Advanced Review
Engineered skeletal muscle tissue for soft robotics: fabrication strategies, current applications, and future challenges Rebecca M. Duffy1 and Adam W. Feinberg1,2∗ Skeletal muscle is a scalable actuator system used throughout nature from the millimeter to meter length scales and over a wide range of frequencies and force regimes. This adaptability has spurred interest in using engineered skeletal muscle to power soft robotics devices and in biotechnology and medical applications. However, the challenges to doing this are similar to those facing the tissue engineering and regenerative medicine fields; specifically, how do we translate our understanding of myogenesis in vivo to the engineering of muscle constructs in vitro to achieve functional integration with devices. To do this researchers are developing a number of ways to engineer the cellular microenvironment to guide skeletal muscle tissue formation. This includes understanding the role of substrate stiffness and the mechanical environment, engineering the spatial organization of biochemical and physical cues to guide muscle alignment, and developing bioreactors for mechanical and electrical conditioning. Examples of engineered skeletal muscle that can potentially be used in soft robotics include 2D cantileverbased skeletal muscle actuators and 3D skeletal muscle tissues engineered using scaffolds or directed self-organization. Integration into devices has led to basic muscle-powered devices such as grippers and pumps as well as more sophisticated muscle-powered soft robots that walk and swim. Looking forward, current, and future challenges include identifying the best source of muscle precursor cells to expand and differentiate into myotubes, replacing cardiomyocytes with skeletal muscle tissue as the bio-actuator of choice for soft robots, and vascularization and innervation to enable control and nourishment of larger muscle tissue constructs. © 2013 Wiley Periodicals, Inc.
How to cite this article:
WIREs Nanomed Nanobiotechnol 2014, 6:178–195. doi: 10.1002/wnan.1254
INTRODUCTION
S
keletal muscle serves as the primary actuator system in animals and can perform tasks over a wide range of length and time scales, spatial configurations, and force regimes. The scalability ∗
Correspondence to: [email protected]
1
Department of Biomedical Engineering, University, Pittsburgh, PA, USA
Carnegie
Mellon
2
Department of Materials Science and Engineering, Carnegie Mellon University, Pittsburgh, PA, USA Conflict of interest: The authors have declared no conflicts of interest for this article.
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of skeletal muscle makes it a good candidate for soft robotics and related applications because it achieves a unique set of performance metrics. Specifically, skeletal muscle offers several advantages over traditional mechanical actuators including superior power-to-weight ratio, force-to-weight ratio, mechanical compliance, and systems control.1 For example, selective activation of muscle fibers via innervation and spatial or temporal summation allows dynamic control and modulation over force generation.2 Compared with traditional, rigid robots, soft robotics can have greater degrees of freedom and can be modeled after specific examples in nature,
© 2013 Wiley Periodicals, Inc.
Volume 6, March/April 2014
WIREs Nanomedicine and Nanobiotechnology
Engineered skeletal muscle tissue for soft robotics
such as the elephant trunk or octopus arm.3 This flexibility in configuration and force provides the potential for the same system to perform diverse functions from high-precision tasks such as wound suturing to high-strength tasks such as weight lifting. In addition, the energy source for skeletal muscle is adenosine triphosphate (ATP), generated from the energy stored within chemical bonds in amino acids, polysaccharides, and fatty acids and with approximately two orders-of-magnitude higher energy density than conventional energy sources used by nonbiological machines. Lastly, another advantage of biological actuators over man-made actuators is the potential ability to self-repair. At the molecular scale, the protein machinery that makes up the contractile apparatus is constantly being replaced while the muscle is functioning, enabling contraction for decades through billions of cycles. At the tissue scale, skeletal muscle in vivo can repair small injuries, preventing loss of muscle mass.4 While current tissue-engineered muscle constructs cannot heal at the tissue scale, enabling this ability to self-repair could revolutionize the field of robotics by creating damage-tolerant systems, and directly coupled to repair is adaptive growth in response to moderate mechanical overloading. Similar to exercise-induced muscle growth in vivo, engineered muscle constructs may eventually be able to grow and become stronger when the load requires more power, or even atrophy and reduce energy requirements when the load is decreased. Engineering skeletal muscle for use in soft robotics requires the ability to guide muscle precursor cells to differentiate into skeletal muscle cells (myotubes) and then to organize these cells into a functional tissue. This muscle formation process (termed myogenesis) is well documented in vivo,4–6 and has been achieved at a basic level in vitro in terms of myoblast fusion into myotubes. However, in vitro it has proven difficult to engineer the formation of dense, innervated, and vascularized muscle in 3D at the tissue scale, a challenge that also faces the field of tissue engineering and regenerative medicine. Advances in engineered skeletal muscle for tissue repair and as in vitro systems for drug discovery have been critical to our growing understanding of how to direct myogenesis in vitro.7–9 Here we will focus on current research to elucidate the physical, mechanical, and chemical cues necessary to guide myogenesis and how to recapitulate them. Initial results demonstrate that the extracellular matrix (ECM) is a key component to the myogenesis process and can serve as an instructive template for engineering skeletal muscle. Major factors include (1) substrate stiffness Volume 6, March/April 2014
and the mechanical environment, (2) the spatial organization of biochemical cues, (3) physical and topographical cues, and (4) electrical and mechanical conditioning. Applied to the engineering of skeletal muscle, controlling some of these parameters has enabled the fabrication of 2D and 3D constructs suitable as actuators in soft robotics. Furthermore, researchers have integrated these engineered muscle tissues into various devices from simple grippers to semi-autonomous walking and swimming soft robots. These proof-of-concept examples highlight the unique capabilities of engineered muscle and the fact that current technology can start to leverage muscle as an actuator system in engineered devices. However, challenges remain and still need to be addressed before engineered muscle is used more widely. Tissue size is limited by the lack of a vascular system for nutrient transport, long-term viability in a nonsterile environment is poor because of lack of an immune system to fight infection, and long-term stability of the muscle is inadequate as the cells slowly die due to removal from the in vivo environment. Looking forward, research in skeletal muscle tissue engineering and regenerative medicine coupled with the growing field of muscle-based soft robotics suggests that many of these problems will be overcome and that wide spread use of muscle as an actuator in a range of biotechnologies from soft robotics, to bioprosthetics to bio-microelectromechanical systems (bio-MEMS) is rapidly approaching.
