3D microfibrous bundle structure fabricated using an electric-field

Article Cite This: ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

Three-Dimensional Microfibrous Bundle Structure Fabricated Using an Electric Field-Assisted/Cell Printing Process for Muscle Tissue Regeneration Miji Yeo and GeunHyung Kim* Department of Biomechatronic Engineering, College of Biotechnology and Bioengineering, Sungkyunkwan University, Suwon 16419, South Korea S Supporting Information *

ABSTRACT: In tissue engineering, biomimetic scaffolds are developed to provide cells with a microenvironment that promotes cellular activities. In this study, we present a three-dimensional (3D) fibrous bundle structure fabricated using an electrohydrodynamic process and a cell printing process using myoblast-laden collagen bioink. An anisotropic topographical cue in a 3D structure is an important factor for muscle tissue regeneration, and therefore, the fibrous bundle structure was uniaxially stretched using optimized conditions for fiber alignment. In addition, for stable cell attachment to facilitate the effect of topological cues, the myoblasts were efficiently released from the collagen bioink. We observed that the 3D fibrous bundle structure was an effective in vitro platform that induced cell proliferation and the formation of myotubes. The synergistic combination of the aligned topological cues and high biocompatibility of collagen enhanced the formation of myotubes, which was represented by the relative expression of myogenic genes (Myf5, Myh2, MyoD, and Myogenin). Therefore, we could confirm the feasibility of the 3D fibrous bundle structure for the regeneration of skeletal muscle tissues. KEYWORDS: skeletal muscle, cell-laden fibrous structure, topological cue, electrohydrodynamic process, tissue engineering

■

INTRODUCTION

Various technologies and biomaterials have been developed for building a 3D structure that is favorable for myoblast regeneration. The first trial of myoblast cultivation in a 3D environment was performed with collagen gel.9 The possibility of 3D cultivation was demonstrated; however, the collagen construct allowed myoblasts to form neonatal but not mature myofibers. For the mature differentiation stage, a structural support was required for the topological cues and physiological properties like electrical stimulation.10 Wang et al. provided a structural support to the myoblasts using electrospun nanofiber yarn and hydrogel coating.11 The myosin heavy chains (MHCs) covered the peripheral surface of the yarn, were highly aligned, and were elongated twice as much as those in a 2D culture. Furthermore, the hydrogel was cross-linked using ultraviolet (UV) light, which provided physical protection but not biocompatibility. In addition, the collagen fibrous scaffold was fabricated in the shape of a string using electrospinning in the aligned manner and then cross-linked with glutaraldehyde.12 The sarcomeric structures indicating mature differentiation were observed after 14 days of culture, but the scaffold was composed of a single string. To address the issue, we devised a scaffold composed of aligned fibrous bundles with

Tissue engineering has emerged as an approach for treating damaged tissues/organs by developing a biomimetic scaffold using living cells and biomaterials.1 During the construction of the microenvironment of the scaffolds, the conditions vary depending on the preference of the diverse cell types. For skeletal muscle regeneration, an aligned topological cue in the three-dimensional (3D) scaffold is an important factor. Anisotropic, densely packed myotubes are formed during skeletal muscle regeneration,2 and this aligned 3D structure enables a muscle to contract and generate force synergistically.3 As compared to a 2D environment, a 3D structure can induce more realistic cellular activities by increasing autocrine/ paracrine activities or proteins involved in mechanotransduction,4,5 which affects the performance of electrically stimulated tissues like nerves or muscles.6,7 Specifically, for the transduction of the stimulus in the muscle system, the expression of integrin-linked kinase and integrin β1 are enhanced in 3D culture, resulting in a more rapid response to mechanical stimulation than that obtained in a 2D culture.5 Furthermore, 2D scaffolds have a critical limitation that cannot serve as a standalone scaffold for volumetric muscle loss.8 It insists that realistic clinical applications using a 2D scaffold may be limited, and the development of a 3D scaffold for muscle regneneration is inevitable. © XXXX American Chemical Society

Received: December 14, 2017 Accepted: January 3, 2018 Published: January 3, 2018 A

DOI: 10.1021/acsbiomaterials.7b00983 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

Article

ACS Biomaterials Science & Engineering

eventually, cell attachment and proliferation benefit.23,24 As controls, we used randomly distributed fibrous bundles and aligned them without the collagen solution coating. Cell (C2C12)-laden collagen/poly(ethylene oxide) (PEO) was printed on the three scaffolds. For optimal printing conditions and cell-releasing behavior of the cell-laden collagen/PEO bioink to be realized, the initial cell viability, viscosity, and degradability of several of the collagen/PEO bioinks were examined. Finally, the optimally selected collagen/PEO bioink was printed for the three fibrous scaffolds, and the factors affecting the feasibility as a muscle tissue regenerative scaffold, such as myoblast alignment and differentiation, were observed using the expression levels of several myogenic genes.

hydrogel inducing myoblast proliferation and differentiation. Furthermore, the terminal stage of differentiation was represented by sarcomeric actin and myogenic gene expression. For a fibrous structure to be obtained, electrospinning has been a versatile tool that allows for controlling arrangements and structural and biochemical properties using synthetic/ natural polymers. For instance, the mixture of polyaniline (PANi) and gelatin were electrospun, and H9C2 cells were cultured.13 The cell morphology was random until 7 days of culture but then arranged in a confluent shape. Furthermore, myoblasts were cultured on protein-coated fibers in random/ aligned arrangements.14 Then, compared to the flat substrate or randomly oriented fibers, the aligned fibers showed the highest myogenic gene expression. The cellular activities on a twodimensional (2D) matrix are becoming clarified; however, the investigation into cellular activities on a 3D matrix is still challenging due to limited thickness or inhomogeneous distribution of cells. For these reasons, we suggest a technique to fabricate an aligned 3D structure with biochemical cues. Collagen, a main structural protein of the extracellular matrix (ECM), is widely used for supporting initial cell adhesion, migration, and proliferation.15 However, the effects of collagen toward myogenesis have not been clarified, and several factors such as chemical interactions or surface topography required elucidation for using collagen effectively for skeletal muscle regeneration.16 In tissue engineering, type I collagen has been known as a gold standard for inducing skeletal muscle regeneration by inducing cell adhesion, proliferation, and differentiation.17 Specifically, type I collagen induces cells to synthesize ECM by a similar structure to that of natural ECM and triggers myogenesis by the arginine-glycine-aspartic acid (RGD) peptide. The RGD peptide, present in fibronectin, plays an important role in myogenesis responding to complex, dynamic cellular activities.17 For this purpose, a technical approach is required to develop a biomimetic in vitro platform using collagen and to overcome challenges like homogeneous cell distribution in the 3D collagen structure.11 Cell printing is a technique that enables the placement of cells with bioink in the desired location homogeneously.18 Collagen is considered as an ideal bioink, and its versatility and effects on cellular activities are being explored. High viability should be achieved and degradability should also be considered for various cell activities like attachment, migration, and proliferation.18 As the concentration of bioink increases, the viability decreases owing to the high shear stress, and the degradability also decreases owing to the high degree of crosslinking sites.19 In contrast, although high viability is guaranteed, the low concentration of the bioink does not sustain its shape for long, and the cells in the bioink may not have enough time to adhere to a scaffold’s surface.20,21 In the present study, we propose a scaffold composed of 3D microfibrous bundles laden with a mixture of myoblasts and collagen. To build the microfibrous bundles, an electrohydrodynamic (EHD) jet process combined with wet electrospinning was used.22 The microfibrous bundle in which the fibers were randomly entangled was uniaxially stretched to obtain a fully aligned 3D structure. The aligned structure provided the myoblasts with physical support and aligned the cues affecting the myoblast alignment and differentiation. In addition, collagen (0.5 wt %) was coated on the uniaxially stretched fibrous bundles to provide more efficient cellular activities. Specifically, the collagen coating changes the surface hydrophilicity that enhances cell-surface interactions, and

