NANO LETTERS
Myotube Assembly on Nanofibrous and Micropatterned Polymers
2006 Vol. 6, No. 3 537-542
Ngan F. Huang,†,‡ Shyam Patel,†,‡ Rahul G. Thakar,†,‡ Jun Wu,§ Benjamin S. Hsiao,§ Benjamin Chu,§ Randall J. Lee,†,| and Song Li*,†,‡ Joint Graduate Program in Bioengineering, UniVersity of California San Francisco/ UniVersity of California Berkeley, California 94720-1762, Department of Bioengineering and Center for Tissue Bioengineering, UniVersity of California Berkeley, California 94720, Department of Chemistry, Stony Brook UniVersity, Stony Brook, New York 11794-3400, and Department of Medicine, UniVersity of California San Francisco, San Francisco, California 94143 Received January 10, 2006; Revised Manuscript Received January 27, 2006
ABSTRACT Skeletal muscle consists of parallel bundles of myotubes formed by the fusion of myoblasts. We fabricated nanofibrous and micropatterned polymers as cell culture substrates to guide the morphogenesis of muscular tissue. The nanoscale and microscale topographic features regulate cell and cytoskeleton alignment, myotube assembly, myotube striation, and myoblast proliferation. This bottom-up approach from nanoscale to tissue level demonstrates the potential of nanofibrous polymers for engineering the assembly of cell and tissue structure.
Spatial and geometric signals regulate numerous aspects of cellular and tissue behavior, including proliferation, migration, and regeneration.1 In the case of skeletal muscle, its structure is highly organized and consists of long parallel bundles of multinucleated myotubes that are formed by differentiation and fusion of myoblast satellite cells.2 When cultured in vitro, however, myoblasts and myotubes lose their native organization and adopt random distributions, which do not resemble physiological muscle architecture. To understand the role of spatial cues in regulating skeletal muscle development and to engineer muscle for therapeutic applications, in vitro systems that mimic the aligned physiological architecture need to be developed. Electrospun nanofibers can provide submicrometer-scale features that resemble the physical structure of native collagen fibrils or extracellular matrixes. The nanofibers can be fabricated from biocompatible polymers such as poly(lactide) (PLA) or poly(glycolide-co-lactide) (PGLA),3,4 which have therapeutic potential for implantation applications. To modulate the organization of nanofibers into aligned fibrous structure, the use of patterned electrodes, wire drum collectors, or postprocessing methods such as uniaxial * To whom correspondence may be addressed: 471 Evans Hall, #1762, University of California Berkeley, Berkeley, CA 94720. Tel: (510) 6653598. Fax: (510) 665-3599. E-mail:
[email protected]. † Joint Graduate Program in Bioengineering, University of California San Francisco/University of California Berkeley. ‡ Department of Bioengineering and Center for Tissue Bioengineering, University of California Berkeley. § Department of Chemistry, Stony Brook University. | Department of Medicine, University of California San Francisco. 10.1021/nl060060o CCC: $33.50 Published on Web 02/16/2006
© 2006 American Chemical Society
stretching have been successful.4-6 Recently, the feasibility of nanofiber substrates for the guidance of cell growth, function, and organization has been demonstrated for fibroblasts, vascular cells, and mesenchymal stem cells.7-11 However, the potential of nanofibers for guiding myoblast assembly and the morphogenesis of skeletal muscle has not been explored. Another method for modulating spatial and geometric organization is microfabrication, which uses soft lithography to topographically or chemically create micrometer-scale features on substrate surfaces.12-14 This technique can be used to control many aspects of cellular behavior, including cell size, shape, spatial organization, proliferation, and survival.15-19 Elastomers such as poly(dimethylsiloxane) (PDMS) can be micropatterned with high reproducibility and provide a flexible substrate for cell attachment.20 Here we will investigate skeletal muscle morphogenesis on deformable micropatterned substrates, and compare the effects of micropatterned and nanopatterned substrates on cell guidance and the morphogenesis of skeletal muscle tissue. Murine C2C12 myoblasts (ATCC, Manassas, VA) were used to study cell organization and assembly. The myoblasts were cultured in growth media that consisted of Dulbecco’s Modified Eagle’s Medium (DMEM), 10% fetal bovine serum, and 1% penicillin/streptomycin. To initiate myoblast differentiation and fusion, the growth medium was replaced with differentiation media that consisted of DMEM, 5% horse serum, and 1% penicillin/streptomycin after 24 h when samples were confluent. In all experiments, time points were denoted by the incubation time in differentiation media.