ENGINEERING THE MICROENVIRONMENT TO GUIDE SKELETAL MUSCLE FORMATION The Role of Substrate Stiffness and the Mechanical Environment Skeletal muscle is a soft tissue composed of myotubes, blood vessels, nerves, and an interconnected network of ECM. The elastic modulus of muscle tissue depends on the specific type, but is reported to be in the range of 10–40 kPa.10 This is important, because tissue compliance is required in order for the muscle to shorten up to 40% during contraction.11 For muscle cells to be able to deform an engineered muscle tissue, the elastic modulus of the scaffold must be a primary consideration, and in fact, studies have shown that stiffness dramatically influences cell behavior.12,13 For example, myoblasts cultured directly on glass (E ∼70 GPa) fuse into myotubes, however, while positive for myosin heavy chain, they do not form sarcomeres (Figure 1(a) and (b)).12 In contrast, when a second layer of myotubes is grown on top of this first layer,
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FIGURE 1 | Myoblast fusion into myotubes is sensitive to the mechanical properties of the underlying substrate. (a) A schematic showing a side view of a 1st layer of myotubes on a glass substrate with actin filaments and a 2nd layer of striated myotubes with well-formed sarcomeres, growing on the 1st layer. (b) Immunofluorescent images of the 1st and 2nd layer myotubes where actin and myosin only show striations in the 2nd layer of myotubes. The nucleus (asterisk) is only visible in the 1st layer myotube, confirming that there are two, distinct layers of myotubes. (c) Myotubes differentiated for 4 weeks on polyacrylamide hydrogels with varying elastic moduli of 1, 8, 11, and 17 kPa only form striations and sarcomeres on substrates with intermediate elastic moduli of 8 and 11 kPa comparable with muscle tissue. (d and e) C2C12 myoblasts differentiated into myotubes on polydimethylsiloxane (PDMS) with elastic moduli of 5 kPa and 1.72 MPa showed different responses, with myotubes on the soft PDMS preferring to clump together versus forming parallel myotubes (green = myosin heavy chain, blue = nuclei). (f) Myotubes grown on stiffer PDMS substrates (130, 830, and 1.72 MPa) were significantly longer than myotubes grown on softer 5 and 50 kPa substrates (* indicates p < 0.0001, # indicates p < 0.001). Scale bars are (a and b) 10 μm; (c) 20 μm; (d and e) 200 μm. (a, b: Reprinted with permission from Ref 12. Copyright 2004 The Company of Biologists Ltd; c: Reprinted with permission from Ref 13. Copyright 2004 Rockefeller University Press; d–f: Reprinted with permission from Ref 14. Copyright 2012 Public Library of Science)
the second layer is able to form striated myotubes with well-organized sarcomeres. This implies that having a softer substrate with stiffness comparable with muscle tissue yields myotubes with more developed and organized sarcomeres. To rule out the possible role of cell–cell signaling in this effect, myotubes were cultured on polyacrylamide gel substrates with elastic moduli similar to muscle tissue (1–23 kPa) and on glass controls.13 Well-organized sarcomeres were observed in myotubes on substrates closest to the elastic modulus of skeletal muscle cells (8 and 11 kPa), but not the softer or harder surfaces (Figure 1(c)). While the force generated by these myotubes was not measured, more highly organized sarcomeres typically indicate more mature myotubes capable of higher force production than those with little to no sarcomere organization. Similar results have been found with 180
stem cells, where specification of mesenchymal stem cells toward a myogenic lineage was enhanced on substrates with an elastic modulus of ∼10 kPa.15 Thus, when engineering skeletal muscle, the literature would suggest that optimal scaffold stiffness on the order of ∼10 kPa is necessary to maximize differentiation into functional muscle. However, other data suggests that a wider range of scaffold mechanical properties may benefit additional aspects of myotube maturation. For example, myotubes cultured on polydimethylsiloxane (PDMS) substrates with elastic moduli ranging from 5 kPa (Figure 1(d)) to 1.7 MPa (Figure 1(e)) were found to be longer on the stiffer formulations (Figure 1(f)), indicating more myoblasts were fused into each myotube.14 Longer myotubes in vitro are often larger in cross-sectional area with more sarcomeres, suggesting they will generate greater force. Thus,
© 2013 Wiley Periodicals, Inc.
Volume 6, March/April 2014
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Engineered skeletal muscle tissue for soft robotics
while the ideal mechanical properties to engineer skeletal muscle are still under research, what the data does demonstrate is that myotube fusion index, length, and sarcomere organization can be controlled by modulating the mechanical properties of the scaffold.