■

MATERIALS AND METHODS

Materials. Polycaprolactone (PCL; Mn = 45,000; Sigma-Aldrich, St. Louis, MO) was dissolved in a methylene chloride (MC) and dimethylformamide (DMF) mixture in the ratio 4:1 and used for the EHD process. The ratio 4:1 was chosen because the increase of DMF volume fraction leads to the formation of beads.25 The bioink was fabricated with collagen type I (Matrixen-PSP; SKBioland, South Korea), PEO (Sigma-Aldrich, St. Louis, MO), and C2C12 myoblast cells (ATCC number CRL-1772, ATCC, Manassas, VA). Fabrication of Bioink and Rheological Measurements. For the cell-laden bioink, collagen (2 wt %) solution was mixed with PEO powder at 1, 2, and 3 wt % and added to C2C12 cells at a density of 1 × 107 cells/mL. A rotational rheometer (Bohlin Gemini HR Nano; Malvern Instruments, Surrey, UK) was used to test the storage modulus (G′) and complex viscosity (n*) of the fabricated bioink. The results were obtained using the setting of cone-and-plate geometries with a 4° cone angle, 40 mm diameter, and 150 μm gap. A dynamic frequency sweep (0.1−10 Hz) was conducted at 25 °C and 2% strain in the linear viscoelastic region. Fabrication of a 3D Fibrous Bundle Structure via the EHD Process. The electric fields of 8 kV and flow rates of 0.28 mL/h were used. For drawing fibrous bundle structures, a three-axis robot (DTR32210-T-SG, DASA Robot, South Korea) and wet electrospinning system were connected. The machine automatically moved through the designed path of a CAD model. It allowed for building a desired 3D structure composed of fibrous bundles that can serve as an interconnected, stand-alone structure. The nozzle (inner diameter (ID): 180 μm) was at 40 mm from the grounded copper filled with EtOH media and moved at a speed of 10 mm/s. EtOH was used for a target solution, and the mass transfer between EtOH and the solvents (MC and DMF) occurred until the concentrations of solvent reached equilibrium with the concentration of EtOH.22 In this process, the entangled fibrous structure was created with the appropriate conditions of mass exchange time, PCL supply, and the solvent concentration. The flow rate of the PCL solution was controlled using a syringe pump (KDS 230; KD Scientific, Holliston, MA, U.S.A.) and ejected using an electric field created with a power supply (SHV300RD-50K; Convertech, Seoul, South Korea). For the contact angle of the fibrous bundle structure to be observed, a droplet of 2 wt % bioink (40 μL) was placed on the surface. The droplet was captured using a digital camera (Canon EOS 450D; Canon, Japan) at 25 °C for 20 min. Uniaxial Stretching and Scaffold Characterization. The samples were stretched uniaxially with a stretching speed of 0.01 mm/s using a microtensile tester (Toptech 2000; Chemilab, Suwon, South Korea) in tensile mode. The surrounding temperature was increased to 45−50 °C using a radiant heater. The fiber orientations were characterized by different stretching percentages of 0, 20, 40, 60, 80, and 100%. After sputter coating with Au, the samples stretched by various stretching percentages were visualized with a scanning electron microscope (SEM; SNE-3000M; SEC Inc., Suwon, South Korea). For the degree of fibers to be measured, the horizontal axis was set to 0°. The angle range was set from −90° to 90°, which were the lower and upper perpendicular points to the horizontal axis, respectively. The B


Article

ACS Biomaterials Science & Engineering fibers were arbitrarily chosen to obtain 100 individual angles, and the measurements were represented in a distribution graph. The full width at half-maximum (fwhm) was calculated by the addition of absolute x values, which correspond to half of the maximum y value. Oxygen Plasma Treatment. Low frequency plasma (CUTE− MP/R, Femto-Science Inc., Korea) was used to treat the fibrous bundle structures with oxygen plasma. The fibrous bundle structures were put in a chamber and subjected to plasma at a low frequency of 50 Hz, pressure of 5.41 × 10−1 Torr, oxygen flow rate of 5 sccm (standard cubic centimeter per minute), and power of 30 W for 5 min. Cell Printing with C2C12-Laden Collagen Bioink and Cell Seeding. The bundle structures were each sanitized with 100% ethanol and distilled water three times. The cell-laden bioink (collagen/PEO/C2C12 cells) was printed on the fibrous bundle structures using a pressure of 100 kPa, a nozzle with a 150 μm inner diameter, and nozzle moving speed of 10 mm/s. After dispensing the cell-laden bioink, the samples were placed in an incubator at 37 °C for gelation.20 For cellular activities between cell printing and cell seeding to be compared, C2C12 cells were seeded at a density of 1 × 106 cells/mL on the scaffold measuring 10 × 15 × 0.1 mm3. The cell density was determined to adjust the number of cells between printing and seeding method per specimen. C2C12 Myoblast Cell Culture. Myoblasts were cultured using Dulbecco’s modified Eagle’s medium (DMEM) with high glucose (Sigma-Aldrich, St. Louis, MO) supplemented with 10% fetal bovine serum (FBS; Gemini Bio-Products, Sacramento, CA, USA) and 1% antibiotic (Antimycotic; Cellgro, Manassas, VA). The samples tested for MHC staining and real-time polymerase chain reaction (RT-PCR) were cultured in DMEM with high glucose supplemented with 2% horse serum and 1% antibiotic after 5 days of culture. The samples were incubated at 37 °C in 5% CO2, and the medium was changed every 2 days. In Vitro Myoblast Cell Responses. For cell viability and initial cell distribution, the live/dead assay kit (2 mM ethidium homodimer-1 and 0.15 mM calcein AM) was used to stain myoblasts after 4 h (in situ) and 1, 3, and 7 days of culture. The myoblasts were immersed into the solution of the live/dead assay and incubated for half an hour. The images captured using light microscopy (CKX41, Olympus) indicated the live cells in green and dead cells in red. The initial cell distribution was represented after 4 h of cell printing/seeding, and the initial cell viability was measured using ImageJ software from the ratio of live cells to the total number of cells. To estimate the cell seeding efficiency of the cell seeding process, we used MTT assay (Cell Proliferation Kit I, Boehringer Mannheim, Mannheim, Germany). The samples (n = 4) were incubated at 37 °C with 0.6 mg/mL of MTT. The absorbance was calibrated using a microplate reader (EL800, Bio-Tek Instruments, VT, USA) at 570 nm. The obtained data were normalized with the optical density value at day 1. We referred to the method of Sobral et al. for calculating cell seeding efficiency.26 Concisely, the seeded cells remained on the scaffolds for 4 h to have sufficient time for cell adhesion. Then, the scaffolds were removed from the well. The efficiency of the seeded scaffold was calculated by the cells on the scaffold and the residual cells on the well after 4 h.21 The following equation was used for seeding efficiency: seeding efficiency (%) = {[(cells added to scaffold) − (cells in wells)]/[(cells added to scaffold) + (cells in wells)]} × 100. PicoGreen assays (Molecular Probes) were performed with samples at 1, 3, and 7 days for calculating cell proliferation in DNA quantity. The samples were rinsed twice using PBS, and 300 μL of cell lysis buffer (Tris-EDTA (TE) buffer) composed of 0.2% Triton X-100, 10 mM Tris-HCl, and 10 mM EDTA were added. The samples were vortexed for every 5 min and stored at 4 °C, and this lysis process was performed for 30 min. The working solution (Quanti-iT PicoGreen reagent) was diluted to 1:200 in dimethyl sulfoxide (DMSO) solution and mixed with the lysed solution in the ratio of 1:1. A spectrofluorometer (Synergy H1; BioTek Instruments) was used to calculate the fluorescence of samples at 480 nm (excitation) and 520 nm (emission). The standard curve was obtained as suggested by the manufacturer’s protocol.

