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WIREs Nanomed Nanobiotechnol
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Engineered skeletal muscle tissue for soft robotics: fabrication strategies, current applications, and future challenges

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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 cantilever‐based 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.

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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 . Copyright 2012 American Chemical Society; e–h: Reprinted with permission from Ref . Copyright 2013 Elsevier)
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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 . Copyright 2004 The Company of Biologists Ltd; c: Reprinted with permission from Ref . Copyright 2004 Rockefeller University Press; d–f: Reprinted with permission from Ref . Copyright 2012 Public Library of Science)
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Examples of muscle‐powered soft robots that can walk and swim. (a) Schematic of a microfabricated walker driven by cardiac muscle. Cardiomyocytes are seeded on the Au film (yellow) and form an integrated, microscale muscle tissue that is subsequently released by dissolving the PIPAAm. This produces a free‐standing microdevice capable of walking when muscle cells are field stimulated to contract. (b) A schematic and corresponding microscopic images of a muscular thin film (MTF) with integrated cardiomyocytes formed into a construct termed a ‘myopod’. When field stimulated to contract the myopod is capable of walking (scale bars are 1 mm). (c) Top‐down and (d) side view schematics of an MTF with integrated cardiomyocytes in the form of a tissue‐engineered jellyfish termed a ‘medusoid’. The medusoid has a micropatterned muscle structure and body plan based on the actual jellyfish and when field stimulated to contract is capable of swimming. (e) A microscopic image of the medusoid floating in the Tyrode's solution in which it swims. (f) A top‐down schematic of a swimming robot that mimics a fish and is powered by explanted frog muscles that are sutured onto the device. Integrated batteries supply the electric charge to alternately stimulate the muscles on either side of the robot to contract, thus actuating the tail in a side‐to‐side motion and enabling it to swim. (a: Reprinted with permission from Ref . Copyright 2005 Macmillan Publishers Ltd; b: Reprinted with permission from Ref . Copyright 2007 AAAS; c–e: Reprinted with permission from Ref . Copyright 2012 Macmillan Publishers Ltd; f: Reprinted with permission from Ref . Copyright 2004 Herr and Dennis)
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Examples of basic muscle‐powered actuators and devices. (a) A skeletal muscle gel attached to a polydimethylsiloxane (PDMS) gripper (i.e., micro hand) that opens when the skeletal muscle is stimulated to contract. (b) Schematic of a muscle‐powered microfluidic pump where a sheet of cardiomyocytes are cultured on the outside of a thin, hollow chamber. (c) The actual device where spontaneous or stimulated contraction of the cardiomyocytes causes the chamber to decrease in volume and pump fluid. (d) Optogenetically engineered skeletal muscle microtissue suspended between two PDMS posts is stimulated with blue light (red circle) by depolarizing the cells and results in contraction of the entire construct (yellow arrows). (e) Selective stimulation of a small region of the construct (red circle) results in contraction of just that region of the construct (yellow arrows), demonstrating light‐based local control of activation. (a: Reprinted with permission from Ref . Copyright 2010 IEEE; b–c: Reprinted with permission from Ref . Copyright 2007 The Royal Society of Chemistry; d: Reprinted with permission from Ref . Copyright 2012 The Royal Society of Chemistry)
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Examples of different types of 3D engineered skeletal muscle tissue. (a) Miniature bioartificial muscle (mBAM) is formed by casting an extracellular matrix (ECM) hydrogel with embedded muscle cells into a specially designed, 7 mm diameter culture well with two, 700 µm diameter polydimethylsiloxane (PDMS) posts. (b) After 4–5 days of differentiation primary mouse myoblasts have begun to compact the gel around the posts. (c) mBAMs differentiated for 7–8 days were stained for sarcomeric tropomyosin (dark gray) showing aligned myotubes. (d) Contractile force generation was calculated from the bending of the PDMS posts during stimulation. (e) Sheets of engineered skeletal muscle consisting of a porous gel network were formed by casting ECM hydrogels with embedded muscle cells around a PDMS mold with rectangular posts. (f) The engineered muscle tissue network is attached to a Velcro frame and can be removed from the PDMS mold. (g) A top‐down view of the skeletal muscle tissue showing the rectangular posts that guide anisotropic organization of the ECM as it is compacted by the cells. (h) Immunofluorescent staining of the sarcomeric α‐actinin (green) and nuclei (red) of primary rat skeletal myoblasts differentiated into myotubes, demonstrating the uniaxial alignment and formation of sarcomeres. (i) Engineered skeletal muscle termed Myooids are formed by the self‐organization of muscle cells from a monolayer on the bottom of a Petri dish around two ECM‐coated anchoring structures (arrows indicate where the cell layer is rolling up). (j) The myooid construct once it has full organized into a linear muscle tissue in between the two sutures. (k) A cross‐section of a myooid showing myotubes in the center with a flattened layer of fibroblasts on the periphery (stained with toluidine blue). (l) A 10‐layer engineered muscle tissue created by stacking myoblasts sheets released from temperature‐responsive PIPAAm culture dishes. These are termed scaffold‐free because the cells make their own ECM and the alternating layers were labeled with red or green CellTracker for verification. (m) C2C12 myoblasts labeled with magnetite cationic liposomes were seeded around a silicone ring with a magnet placed underneath the culture dish to produce a scaffold‐free, multi‐layered muscle construct. Scale bars are (a) and (b) 4 mm, and otherwise as labeled. (a–d: Reprinted with permission from Ref . Copyright 2008 Wiley; e–h: Reprinted with permission from Ref . Copyright 2009 Elsevier; i–k: Reprinted with permission from Ref . Copyright 2000 Springer; (l) Reprinted with permission from Ref . Copyright 2012 Macmillan Publishers Ltd; (m) Reprinted with permission from Ref . Copyright 2009 Elsevier)
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Engineered skeletal muscle tissue in the format of 2D cantilever‐based actuators where the myotubes are integrated along the length of the beam. (a) An embryonic rat myotube differentiated on a DETA‐modified silicon cantilever (phase‐contrast image). (b) A C2C12 myotube on a vitronectin‐modified silicon cantilever (α‐actinin = green). Muscular thin films (MTFs) consisting of C2C12 myotubes of polydimethylsiloxane (PDMS) cantilevers showing a side view in (c) a relaxed state and (d) at peak contraction generating a twitch stress of ∼2 kPa. The yellow lines indicate the position of the tip of the muscular thin films (MTF) between the relaxed and full contracted states (scale bars are 1 mm). (a: Reprinted with permission from Ref . Copyright 2006 Elsevier; b: Reprinted with permission from Ref . Copyright 2004 The Royal Society of Chemistry; c–d: Reprinted with permission from Ref . Copyright 2013 Elsevier)
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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 . Copyright 2010 Mary Ann Liebert, Inc.; b Reprinted with permission from Ref . Copyright 2012 The Royal Society of Chemistry; c–d: Reprinted with permission from Ref . Copyright 2011 The Royal Society of Chemistry)
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Myotube alignment can be controlled using engineered 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 . Copyright 2012 Mary Ann Liebert, Inc.; b: Reprinted with permission from Ref . Copyright 2009 Elsevier; c–d: Reprinted with permission from Ref . Copyright 2009 Mary Ann Liebert, Inc.)
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Nanotechnology Approaches to Biology > Nanoscale Systems in Biology
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