Extracellular Matrix and the Spatial Organization of Biochemical Cues The composition and structure of the ECM in vivo are key regulators of cell behavior, and are equally important factors when engineering skeletal muscle constructs in vitro. Recreating the complex 3D architecture of the ECM within muscle tissue is beyond current technology, but scaffolds composed of the ECM proteins collagen type I (COLI), collagen type IV (COL4) and laminin (LAM) can still be engineered to control myogenesis. For example, Matrigel™ (primarily LAM and COL4) mixed with varying amounts of COLI and fibrin were cast as isotropic gels and used to engineer 3D skeletal muscle from C2C12 myoblasts or adult primary rat skeletal myoblasts.16 In these cases the embedded cells remodel the ECM hydrogels over time and compact it into a cell-dense tissue construct. The C2C12 myoblasts compacted high fibrin content hydrogels more than those with high collagen content, indicating the rate of remodeling is ECM dependent. Furthermore, neonatal primary rat skeletal myoblasts grown in ECM hydrogels comprised of varying ratios of Matrigel™, fibrinogen, and/or collagen I, showed that the hydrogel composition can have a larger effect on contractility and peak stress generation than other factors such as electrical stimulation, coculture with other cell types, or added growth factors.17 These examples demonstrate that the interaction of cells with the extracellular environment plays a key role in skeletal muscle cell differentiation and that ECM protein composition is an important factor. The spatial organization (i.e., patterning) of ECM proteins can be used to control skeletal muscle formation and direct myotube organization. Most patterning techniques are optimized for 2D, where photolithography-based approaches can be used to define microscale and nanoscale features on the surface. For example, geometrically patterning ECM proteins by microcontact printing (μCP) has been used to guide cell attachment and probe the influence of spatial organization of chemical cues on cell behavior (see here for a general review of μCP18 ). The μCP of FN and/or LAM has been used to engineer all main muscle types including smooth,19,20 cardiac,21,22 and skeletal.23,24 For skeletal muscle, one of the main uses of μCP is the engineering of Volume 6, March/April 2014
uniaxially aligned myotubes using patterned ECM lines to better mimic myotube organization in vivo. Researchers have shown that increasing the width of LAM lines from 100 to 300 to 500 μm (Figure 2(a) and (b)) decreased C2C12 proliferation but increased differentiation into myotubes (Figure 2(c) and 2(d)).23 Parallel FN lines an order-of-magnitude narrower (10, 20, and 50 μm wide) are also effective at aligning C2C12 myoblasts and controlling orientation upon differentiation into myotubes.24 Furthermore, both line spacing and line width are important factors. For example, on uniform FN surfaces myotubes are isotropic (Figure 2(e)), but when patterned on 50 μm wide, 10 μm spaced (50 × 10) lines the myotubes become aligned, but at approximately a 30◦ angle to the underlying pattern25 (Figure 2(f)). Increasing the line spacing to 20 μm keeps the myotubes on the 50 μm wide FN lines and creates uniaxially aligned muscle tissue, but the overall myotube density on the surface is decreased (Figure 2(g)). Increasing the line width to 200 μm increases the myotube density on the surface, but the myotubes align at an approximately 10◦ angle to the underlying pattern (Figure 2(h)). What this demonstrates is that μCP of ECM proteins is a useful tool to control and probe myoblast proliferation, differentiation, and alignment at the cellular level. Future investigations need to examine differences in myoblast or myotube behavior among varying ECM proteins in addition to the influence of geometric cues. By doing this, we will have a better understanding of how to control the microenvironment when engineering skeletal muscle actuators for soft robotic devices.
Physical and Topographical Cues that Guide Muscle Alignment Physical and topographical features within the cell microenvironment are able to guide myoblasts and direct their fusion into aligned myotubes. Termed contact guidance, this process has been studied in many cell types with similar results.26 Grooves (i.e., channels) and fibers on the nanometer to micrometer scale are able to direct uniaxial alignment of skeletal muscle and are similar to the diameter of many of the protein filaments within the native muscle ECM, suggesting that cells are intrinsically sensitive to physical features in this size range. Typical microtopographies include parallel microgrooves, posts, and holes in substrates composed of polymers such as PDMS, polystyrene, gelatin methacrylate (gelMA), and poly(l-lactic acid) (PLA).27–31 Groove depth, width, and spacing (Figure 3(a)) can range from hundreds of nanometers30 to tens of micrometers.31
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FIGURE 3 | Myotube alignment can be controlled using engineered (g)
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FIGURE 2 | Micropatterned extracellular matrix (ECM) proteins guide myoblast differentiation and alignment. Examples of microcontact printing (μCP) lines of laminin (LAM) immunofluorescently labeled with widths of (a) 100 μm and (b) 300 μm. Phase-contrast images of myoblasts seeded onto (c) the 100 μm and (d) the 300 μm wide lines, demonstrating cell confinement to the pattern. (e) C2C12 myoblasts seeded onto a polydimethylsiloxane (PDMS) surface with a uniform FN coating differentiate into myotubes (myosin heavy chain = red) with isotropic (random) alignment. (f) The μCP of FN lines 50 μm wide and 10 μm spaced (50 × 10) induces myotube alignment, but not in the direction of the underlying pattern because the myotubes can bridge the spacing (pattern runs left to right). (g) Increasing the line spacing to 20 μm (50 × 20) keeps myotubes on the FN lines and forms a myotubes with uniaxial alignment but fewer overall myotubes. (h) Increasing the FN line width to 200 μm increase the number of myotubes, but decrease alignment. Scale bars are (a–d) 100 μm and (e–h) 200 μm. (a–d: Reprinted with permission from Ref 23. Copyright 2012 American Chemical Society; e–h: Reprinted with permission from Ref 25. Copyright 2013 Elsevier)
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substrate microtopography. (a) Schematic of microgrooves (i.e., channels) in a surface with controlled groove width, depth, and spacing that can induce the alignment of myotubes differentiated on the surface. (b) DIC image of myotubes grown on a surface with parallel microgrooves. (c) Myotubes differentiated for 10 days on a flat surface and stained for Troponin I (red) and DAPI (blue) are isotropic with no orientation preference. (d) In contrast, myotubes grown on the poly(L-lactic acid) (PLA) membrane patterned with rectangular holes induced uniaxial myotube alignment. Scale bars are (b) 20 μm and (c and d) 50 μm. (a: Reprinted with permission from Ref 31. Copyright 2012 Mary Ann Liebert, Inc.; b: Reprinted with permission from Ref 29. Copyright 2009 Elsevier; c–d: Reprinted with permission from Ref 27. Copyright 2009 Mary Ann Liebert, Inc.)