Figure 1. Myoblast alignment and myotube assembly on an aligned PLLA nanofibrous scaffold. SEM images showing structure of (A) randomly oriented and (B) aligned nanofibrous scaffolds, followed by F-actin immunofluorescent staining of myoblasts on (C) randomly oriented and (D) aligned nanofibrous scaffolds after 3 days in differentiation media. Immunofluorescence staining of skeletal MHC was performed to show myotubes on random (E, G) and aligned (F, H) nanofibrous scaffolds at 3 days (E, F) and 7 days (G, H). Low magnification merged images of skeletal MHC staining on (I) randomly oriented and (J) aligned substrates after 7 days showing the global alignment and length of myotubes. Arrows indicate direction of nanofibers. Arrowheads in E and F indicate nuclei. Scale bars are 50 µm (A-H) and 100 µm (I, J), respectively.
Biodegradable poly(L-lactide) (PLLA) (Lactel Absorbable Polymers, Pelham, AL, 1.09 dL/g inherent viscosity) was used to fabricate nanofiber scaffolds by electrospinning.7 Briefly, the PLLA solution (10% w/v in chloroform) was delivered by a programmable pump to the exit hole of the electrode at a flow rate of 25 µL/min. A high-voltage supply (Glassman High Voltage Inc., High Bridge, NJ) was used to apply the voltage at 20 kV. The collecting plate was on a rotating drum that was grounded and controlled by a stepping motor. To align the nanofibers, the electrospun scaffold was stretched uniaxially to 200% deformation in length at 60 °C. Nanofibrous scaffolds were approximately 150 µm in thickness. The surface of the nanofibrous scaffold was coated with 2% gelatin or fibronectin (5 µg/cm2) before cell seeding. No significant difference in cell adhesion and morphology was detected between gelatin and fibronectin coating. Randomly oriented scaffolds were used as controls. Scanning electron microscopy (SEM) was used to visualize nanofiber alignment after uniaxial stretching. SEM images show that uniaxial stretching resulted in aligned nanofibers (Figure 1A,B). The average nanofiber diameter in the scaffold was approximately 500 nm with an average gap size of approximately 4 µm. Our micropatterning experiments showed that spacing up to 20 µm had a similar effect on cell alignment (data not shown). 538
Confluent myoblasts were grown on the nanofibrous scaffolds in differentiation media for up to 7 days. To examine cell organization and cytoskeletal structure on random-oriented and aligned nanofibrous scaffolds, fluorescence staining of F-actin, myosin heavy chain (MHC), and cell nuclei were performed (Supporting Information). F-actin was stained using fluorescein (FITC)-conjugated phalloidin (Molecular Probes, Eugene, OR). MHC was immunostained with a mouse anti-skeletal fast MHC antibody (Sigma, St. Louis, MO). Cell nuclei were stained with ToPro dye (Molecular Probes). Fluorescence microscopy was performed by using a Nikon TE300 microscope and Leica TCS SL confocal microscope. Confocal microscopy of fluorescently stained F-actin stress fibers after 3 days of differentiation media treatment revealed that the actin assembly appeared disordered on the randomly oriented nanofibrous substrate, whereas on aligned nanofibers the F-actin stress fibers were oriented along the nanofiber direction (Figure 1C,D). For the visualization of myoblasts that fused to form myotubes, samples were stained for fast skeletal MHC, a marker for mature differentiated myofibers,21,22 after 3 and 7 days in differentiation media. On the randomly oriented nanofibrous substrate (Figure 1E,G,I), myotubes were generally scattered in a wide range of directions. As myoblasts attached and formed multinucleated Nano Lett., Vol. 6, No. 3, 2006
Figure 3. Quantification of myoblast proliferation and myotube striation on aligned nanofibrous scaffolds. (A) BrdU incorporation for cell proliferation (R, ran; A, align). (B) Immunofluorescence staining of anti-MHC showing a striated myotube on aligned nanofibrous scaffold (scale bar, 20 µm). (C) Quantification of the percentage of striated cells after 7 days. Asterisks indicate statistically significant difference (P < 0.05). Figure 2. Quantification of myotube organization and morphology on aligned nanofibrous substrates. (A) Angle of myotube alignment in reference to nanofiber direction. (B) Myotube length after 7 days. (C) Myotube width after 7 days. Asterisks indicate statistically significant difference (P < 0.05).
