Skeletal Muscle Tissue Engineering: Methods to Form Skeletal Myotubes and Their Applications

Cell alignment by topography

It has long been known that cell behavior is influenced by surface features. ³⁴ Thus, numerous studies have focused on the effects of different topographical features, such as size and geometry, on the cellular response.^35–38 Among these topographical features, parallel grooves are among the most-studied patterns to elongate muscle cells in one direction. The first studies aimed to determine how the cells sense their environment and what causes the cells to undergo alignment. Thus, grooved patterns with different widths and depths were tested. For example, Evans et al. generated micropatterned grooves with depths ranging from 40 nm to 6 μm and widths ranging from 5 to 100 μm on silicon substrates by etching with conventional photolithographic methods and studied myoblast direction and alignment along the grooves. ³⁹ They showed that shallow grooves with a depth of 40–140 nm did not significantly affect myoblast alignment, whereas significant cell alignment was achieved with deep grooves that had a width of 5–12 μm and a depth of 2–6 μm. Additionally, Clark et al. showed that nanosized grooves with a width of 130 nm and a depth of 210 nm also induced myoblast alignment. ⁴⁰ In addition, because they observed that myotubes with identical diameters formed in grooves with different widths, Clark et al. hypothesized that lateral fusion of myoblasts was not a possible mechanism in myotube formation. Therefore, they cultured myoblasts on ultrafine grating (grooves with a width of 130 nm and a depth of 210 nm and ridges with a width of 130 nm) that strongly aligned the myoblasts, and showed that myoblasts fused in end-to-end configurations. ⁴¹

To easily fabricate groove/ridge micro- and nanopatterns without requiring a clean room, alternative methods to photolithography have also been used. Thus, since they contain nano/microgrooves, commercially CD-R and DVD-R in polycarbonate have been used for directing cell alignment or for patterning polymers.^42,43 Abrasive paper has also been proposed to easily produce parallel grooves on a surface at low cost to direct the alignment of myoblasts. ⁴⁴ Similarly, Jiang et al. fabricated sinusoidal-wavy-grooved (size ranging between 0.1 and 10 μm) micropatterns on a PDMS surface by stretching a PDMS slab and then subjecting it to extended oxidation under low pressure before relaxing it. For this continuous topography without sharp edges, they showed that sharp-edge features were not necessary to induce contact guidance. ⁴⁵ Another study by Lam et al. focused on the effects of wave periodicity on C2C12 cells and showed that a wavelength of 6 μm was optimal to induce myoblast and myotube alignment. ⁴⁶ These topography–cell interaction studies opposed the theory proposed by Curtis and Clark, who suggested that cell guidance on groove-ridge patterns is mostly governed by groove depth.^37,47 Although numerous studies have suggested that cells sense and grow on predefined topography, the mechanism by which the cells sense the topography is not well understood. However, filopodia are involved in this detection because they extend in front of the cells and probe the topographic features. ⁴⁸ This topographical surface guidance is the foundation of several approaches used for designing scaffolds in 2D and 3D. For instance, Neumann et al. used arrays of parallel polymer fibers with thicknesses of 10 to 50 μm and spacings of 30 to 95 μm to generate a scaffold for engineering a C2C12 myoblast sheet. They showed that by using this method, it was possible to generate a continuous contractile aligned muscle sheet with fiber spacing of up to 55 μm ⁴⁹ (Fig. 3).

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FIG. 3.

C2C12 cells cultured on an array of large fibers. (A) Thirty minutes after seeding. (B) Gaps between fibers were closed after 5 weeks of culture and a cell sheet was formed. (C) After 10 weeks in culture, myotubes were striated (DIC image). (D) Cross-section of the muscular tissue fixed and stained by hematoxylin and eosin. The fiber plane is in the upper section (scale bars for [A–D] 50 μm). Reprinted with permission from Neumann et al. ⁴⁹ Copyright © 2003, Mary Ann Liebert, Inc.

In another example, to build a muscle-tendon-bone tissue in one step, Ker et al. used a spinneret-based tunable engineered parameter technique to fabricate polystyrene fiber (655-nm diameter) arrays that they coated with extracellular matrix (ECM) proteins, such as fibronectin, and bioprinted with patterns of two different growth factors. By combining topographical and chemical cues to mimic the in vivo environment, they showed that although the fibers induced C2C12 cell alignment by topography, the localized presence of growth factors induced myoblast differentiation in tenocytes and osteoblasts, and the absence of growth factors enabled differentiation in myotubes ⁵⁰ (Fig. 4).

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FIG. 4.

Fiber-fabrication procedure by Polystyrene Spinneret-based Tunable Engineered Parameter (STEP). (A) Schematic showing the building of fibers by STEP. (B) STEP fiber types that can be fabricated by one set of fibers running in a parallel manner (left), two sets of fibers running perpendicular to each other (middle), and one set of fibers running in a parallel manner with a hollowed-out support base (right). (C) SEM and TEM pictures used to quantify the STEP fiber diameter and length. (D) SEM image showing the cell attachment to the polystyrene STEP fibers. (E) Images in fluorescence of polystyrene STEP fibers coated with Alexa649-conjugated fibrin (right) and uncoated (left). (F) Polystyrene STEP fibers can be printed on by inkjet printing. (Scale bars: B, 2 mm; C, 100 μm and right photo 2 μm; D, 100 μm; E, 200 μm). Reprinted with permission from Ker et al. ⁵⁰ Copyright © 2011, Elsevier.