In general, myotubes organize in parallel to the groove direction (Figure 3(b)), which is enhanced with increasing groove depth to width ratio and is less sensitive to the groove width itself.28,30 Parallel grooves are also more effective at aligning myotubes than square posts,29 presumably because of the increased contact area between the myotube and the topographical feature. Compared with flat controls (Figure 3(c)), microtopographies are effective at guiding myotube alignment (Figure 3(d)), with comparable results to micropatterned ECM protein lines.27 Nanometer and micrometer scale fibers, when organized into aligned scaffolds, are also effective
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at guiding myotube formation and alignment. Electrospinning is the most common technique to create fibers at this scale, which uses an electric field applied between an extrusion needle and a collector to draw out the polymer fibers. Electrospinning techniques have been optimized to give control over fiber alignment and diameter. For example, uniaxially aligned polypropylene fibers coated with LAM were used to form aligned sheets of C2C12 myoblasts,32 with a fiber spacing of 55 μm or less found to be optimal to form continuous myoblast sheets. Fibers at the nanometer scale are also effective at guiding muscle alignment. For example, electrospun polycaprolactone and polyaniline nanofibers have been used to direct the differentiation of C2C12 myoblasts into uniaxially aligned myotubes.33 Other fiber spinning techniques such as spinneret-based tuneable engineered parameter (STEP) polystyrene nanofibers 668 ± 63 nm in diameter were used similarly to direct the differentiation of C2C12 myoblasts into uniaxially aligned myotubes.34 In total, these examples demonstrate that nanometer and micrometer scale topographic features can be used to direct the uniaxial alignment of myotubes. However, more research is necessary to determine whether there is a specific size or morphology that is optimal. For soft robotics, a key limitation is that these techniques are primarily 2D, limiting the thickness of the engineered muscle tissue to < 100 μm.
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Once muscle tissue forms, it needs to actively contract and work against a load in order to continue to mature. For this reason many researchers are engineering bioreactors that incorporate electrical stimulation and/or mechanical loading in order to engineer improved skeletal muscle constructs. Field stimulation is used to excite skeletal muscle constructs in vitro without requiring innervation35 and is effective for conditioning and testing force generation capabilities. Typically, electrodes (platinum or carbon) are placed on either side of the muscle tissue construct during culture (Figure 4(a)) and a square wave in the range of 0 to 100 V in amplitude, 10–20 milliseconds in duration and up to 100 Hz or greater is applied. The electrical stimulation depolarizes the myotubes causing contraction and mechanical stress that has been shown to increase differentiation, alignment, and force generation (as reviewed in Ref 39). For example, electrical pulse stimulation (EPS) of C2C12 myotubes at 1 Hz increased sarcomere assembly Volume 6, March/April 2014
2.5 s
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FIGURE 4 | Examples of different bioreactor designs and microfabricated electrodes for the electrical stimulation of engineered skeletal muscle constructs. (a) A bioreactor based on a modified six-well plate with platinum wire electrodes integrated into the lid capable of stimulating cultures of 3D engineered skeletal muscle tissue. (b) A schematic of contactless electrodes showing a top view of the design (black), which are positioned underneath a glass coverslip to isolated them from the cells and media. Myoblasts (yellow) cultured on top of microtopographically patterned gelMA (green) uniaxially align during culture. (c) Micropatterned lines of C2C12 myotubes can be selectively stimulated using microfabricated platinum electrode arrays. (d) Plots of displacement of the myotubes strands in (c) demonstrating the ability to independently stimulate the closely spaced muscle tissues (amplitude, 2 V; frequency, 1 Hz; duration, 10 milliseconds). (a: Reprinted with permission from Ref 36. Copyright 2010 Mary Ann Liebert, Inc.; b Reprinted with permission from Ref 37. Copyright 2012 The Royal Society of Chemistry; c–d: Reprinted with permission from Ref 38. Copyright 2011 The Royal Society of Chemistry)
within as little as 2 hours.40 Similarly, EPS of C2C12 myotubes on deformable collagen strips resulted in significantly higher force generation compared with no EPS controls.41 In addition to more organized sarcomeres, electrically stimulating C2C12 myotubes resulted in increased glucose uptake.28
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Researchers have also tried replicating the electrical stimulation patterns experienced by muscle cells during development in order to produce more mature myotubes (Figure 4(a)).36 The peak stress generated by these constructs was 60–120 Pa, considerably less than the >100 kPa of native skeletal muscle.42 However, these constructs did show significantly higher force generation and increased excitability compared with constructs that were not stimulated or conditioned with a suboptimal stimulation pattern.42 A limitation of field stimulation is that continuously exciting cell cultures with electrodes (platinum or carbon) can cause electrolysis of the media, leading to heating, changes in pH, and electrode corrosion. Contactless electrodes are one potential solution, and recent work has demonstrated excitation of C2C12 myoblasts was possible without causing electrolysis of the media or other adverse effects (Figure 4(b)).43 Electrodes have also been miniaturized to stimulate small, micropatterned lines of myotubes in order to selectively actuate a subset of cells, similar to innervated motor units (Figure 4(c)).38 With this type of approach it is possible to control stimulation of individual myotube strands separated by as few as 200 μm (Figure 4(d)). Electrically conductive scaffolds have also been explored as a means to continuously stimulate cardiac constructs44–46 as well as skeletal muscle.47 These electrically conductive scaffolds are another potential solution to electrical stimulation of skeletal muscle in vitro. Mechanical stress can also be created by stretching skeletal muscle constructs using an externally applied load, and has also resulted in increased force generation.48 For example, cyclical, mechanical strain of human bioartificial muscles (BAMs) consisting of collagen I, Matrigel™, and human primary skeletal muscle cells resulted in a significant increase in the myofiber diameter and relative cross-sectional area compared with controls.49 Similar studies have investigated the electrical and mechanical stimulation of engineered cardiac muscle constructs and shown significant improvement to maturation of the sarcomere structure and increased force generation.50 Thus, for soft robotics, control of electrical simulation is important both in terms of the maturation of skeletal muscle constructs during culture as well as for the modulation of contraction force and temporal actuation.
ENGINEERED SKELETAL MUSCLE FOR SOFT ROBOTIC ACTUATORS The approaches presented thus far have focused on guiding the formation of aligned skeletal muscle tissue and working toward maturing its electromechanical 184
properties. For soft robotics, equally important is engineering a skeletal muscle construct in a physical format that is adaptable as a component within soft robotic devices, or is the soft robotic device itself. While primarily investigated in the context of tissue engineering, a number of advances have been made that enable the engineering of 2D and 3D engineered skeletal muscle as functional tissue suitable for use as actuators and can be further modified for specific applications.