Analysis on in Vitro Cell Morphology and Differentiation. The samples were treated with 2.5% glutaraldehyde for 4 h after rinsing with PBS twice. For every 10 min, the various samples were washed with 50, 60, 70, 80, 90, and 100% ethanol, respectively. The samples were then blocked with hexamethyldisilazane and air-dried. A pedestal attached to the dried samples via double-sided carbon tape was sputter coated with Au. The surface of the cell-laden scaffolds was captured using SEM. For capturing the morphology of F-actin and nuclei, the samples were rinsed twice with PBS and treated with 3.8% paraformaldehyde for half an hour at room temperature. After the treatment of 0.1% Triton X-100 for 5 min, fluorescence staining was performed with 15 U/ml phalloidin (Invitrogen, Carlsbad, CA) conjugated to Alexa Fluor 568 and 5 μM diamidino-2-phenylindole (DAPI; Invitrogen, Carlsbad, CA). The images were captured using a fluorescence microscope (LSM700, Zeiss, Germany). For the immunofluorescence staining of MHC, the samples were washed with PBS twice and fixed using 2.5% glutaraldehyde at room temperature for 15 min. The samples were treated with Triton X-100 for half an hour after rinsing twice with PBS. Bovine serum albumin (BSA; 1%) was used to block nonspecific background for 1.5 h, and rabbit anti-MHC antibody (1:50 Santa Cruz) was treated at 4 °C overnight. The samples were dyed with 5 μM DAPI and Alexa Fluor 488-conjugated secondary antibody (1:500; Molecular Probes). The images were captured with the fluorescence microscope. Real-Time PCR. A real-time polymerase chain reaction (PCR) was conducted to measure the relative expression levels of Myf5, Myh2, MyoD, and Myogenin. For isolating the RNA from the samples, the samples were treated with TRI reagent (Sigma-Aldrich, St. Louis, MO) as per the instructions provided by the manufacturer. A spectrophotometer (Optizen Pop; K Lab, Daejeon, South Korea) was used to measure the purity and concentration of the RNA at 260 nm absorbance. The RNase-free DNase-treated total RNA (1 μg) was synthesized into cDNA. Taqman assays were added to the DNA (Table 1) and were treated with holding stage at 50 °C (2 min) and 95

Table 1. Taqman Myogenic Gene Markers commercial taqman probes gene

catalog number

β-actin Myf5 MyoD Myh2 Myogenin

Mm00607939_s1 Mm00435125_m1 Mm00440387_m1 Mm01332564_m1 Mm00446194_m1

company Thermo Thermo Thermo Thermo Thermo

Fisher Fisher Fisher Fisher Fisher

Science Science Science Science Science

°C (10 min). Then, polymerase denaturation at 95 °C (15 s) and annealing/extension at 60 °C (1 min) for 40 cycles were performed. The comparative threshold cycle (Ct) method was applied to analyze the data, and the gene markers were presented as a comparison with βactin and were normalized by the expression of cells on the R-scaffold. Statistical Analysis. At least three replicates were conducted for all experiments and expressed in the form of mean ± SD. Statistical analyses were performed using SPSS software (SPSS, Inc., Chicago, IL, USA), and the single-factor analysis of variance was used. A value of p < 0.05 was considered to indicate the statistical significance.

■

RESULTS AND DISCUSSION The EHD-printing, uniaxial-stretching, and cell-printing process for obtaining a random (R), aligned (A), and collagen-coated (C) scaffold are presented in Figure 1a−c, respectively. Recently, we developed the process of EHD printing with the assistance of a wet electrospinning process to realize 3D fibrous structures consisting of microsized poly(ε-caprolactone) (PCL) or cellulose fibers.27 By manipulating the height of ethanol in the target bath, surface tension of the bath media, flow rate of the polymeric solution, and various electric field conditions, the C


Article


Figure 1. Schematic of the fabrication of random (R), aligned (A), and (c) collagen-coated C-scaffolds with different processing conditions. (a) Rscaffold consisted of randomly oriented fibrous bundles in parallel structure fabricated via the EHD process. (b) A-scaffold consisted of aligned fibrous bundles fabricated via the EHD process and uniaxial stretching. (c) C-scaffold consisted of aligned, collagen-coated bundles fabricated via the EHD process, uniaxial stretching, oxygen plasma, and collagen coating. After all fabricating processes, the structures were printed with C2C12-laden bioink under the same conditions. (d) Schematic of native muscle scaffold and (e) optical/SEM images of the 3D fibrous bundle scaffold.

