johnson Johnson, Somerville, NJ, USA). At the 4-week timepoint, n=3 per group were tested and sacrificed and the rest of the mice were tested at the 8-week timepoint.

Ankle Plantar Flexion Torque Testing

4- and 8-weeks post-surgery, an in-vivo strength test was performed on all the animals to assess ankle plantar flexion torque using a servomotor-based apparatus (Model 305C, Aurora Scientific, Aurora, Ontario, Canada). Briefly, under general anesthesia, the right knee of the animal was stabilized to the knee clamp, and the right foot was fixed on a mouse footplate attached to the force transducer machine. The lower leg was aligned ensuring that the ankle was at a 90-degree angle and the tibia was in line with the pelvis. One needle electrode was then placed subcutaneously near the belly of GA muscle, while the second electrode was placed next to the first electrode. After achieving the resting tension close to 0 N, the peak isometric plantar flexion torque defined as the greatest torque recorded during a 500-ms stimulation using 1-ms square-wave pulses at 100 Hz and increasing amperage from 10mA to 1A was measured. Torques (Nmm) were normalized according to the body mass of animals (Kg).

In situ Muscle Strength Testing

Eight weeks post-surgery, right hind limb was depilated, and the skin and fascia were incised to expose the right GA muscle under general anesthesia. The distal part of the tendon was severed and fixed onto the lever arm of the force transducer instrument. The soleus muscle was dissected from the calcaneal tendon. A needle was inserted through the knee and locked in place with a set screw to secure the leg in place. During the procedures, the exposed tissue was kept moist using Ringer solution. Electrodes were placed directly into the GA muscle. The stimulation current and resting tension were adjusted to maximize twitch force produced by a single stimulus pulse with pulse width of 0.2 ms (610A Dynamic Muscle Control LabBook v6, Aurora Scientific, Aurora, Ontario, Canada). The optimal length (Lo) at which muscle produced its greatest force was measured using a ruler as the distance between the knee and distal insertion of muscle to the calcaneal tendon. This optimum stimulation current and resting tension was then applied to measure tetanic force for 500 ms duration of stimulation using pulses given at 100 Hz with increasing amperage from 10mA to 1A. The peak twitch and tetanic forces of right GA muscle of each animal were normalized to the animal’s body weight. At the end of in situ force measurements at 8-week timepoint, the GA muscles were harvested from both legs, weighed, and snap-frozen in liquid nitrogen for subsequent histological evaluations. The animals were then euthanized in carbon dioxide chamber.

Characterization of in vivo cultured samples

After implantation, the constructs were incubated in 10% formalin solution for 48 hours followed by storage in 70% alcohol in 4 °C. Histological analysis and immunostaining of paraffin sectioned samples were performed afterwards by iHisto.io company. For cryosectioned samples, frozen sections of samples were stained with H&E and Masson trichrome. Next, cryostat muscle cross-sections were thawed at room temperature for 10–20 minutes followed by two PBS washing steps and 5-min incubation in 0.05% TX-100 in PBS for permeabilization. The washed sections were incubated for 1h in blocking buffer (i.e., 1% BSA and 5% Goat normal serum in tris buffered saline) at room temperature, and then in primary antibodies diluted in blocking buffer overnight at 4 °C. Slides were incubated with secondary antibodies for 1h at room temperature following three PBS wash steps and were mounted on ProLong^™ Diamond Antifade Mountant (Invitrogen^™) and glass coverslips.

For volumetric muscle loss studies, MyHC (1:100), Pax7 (1:80), DAPI, MYH3 (1 μg/ml), TUNEL, βIII-tubulin (1:500), mCD31 antibody (1:100) and hCD31 antibody(1:100), HLA-A (1:100), hSpectrin(1:100), hLamin A/C (1:500, ab108595 / 1:50, ab40567) and hDystrophin (1:25) were used. For subcutaneous studies, anti-mouse and anti-human CD31 antibodies were used, as provided by iHisto.io.

QUANTIFICATION AND STATISTICAL ANALYSIS

Statistical analysis of the data was performed using GraphPad Prism. One-way ANOVA Tukey’s multiple comparisons, Two-way ANOVA Sidak’s multiple comparisons, Mann-Whitney test and unpaired t-test were used to compare the data groups of in vitro and in vivo experiments. The p-values, number of samples (n), and the type of each analysis are provided in the figure captions of the corresponding graphs.

Supplementary Material

1

Figure S1. Rheological characterization of inks and the supporting matrix.

Figure S2. Charactarization of muscle bioink and supporting matrix.

Figure S3. Characterization of the supporting matrix without cells.

Figure S4. Various types of printed constructs.

Figure S5. Permeability Analysis of printed constructs with perfusable channels.

Figure S6. Differentiation of hiPSC-MPCs in optimized bioink.

Figure S7. Encapsulation of iPSC-MPCs in 7.5% GelMA hydrogel.

Figure S8. Live (green) and dead (red) staining of bioprinted human hiPSC-MPCs 6 days after differentiation induction

Figure S9. Mechanical property of printed constructs

Figure S10. Optimizing distance between cell fibers and perfusable channels.

Figure S11. FFT analysis of the 3D bioprinted iPSC-MPC laden channels.

Figure S12. Immnunostaining of myogenic markers in bio-printed iPSC-laden construct.

Figure S13 Characterization of the control groups for the subcutaneous study.