2D Cantilever-Based Skeletal Muscle Actuators Skeletal muscle is a scalable actuator system that can potentially be used for a wide range of applications. Where relatively low force is required, 2D cantileverbased actuators are a good choice because the muscle can be integrated as a thin layer (
Engineered skeletal muscle tissue for soft robotics: fabrication strategies, current applications, and future challenges Rebecca M. Duffy1 and Adam W. Feinberg1,2∗ Skeletal muscle is a scalable actuator system used throughout nature from the millimeter to meter length scales and over a wide range of frequencies and force regimes. This adaptability has spurred interest in using engineered skeletal muscle to power soft robotics devices and in biotechnology and medical applications. However, the challenges to doing this are similar to those facing the tissue engineering and regenerative medicine fields; specifically, how do we translate our understanding of myogenesis in vivo to the engineering of muscle constructs in vitro to achieve functional integration with devices. To do this researchers are developing a number of ways to engineer the cellular microenvironment to guide skeletal muscle tissue formation. This includes understanding the role of substrate stiffness and the mechanical environment, engineering the spatial organization of biochemical and physical cues to guide muscle alignment, and developing bioreactors for mechanical and electrical conditioning. Examples of engineered skeletal muscle that can potentially be used in soft robotics include 2D cantileverbased skeletal muscle actuators and 3D skeletal muscle tissues engineered using scaffolds or directed self-organization. Integration into devices has led to basic muscle-powered devices such as grippers and pumps as well as more sophisticated muscle-powered soft robots that walk and swim. Looking forward, current, and future challenges include identifying the best source of muscle precursor cells to expand and differentiate into myotubes, replacing cardiomyocytes with skeletal muscle tissue as the bio-actuator of choice for soft robots, and vascularization and innervation to enable control and nourishment of larger muscle tissue constructs. © 2013 Wiley Periodicals, Inc.
How to cite this article:
WIREs Nanomed Nanobiotechnol 2014, 6:178–195. doi: 10.1002/wnan.1254
INTRODUCTION
S
keletal muscle serves as the primary actuator system in animals and can perform tasks over a wide range of length and time scales, spatial configurations, and force regimes. The scalability ∗
Correspondence to: [email protected]
1
Department of Biomedical Engineering, University, Pittsburgh, PA, USA
Carnegie
Mellon
2
Department of Materials Science and Engineering, Carnegie Mellon University, Pittsburgh, PA, USA Conflict of interest: The authors have declared no conflicts of interest for this article.
178
of skeletal muscle makes it a good candidate for soft robotics and related applications because it achieves a unique set of performance metrics. Specifically, skeletal muscle offers several advantages over traditional mechanical actuators including superior power-to-weight ratio, force-to-weight ratio, mechanical compliance, and systems control.1 For example, selective activation of muscle fibers via innervation and spatial or temporal summation allows dynamic control and modulation over force generation.2 Compared with traditional, rigid robots, soft robotics can have greater degrees of freedom and can be modeled after specific examples in nature,
© 2013 Wiley Periodicals, Inc.
Volume 6, March/April 2014
WIREs Nanomedicine and Nanobiotechnology
Engineered skeletal muscle tissue for soft robotics
such as the elephant trunk or octopus arm.3 This flexibility in configuration and force provides the potential for the same system to perform diverse functions from high-precision tasks such as wound suturing to high-strength tasks such as weight lifting. In addition, the energy source for skeletal muscle is adenosine triphosphate (ATP), generated from the energy stored within chemical bonds in amino acids, polysaccharides, and fatty acids and with approximately two orders-of-magnitude higher energy density than conventional energy sources used by nonbiological machines. Lastly, another advantage of biological actuators over man-made actuators is the potential ability to self-repair. At the molecular scale, the protein machinery that makes up the contractile apparatus is constantly being replaced while the muscle is functioning, enabling contraction for decades through billions of cycles. At the tissue scale, skeletal muscle in vivo can repair small injuries, preventing loss of muscle mass.4 While current tissue-engineered muscle constructs cannot heal at the tissue scale, enabling this ability to self-repair could revolutionize the field of robotics by creating damage-tolerant systems, and directly coupled to repair is adaptive growth in response to moderate mechanical overloading. Similar to exercise-induced muscle growth in vivo, engineered muscle constructs may eventually be able to grow and become stronger when the load requires more power, or even atrophy and reduce energy requirements when the load is decreased. Engineering skeletal muscle for use in soft robotics requires the ability to guide muscle precursor cells to differentiate into skeletal muscle cells (myotubes) and then to organize these cells into a functional tissue. This muscle formation process (termed myogenesis) is well documented in vivo,4–6 and has been achieved at a basic level in vitro in terms of myoblast fusion into myotubes. However, in vitro it has proven difficult to engineer the formation of dense, innervated, and vascularized muscle in 3D at the tissue scale, a challenge that also faces the field of tissue engineering and regenerative medicine. Advances in engineered skeletal muscle for tissue repair and as in vitro systems for drug discovery have been critical to our growing understanding of how to direct myogenesis in vitro.7–9 Here we will focus on current research to elucidate the physical, mechanical, and chemical cues necessary to guide myogenesis and how to recapitulate them. Initial results demonstrate that the extracellular matrix (ECM) is a key component to the myogenesis process and can serve as an instructive template for engineering skeletal muscle. Major factors include (1) substrate stiffness Volume 6, March/April 2014
and the mechanical environment, (2) the spatial organization of biochemical cues, (3) physical and topographical cues, and (4) electrical and mechanical conditioning. Applied to the engineering of skeletal muscle, controlling some of these parameters has enabled the fabrication of 2D and 3D constructs suitable as actuators in soft robotics. Furthermore, researchers have integrated these engineered muscle tissues into various devices from simple grippers to semi-autonomous walking and swimming soft robots. These proof-of-concept examples highlight the unique capabilities of engineered muscle and the fact that current technology can start to leverage muscle as an actuator system in engineered devices. However, challenges remain and still need to be addressed before engineered muscle is used more widely. Tissue size is limited by the lack of a vascular system for nutrient transport, long-term viability in a nonsterile environment is poor because of lack of an immune system to fight infection, and long-term stability of the muscle is inadequate as the cells slowly die due to removal from the in vivo environment. Looking forward, research in skeletal muscle tissue engineering and regenerative medicine coupled with the growing field of muscle-based soft robotics suggests that many of these problems will be overcome and that wide spread use of muscle as an actuator in a range of biotechnologies from soft robotics, to bioprosthetics to bio-microelectromechanical systems (bio-MEMS) is rapidly approaching.