Nanotopography also greatly influences cell contact guidance. ⁵¹ In groove/ridge patterns, the contact guidance cues are efficient in aligning cells for groove sizes up to 100 μm. With the development of methods to fabricate materials from micro- to nanoscale, new advancements of SMTE became available and materials with nanofeatures are of great interest. Indeed, cells in vivo evolve in an ECM environment, which comprises a material with nanoscale features that is composed of different proteins. For example, collagen fibrils found in the ECM usually have a length of ∼10 μm and a width of 260–410 nm.^52,53 The fabrication of materials with nano-cues and nano-signals that are able to interact with cells and mimic the natural environment of the ECM can have tremendous applications in tissue engineering. Several studies have used columns, protrusions, pits, nodes, or nano-islands as substrates and have shown that small features, such as 11-nm columns, 35-nm pits, nanoposts (pointed columns), or 20-nm islands, promote cell adhesion, whereas increases in the size of these features decreases cell adhesion. ⁵⁴ Moreover, it has been shown that the symmetry of these features as well as the surface roughness also affects the adhesion of cells.^55–58 Surface topography affects not only cell orientation and elongation but also the cell differentiation. For example, adult rat hippocampal progenitor cells cultured on a patterned polystyrene dish with 16-μm-wide grooves overexpressed neuronal marker (class III β-tubulin) when compared with smooth substrates. ⁵⁹ Electrospinning has also been used to fabricate aligned nanofiber scaffolds to induce the alignment of myoblasts.^29,60–62 The 3D structure of the electrospun fibers resembles the physical structure of native collagen fibrils or ECM. ⁶³ However, although electrospun scaffolds are 3D structures, in many studies, the dense packing of fibers inhibits cell infiltration into the fiber network, and cells proliferate mostly on the top side of the electrospun fiber to generate a tissue similar to that formed using other 2D topographic substrates. ⁶⁴ Recently, direct electrospinning of a 3D aligned nanofibrous tube has been realized, promoting cell alignment and myotube formation. ⁶⁵ In another attempt to scale down the topography features, Dugan and coworkers employed oriented cellulose nanowhiskers with a diameter of 10–15 nm on a glass surface. They showed (Fig. 5) that myoblasts effectively sense the topography of such a surface and that myotubes resulting from myoblast fusion were nearly oriented in line with the direction of the cellulose nanowhiskers. ⁶⁶ This study clearly shows that cells are sensitive to topographical features, which affect cell orientation, shape, and differentiation.

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FIG. 5.

Oriented cellulose nanowhiskers of 10–15-nm diameter were produced by spin-coating. The response of myoblasts to the surfaces was assessed by atomic force microscopy 12 h after seeding. The inset picture shows a large-scale image of the whole cell (inset scale bar=20 μm), whereas the yellow arrow indicates the direction of the nanowiskers. The main image shows a closeup scan of the area bounded by the dashed box in the inset picture (scale bar=5 μm), whereas the white arrows indicate filopodia. Reprinted with permission from Dugan et al. ⁶⁶ Copyright © 2010, American Chemical Society.

To mimic in vivo muscle tissue, engineering of a 3D structure from aligned myotubes is needed. Zhao et al. have shown that a multilayered construct of aligned myotubes can be obtained by seeding additional myoblasts on a first layer of aligned myotubes formed in a groove (2-μm width and depth)/ridge micropattern. ³⁸ In addition, Hume et al. recently showed that if C2C12 cells aligned well in small grooves (≤100 μm) and did not align in large grooves in 2D culture, then their behavior changed in 3D culture. ⁶⁷ Thus, C2C12 cells were able to align in larger grooves (width of 200 μm and depth of 200 μm) when layered into the grooves. This study highlights the importance of 3D cultures in tissue engineering applications.

In an attempt to generate 3D tissue cultures in an environment that allows cells to assume a shape and exhibit matrix adhesion similar to that of native tissues, hydrogels have been extensively studied. ⁶⁸ Hydrogels can be generated from synthetic (e.g., poly(ethylene glycol) [PEG]) or natural polymers (e.g., collagen, chitosan, and hyaluronic acid). To generate skeletal muscle tissue, myoblasts must proliferate, migrate, align, and fuse to form a functional construct. ECM-derived hydrogel-like collagen or fibrin gel contains fibrils. These tiny protein nanofibers play the role of natural 3D topographical cues to guide the cells.^69,70 Thus, Lanfer et al. used a microfluidic device to create highly aligned type I collagen matrices and showed that myotube assembly and alignment were influenced by the topographical feature of collagen fibrils. ⁷¹ Hydrogel molding was also considered for guiding myoblasts in a method developed by Bian and coworkers. In their work, a PDMS mold was used to cast cell-laden hydrogels resulting in myotube alignment depending on the geometry and size ⁷² (Fig. 6).

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FIG. 6.

(a) A Silicon wafer coated with SU-8 was patterned by UV through (b) a photomask (scale bars=2 mm; inset=500 μm). (c) Optical profile of the master mold. (d) Polydimethylsiloxane (PDMS)-negative replica (scale bars=1 mm; inset vertical cross-section 500 μm). (e) PDMS-positive replica (scale bars=1 mm; inset vertical cross-section 500 μm). (f) Cells in hydrogel prepolymer solution were poured in the PDMS mold and incubated at 37°C to allow (g) hydrogel polymerization. (h) The culture medium was then added and the cells were cultured for 2 weeks. The hydrogel was fixed on a Velcro frame (scale bars in f–h 5 mm). Reprinted with permission from Bian et al. ⁷² Copyright © 2009, Nature Publishing Group. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

A composite 3D scaffold made of parallel glass fibers within a collagen gel has been used to direct the growth and differentiation of primary human masseter muscle derived cells.^73,74 The effect of the cell density on the maturation and contractile ability of an engineered muscle in a collagen gel was also studied by the same group. ⁷⁵ In another study, gelatin, which is a hydrolyzed derivative of collagen, has been methacrylated to become photocrosslinkable. This acrylated gelatin showed promising aspects for supporting cell proliferation, and cell-laden photopatterned methacrylated gelatin was successfully used to direct, elongate, and align myoblasts in a 3D hydrogel environment^76,77 (Fig. 7). These techniques rely on the limitation of cell migration by molding them in groove-like structures, which induced their alignment in a 3D environment improving their functionalities.

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FIG. 7.