Figure S14. PGS/PCL scaffold

Figure S15. Characterization of sham muscle harvests 8 weeks post-surgery (no VML injury, V0)

Figure S16. Human marker staining on NOD-scid IL2Rgammanull mice

Figure S17. Human marker staining on RAG2−/−γc−/− mice.

Figure S18. Vessel marker staining on RAG2−/−γc−/− mice.

Table S1. Recent iPSC-therapy for muscle regeneration and mechanism for regeneration.

2

Movie S1. Injection of dye into the perfusable channel.

3

Movie S2. Calcium transition analysis in contraction behavior of encapsulated iPSC-MPC in 7.5% GelMA hydrogel within 10–15 days post-induction.

KEY RESOURCES TABLE

Reagents and resources	Source	Identifier
Antibodies
MyoG	DHSB	F5D-C
Pax7	DHSB	Pax3
CD31	DHSB	P2B1
MYHC	Sigma-aldrich	MY-32
hDystrophin	Sigma-aldrich	MABT827
MYH3	Abcam	Ab123205
βIII-tubulin	Abcam	Ab18207
HLA-A	Abcam	Ab52922
hLaminA/C	Abcam	Ab40567, ab108595
mCD31	Invitrogen	14–0311-82
hCD31	Fortis	A700–127
hSpectrin	Leica	NCL-SPEC1
VE-cadherin	LSBio	LSC75974550
Goat anti-mouse IgG Alexa Fluor 594	Invitrogen	A-11005
Goat anti-rabbit IgG Alexa Fluor 594	Invitrogen	A-11012
Goat anti-mouse IgG Alexa Fluor 488	Invitrogen	A-10680
Goat anti-rabbit IgG Alexa Fluor 488	Invitrogen	A-11008
Goat anti-rat IgG Alexa Fluor 488	Invitrogen	A-11006
Goat anti-mouse IgG Alexa Fluor 647	Invitrogen	A-21235
Chemicals and recombinant proteins
Dulbecco’s modified Eagle medium	Gibco	11965092
Horse serum	Gibco	26050088
Goat serum	Gibco	16210072
DMEM/F-12, GlutaMAX™	Gibco	10565018
Fatal bovine serum	Gibco	26140079
Pen/Strep	Gibco	10378016
Insulin-Transferrin-Selenium	Gibco	41400045
Phosphate buffer saline	Gibco	10010023
Hanks’ balanced salt solution	Gibco	14175095
Skeletal muscle growth medium	Lonza	CC-3245
Endothelial cell growth medium	Lonza	CC-3162
Materigel	Corning	CLS354234
Knock-out serum replacement	Invitrogen	10828028
ProLong™ Diamond Antifade Mountant	Invitrogen	P36965
Glycerol	Invitrogen	15514011
TGF-b inhibitor (SB431542)	Tocris	1614
CHIR99021	Sigma-aldrich	SML1046
Gelatin from porcine skin	Sigma-aldrich	G6144
2-hydroxy-4′-(2-hydroxyethoxy)-2-methylpropiophenone	Sigma-aldrich	410896
methacrylic anhydride	Sigma-aldrich	276685
Triton-X 100	Sigma-aldrich	T8787
Prednisolone	Sigma-aldrich	1555005
Sebacic acid	Sigma-aldrich	8.00753
Polycaprolactone (PCL) Mn 80,000	Sigma-aldrich	440744
Hexafluoroisopropanol (HFIP)	CovaChem	105228
paraformalehyde solution	Thermo Fisher Scientific	0433689M
Cell lines
C2C12	ATCC	CRL-1772
Human Umbilical Vein Endothelial Cells	Lonza	CC-2519
NCRM-1	Harvard Medical School	RRID:CVCL_1E71
GM23338	Harvard Medical School	RRID:CVCL_F182
Mice strains
NOD-scid IL2Rgammanull	Jackson Laboratories	005557
RAG2−/−γc−/−	Jackson Laboratories	014593
NIH-Foxn1rnu Rat	Charles River	316

Highlights.

• Large-scale vessel-integrated muscle-like lattices (VMLs) containing dense and aligned hiPSC-derived myofibers alongside vessel-like microchannels using an advanced bioprinting technology and stem cell-laden extracellular matrix-based bioinks.

• Incorporating vessel-like lattice was of vital importance for enhancing myofiber maturation in-vitro and host vessel invasion in-vivo, improving implant integration.

• Successful de novo muscle formation and muscle function restoration through a combinatorial effect between improved hiPSC-derived VMLs graft-host integration and increased release of paracrine factors at volumetric muscle loss injury.

• Observing the human markers in the implantation area confirmed that the hiPSC-MPCs at the transplantation site play a significant role for improving regenerative capacity of volumetric muscle loss.

GLOSSARY

HiPSC-derived muscle precursor cells (hiPSC-MPCs)

Cells that originate from induced pluripotent stem cells (iPSCs) and are committed to differentiating into myofibers

Unconventional embedded multi-material bioprinting (uEMB) method

An advanced 3D bioprinting process which includes programmatic injection of multiple bioinks inside a non-sacrificial hydrogel supporting bath to form biomaterial-embedded 3D structures

Vessel-integrated muscle-like lattices (VMLs)

An engineered construct containing the co-culture of muscle and endothetial cells. Muscle-like structures were created using the uEMB method by printing muscle cell channels and hollow channels, followed by coating the hollow channels with endothelial cells