ENGINEERING THE MICROENVIRONMENT TO GUIDE SKELETAL MUSCLE FORMATION The Role of Substrate Stiffness and the Mechanical Environment Skeletal muscle is a soft tissue composed of myotubes, blood vessels, nerves, and an interconnected network of ECM. The elastic modulus of muscle tissue depends on the specific type, but is reported to be in the range of 10–40 kPa.10 This is important, because tissue compliance is required in order for the muscle to shorten up to 40% during contraction.11 For muscle cells to be able to deform an engineered muscle tissue, the elastic modulus of the scaffold must be a primary consideration, and in fact, studies have shown that stiffness dramatically influences cell behavior.12,13 For example, myoblasts cultured directly on glass (E ∼70 GPa) fuse into myotubes, however, while positive for myosin heavy chain, they do not form sarcomeres (Figure 1(a) and (b)).12 In contrast, when a second layer of myotubes is grown on top of this first layer,
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FIGURE 1 | Myoblast fusion into myotubes is sensitive to the mechanical properties of the underlying substrate. (a) A schematic showing a side view of a 1st layer of myotubes on a glass substrate with actin filaments and a 2nd layer of striated myotubes with well-formed sarcomeres, growing on the 1st layer. (b) Immunofluorescent images of the 1st and 2nd layer myotubes where actin and myosin only show striations in the 2nd layer of myotubes. The nucleus (asterisk) is only visible in the 1st layer myotube, confirming that there are two, distinct layers of myotubes. (c) Myotubes differentiated for 4 weeks on polyacrylamide hydrogels with varying elastic moduli of 1, 8, 11, and 17 kPa only form striations and sarcomeres on substrates with intermediate elastic moduli of 8 and 11 kPa comparable with muscle tissue. (d and e) C2C12 myoblasts differentiated into myotubes on polydimethylsiloxane (PDMS) with elastic moduli of 5 kPa and 1.72 MPa showed different responses, with myotubes on the soft PDMS preferring to clump together versus forming parallel myotubes (green = myosin heavy chain, blue = nuclei). (f) Myotubes grown on stiffer PDMS substrates (130, 830, and 1.72 MPa) were significantly longer than myotubes grown on softer 5 and 50 kPa substrates (* indicates p < 0.0001, # indicates p < 0.001). Scale bars are (a and b) 10 μm; (c) 20 μm; (d and e) 200 μm. (a, b: Reprinted with permission from Ref 12. Copyright 2004 The Company of Biologists Ltd; c: Reprinted with permission from Ref 13. Copyright 2004 Rockefeller University Press; d–f: Reprinted with permission from Ref 14. Copyright 2012 Public Library of Science)
the second layer is able to form striated myotubes with well-organized sarcomeres. This implies that having a softer substrate with stiffness comparable with muscle tissue yields myotubes with more developed and organized sarcomeres. To rule out the possible role of cell–cell signaling in this effect, myotubes were cultured on polyacrylamide gel substrates with elastic moduli similar to muscle tissue (1–23 kPa) and on glass controls.13 Well-organized sarcomeres were observed in myotubes on substrates closest to the elastic modulus of skeletal muscle cells (8 and 11 kPa), but not the softer or harder surfaces (Figure 1(c)). While the force generated by these myotubes was not measured, more highly organized sarcomeres typically indicate more mature myotubes capable of higher force production than those with little to no sarcomere organization. Similar results have been found with 180
stem cells, where specification of mesenchymal stem cells toward a myogenic lineage was enhanced on substrates with an elastic modulus of ∼10 kPa.15 Thus, when engineering skeletal muscle, the literature would suggest that optimal scaffold stiffness on the order of ∼10 kPa is necessary to maximize differentiation into functional muscle. However, other data suggests that a wider range of scaffold mechanical properties may benefit additional aspects of myotube maturation. For example, myotubes cultured on polydimethylsiloxane (PDMS) substrates with elastic moduli ranging from 5 kPa (Figure 1(d)) to 1.7 MPa (Figure 1(e)) were found to be longer on the stiffer formulations (Figure 1(f)), indicating more myoblasts were fused into each myotube.14 Longer myotubes in vitro are often larger in cross-sectional area with more sarcomeres, suggesting they will generate greater force. Thus,
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while the ideal mechanical properties to engineer skeletal muscle are still under research, what the data does demonstrate is that myotube fusion index, length, and sarcomere organization can be controlled by modulating the mechanical properties of the scaffold.