3T3 fibroblasts encapsulated in 5% GelMA hydrogels patterned into rectangular microconstructs [50 μm (w)×800 μm (l)×150 μm (h) with 200-μm interlines]. (A) Hydrogel stained with Rhodamine B showing initial microconstruct at day 0 and phase-contrast images of cell-laden microconstructs at days 1, 4, and 7 of culture. The red arrows indicate points of contact between neighboring lines at day 4 of culture, with tissue convergence at day 7 of culture. (B) Three-dimensional (3D) tissue construct (1×1 cm²) at day 7 of culture. (C) F-Actin staining showing the middle of the 3D tissue construct with aligned actin fibers. Reprinted with permission from Aubin et al. ⁷⁷ Copyright © 2010, Elsevier. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

Cell alignment by surface patterning

Surface patterning is a general term used to describe the modification of a biomaterial's surface by patterning techniques. Soft lithography, which was introduced by the Whitesides group in the late 1990s to facilitate microfluidic device fabrication and fast prototyping, has also been used to pattern surfaces. ⁷⁸ This technique is based on the use of an elastomeric master that is easy to mold or emboss and can be used directly as substrate for biological applications or as mold. Among the elastomers used, PDMS is the most popular elastomer for biological applications, and the construction of a PDMS master is related to another mold prepared by conventional photolithography approaches.^79–81 Soft lithography is widely used for the patterning of cells and proteins through using patterning techniques such as microcontact printing, microfluidic patterning, and stencil micropatterning.^23,82,83 To guide cells on a surface, patterning of ECM proteins, such as collagen, fibronectin, or laminin, is widely used, as is the printing of self-assembled monolayers (SAMs) with cell-repellant molecules, such as PEG derivatives, poly(ethylene oxide)-b-poly(propylene oxide) (PEO-PPO) block copolymer, or copolymer surfactant with primary hydroxyl groups, thereby limiting the area where cells can settle. A combination of printed cell repellant and cell-adhesive molecules could also be used^84–86 (Fig. 8). Another technique is showed by Shimizu et al., who used a stencil membrane to micropattern myoblasts and form a pattern of single myotubes ⁸⁷ (Fig. 9). Direct patterning of myoblasts by inkjet printing techniques has been also done to improve the cell alignment and tissue formation. ⁸⁸

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FIG. 8.

Micropatterned substrate building process and cell culture. (A) Schematic showing the preparation of the micropatterned substrate. (B) Phase-contrast images of the different micropatterns with cells: lines of different widths (300 μm, 150 μm, 80 μm, 40 μm, 20 μm, and 10 μm), tori of different inner diameters (40 μm, 100 μm, and 200 μm), and hybrid patterns of different arc degrees (30°, 60°, and 90°), (scale bar=100 μm). Reprinted with permission from Bajaj et al. ⁸⁴ Copyright © 2011, Royal Society of Chemistry. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

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FIG. 9.

Schematic showing the micropatterning of myotubes by the use of a thin PDMS stencil membrane. A BSA-coated membrane was attached on the surface of a Petri dish and C2C12 were seeded on the membrane. After culturing in differentiation medium, the membrane was peeled off under a microscope to free the micropatterned myotubes, which can be removed. Reprinted with permission from Shimizu et al. ⁸⁷ Copyright © 2010, Elsevier. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

Cell patterning has been mostly used to study cell behavior, such as cell migration, proliferation, cell–cell interactions, and drug screening, in a 2D environment. However, this approach is also appealing for the creation of 3D tissue-like constructs via cell-sheet-based tissue engineering. Indeed, various methods exist for the harvesting of prepatterned cell sheets. For example, Nagamine et al. used a fibrin gel to embed aligned myotubes into a 3D hydrogel system. ⁸⁹ Similarly, Huang et al. transferred aligned myotubes from a parallel micropattern of poly(2-hydroxyethyl methacrylate) (pHEMA) to a type I collagen gel overlaid on the micropattern. After 3 days of culture, the collagen sheet was rolled around a biodegradable polymeric mandrel to fabricate a tubular muscle-like construct with aligned myotubes. ⁹⁰ Polymeric nanomembranes, with exceptional flexibility were also micropatterned by microcontact printing with lines of fibronectin plus multiwalled carbon nanotubes (MWNTs) to enhance the cell alignment and myotube formation and then rolled up to fabricate a tubular structure. ⁹¹ In an interesting study, prevascularized 3D muscle tissue was constructed by stacking multiple layers of endothelial cells sandwiched by myoblast sheets. ⁹² Petri dishes covalently grafted with the temperature-responsive polymer poly(N-isopropylacrylamide) were used to harvest the different cell sheets, and the handling of the cell sheets was secured by a plunger coated with a hydrogel matrix. By patterning hydrophilic polymer on thermoresponsive surface and by using a plunger coated with gelatin to harvest the different cell sheets of human skeletal myoblasts, Takahashi et al. showed that an anisotropic cell sheet placed on the top of four random cell sheets stacked together induced the myoblasts and the ECM alignment in the whole construct. ⁹³ Recently, Guillaume-Gentil et al. described a method to fabricate and harvest micropatterned heterotypic cell sheets by local electrochemical dissolution of a polyelectrolyte coating ⁹⁴ (Fig. 10). Such methods introduce the possible creation of cocultured harvestable cell sheets and the growth of more complex tissue constructs via cell-sheet-based tissue engineering.

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FIG. 10.

Micropatterning of heterotypic cell sheets. (a) Patterning of an SU-8 layer spin coated on an indium tin oxide (ITO) electrode by UV through a photomask. (b) After development, SU-8 micropatterns are formed on the ITO electrode. (c) The substrate is then coated with a weak cell-adhesive polyelectrolyte and (d) subjected to electrochemical polarization, which induced the dissolution of the polyelectrolyte only from the ITO regions. (e) The ITO regions are backfilled with a cell-repellent polymer PLL-g-PEG and (f) a first cell type is seeded whereas the nonadherent cells are washed away. (g) The PLL-g-PEG monolayer is then removed by a second electrochemical step. (h) The ITO regions are backfilled with PLL-g-PEG/PEG-RGD, which is a cell-adhesive monolayer. (i) The second cell type is seeded and the nonadherent cells are washed away. (j) After 1 day of culture, the PLL-g-PEG/PEG-RGD monolayer is dissolved by a final electrochemical step, which allows the whole cell sheet to detach. Indeed, due to their weak interaction with the substrate, the cells on weak cell-adhesive regions also detach easily. PEG, poly(ethylene glycol). Reprinted with permission from Guillaume-Gentil et al. ⁹⁴ Copyright © 2010, Springer Science+Business Media, LLC. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