myofibers along the nanofibers, myotubes on randomly oriented scaffolds only showed local areas of alignment and were not aligned globally. In stark contrast, myotubes on the aligned nanofiber substrate organized within close proximity to the direction of the nanofiber direction and formed globally aligned myotubes (Figure 1F,H,J). SEM images confirmed that the surfaces were covered by random and well-organized myotubes on randomly oriented and aligned nanofibrous scaffolds, respectively, after 7 days in differentiation media (Figure 1, Supporting Information). To quantify the effect of aligned nanofibrous structure on myoblast assembly into myotubes, we measured the angle, length, width, proliferation rate, and percent striation of myotubes by using immunofluorescent images of skeletal MHC staining (Supporting Information). The minimum myotube alignment value of 0° denoted parallel alignment from the axis of the nanofibers and the maximum of 90° represented perpendicular alignment. For alignment analysis of randomly oriented nanofibrous scaffolds, an arbitrary axis of alignment was used. Serial images under low magnification were merged to quantify myotube length. All data were expressed as mean ( standard deviation (n g 3). The aligned nanofibers promoted skeletal muscle morphogenesis into parallel-oriented myotubes. On aligned nanofibrous scaffolds the myotubes were highly organized. They formed angles of only 7 ( 2° from the axis of the nanofiber direction after 3 days, and the alignment remained at similar levels (7 ( 3°) after 7 days (Figure 2A). In contrast, Nano Lett., Vol. 6, No. 3, 2006
myotubes on randomly oriented nanofibrous scaffolds had a wide distribution of myotube orientations, with averaged alignments of 36 ( 11° and 54 ( 16° after 3 and 7 days, respectively. Deviations from a perfectly random alignment (45°) could be attributed to regions of local alignment on the random-oriented nanofibrous scaffolds. Myotubes on aligned nanofibers were not only highly organized but also significantly longer than those on randomly oriented nanofibrous scaffolds, with 30-40 nuclei in each myotube. The average length of myotubes (1.8 ( 0.3 mm) after 7 days on the aligned scaffold was more than twice the length on the randomly oriented one (0.8 ( 0.1 mm), suggesting that aligned nanofibrous scaffolds promote the assembly of myotubes (Figure 2B). In addition, myotube widths increased for both groups from 3 to 7 days, but the comparative widths at day 3 (14.9 ( 0.9 µm random-oriented vs 14.3 ( 0.3 µm aligned) and day 7 (18.6 ( 0.5 µm randomly oriented vs 18.1 ( 0.8 µm aligned) were not significantly different at either time point (Figure 2C). Together, these results demonstrate that aligned nanofibrous scaffolds can guide the self-assembly of longer and parallel myotubes that mimic the native muscular tissue. Myoblast differentiation and fusion into myotubes are usually correlated with the decrease of proliferation. To determine whether the aligned nanofibers affect cell cycle progression, time course analysis of cell proliferation was performed by using bromodeoxyuridine (BrdU) incorporation assay (Supporting Information). As expected, the rate of cell proliferation decreased during the first 3 days in differentiation media on both aligned and random nanofibers (Figure 3A). In addition, the proliferation rate was lower on the aligned than on the randomly oriented nanofibers. These results suggest that aligned nanofibrous scaffolds promote 539
Figure 4. Myoblast alignment and myotube organization on a micropatterned PDMS substrate. A micropatterned PDMS substrate is shown by (A) SEM (side view) and (B) phase contrast microscopy. F-actin distribution after 2 days in differentiation media is shown on (C) nonpatterned and (D) micropatterned substrates. Immunofluorescent staining of skeletal MHC was performed to show cell fusion on nonpatterned (E, G) and micropatterned (F, H) membranes after 2 days (E-F) and 7 days (G-H). Arrows indicate direction of microgrooves. Scale bars are 5 µm (A), 20 µm (B), and 50 µm (C-H), respectively.