Extracellular Matrix and the Spatial Organization of Biochemical Cues The composition and structure of the ECM in vivo are key regulators of cell behavior, and are equally important factors when engineering skeletal muscle constructs in vitro. Recreating the complex 3D architecture of the ECM within muscle tissue is beyond current technology, but scaffolds composed of the ECM proteins collagen type I (COLI), collagen type IV (COL4) and laminin (LAM) can still be engineered to control myogenesis. For example, Matrigel™ (primarily LAM and COL4) mixed with varying amounts of COLI and fibrin were cast as isotropic gels and used to engineer 3D skeletal muscle from C2C12 myoblasts or adult primary rat skeletal myoblasts.16 In these cases the embedded cells remodel the ECM hydrogels over time and compact it into a cell-dense tissue construct. The C2C12 myoblasts compacted high fibrin content hydrogels more than those with high collagen content, indicating the rate of remodeling is ECM dependent. Furthermore, neonatal primary rat skeletal myoblasts grown in ECM hydrogels comprised of varying ratios of Matrigel™, fibrinogen, and/or collagen I, showed that the hydrogel composition can have a larger effect on contractility and peak stress generation than other factors such as electrical stimulation, coculture with other cell types, or added growth factors.17 These examples demonstrate that the interaction of cells with the extracellular environment plays a key role in skeletal muscle cell differentiation and that ECM protein composition is an important factor. The spatial organization (i.e., patterning) of ECM proteins can be used to control skeletal muscle formation and direct myotube organization. Most patterning techniques are optimized for 2D, where photolithography-based approaches can be used to define microscale and nanoscale features on the surface. For example, geometrically patterning ECM proteins by microcontact printing (μCP) has been used to guide cell attachment and probe the influence of spatial organization of chemical cues on cell behavior (see here for a general review of μCP18 ). The μCP of FN and/or LAM has been used to engineer all main muscle types including smooth,19,20 cardiac,21,22 and skeletal.23,24 For skeletal muscle, one of the main uses of μCP is the engineering of Volume 6, March/April 2014
uniaxially aligned myotubes using patterned ECM lines to better mimic myotube organization in vivo. Researchers have shown that increasing the width of LAM lines from 100 to 300 to 500 μm (Figure 2(a) and (b)) decreased C2C12 proliferation but increased differentiation into myotubes (Figure 2(c) and 2(d)).23 Parallel FN lines an order-of-magnitude narrower (10, 20, and 50 μm wide) are also effective at aligning C2C12 myoblasts and controlling orientation upon differentiation into myotubes.24 Furthermore, both line spacing and line width are important factors. For example, on uniform FN surfaces myotubes are isotropic (Figure 2(e)), but when patterned on 50 μm wide, 10 μm spaced (50 × 10) lines the myotubes become aligned, but at approximately a 30◦ angle to the underlying pattern25 (Figure 2(f)). Increasing the line spacing to 20 μm keeps the myotubes on the 50 μm wide FN lines and creates uniaxially aligned muscle tissue, but the overall myotube density on the surface is decreased (Figure 2(g)). Increasing the line width to 200 μm increases the myotube density on the surface, but the myotubes align at an approximately 10◦ angle to the underlying pattern (Figure 2(h)). What this demonstrates is that μCP of ECM proteins is a useful tool to control and probe myoblast proliferation, differentiation, and alignment at the cellular level. Future investigations need to examine differences in myoblast or myotube behavior among varying ECM proteins in addition to the influence of geometric cues. By doing this, we will have a better understanding of how to control the microenvironment when engineering skeletal muscle actuators for soft robotic devices.
Physical and Topographical Cues that Guide Muscle Alignment Physical and topographical features within the cell microenvironment are able to guide myoblasts and direct their fusion into aligned myotubes. Termed contact guidance, this process has been studied in many cell types with similar results.26 Grooves (i.e., channels) and fibers on the nanometer to micrometer scale are able to direct uniaxial alignment of skeletal muscle and are similar to the diameter of many of the protein filaments within the native muscle ECM, suggesting that cells are intrinsically sensitive to physical features in this size range. Typical microtopographies include parallel microgrooves, posts, and holes in substrates composed of polymers such as PDMS, polystyrene, gelatin methacrylate (gelMA), and poly(l-lactic acid) (PLA).27–31 Groove depth, width, and spacing (Figure 3(a)) can range from hundreds of nanometers30 to tens of micrometers.31
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FIGURE 2 | Micropatterned extracellular matrix (ECM) proteins guide myoblast differentiation and alignment. Examples of microcontact printing (μCP) lines of laminin (LAM) immunofluorescently labeled with widths of (a) 100 μm and (b) 300 μm. Phase-contrast images of myoblasts seeded onto (c) the 100 μm and (d) the 300 μm wide lines, demonstrating cell confinement to the pattern. (e) C2C12 myoblasts seeded onto a polydimethylsiloxane (PDMS) surface with a uniform FN coating differentiate into myotubes (myosin heavy chain = red) with isotropic (random) alignment. (f) The μCP of FN lines 50 μm wide and 10 μm spaced (50 × 10) induces myotube alignment, but not in the direction of the underlying pattern because the myotubes can bridge the spacing (pattern runs left to right). (g) Increasing the line spacing to 20 μm (50 × 20) keeps myotubes on the FN lines and forms a myotubes with uniaxial alignment but fewer overall myotubes. (h) Increasing the FN line width to 200 μm increase the number of myotubes, but decrease alignment. Scale bars are (a–d) 100 μm and (e–h) 200 μm. (a–d: Reprinted with permission from Ref 23. Copyright 2012 American Chemical Society; e–h: Reprinted with permission from Ref 25. Copyright 2013 Elsevier)
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substrate microtopography. (a) Schematic of microgrooves (i.e., channels) in a surface with controlled groove width, depth, and spacing that can induce the alignment of myotubes differentiated on the surface. (b) DIC image of myotubes grown on a surface with parallel microgrooves. (c) Myotubes differentiated for 10 days on a flat surface and stained for Troponin I (red) and DAPI (blue) are isotropic with no orientation preference. (d) In contrast, myotubes grown on the poly(L-lactic acid) (PLA) membrane patterned with rectangular holes induced uniaxial myotube alignment. Scale bars are (b) 20 μm and (c and d) 50 μm. (a: Reprinted with permission from Ref 31. Copyright 2012 Mary Ann Liebert, Inc.; b: Reprinted with permission from Ref 29. Copyright 2009 Elsevier; c–d: Reprinted with permission from Ref 27. Copyright 2009 Mary Ann Liebert, Inc.)