Cell alignment by mechanical stimulation

Lack of stimulation and mechanical force causes muscle degeneration, as occurs in disabled individuals or during skeletal muscle atrophy in the microgravity of spaceflight. Although the role of mechanical stimulation has been widely studied in gene regulation, endogenous protein regulation, accumulation, and metabolic products,^95,96 it has been less studied as a tool in SMTE. However, it has been reported that under continuous uniaxial strain, avian myoblasts and L6 rat skeletal muscle cells cultured on an elastic substratum differentiated into myotubes oriented parallel to the direction of strain, whereas under stretching/relaxation cycles, the myotubes were aligned perpendicular to the stretch direction.^31,97,98 Other studies of myoblasts encapsulated in a collagen hydrogel and treated by continuous uniaxial strain also showed the formation of myotubes parallel to the direction of the strain.^5,99,100 One hypothesis to explain the difference in the angle of cell and myotube orientation between cells cultured under continuous strain or under stretching/relaxing cycles is the appearance of micro-ripples or micro-cracks in the matrix or in the elastomer surface perpendicular to the stretch direction, as observed for PDMS surfaces treated with stretching/relaxing cycles. ¹⁰¹ The passive tension observed during the coalescence of a collagen gel has also been used to align myoblasts between two posts to form aligned myotubes. ³

Cell alignment by magnetic or electrical fields

Electrical fields are often used in single-cell manipulation and cell sorting techniques but less in the engineering of a whole tissue. Indeed, a cell placed in an alternating electrical field polarizes into a dipole and is subjected to a dieletrophoretic force F ¹⁰² given by the formula: \documentclass{aastex}\usepackage{amsbsy}\usepackage{amsfonts}\usepackage{amssymb}\usepackage{bm}\usepackage{mathrsfs}\usepackage{pifont}\usepackage{stmaryrd}\usepackage{textcomp}\usepackage{portland, xspace}\usepackage{amsmath, amsxtra}\pagestyle{empty}\DeclareMathSizes{10}{9}{7}{6}\begin{document} \begin{align*}F = 2 \pi r^3 \varepsilon_m \mid ( \varepsilon_c - \varepsilon_m ) / ( \varepsilon_c + 2 \varepsilon_m ) \mid \nabla E^2 ,\end{align*} \end{document}

Where r is the radius of the cell, ɛ_m is the medium permittivity, ɛ_c is the cell permittivity, ∇E is the magnitude of the electrical field, and |(ɛ_c−ɛ_m)/(ɛ_c+2ɛ_m)| is the real part of the Clausius-Mossotti factor. ¹⁰³ If |(ɛ_c−ɛ_m)/(ɛ_c+2ɛ_m)|>0, then a positive dielectrophoresis (DEP) force F exists that induces the cell to move toward regions with a high electrical field. If |(ɛ_c−ɛ_m)/(ɛ_c+2ɛ_m)|<0, a negative DEP force F exists that repels the cell toward regions of low electrical field. As the magnitude of the DEP force F is inversely proportional to the electrode gap, the electrodes are usually designed to be close to each other to allow the induction of an electrical field of several hundred V/cm that is suitable for cell manipulation. ¹⁰⁴ Thus, DEP has been used to pattern several cell types for coculture and tissue engineering applications^105–107 (Fig. 11).

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FIG. 11.

Patterning of two different cell populations by dielectrophoresis (DEP). (A) An interdigitated array of four-electrode subunit was used to pattern cells. (B) By applying an AC voltage, the n-DEP force was induced and cells were directed toward weaker region of electric field strength. (C) Cells in excess were removed. (D) A second cell type was loaded into the device and guided to other areas by changing the AC voltage mode. Reprinted with permission from Suzuki et al. ¹⁰⁷ Copyright © 2008, Elsevier. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

Ramon-Azcon and colleagues used DEP to pattern C2C12 myoblasts in a hydrogel matrix, which resulted in a highly aligned muscle tissue construct.^32,33 MWNTs have also been included into hydrogel and aligned by DEP improving the hydrogel electrical conductivity and favoring the cell alignment and the myotube formation. ¹⁰⁸ Some notable characteristics of the DEP method include accuracy, high cell manipulation speed, and the ability to scale-up. ¹⁰⁹ It has also been shown that a static magnetic field alone can induce the alignment of L6 myoblasts. ¹¹⁰ However, the mechanism underlying this phenomenon is not well understood. Yamamoto et al. reported that C2C12 cells were elongated along the axis of a magnetic field after endocytosis of magnetic nanoparticles. ¹¹¹ By using this method of magnetic-force-based tissue engineering, ¹¹² which promotes tissue organization under a magnetic field, Akiyama et al. fabricated 3D tissue architecture, ¹¹³ whereas Yamamoto et al. fabricated 200-μm-thick skeletal muscle tissue.^111,114 To fabricate a 1.9-mm-thick skeletal muscle tissue, Yamamoto et al. combined the application of a magnetic field to C2C12 cells loaded with magnetic nanoparticles to induce tissue organization with the use of cell culture in a perfused hollow fiber reactor that allowed the maintenance of high cell density by supplying oxygen and nutrients. ¹¹⁵

Cell differentiation

Myogenesis is the differentiation process that drives the formation of multinucleated myotubes. However, cell proliferation and phenotypic differentiation are mutually exclusive events. Therefore, myoblasts have to exit the cell cycle to enter into myogenesis. During this process, mononucleated myoblasts, which have a spindle or a polygonal shape, migrate toward each other and aggregate. Following this cell adhesion, the myoblasts align in an end-to-end configuration ⁴¹ by the parallel apposition of their membranes. ¹¹⁶ Membrane fusion then occurs, and the cells fuse together to generate a multinucleated structure. ¹¹⁷ Initially, myoblasts fuse with each other to form small nascent myotubes, which subsequently fuse with additional myoblasts to form large and mature myotubes.^118,119 All of these successive events are regulated by numerous factors, such as transcription factors, and involve several protein regulatory mechanisms, which are discussed in the following subsections.