3D fibrous bundles can be obtained.22 To obtain the 3D PCL fibrous bundles, we used the various processing conditions shown in Figure 1a. Next, the C2C12-laden bioink (collagen/PEO) with a cell density of 1 × 107 cells/mL was printed on the fabricated random fibrous bundles, as shown in the Figure 1a, to obtain the cell-laden structure (random scaffold, R). Cell release is important as it determines the moment when the cells initially interact with the surface of the scaffold, and the cellular activities like proliferation and differentiation are influenced.17 Furthermore, the remnant collagen bioink may benefit cellular activities, so R- and A-scaffolds may provide both physical and biochemical cues. The microfibrous bundles, which consisted of randomly distributed microfibers in the SEM image of Figure 1c, were realized using the EHD process. For A-scaffold, the 3D microfibrous bundles were stretched in the uniaxial direction with a surrounding temperature of ∼45−50 °C and stretching speed of 0.01 mm/s (Figure 1b, c). As the PCL has a melting point of ∼60 °C, we set the stretching temperature at close to 50 °C to make the PCL fibrous bundle more easily elongated.28−30 In addition, a low strain rate (0.01 mm/s) was applied to reduce the breakage percentage of the microfibers. As shown in the schematic image of Figure 1b, the microfibers were arranged in the stretching direction, and their orientation was in an anisotropic manner compared with that of the nonstretched fibers. After the stretching of the microfibers, the cell-printing process was conducted with the same condition of the R-scaffold. To obtain the C-scaffold, all the procedures until the stretching process were completely same as that of the fabricating procedure of the A-scaffold; however, in addition, collagen (0.5 wt %) was coated on the stretched fibrous

bundles after the oxygen plasma treatment (Figure 1c). After the coating of collagen, the same cell printing process was applied. The final structures of the scaffolds were derived from the native muscle structure, which provides evidence of structural guidance for muscle tissue regeneration (Figure 1d). As myoblasts grow, they fuse to become a highly arranged, cylindrical structure named muscle fiber.31 A group of muscle fibers form a fascicle, which eventually composes the whole muscle structure (Figure 1d). On the basis of this concept, muscle regeneration can be induced with aligned cues in 3D cylindrical structure to mimic the unit of muscle structure, myofibers.11 Specifically, we developed a fibrous bundle structure with 100 μm for mimicking a myofiber, which is observed between 20 and 100 μm. Although there is a curvature on a structure, we cannot ensure whether a cell or tissue interacts with the surface in 2D or 3D. However, the fibrous bundles with various bundle diameters affected cellular activities, which were distinct from the 2D substrate.11 In this concern, we used the expression “3D structure”, which would result in distinct cellular activities from a 2D structure. Hence, we designed the scaffold composed of uniaxially arranged fibrous bundles in a 3D cylindrical structure, and for a possible application of multilayered fibers, the structure was rolled (Figure 1e). Uniaxial Stretching of Microfibrous Bundles for Fiber Alignment. To achieve the optimal stretching conditions for the 3D fibrous bundles, we simply stretched the randomly distributed fibers. The samples for the stretching test were fabricated in 10 parallel bundles in a wide gap (∼1 mm) to observe the individual breakage, whereas the samples for in vitro cell culture were fabricated in dozens of parallel bundles in D


Article


Figure 2. (a) Optical and (b) SEM images of the bundle structure applied with various strains (0, 40, 80, and 100%). (c) Fiber alignment for different strains represented by the fwhm. (d) Stretching analysis in the aspect of fwhm and breakage percentage of stretched fibrous bundle (n = 100). (e) Stress−strain curve of the bundle structure.

small gaps (∼50 μm) to provide contact space for bioink to be printed. To minimize the breakage of the fibers, we applied a low stretching speed (0.01 mm/s) at a temperature of 45−50 °C. Panels a−c in Figure 2 show the optical and SEM images and fwhm, respectively, for the various strains (0, 40, 80, and 100%). The optical images of Figure 2a show the parallel fibrous bundles with a couple of supporting perpendicular fibrous bundles. The bundle structure was stretched up to 100% of its original length, and breakage of the bundles was observed and quantitively analyzed. Figure 2b shows the SEM images of the stretched fibrous bundles, and the degree of the alignment was represented in the fwhm values (Figure 2c). As shown in the images and fwhm, the alignment of the microfibers became increasingly arranged in the uniaxial direction as the stretching strain increased. The stretching analysis in the aspect of the fwhm and breakage percentage of the stretched fibers is shown in Figure 2d. The breakage percentage was calculated from the ratio of the number of the broken bundle to the total number of stretched bundles (n = 100). The breakage percentage increased linearly and reached approximately 4 and 6% at 80 and 100 stretching percentage, respectively. As our aim is to fabricate a highly aligned fibrous bundle structure to mimic the original muscle structure, a slight breakage can be allowable, and therefore, the stretching percentage of 80% was chosen for

the stretching condition. Figure 2e shows the stress−strain curve for the fibrous bundle structure, and its Young’s modulus was 279.4 ± 41.6 kPa. It can be considered as an adequate mechanical strength for regenerating muscle that endures from 40 to 462 kPa in different body parts.32 Optimal Mixture Ratio of Collagen/PEO Bioink for C2C12 Printing. Collagen-based bioink has been actively investigated in various tissue engineering applications, specifically for cell printing processes.33−35 In this work, we propose a collagen bioink supplemented with PEO for high printability, biocompatibility, and appropriate cell release. In general, poly(ethylene glycol) (PEO) of high molecular weight degrades faster than collagen when immersed into a PBS solution, and therefore, it can be used as a leaching material to aid rapid cell release.36 Unfortunately, as the concentration of the PEO in the bioink increases, the initial cell viability after printing could decrease owing to the increased shear wall stress in a printing nozzle, and if the cell release is too rapid, the time available for the released cells to attach to the substrates will be insufficient.37 In addition, the bioink with a high concentration of PEO may not retain its printed shape for the culture period owing to its rapid degradation. The disruption of collagen bioink collapses the cytoplasmic actin filament bundles, so cells lose the surface fibronectin for adhesion.38 If the cells do not E


Article


Figure 3. (a) Optical images of collagen (2 wt %) and collagen/PEO 1, 2, and 3 wt % captured after 4 h and 1 day of cell printing. Rheological analysis on (b) complex viscosity and (c) storage modulus (G′) of pure cell-laden collagen and cell-laden collagen/PEO mixture. Quantitative results of (d) the storage modulus at 5 Hz and (e) initial cell viability. NS refers to no significant difference.