In general, myotubes organize in parallel to the groove direction (Figure 3(b)), which is enhanced with increasing groove depth to width ratio and is less sensitive to the groove width itself.28,30 Parallel grooves are also more effective at aligning myotubes than square posts,29 presumably because of the increased contact area between the myotube and the topographical feature. Compared with flat controls (Figure 3(c)), microtopographies are effective at guiding myotube alignment (Figure 3(d)), with comparable results to micropatterned ECM protein lines.27 Nanometer and micrometer scale fibers, when organized into aligned scaffolds, are also effective
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at guiding myotube formation and alignment. Electrospinning is the most common technique to create fibers at this scale, which uses an electric field applied between an extrusion needle and a collector to draw out the polymer fibers. Electrospinning techniques have been optimized to give control over fiber alignment and diameter. For example, uniaxially aligned polypropylene fibers coated with LAM were used to form aligned sheets of C2C12 myoblasts,32 with a fiber spacing of 55 μm or less found to be optimal to form continuous myoblast sheets. Fibers at the nanometer scale are also effective at guiding muscle alignment. For example, electrospun polycaprolactone and polyaniline nanofibers have been used to direct the differentiation of C2C12 myoblasts into uniaxially aligned myotubes.33 Other fiber spinning techniques such as spinneret-based tuneable engineered parameter (STEP) polystyrene nanofibers 668 ± 63 nm in diameter were used similarly to direct the differentiation of C2C12 myoblasts into uniaxially aligned myotubes.34 In total, these examples demonstrate that nanometer and micrometer scale topographic features can be used to direct the uniaxial alignment of myotubes. However, more research is necessary to determine whether there is a specific size or morphology that is optimal. For soft robotics, a key limitation is that these techniques are primarily 2D, limiting the thickness of the engineered muscle tissue to < 100 μm.
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Once muscle tissue forms, it needs to actively contract and work against a load in order to continue to mature. For this reason many researchers are engineering bioreactors that incorporate electrical stimulation and/or mechanical loading in order to engineer improved skeletal muscle constructs. Field stimulation is used to excite skeletal muscle constructs in vitro without requiring innervation35 and is effective for conditioning and testing force generation capabilities. Typically, electrodes (platinum or carbon) are placed on either side of the muscle tissue construct during culture (Figure 4(a)) and a square wave in the range of 0 to 100 V in amplitude, 10–20 milliseconds in duration and up to 100 Hz or greater is applied. The electrical stimulation depolarizes the myotubes causing contraction and mechanical stress that has been shown to increase differentiation, alignment, and force generation (as reviewed in Ref 39). For example, electrical pulse stimulation (EPS) of C2C12 myotubes at 1 Hz increased sarcomere assembly Volume 6, March/April 2014
2.5 s
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FIGURE 4 | Examples of different bioreactor designs and microfabricated electrodes for the electrical stimulation of engineered skeletal muscle constructs. (a) A bioreactor based on a modified six-well plate with platinum wire electrodes integrated into the lid capable of stimulating cultures of 3D engineered skeletal muscle tissue. (b) A schematic of contactless electrodes showing a top view of the design (black), which are positioned underneath a glass coverslip to isolated them from the cells and media. Myoblasts (yellow) cultured on top of microtopographically patterned gelMA (green) uniaxially align during culture. (c) Micropatterned lines of C2C12 myotubes can be selectively stimulated using microfabricated platinum electrode arrays. (d) Plots of displacement of the myotubes strands in (c) demonstrating the ability to independently stimulate the closely spaced muscle tissues (amplitude, 2 V; frequency, 1 Hz; duration, 10 milliseconds). (a: Reprinted with permission from Ref 36. Copyright 2010 Mary Ann Liebert, Inc.; b Reprinted with permission from Ref 37. Copyright 2012 The Royal Society of Chemistry; c–d: Reprinted with permission from Ref 38. Copyright 2011 The Royal Society of Chemistry)
within as little as 2 hours.40 Similarly, EPS of C2C12 myotubes on deformable collagen strips resulted in significantly higher force generation compared with no EPS controls.41 In addition to more organized sarcomeres, electrically stimulating C2C12 myotubes resulted in increased glucose uptake.28
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Researchers have also tried replicating the electrical stimulation patterns experienced by muscle cells during development in order to produce more mature myotubes (Figure 4(a)).36 The peak stress generated by these constructs was 60–120 Pa, considerably less than the >100 kPa of native skeletal muscle.42 However, these constructs did show significantly higher force generation and increased excitability compared with constructs that were not stimulated or conditioned with a suboptimal stimulation pattern.42 A limitation of field stimulation is that continuously exciting cell cultures with electrodes (platinum or carbon) can cause electrolysis of the media, leading to heating, changes in pH, and electrode corrosion. Contactless electrodes are one potential solution, and recent work has demonstrated excitation of C2C12 myoblasts was possible without causing electrolysis of the media or other adverse effects (Figure 4(b)).43 Electrodes have also been miniaturized to stimulate small, micropatterned lines of myotubes in order to selectively actuate a subset of cells, similar to innervated motor units (Figure 4(c)).38 With this type of approach it is possible to control stimulation of individual myotube strands separated by as few as 200 μm (Figure 4(d)). Electrically conductive scaffolds have also been explored as a means to continuously stimulate cardiac constructs44–46 as well as skeletal muscle.47 These electrically conductive scaffolds are another potential solution to electrical stimulation of skeletal muscle in vitro. Mechanical stress can also be created by stretching skeletal muscle constructs using an externally applied load, and has also resulted in increased force generation.48 For example, cyclical, mechanical strain of human bioartificial muscles (BAMs) consisting of collagen I, Matrigel™, and human primary skeletal muscle cells resulted in a significant increase in the myofiber diameter and relative cross-sectional area compared with controls.49 Similar studies have investigated the electrical and mechanical stimulation of engineered cardiac muscle constructs and shown significant improvement to maturation of the sarcomere structure and increased force generation.50 Thus, for soft robotics, control of electrical simulation is important both in terms of the maturation of skeletal muscle constructs during culture as well as for the modulation of contraction force and temporal actuation.
ENGINEERED SKELETAL MUSCLE FOR SOFT ROBOTIC ACTUATORS The approaches presented thus far have focused on guiding the formation of aligned skeletal muscle tissue and working toward maturing its electromechanical 184
properties. For soft robotics, equally important is engineering a skeletal muscle construct in a physical format that is adaptable as a component within soft robotic devices, or is the soft robotic device itself. While primarily investigated in the context of tissue engineering, a number of advances have been made that enable the engineering of 2D and 3D engineered skeletal muscle as functional tissue suitable for use as actuators and can be further modified for specific applications.
2D Cantilever-Based Skeletal Muscle Actuators Skeletal muscle is a scalable actuator system that can potentially be used for a wide range of applications. Where relatively low force is required, 2D cantileverbased actuators are a good choice because the muscle can be integrated as a thin layer (