Molecular induction of myoblast differentiation

Electric and magnetic induction of myoblast differentiation

Mechanical and electrical stimulation are linked to muscle tissue formation. In the laboratory, an electrical stimulator can be used to easily mimic neuronal muscle stimulation with controlled parameters. ²³⁸ In one example, Flaibani et al. applied 3-ms pulses with an amplitude of 70 mV/cm for 30 s to muscle precursor cells cultured on a micropatterned poly(L-lactic acid) membrane. ²³⁹ They observed an increase in myotube density with a 30% increase in the release of nitrite (NO₂⁻), which is a signaling molecule that is involved in myoblast fusion and myotube growth. ¹⁵⁷ Kawahara et al. applied 2-ms pulses of 50 V for 5 min/day to L6 rat myoblast cultures and observed accelerated myotube differentiation with the formation of thick myotubes followed by contracting striated muscle cells. ²⁴⁰ The electrical stimulation was achieved by plunging electrodes into the culture medium. The drawback of this method is that most of the current lines cross under the culture medium rather than under the cells. In addition, electrolysis and the release of toxic products in the medium may result from this type of setup and should be avoided. Thus, to focus the current toward the myotubes, Kaji et al. cultured C2C12 cells on a conductive porous membrane and adopted a vertical setup for the electrodes. ²⁴¹ To protect the cells, Nagamine et al. transferred cultured myotubes to a fibrin gel, which they placed on microelectrode arrays for electrical stimulation (amplitude 2 V, duration 3 ms, frequency 10 Hz, train 1 s, and interval 10 s). The fibrin gel had been previously coated with a conductive polymer to improve interfacial electrical capacity. ⁸⁹ Other groups have also opted to use electrically conductive polymers to provide a matrix environment with safer electrical stimulation for the cells. In such cases, the cells can be encapsulated in the conductive polymer, seeded on the conductive polymer, or encapsulated in a polymer placed on electrodes. Thus, Sirivisoot and Harisson studied the formation of myotubes on electrically inductive composite scaffolds generated from electrospun polyurethane (PU) and carbon nanotubes. ²⁴² They showed that increased myotube formation was correlated with increased electrical conductivity because of the presence of carbon nanotubes. Also, Sekine et al. printed poly(3,4-ethylenedioxythiophene) (PEDOT) electrodes on an agarose hydrogel and electrically stimulated contractile myotubes embedded in a fibrin gel deposited on the PEDOT/agarose sheet, ²⁴³ whereas Ido et al. embedded PEDOT electrodes in a hydrogel. ²⁴⁴ In another strategy, Mawad et al. combined the advantages of hydrogels (e.g., mechanical properties, hydrated environment, and biocompatibility) and conducting polymers to build a conducting polymer hydrogel on which C2C12 cells proliferated. ²⁴⁵ Ku et al. electrospun a blend of polycaprolactone with polyaniline to combine the topographical constraint of aligned fibers with electrical conductivity and observed a synergistic effect on myotube formation. ²⁴⁶

Electrical stimulation not only increases the myotube density by increasing the speed of formation but also changes the nature of the muscle fibers and acts at the molecular level of muscle fiber formation. For example, after 8 days of conventional differentiation, C2C12 myotubes lacked spontaneous contractibility because of negligible sarcomere architecture. Fujita et al. used electrical pulses to study the effects of Ca²⁺ oscillation on the assembly of functionally active sarcomeres. ²⁴⁷ They observed an increase in striated myotubes that peaked at 2 h and then decreased during stimulation with a 24-ms electrical pulse of 40 V/mm at 1 Hz. When they applied a lower-frequency signal (0.1 Hz), the striation was delayed and peaked at 12 h of stimulation. When they applied a high-frequency signal (10 Hz) for 2 h, they did not induce sarcomere assembly and did not observe contractile activity. This electrically induced contractile activity appeared to be mediated through the protease activity of calpain and also involved ECM-integrin engagement. Another study ²⁴⁸ (Fig. 12) by the same group showed that electrical stimulation induced a switch from fast MyHC chain (type II) to slow MyHC chain (type I); consequently, the muscle fiber phenotype changed under stimulation. ²⁴⁹

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FIG. 12.

Effects of electrical stimulation on muscular protein and sarcomere development. C2C12 myotubes were stimulated 24 h with electrical pulses (40 V, 1 Hz, 2 ms duration). The cell lysates were analyzed by western blot (left) and cells were fixed (right) and stained with DAPI for nucleus (blue), anti-sarcomeric α-actinin antibody for sarcomeric α-actinin (red), and phalloidin for MHC (green). Reprinted with permission from Nedachi et al. ²⁴⁸ Copyright © 2008, The American Physiological Society. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

Some studies have also focused on the effects of combinations of stimulation types applied to cells. For example, Liao et al. combined mechanical and electrical stimulation by culturing C2C12 cells on PU mats with different diameters and fiber orientations under stretching and 10-ms electrical pulses of 20 V. ²⁵⁰ They showed a net improvement of myotube striation and contractile protein (myosin, MyHC, and α-actinin) secretion under electromechanical stimulation but did not observe a significant benefit of bistimulation over monostimulation (mechanical or electrical only). Magnetic induction has also been used to induce myoblast differentiation. For example, Yuge and Kataoka introduced magnetic microparticles (0.05–0.1 μm Ø) into the cytoplasm of rat myoblast L6 cells by electroporation and then cultured the cells in magnetic fields of 0.01, 0.03, or 0.05 T. ²⁵¹ They observed that cells aligned and elongated along the N-S direction of the magnetic lines and that the formation of myotubes accelerated with the intensity of the magnetic field. Complete differentiation was obtained with striated myotube formation, and the myotube size also increased. Another study showed that a magnetic field of 80 mT orthogonal to the cell plane promoted myogenic differentiation and myotube hypertrophy without requiring other treatments, such as the introduction of magnetic particles into cells. ¹¹⁰ Interestingly, when cells were treated with 5 μg/mL of TNFα, which is an inhibitor of myoblast differentiation, ²⁵² the exposure of cells to the magnetic field restored the myogenic differentiation. Clearly, electrical stimulation with or without other types of stimulation has important effects on myoblast differentiation and allows faster myotube formation, higher myotube density, increased myotube size, and a higher degree of myotube maturation.

Mechanical induction of myoblast differentiation

It is known that in body-building, muscular work through resistance training results in muscles with increased diameter. Therefore, it is interesting to analyze the effects of mechanical stimulation on myogenesis. Cells can be mechanically stimulated by topography, the stiffness of the material, and stretching. We previously described (“Cell alignment by topography” section) the degree to which topography can influence cellular alignment. Because myoblasts fuse mainly in an end-to-end configuration, cellular alignment seems to be required to obtain myotubes. Curiously, when Charest et al. analyzed the differentiation of C2C12 cells cultured on topographic patterns consisting of embossed ridges and grooves or arrays of holes with sizes ranging from 5 to 75 μm on polycarbonate coated with SAM and fibronectin, they concluded that the topography strongly influenced myoblast alignment but had no effect on the differentiation of the myoblasts. ²⁵³ Their analysis, however, was mainly based on the measurement of sarcomeric myosin expression, and no images of myotubes were presented. In contrast, Bajaj et al. showed that the geometrical cues of a substrate significantly affect myoblast differentiation. ⁸⁴ They cultured C2C12 cells on dishes with different geometrical fibronectin coatings for 1 week and observed that myotubes with a hybrid line-torus shape had a two- and threefold increase in their fusion index when compared with myotubes on line and torus patterns, respectively (Fig. 13).