Figure 4. Schematic, optical, SEM, and live/dead (in situ) images captured for fabricated cell-laden (a) R-, (b) A-, and (c) C-scaffolds.

bioink. In this process, initial cellular events like cell adhesion and proliferation are affected. The collagen without PEO and that with 1 wt % PEO sustained their shape after 1 day, although slight degradation occurred at their boundaries. More specifically, myoblasts did not migrate out of Col/PEO 1 wt % after 1 day of culture, and initial cell adhesion and proliferation would occur after 1 d or later. In contrast, the collagen with 3 wt % PEO degraded too rapidly even with a culture period of 4 h in which encapsulated myoblasts were dispersed to

have sufficient time to attach on the target site by rapid degradation of bioink, the efficiency of bioink would be considered inadequate to build an in vitro cell culture platform. Figure 3a shows the optical images of the cell-printed bioink (2 wt % collagen with C2C12 cells) without PEO and with 1, 2, and 3 wt % PEO for culture periods of 4 h and 1 day to compare the degradability of the bioink. Because the cells were encapsulated in bioink, the interaction between cells and the scaffold surface occurred when the cells migrated out of the F


Article


Figure 5. (a) Live/dead images were captured after 3 and 7 days of cell culture (live cells in green; dead cells in red) with quantitative analysis on (b) initial cell viability and (c) DNA concentration at 1, 3, and 7 d. Significant differences are indicated by an asterisk (*), and NS refers to no significant difference.

bioink. The geometry of the fabricated scaffolds is roughly 10 mm × 15 mm × 90 μm, and the diameters of fibrous bundles (n = 100) and fibers (n = 100) were measured using SEM images. The fibrous bundle diameters were approximately 95.2 ± 8.4, 85.0 ± 7.0, and 98.5 ± 9.2 μm for the R-, A-, and Cscaffolds, respectively. In addition, the average diameters of the constituent fibers in the bundles were approximately 2.4 ± 0.2 μm for the R-scaffold, 1.1 ± 0.2 μm for the A-scaffold, and 1.6 ± 0.3 μm for the C-scaffold. The fiber diameter decreased from the R-scaffold to the A- and C-scaffolds owing to the uniaxial stretching process. Furthermore, the bioink was printed on the couple of bundles, and there were couple of bundles in the gap between the printed bioink as shown in the schematic images (Figure 4a−c). In the live/dead images, the in situ cell viability was greater than 93%, indicating that the printing process using the C2C12-laden collagen bioink was an extremely safe process. For the feasibility of the bundle structure to be evaluated further, the wetting ability of the bioink (mixed with rhodamine) was observed using the R-scaffold. Panels a and b in Figure S1 reveal the drastic decrease in the contact angle of the droplet from 106.6° to 78.2° for 0 and 60 s, respectively. At 120 s, a physically mixed region was observed from the crosssectional optical image, revealing the wetted interface between the bioink and fibrous structure; therefore, the infiltration of bioink facilitated successful cell attachment and proliferation resulting in a homogeneous cell−matrix interaction (Figure S1c). In addition, the cell seeding and cell printing methods using the bioink (Col/PEO 2 wt %) were compared. The cells were not evenly distributed by cell seeding, whereas cell printing showed a homogeneous cell distribution (Figure S2a). For the cell seeding efficiency, the R-, A-, and C-scaffolds resulted in

nonspecific binding sites. Collagen with 2 wt % PEO showed the released C2C12 cells after 1 day, and it represented appropriate cell release compared to that of the collagen with different weight fractions of PEO. Thus, we used collagen with 2 wt % PEO in this paper. To observe the modulus of the collagen/PEO bioink, we measured the complex viscosity and storage modulus (G′) (Figure 3b, c). As shown in the figures, as the concentration of PEO increased, the modulus of the bioinks increased significantly compared to that of the pure collagen (27 ± 7 Pa at 5 Hz) (Figure 3d). Although the high modulus can aid in constructing porous cell-laden structures during the printing, as the increased storage modulus can cause low cell viability after the printing owing to the high wall shear stress, we should consider the initial cell viability. Figure 3e shows the cell viability after printing the bioinks using the printing process (cell density: 1 × 107 cells/mL, pressure: 100 kPa, nozzle size: 150 μm, and nozzle moving speed: 10 mm s−1). Fortunately, the cell viability of the bioink was greater than 96% for all scaffolds. However, with respect to the cell release, we selected the cell-laden collagen/PEO (2 wt %) bioink that provides an appropriate cell release, safe cell viability, and reasonable mechanical properties. Fabrication of a Fibrous Scaffold Printed with C2C12 Cells. By using the selected cell-laden bioink (collagen/PEO-2 wt %) and the combined processes of EHD printing, uniaxial stretching, and cell printing, we can fabricate three cell-laden fibrous scaffolds (R-, A-, and C-scaffolds). Panels a−c in Figure 4 show the optical, SEM, and fluorescence images (live/dead) for the three fabricated scaffolds, R-scaffold, A-scaffold, and C-scaffold, respectively. In the optical images, the pink struts indicate the cell-laden G


Article


Figure 6. Fluorescence images stained by DAPI/phalloidin assays (nuclei in blue; F-actin in red) and SEM images for (a) R-, (b) A-, and (c) Cscaffolds after 1, 3, and 7 days of culture.

Figure 7. (a) MHCs were captured (nuclei in blue; MHC in green) at 3 and 7 d, and the SEM images at 7 d were represented. Sarcomeric α-actin (green) was visualized with nuclei (blue) by immunofluorescence staining at 14 d. Quantitative results of (b) MHC positive area and growth of (c) fusion index and (d) maturation index at 7 d were measured and normalized by the results of 3 d. The MHCs were also measured by (e) myotube length. Significant differences are indicated by an asterisk (*), and NS refers to no significant difference.

28.8 ± 4.2, 28.4 ± 3.0, and 46.6 ± 3.4%, respectively (Figure S2b). For the effect to be extended on differentiation, MHCs were homogeneously distributed in greater number with the cell printing method (Figure S2c). By these observations, we confirmed that cell printing is an efficient method for locating cells in a 3D environment and successively promoting various cellular activities.

Cell Viability and Proliferation on Scaffolds. The printed C2C12 cells were stained by live/dead assays, and their fluorescence images were captured after 4 h (in situ) and 3 and 7 days of cell culture (Figure 5a). Live cells were represented in green, and dead cells were represented in red. As shown in the images, the printed cells on the fibrous structure lived and proliferated well. The cell viability was measured at 7 H