cell cycle exit at early time points, which may be one of the factors contributing to myoblast differentiation and fusion into myotubes. The assembly of striated sarcomeres in myotube is the characteristic of contractile and functional myotubes. The striated sarcomeres (stained for MHC) in myotubes could be visualized at higher magnification (Figure 3B). The percentages of striated myotubes on aligned and random nanofibers were similar (65 ( 13% randomly oriented vs 66 ( 7% aligned) (Figure 3B,C), suggesting that either random or aligned topographic features at nanoscale can promote myotube striation. To determine whether myotube assembly depends on micrometer-scale topographic features, we employed soft lithography to create parallel microgrooves on PDMS membranes.23 The microgrooves were 10 µm wide, 10 µm apart, 2.8 µm deep, and 1 cm in length (Figure 4A,B). On micropatterned substrates, the grooves were selected to be 540
Figure 5. Quantification of myotube organization and morphology on micropatterned membranes. (A) Angle of myotube alignment in reference to the microgroove direction on nonpatterned (Con) and micropatterned (Pat) membranes. (B) Myotube length after 7 days. (C) Myotube width after 7 days. Asterisks indicate statistically significant difference (P < 0.05).
10 µm wide based on our prior experience with groove widths up to 100 µm, in which we found myoblast alignment decreased with increasing groove width. After oxygen plasma treatment, the PDMS substrates were coated with 2% gelatin before cell seeding. Nonpatterned PDMS membranes were used as controls. Confluent myoblasts were grown on the micropatterned or control surfaces in differentiation media for up to 7 days. Quantitative analysis of cell organization and proliferation was carried out as previously described. The widths of the microgrooves were similar to the dimension of myoblasts and restricted cell spreading such that the myoblasts adopted an elongated morphology and both cell and actin fibers aligned with the microgrooves (Figure 4C,D). Similar to observations on nanofibrous substrates, the myotubes on the nonpatterned PDMS surface appeared randomly oriented (Figure 4E,G), while on micropatterned microgrooves the myotubes aligned in the direction of the microgrooves after 2 and 7 days (Figure 4F,H). The alignment was 5 ( 2° after 7 days, while on the nonpatterned substrate the corresponding value was 44 ( 7° (Figure 5A). We also successfully created micropatterned membranes Nano Lett., Vol. 6, No. 3, 2006
Figure 6. Quantification of myoblast proliferation and striation on micropatterned PDMS membranes. (A)BrdU incorporation for cell proliferation at the early stage of fusion on nonpatterned (Con) and micropatterned (Pat) membranes. (B) Quantification of the percentage of striated cells after 7 days. Asterisks indicate statistically significant difference (P < 0.05).
using novel biodegradable poly(L-lactide-co-glycolide-co-caprolactone) (PLGC) triblock copolymers and found similar trends in cell alignment as on PDMS (Figure 2, Supporting Information). On microgrooves, myotube length was about 40% longer than that on nonpatterned membranes after 7 days (0.7 ( 0.1 mm vs 0.5 ( 0.1 mm) (Figure 5B), which was similar to the myotube length on random nanofibers. In comparison, the myotube length on aligned nanofibers was more than twice the length on micropatterned substrates. These results suggest that the nanoscale feature is more efficient in promoting myotube assembly. The width on both PDMS substrates increased from 2 to 7 days, similar to the finding on nanofibrous scaffolds, but the temporal trends differed from nanofibrous substrates in that the widths were consistently higher on nonpatterned than on micropatterned PDMS (Figure 5C). Cell proliferation rates declined over the first 2 days on both micropatterned and nonpatterned samples upon the initiation of myoblast differentiation (Figure 6A). The proliferation rate of myoblasts on micropatterned surfaces at day 2 was 35% lower than that on nonpatterned surfaces (19 ( 4% nonpatterned vs 12 ( 2% micropatterned), suggesting that the micropatterned surfaces may promote cell cycle exit of myoblasts. On comparison of the proliferation rate between nanofibers and PDMS substrates, the proliferation rates were generally lower on nanofibers, further supporting that aligned nanofibrous substrates promote exit out of cell cycle, which is necessary for differentiation and fusion into myotubes. In addition, the percentage of cells with striated sarcomere assembly on microgrooves was also significantly higher (58 ( 7% on micropatterned PDMS vs 37 ( 3% on nonpatterned Nano Lett., Vol. 6, No. 3, 2006