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FIG. 13.

Topographical effects on C2C12 differentiation and myotube formation at different days in differentiation culture medium. Cells were cultured on unpatterned (control) and patterned substrates with different shapes (scale bar=100 μm) and stained with DAPI for nucleus (blue) and anti-MHC (green). Reprinted with permission from Bajaj et al. ⁸⁴ Copyright © 2011, Royal Society of Chemistry. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

Similarly, when Aviss et al. prepared a scaffold of aligned poly(lactic-co-glycolic) acid (PLGA) fibers by electrospinning and cultured C2C12 cells for 14 days, they initially observed cell alignment with the fiber axis after 30 min of culture, followed by cell differentiation into long multinucleated myotubes aligned along the fibers. ²⁹ Our group also showed that topographical features can favor myoblast differentiation into myotubes. ²⁵⁴

The stiffness of the material is also an important parameter in biomaterial and tissue engineering. Indeed, many studies have shown that on a material with different degrees of stiffness, cells migrate toward locations with the preferred stiffness. ²⁵⁵ Engler et al. studied the effect of material stiffness on myoblast differentiation by culturing C2C12 cells on collagen-micropatterned polyacrylamide (PA) gels with different degrees of stiffness for 2 and 4 weeks. ²⁵⁶ Although all cultures formed myotubes, myotubes only fully differentiated and reached myosin striation on gels of intermediate stiffness (8–11 kPa) (Fig. 14). The plot of material stiffness versus myotube striation fitted a Gaussian curve with an optimal modulus of 12 kPa that maximized myosin striation, whereas myotubes on gels with low (<5 kPa) or high stiffness (>17 kPa) had only poor or no striation. This optimal modulus value closely matched the elasticity of C2C12 myotubes, which is 12–15 kPa, ²⁵⁷ and the native skeletal muscle tissue stiffness value, which is 12±4 kPa, ²⁵⁸ as measured by atomic force microscopy on extensor digitorum longus muscles harvested from C57 mice.

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FIG. 14.

Stiffness effects on myotube striation at 4 weeks in differentiation culture medium. C2C12 were cultured on collagen-patterned substrates with different stiffness (scale bar=20 μm) and stained with DAPI for nucleus (blue) and anti-MHC (green). Myosin striation occurred only when cells were cultured on intermediated stiffness substrate. Reprinted with permission from Engler et al. ²⁵⁶ Copyright © 2004, Rockefeller University Press. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

Other studies also showed that the degree of myoblast differentiation depends on material stiffness. For example, Ren et al. generated biopolymeric films of poly(L-lysine)/hyaluronan with a controlled stiffness ranging from 3 kPa (native film) to 100 kPa (low-cross-linked films) and 400 kPa (high-cross-linked films). ¹¹⁸ After culturing C2C12 cells on these films for 1 week in differentiation medium, they observed that myotubes were short and thick on soft films, whereas they were elongated and thin on stiff films and on a polystyrene dish, which was used as a control. Myotube striation increased with the stiffness of the film, from 14% (with a ∼350-kPa film) to 43% (with a ∼400-kPa film) and 69% (with the polystyrene plastic dish; 1 MPa). By using surfaces with different concentrations of silk and tropoelastin, Hu et al. studied the impact of surface roughness and stiffness on the differentiation of myogenic and osteogenic lineages. ²⁵⁹ They showed that C2C12 cells preferred low surface roughness with high stiffness. Regarding myoblast differentiation under stretching, the recent literature is controversial. Globally, the consensus is that stretching stimulation increases myoblast proliferation but decreases myoblast differentiation.^260–264 In SCs, it also seems that stretching induces NO production and hepatocyte growth factor secretion, both of which are involved in SC activation.^265–267 However, the literature diverges concerning MRF expression by myoblasts under stretching. Kook et al. found that mechanical stretching stimulated C2C12 cell proliferation but inhibited differentiation into myotubes through the continuous phosphorylation of p38 MAP kinase, which decreases the level of MyoD expression. ²⁶³ Akimoto et al. cultured C2C12 cells on a silicon membrane (BioFlex) that was stretched with a 20% elongation for 24 h at a frequency of 10 cycles/min (2-s on time, 4-s off time) and observed a decrease in the expression of MyoD and myocyte nuclear factor-α when compared with the nonstretched culture. ²⁶⁸ Similarly, Kuang et al. observed a reduction of myogenin expression in C2C12 cells cultured under stretching stimulation. ²⁶¹ In contrast, another study by Abe et al. used a similar system (Flexercell) to culture C2C12 cells stretched to 15% elongation with cycles of 1-s on stretch and 1-s off stretch and observed an increase of MyoD and other myogenic factors after stretching for 12 and 24 h. ²⁶⁹ Similarly, Gomes et al. reported increased MyoD expression 24 h after a passive stretching session on in vivo rat muscles. ²⁷⁰ Another study using adipose-derived stem cells demonstrated an upregulation of MyoD when the cells were stimulated by stretching. ²⁷¹ Similarly, by using myoblasts loaded in an anchored mixture of collagen-Matrigel and submitted to frequent strain, Powell et al. demonstrated a 12% and 40% increase in myotube diameter and density, respectively. ²⁷² Although several studies have established the role of mechanotransduction in myogenesis activation, notably through the p38 MAPK pathway,^273,274 the effects of stretching on myoblast differentiation are not yet well understood. A standardized stretching process may be beneficial to better compare results from different research groups.