Article


on day 7 (Figure 7d). However, the R-scaffold did not improve the level of the maturation index, and the result implied that myoblasts may be prevented from forming a multinucleated structure by random topological cues. The quantitative analysis of the cell morphology was reported in Figure 7e. The myotube length showed a significant growth on day 7 for all the scaffolds: 136 ± 19, 186 ± 39, and 132 ± 32 μm for the R-, A-, and C-scaffolds, respectively. The increase in the myotube length is greatly affected by the guidance of the aligned cues, and the A-scaffold allowed the cells to directly interact and expand through uniaxially aligned fibers.40 Although the ideal morphology for myotube was represented with the A-scaffold, the quantity was greater in the C-scaffold. Collagen naturally forms fibrils and triggers various cellular activities like cell adhesion and proliferation. The fibrils provide micro/nanoscale topological and biochemical cues, but their orientation would be random owing to sporadic aggregation among collagen molecules. In this concept, on the fibril-coated aligned fibers, the myotube length of C-scaffold can be not distinct from that of the R-scaffold. However, cell− cell interactions on the C-scaffold can be perceived to be more active compared to those of the R-scaffold because collagen contains RGD peptide and helps to synthesize type I collagen in cells. For these reasons, the C-scaffold showed a greater quantity compared to that of the A-scaffold and greater confluency of myoblasts compared to that of the R-scaffold. MHCs on the C-scaffold were greater in fluorescent area and evenly distributed. These observations support that the myoblasts grow through the pattern of topological cues. The R-scaffold showed a significant increase in myotube length from day 3 to 7, but the confluence of myoblasts was lower than those of the A- and C-scaffolds. The ideal morphology for myotubes may be assumed with MHCs on the A-scaffold, but the quantity was much greater in the Cscaffold. The sarcomeric structure (Figure 8a) staining shows that the degree of differentiation between A- and C-scaffolds was not significantly different. However, the homogeneously distributed myotubes on the C-scaffold may affect much higher myogenic gene expression than that of the A-scaffold. Blebbistatin, a small molecule, binds to myosin heads and hinders cell contractility by creating a complex for actin with low affinity. We referred to Mnatsakanyan et al. and used blebbistatin to examine the role of contractility against the capability of scaffolds to trigger differentiation.41 As shown in Figure 8a, the contractility of myoblasts on the scaffolds was examined using the sarcomeric structure cultured with and without blebbistatin. The R-scaffold showed a fraction of 1.7 between the sarcomeric structure cultured without and with blebbistatin. The A-scaffold showed an increase in the formation of sarcomeric structure by 3.6-fold and the Cscaffold increased 4.6-fold, which was significantly different from that of the R-scaffold (Figure 8b). This represents that the formation of sarcomeric structure, which has an interrelation with actin-myosin contractility, on the A- and C-scaffolds was critically affected by the presence of blebbistatin. Real-time PCR was used to obtain the result after 3 weeks of cell culture. In the results, the myogenic gene markers were relatively expressed with Myf5, MyoD, Myh2, and Myogenin (Figure 8c). Myf5 and MyoD act as regulatory factors of myogenesis and help to determine and differentiate specifically into skeletal muscle.42 Myh2, referred to as myosin heavy chain 2 gene, is a protein that comprises myosin heavy chains and helps skeletal muscle contraction.43 Myogenin is a factor that is

d using the live/dead images, which showed high viability of ∼94% for all scaffolds (Figure 5b). In addition, the proliferation rate of the printed cells was measured using PicoGreen assay (Figure 5c). A similar DNA concentration was observed on the R- and A-scaffolds from 1 to 7 d, whereas the C-scaffold revealed significantly higher DNA concentration throughout the 7 d. These results support the notion that the fibrous scaffolds could act as an outstanding platform that can promote both proliferation and differentiation. Morphological Analysis and Differentiation of C2C12 Cells. Panels a−c in Figure 6 display the nuclei (blue)/F-actin (red) images and SEM images after cell culture days 1, 3, and 7 for the R-, A-, and C-scaffolds, respectively. As shown in the results, we can confirm that the C2C12 cells in the A- and Cscaffolds were more proficiently aligned than in the R-scaffold (control). In particular, the C-scaffold provided more rapid alignment of the cells/ECM than that of the A-scaffold. The SEM images further reveal that the ECM is elongated along the fibrous cue, especially on the C-scaffold, and it can be elaborated by the synergistic effect of the topological cue of the fibrous structure and thin-coated collagen component of the C-scaffold. Collagen molecules intrinsically regulate themselves tightly, so this aggregation leads to the formation of nanoscale fibrils. 39 These fibrils provide nanoscale topological cues and benefit cell adhesion and myoblast differentiation.16 On the basis of this result, it can carefully be estimated that the C-scaffold can provide a much more stable microenvironmental physiochemical condition to induce myoblast differentiation. The myosin heavy chain (MHC) was stained to show myotube formation of C2C12 cells cultured on the R-, A-, and C-scaffolds. Figure 7a shows fluorescence images of DAPI/ MHC on days 3 and 7. The morphology of the MHC at day 3 was mostly circular around the nuclei on the R-scaffold, whereas the A- and C-scaffolds showed an aligned morphology of MHC. At day 7, the MHC formation of all scaffolds was much larger in number and broader on the scaffold surface as compared to that of day 3 and, for the A- and C-scaffolds, a multinucleated/confluent structure was observed. However, as shown in the DAPI/MHC images of day 7, the C-scaffold provides much more homogeneous cell proliferation compared to those of the R- and A-scaffolds due to the coated collagen layer. Furthermore, the sarcomeric structure at day 14 for the scaffolds was stained to show the mature differentiation stage of myoblasts (Figure 7a). For quantitative analysis of the MHC, three fluorescence images of each scaffold were used, and the myotubes with more than two nuclei were evaluated (n = 50). Furthermore, MHC was represented by a positive area and growth percentage of fusion and maturation index normalized by the result on day 3 (Figure 7b−d). The growth of the MHC positive area was increased ∼1.5-fold in the A-scaffold and ∼1.7-fold in the Cscaffold, whereas no significant difference was observed in the R-scaffold. The fusion index indicating MHCs with two or more nuclei was obtained, and a significant difference was observed between days 3 and 7 on the A- and C-scaffolds (Figure 7c). On the basis of an analysis of the results, it can be observed that the topological cue affects the myoblasts, i.e., triggers differentiation, such as the formation of neonatal myotubes. The maturation index is used to indicate a mature MHC that has five or more nuclei in an MHC. Both the A- and C-scaffolds revealed that the myotube maturation was actively conducted I


Article


appropriate in vitro platform for high cell viability and proliferation; however, the progress of differentiation was distinct among the scaffolds. The aligned cues greatly benefited the formation of myotubes, and furthermore, the collagencoated surface with aligned/biochemical cues enhanced the myotubes to develop into mature myotubes. This study presents a 3D fibrous bundle structure with topological and biochemical cues that induce myogenesis, and an estimation of the clinical relevance would proceed further through studies on muscle functionality.

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsbiomaterials.7b00983. (Figure S1) Water contact angles and SEM/fluorescence images to demonstrate cell infiltration and (Figure S2) live/dead images and MHC formation at 7 days on the scaffolds fabricated via cell seeding and cell printing methods (PDF)

■

AUTHOR INFORMATION

Corresponding Author

*Tel.: +82-31-290-7828. E-mail: [email protected]. ORCID

GeunHyung Kim: 0000-0002-2965-2171 Notes

The authors declare no competing financial interest.

■ ■

Figure 8. (a) Fluorescence images of sarcomeric α-actin (green) and nuclei (blue) cultured without and with blebbistatin at 7 d. (b) The fraction of sarcomeric structure area cultured without/with blebbistatin at 7 d. (c) Relative gene expression of myogenic gene markers (Myf5, Myh2, MyoD, and Myogenin) after 21 days of cell culture. The gene expression was compared with β-actin and normalized with that of the R-scaffold. Significant differences are indicated by an asterisk (*).