Regenerative medicine and stem cells

One of the main goals of muscle engineering is to develop muscle tissue for medical applications. Thus, the replacement of damaged, wounded, or nonfunctional muscle tissue is an important goal of most research in this field and has been extensively reviewed.^1,309 Traditionally, regenerative strategies are based on ex-vivo-engineered constructs with autologous cells that can be reimplanted into the patient. ³¹⁰ SCs are stem cells that directly involve in muscle regeneration. ³¹¹ They are located between the sarcolema of the muscle fibers and the basal lamina under a quiescent status (G0 phase) and are characterized by the expression of the transcription factors Pax7 and Myf5 but not MyoD or Myogenin. ³¹² When the muscle fibers are damaged, SCs are activated and start to proliferate and to express MyoD. A symmetric/asymmetric mechanism of cell division induces the generation of SCs expressing Pax7⁺/Myf5⁻ (basal cell in contact with the muscle fiber) that will contribute to the maintenance of the pool of SCs, and other SCs expressing Pax7⁺/Myf5⁺ (apical cell in contact with the other cell) that will differentiate and will fuse together into myotubes to regenerate the muscle fibers.^313,314 Freshly harvested SCs have been used in injured or disease mice model that shows efficient myofiber regeneration. ³¹⁵ However, SCs cultured in vitro and then transplanted have shown decrease in cell proliferation and in their myogenic potential. ³¹⁶ In addition to their potential of generating different tissues, a large diversity and heterogeneity exist among stem cells; therefore, other cell types may also be used for muscle regeneration. ³¹⁷ Bone marrow stem cells, which contain hematopoietic stem cells and mesenchymal stem cells, have been shown to restore the SC pool and to generate myofibers in injured mice.^318,319 However, bone marrow transplantation has not been reported efficient in therapy for muscle and further studies are required to define the subpopulation of bone marrow cells with myogenic potential in an animal model of muscle disease.^312,320 Muscle side population cells (SPs) are localized in the interstitium between the muscle fibers, near the blood vessels, and are characterized by the expression of Sca-1⁺, ABCG2⁺, CD45⁻, CD43⁻, c-kit⁻, and Pax7⁻. SPs are able to differentiate into SCs and to form myotubes in monoculture or in coculture with myogenic cells.^321–323 A subgroup of SPs characterized by Sca-1⁺, ABCG2⁺, CD45⁻, Pax7⁺, and Syndecan-4⁺ has been shown to regenerate skeletal muscle efficiently in an injured mouse model. ³²⁴ Moreover, SPs injected intravenously were able to migrate toward the injured muscle for restoring it. ³²⁵ Another group of cells from the interstitium are characterized by PW1⁺/Pax7⁻ and are myogenic in vitro. ³²⁶ When injected in vivo into injured muscle, these cells both proliferated to increase their own pool and differentiated into SCs (Pax7⁺). ³²⁷ Muscle-derived stem cells (MDSCs) are characterized by Sca-1⁺, CD45⁻, CD34⁻, Flk1⁺, Desmin⁺, and M-cadherin⁻ and are multipotent. ³²⁸ Used in a model of dystrophic mice (mdx mice), MDSCs injected in the blood stream were able to migrate into the host muscle tissues and to regenerate myofibers and dystrophin expression. ³²⁹ Platelet-rich plasma has been used in culture medium to improve in vitro the expansion of MDCs while maintaining their stemness. ³³⁰ Mesoangioblast cells are characterized by CD34⁺, c-kit⁻, Flk1⁺, NKX2.5⁻, Myf5⁻, and Oct4⁻ and are multipotent. ³¹² They proliferate well in vitro maintaining their multipotency and have shown a particular efficiency in the treatment of different animal models of muscular dystrophy.^331,332 Pericytes are derived from blood vessel and are characterized in human by CD45⁻, CD34⁻, CD56⁻, CD144⁻, CD146⁺, PDGFR-β1⁺, and NG2 proteoglycans⁺. ³³³ Transplanted in dystrophic animal through blood stream, pericytes repopulated the SC niche and regenerated myofiber. ³³³ AC133 cells or CD133⁺ cells are a subpopulation of hematopoietic cells with myogenic activity when cocultured with myogenic cells or with cells expressing Wnt. ³³⁴ When injected into a muscle of dystrophic mice, cocultured AC133 cells replenished the SC niche and contributed in myofiber regeneration. ³³⁵ An autologous transplantation of AC133 cells in a dystrophic boy demonstrated the safety of the therapeutic strategy used but did not offer substantial functional benefit. ³³⁶ Embryonic stem cells (ESCs) are pluripotent and have great potential in therapy applications. However, immunogenic response, teratoma formation, and ethical concerns are the major hindrances of their use. The derivation of human ESCs into multipotent mesenchymal precursors followed by their differentiation into myoblasts allowed stable engraftment after transplantation into muscle-injured mice without teratoma formation. ³³⁷ The use of ESC-derived embryoid bodies induced to myogenic differentiation has also been used and transplanted myoblasts in muscle-injured mice showed stable engraftment, myofiber regeneration, replenishment of the SC pool, and absence of teratoma. ³³⁸ Induced pluripotent stem cells (iPS cells) are reprogrammed somatic cells into ESC-like state. ³³⁹ The pluripotency is reprogrammed by nuclear transfer of four transcription factors (Oct3/4, Sox2, c-Myc, and Klf4). ³⁴⁰ iPS cells can be generated from many cell types and different methods have been developed to avoid the use of viral vectors for the delivery of the four transcription factors.^341,342 Moreover, iPS cells can be generated from genetic-disease tissues and can be used for specific disease study and drug screening. ³⁴³ Myogenic progenitors can be derived from iPS cells and when transplanted in dystrophic mice have been shown to efficiently engraft, to replenish the SC pool, to regenerate dystrophin-positive myofibers, and to improve contractibility. ³⁴⁴ Human iPS cells have also been generated from disease patient, genetically corrected, and transplanted in dystrophic mice showing specific functional tissue and gene restorations. ³⁴⁵ In summary, stem cell therapy is a developing field and encouraging results have already been obtained in muscular regeneration. Some stem cells, such as mesangioblasts or AC133, are already in clinical-trial phase. Depending on the stem cell type, the main problems encountered are the difficulties of stem cell expansion in vitro and the loss of stemness, the difficulties of systemic delivery via blood stream due to the inability to cross the endothelial wall, the migration of the stem cells to the damaged muscle, and the survival of stem cells after injection. However, the techniques of cell derivation and of genetic modification open the field of stem cell therapy and make it full of potentialities. ³⁴⁶ Recently, a new therapeutic strategy appeared with a recent shift toward an in-vivo-regenerative strategy, also called in situ tissue regeneration or endogenous regeneration, based on the recruitment and stimulation of progenitor cells in situ.^347–350

Drug screening

Engineered skeletal muscle tissue is useful not only for reconstructive surgery but also for applications in drug screening. Indeed, engineered muscle tissue offers a physiological environment that should provide valuable information in multiple signaling pathways about drug efficiency, pharmacology, and toxicity. ³ Such skeletal tissues used as a substrate can be either healthy or a model of human diseases and must be built in a reproducible way to allow standardization and comparison of results. As skeletal muscular tissues are notably involved in glucose homeostasis, any impairment of glucose signal transduction will decrease the glucose uptake by muscles under insulin stimulation and may contribute to type 2 diabetes development. ³⁵¹ Understanding this insulin activation pathway is therefore important. It has been shown that the glucose transporter GLUT4 is translocated from intracellular storage compartments to the plasma membrane under insulin stimulation.^352,353 During this translocation, accelerated GLUT4-containing vesicle exocytosis was observed, as well as the involvement of the PI3K and Akt pathways.^354,355 Thus, several investigators reported glucose uptake by differentiated C2C12 cells.^356–358 However, to provide a model with higher insulin sensitivity, Hayata et al. established a new cell line named C2C12-IS and screened a chemical library in a high-throughput screening (HTS) study to search for compounds that promote glucose uptake. ³⁵⁹ Although HTS is a powerful method that uses 2D-cell-based assays to screen a large amount of potential drugs, the use of cell cultures as a substrate does not necessarily match the in vivo complexity, which results in a great number of hits that fail in clinical trials.^360,361 Indeed, many studies have now shown that cellular behavior in the 2D and 3D environment is different.^67,362 In addition, the use of animal models for testing drug efficacy creates ethical problems, and the results derived from such models are difficult to translate to humans. Therefore, there is a great need for new substrates that mimic the complex biological architecture ³⁶³ and support its physiological and metabolic function. To this end, some investigators have chosen to use simple organisms, such as zebrafish or nematode worms, for drug and genetic screening,^364–366 whereas other groups have chosen to develop engineered muscle tissue ³ (Fig. 17).

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FIG. 17.

Engineering of miniature bioartificial muscles (mBAMs) on PDMS microposts in a 96-well plate for drug screening. PDMS microposts are 7-mm high, 7 mm Ø, and 4-mm apart. The mBAM shown in the well is 5 days postcasting whereas at day 7 the mBAM showed aligned myofibers after staining for sarcomeric tropomyosin. Reprinted with permission from Vandenburgh. ³ Copyright © 2010, Mary Ann Liebert, Inc.

An important aspect of such muscular models that mimic the in vivo structure is to establish control of the contractibility of the engineered muscular tissue. This control can be achieved through neuronal or electrical stimulation. To this end, Nagamine et al. used microelectrode arrays to stimulate myotubes, and Kaji et al. showed that myotube contractibility was positively correlated with the glucose uptake.^241,367 The current transition from 2D to 3D in vitro muscular models is also of great importance for drug testing. ³⁶² Finally, in the future, new directions for tissue drug screening, such as the individual screening of patient tissue through outpatient biopsy procedures, may also be possible.^368,369 Moreover, iPS cells ³⁷⁰ and ESCs^371,372 represent potential sources of cells from which to engineer tissues for drug-screening applications.

Other applications

In addition to the aforementioned medical applications, engineered muscle tissue also has great potential in many other applications. Indeed, muscle is the best natural motility motor; therefore, an actuator made of living muscle would have several advantages regarding efficiency when compared with a synthetic actuator. ³⁷³ Muscle tissue is also built by a hexagonal lattice of tightly packed filaments composed of actin and myosin chains, which has the best-known packing order and the lowest volume when compared with synthetic counterparts. These units of actin chains sliding on myosin chains are among the smallest, most well-organized motors and could be modified to generate miniaturized moving parts for micromechanical devices with several applications in biological microelectromechanical systems. ³⁷⁴ The mechanical force developed during muscle contraction can also be converted into other forms of energy, such as electricity. ³⁷⁵ Currently, researchers have attempted to harvest electricity from muscle contraction mostly for use as an energy source for implanted medical devices and other applications.^376,377 Recently, Wang et al. were issued a patent in which the energy from the contractions of muscle cells cultured on an array of piezoelectric nanowires could be turned into electricity. ³⁷⁸ Ishisaka et al. used pulsating heart cells as a micromechanical actuator to produce electricity by culturing heart muscle cells on a PDMS membrane, which transfers the vibration energy to piezoelectric fibers. ³⁷⁹ These efforts show the use of muscles as electrical generators. In the context of the world energy crisis, there is a strong demand for new energy sources. Therefore, harvesting energy from muscle contractions may lead to important developments and applications in the future.

The food industry is another field in which engineered muscles could have applications. Indeed, meat is the most important source of protein in the human food cycle. Recent studies showed that engineered muscle tissues (cultured meat) as a new source of protein have several advantages over conventionally produced meat. For example, in comparison to conventionally produced meat, cultured meat reduces the use of energy by ∼7% to 45% (only poultry has a lower use of energy), greenhouse gas emissions by 78–96%, land use by 99%, and water use by 82–96% (Fig. 18). Finally, it was concluded that the overall impact on the environment from cultured meat production was substantially lower than those from conventionally produced meat. ⁹ A tissue-engineered meat product and a method for producing such meat were also disclosed in a patent that gives a commercial prospective for such meat sources. ³⁸⁰ Recently, Mark Post made the prototype burger of cultured meat from stem cells.^381,382

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FIG. 18.

Comparison of energy, greenhouse gas (GHG) emissions, and land and water used between different sources of meat production. Reprinted with permission from Tuomisto and Teixeira de Mattos. ⁹ Copyright © 2011, American Chemical Society. Color images available online at www-liebertpub-com.web.bisu.edu.cn/teb

In addition, a patent has also been issued on artificially produced 3D muscle tissue, which shows the value of engineered muscle tissue for applications that may interest investors. ³⁸³ All these examples, and others, clearly show the progress in SMTE toward clinically feasible, functional, engineered muscles, which could improve the life of patients.