ACKNOWLEDGMENTS This paper was supported by 63 Research Fund, Sungkyunkwan University, 2016. REFERENCES

(1) Nerem, R. M.; Sambanis, A. Tissue engineering: from biology to biological substitutes. Tissue Eng. 1995, 1 (1), 3−13. (2) Qazi, T. H.; Mooney, D. J.; Pumberger, M.; Geissler, S.; Duda, G. N. Biomaterials based strategies for skeletal muscle tissue engineering: existing technologies and future trends. Biomaterials 2015, 53, 502− 521. (3) Ostrovidov, S.; Hosseini, V.; Ahadian, S.; Fujie, T.; Parthiban, S. P.; Ramalingam, M.; Bae, H.; Kaji, H.; Khademhosseini, A. Skeletal muscle tissue engineering: methods to form skeletal myotubes and their applications. Tissue Eng., Part B 2014, 20 (5), 403−436. (4) Yoshioka, J.; Prince, R. N.; Huang, H.; Perkins, S. B.; Cruz, F. U.; MacGillivray, C.; Lauffenburger, D. A.; Lee, R. T. Cardiomyocyte hypertrophy and degradation of connexin43 through spatially restricted autocrine/paracrine heparin-binding EGF. Proc. Natl. Acad. Sci. U. S. A. 2005, 102 (30), 10622−10627. (5) Fan, X.; Zou, R.; Zhao, Z.; Yang, P.; Li, Y.; Song, J. Tensile strain induces integrin β1 and ILK expression higher and faster in 3D cultured rat skeletal myoblasts than in 2D cultures. Tissue Cell 2009, 41 (4), 266−270. (6) Padmawar, P.; Yao, X.; Bloch, O.; Manley, G. T.; Verkman, A. K+ waves in brain cortex visualized using a long-wavelength K+-sensing fluorescent indicator. Nat. Methods 2005, 2 (11), 825−827. (7) Cascio, W. E.; Yan, G.-X.; Kleber, A. G. Early changes in extracellular potassium in ischemic rabbit myocardium. The role of extracellular carbon dioxide accumulation and diffusion. Circ. Res. 1992, 70 (2), 409−422. (8) Jana, S.; Levengood, S. K. L.; Zhang, M. Anisotropic Materials for Skeletal Muscle Tissue Engineering. Adv. Mater. 2016, 28, 10588.

expressed at the postmitotic stage.44 All expressed gene markers were significantly high for the C-scaffold. This was because the C-scaffold can provide much wider and homogeneous myogenesis within the 3D structure owing to the synergistic effect of the topographical and biochemical cue (coated collagen) instead of the A-scaffold only having a topological cue. In brief, the regulatory factors initiating the early stage of differentiation are consistently expressed until the myotubes become mature.45−47 The level of Myh2 and Myogenin was also high for the C-scaffold and proved the formation of mature myotubes. These data indicate that the myoblast differentiation was achieved effectively by the aligned topological cues and great biocompatibility of the collagen-coated surface.

■

CONCLUSIONS In this study, a fibrous 3D bundle structure with cell-laden bioink was proposed to induce myoblast regeneration. The bundle structures were fabricated with various surface properties of random, aligned, and aligned/collagen-coated surfaces. Randomly oriented fibrous scaffolds provided myoblasts a 3D platform, and aligned and aligned/collagen-coated scaffolds helped myoblast growth with topological and/or biochemical guidance in a 3D environment. All scaffolds provided an J


Article

ACS Biomaterials Science & Engineering (9) Vandenburgh, H. H.; Karlisch, P.; Farr, L. Maintenance of highly contractile tissue-cultured avian skeletal myotubes in collagen gel. In Vitro Cell. Dev. Biol. 1988, 24 (3), 166−174. (10) Donnelly, K.; Khodabukus, A.; Philp, A.; Deldicque, L.; Dennis, R. G.; Baar, K. A novel bioreactor for stimulating skeletal muscle in vitro. Tissue Eng., Part C 2010, 16 (4), 711−718. (11) Wang, L.; Wu, Y.; Guo, B.; Ma, P. X. Nanofiber yarn/hydrogel core−shell scaffolds mimicking native skeletal muscle tissue for guiding 3D myoblast alignment, elongation, and differentiation. ACS Nano 2015, 9 (9), 9167−9179. (12) Takeda, N.; Tamura, K.; Mineguchi, R.; Ishikawa, Y.; Haraguchi, Y.; Shimizu, T.; Hara, Y. In situ cross-linked electrospun fiber scaffold of collagen for fabricating cell-dense muscle tissue. J. Artif. Organs 2016, 19 (2), 141−148. (13) Li, M.; Guo, Y.; Wei, Y.; MacDiarmid, A. G.; Lelkes, P. I. Electrospinning polyaniline-contained gelatin nanofibers for tissue engineering applications. Biomaterials 2006, 27 (13), 2705−2715. (14) Ricotti, L.; Polini, A.; Genchi, G. G.; Ciofani, G.; Iandolo, D.; Vazao, H.; Mattoli, V.; Ferreira, L.; Menciassi, A.; Pisignano, D. Proliferation and skeletal myotube formation capability of C2C12 and H9c2 cells on isotropic and anisotropic electrospun nanofibrous PHB scaffolds. Biomed. Mater. 2012, 7 (3), 035010. (15) Murr, L. E. Bioprinting and Biofabrication of Organs. In Handbook of Materials Structures, Properties, Processing and Performance; Springer, 2015; pp 629−638. (16) Arnesen, S.; Mosler, S.; Larsen, N. B.; Gadegaard, N.; Purslow, P. P.; Lawson, M. A. The effects of collagen type I topography on myoblasts in vitro. Connect. Tissue Res. 2004, 45 (4−5), 238−247. (17) Rossi, C. A.; Pozzobon, M.; De Coppi, P. Advances in musculoskeletal tissue engineering: moving towards therapy. Organogenesis 2010, 6 (3), 167−172. (18) Rutz, A. L.; Hyland, K. E.; Jakus, A. E.; Burghardt, W. R.; Shah, R. N. A multimaterial bioink method for 3D printing tunable, cellcompatible hydrogels. Adv. Mater. 2015, 27 (9), 1607−1614. (19) Blaeser, A.; Duarte Campos, D. F.; Puster, U.; Richtering, W.; Stevens, M. M.; Fischer, H. Controlling shear stress in 3D bioprinting is a key factor to balance printing resolution and stem cell integrity. Adv. Healthcare Mater. 2016, 5 (3), 326−333. (20) Davidenko, N.; Schuster, C. F.; Bax, D. V.; Farndale, R. W.; Hamaia, S.; Best, S. M.; Cameron, R. E. Evaluation of cell binding to collagen and gelatin: a study of the effect of 2D and 3D architecture and surface chemistry. J. Mater. Sci.: Mater. Med. 2016, 27 (10), 148. (21) You, R.; Li, X.; Luo, Z.; Qu, J.; Li, M. Directional cell elongation through filopodia-steered lamellipodial extension on patterned silk fibroin films. Biointerphases 2015, 10 (1), 011005. (22) Kim, M. S.; Kim, G. Electrohydrodynamic jet process for porestructure-controlled 3D fibrous architecture as a tissue regenerative material: fabrication and cellular activities. Langmuir 2014, 30 (28), 8551−8557. (23) Chen, G.; Ushida, T.; Tateishi, T. A hybrid network of synthetic polymer mesh and collagen sponge. Chem. Commun. 2000, 16, 1505− 1506. (24) Lee, S. B.; Kim, Y. H.; Chong, M. S.; Lee, Y. M. Preparation and characteristics of hybrid scaffolds composed of β-chitin and collagen. Biomaterials 2004, 25 (12), 2309−2317. (25) Khil, M. S.; Bhattarai, S. R.; Kim, H. Y.; Kim, S. Z.; Lee, K. H. Novel fabricated matrix via electrospinning for tissue engineering. J. Biomed. Mater. Res. 2005, 72B (1), 117−124. (26) Sobral, J. M.; Caridade, S. G.; Sousa, R. A.; Mano, J. F.; Reis, R. L. Three-dimensional plotted scaffolds with controlled pore size gradients: effect of scaffold geometry on mechanical performance and cell seeding efficiency. Acta Biomater. 2011, 7 (3), 1009−1018. (27) Kim, M.; Kim, G. 3D multi-layered fibrous cellulose structure using an electrohydrodynamic process for tissue engineering. J. Colloid Interface Sci. 2015, 457, 180−187. (28) Abedalwafa, M.; Wang, F.; Wang, L.; Li, C. Biodegradable polyepsilon-caprolactone (PCL) for tissue engineering applications: a review. Rev. Adv. Mater. Sci. 2013, 34 (2), 123−40.

(29) Bezwada, R. S.; Jamiolkowski, D. D.; Lee, I.-Y.; Agarwal, V.; Persivale, J.; Trenka-Benthin, S.; Erneta, M.; Suryadevara, J.; Yang, A.; Liu, S. Monocryl suture, a new ultra-pliable absorbable monofilament suture. Biomaterials 1995, 16 (15), 1141−1148. (30) Rezgui, F.; Swistek, M.; Hiver, J.; G’sell, C.; Sadoun, T. Deformation and damage upon stretching of degradable polymers (PLA and PCL). Polymer 2005, 46 (18), 7370−7385. (31) Burattini, S.; Ferri, P.; Battistelli, M.; Curci, R.; Luchetti, F.; Falcieri, E. C2C12 murine myoblasts as a model of skeletal muscle development: morpho-functional characterization. Eur. J. Histochem. 2004, 48 (3), 223. (32) Gillies, A. R.; Lieber, R. L. Structure and function of the skeletal muscle extracellular matrix. Muscle Nerve 2011, 44 (3), 318−331. (33) Wilson, W. C.; Boland, T. Cell and organ printing 1: protein and cell printers. Anat. Rec. 2003, 272A (2), 491−496. (34) Ovsianikov, A.; Gruene, M.; Pflaum, M.; Koch, L.; Maiorana, F.; Wilhelmi, M.; Haverich, A.; Chichkov, B. Laser printing of cells into 3D scaffolds. Biofabrication 2010, 2 (1), 014104. (35) Ferris, C. J.; Gilmore, K. J.; Beirne, S.; McCallum, D.; Wallace, G. G. Bio-ink for on-demand printing of living cells. Biomater. Sci. 2013, 1 (2), 224−230. (36) Yeo, M.; Kim, G. Optimal size of cell-laden hydrogel cylindrical struts for enhancing the cellular activities and their application to hybrid scaffolds. J. Mater. Chem. B 2014, 2 (39), 6830−6838. (37) Kolesky, D. B.; Truby, R. L.; Gladman, A.; Busbee, T. A.; Homan, K. A.; Lewis, J. A. 3D bioprinting of vascularized, heterogeneous cell-laden tissue constructs. Adv. Mater. 2014, 26 (19), 3124−3130. (38) Mochitate, K.; Pawelek, P.; Grinnell, F. Stress relaxation of contracted collagen gels: disruption of actin filament bundles, release of cell surface fibronectin, and down-regulation of DNA and protein synthesis. Exp. Cell Res. 1991, 193 (1), 198−207. (39) Reese, S. P.; Underwood, C. J.; Weiss, J. A. Effects of decorin proteoglycan on fibrillogenesis, ultrastructure, and mechanics of type I collagen gels. Matrix Biol. 2013, 32 (7), 414−423. (40) McClure, M. J.; Clark, N. M.; Hyzy, S. L.; Chalfant, C. E.; Olivares-Navarrete, R.; Boyan, B. D.; Schwartz, Z. Role of integrin α7β1 signaling in myoblast differentiation on aligned polydioxanone scaffolds. Acta Biomater. 2016, 39, 44−54. (41) Mnatsakanyan, H.; Rico, P.; Grigoriou, E.; Candelas, A. n. M.; Rodrigo-Navarro, A.; Salmeron-Sanchez, M.; Sabater i Serra, R. Controlled assembly of fibronectin nanofibrils triggered by random copolymer chemistry. ACS Appl. Mater. Interfaces 2015, 7 (32), 18125−18135. (42) Andrés, V.; Walsh, K. Myogenin expression, cell cycle withdrawal, and phenotypic differentiation are temporally separable events that precede cell fusion upon myogenesis. J. Cell Biol. 1996, 132 (4), 657−666. (43) Khodabukus, A.; Baehr, L. M.; Bodine, S. C.; Baar, K. Role of contraction duration in inducing fast-to-slow contractile and metabolic protein and functional changes in engineered muscle. J. Cell. Physiol. 2015, 230 (10), 2489−2497. (44) Ott, M.-O.; Bober, E.; Lyons, G.; Arnold, H.; Buckingham, M. Early expression of the myogenic regulatory gene, myf-5, in precursor cells of skeletal muscle in the mouse embryo. Development 1991, 111 (4), 1097−1107. (45) Olguin, H. C.; Yang, Z.; Tapscott, S. J.; Olwin, B. B. Reciprocal inhibition between Pax7 and muscle regulatory factors modulates myogenic cell fate determination. J. Cell Biol. 2007, 177 (5), 769−779. (46) Mofarrahi, M.; Hussain, S. N. Expression and functional roles of angiopoietin-2 in skeletal muscles. PLoS One 2011, 6 (7), e22882. (47) Luo, W.; Li, E.; Nie, Q.; Zhang, X. Myomaker, regulated by MYOD, MYOG and miR-140−3P, promotes chicken myoblast fusion. Int. J. Mol. Sci. 2015, 16 (11), 26186−26201.

K


3D microfibrous bundle structure fabricated using an electric-field

Recommend Documents