control) (Figure 6B). The level of striated cells on micropatterned samples was comparable to corresponding values on random or aligned nanofibrous substrates, suggesting that both microscale and nanoscale of topographic features enhance the assembly of striated sarcomeres. Our results indicate that myoblasts can sense the nanoscale and microscale topographic features in the extracellular space, which can guide cell orientation, interaction, organization, and assembly. On the basis of our results, we reach five conclusions on topographical regulation of myotube assembly. (1) Aligned nanoscale and microscale topographic features cause the alignment of myoblasts and cytoskeletal proteins and promote myotube assembly along the nanofibers and microgrooves (Figures 1 and 4), which mimic the myotube organization in skeletal muscle. (2) Nanoscale features are more efficient in promoting the assembly of longer myotubes than microscale features. As shown in Figures 2 and 5, while the myotube length on random nanofibers (with local alignment) is similar to that on microgrooves, myotubes formed on aligned nanofibers are much longer than those on aligned microgrooves. (3) Local topographic features at nanoscale and microscale enhance myotube striation. Microgrooves increase myotube striation (Figure 6) to the level induced by nanofibers (Figure 3). The similar level of myotube striation on random and aligned nanofibers suggests that a local topographic feature is sufficient to promote striation and that the global alignment is not required for striation formation. (4) Aligned microscale and nanoscale features may restrict cell spreading and suppress myoblast proliferation during differentiation and cell fusion. Nanofibers are more efficient than microgrooves in suppressing myoblast proliferation (Figures 3 and 6). (5) The topographic features at the microscale but not nanoscale affect the width of myotubes (Figures 2 and 5). This work not only sheds light on the regulation of cell alignment, cytoskeleton organization, myoblast proliferation, and myotube assembly by nanoscale and microscale topographic features but also generates culture platforms for engineering skeletal muscle at the tissue scale. The advantages of using nanofibrous scaffolds for myotube assembly provide a rational basis for muscular tissue engineering. This approach can also be combined with electrical, mechanical, and chemical stimulation factors to engineer skeletal muscle.24 The biodegradable nanofibrous scaffolds can be used to guide the morphogenesis of engineered muscular tissue and gradually degrade after the assembly of myoblasts, myotubes, and skeletal muscle tissue. The degradation rate of the polymers can be easily tailored to match the tissue generation rate. Furthermore, the aligned nanofibers can be used to guide the morphogenesis of other types of tissues with anisotropic structure, e.g., cardiac muscle, blood vessel, tendon, and ligament. This bottom-up approach from nanoscale feature to the assembly of cell and tissue structure opens tremendous opportunity for cell and tissue engineering. Acknowledgment. The authors thank Dr. Dorian Liepmann, Julia Chu, Maelene Wong, Pui Leng Leong, Jonathan Jung, Daniel Kim, and Amy Lam for technical assistance. This work was supported by research grants from the 541
National Institutes of Health (HL078534) and the Hellman Fund to S.L. N.H. was funded by graduate fellowships from the Whitaker Foundation (2003) and National Science Foundation (2004-2006). The Stony Brook group is grateful for the support of an NIH-SBIR Phase II grant (GM6328302) administered by the Stonybrook Technology and Applied Research, Inc. Supporting Information Available: A description of the materials and methods and SEM images of myotubes on nanofibrous scaffolds after 7 days and fabrication and characterization of micropatterned biodegradable PLGC polymer. This material is available free of charge via the Internet at http:/pubs.acs.org. References (1) Raghavan, S.; Chen, C. S. Micropatterned environments in cell biology. AdV. Mater. 2004, 16 (15), 1303-1313. (2) Wigmore, P. M.; Dunglison, G. F. The generation of fiber diversity during myogenesis. Int. J. DeV. Biol. 1998, 42 (2), 117-125. (3) Kim, K.; Yu, M.; Zong, X.; Chiu, J.; Fang, D.; Seo, Y. S.; Hsiao, B. S.; Chu, B.; Hadjiargyrou, M. Control of degradation rate and hydrophilicity in electrospun nonwoven poly(D, L-lactide) nanofiber scaffolds for biomedical applications. Biomaterials 2003, 24 (27), 4977-4985. (4) Zong, X.; Bien, H.; Chung, C. Y.; Yin, L.; Fang, D.; Hsiao, B. S.; Chu, B.; Entcheva, E. Electrospun fine-textured scaffolds for heart tissue constructs. Biomaterials 2005, 26 (26), 5330-5338. (5) Katta, P.; Alessandro, M.; Ramsier, R. D.; Chase, G. G. Continuous electrospinning of aligned polymer nanofibers onto a wire drum collector. Nano Lett. 2004, 4 (11), 2215-2218. (6) Li, D.; Ouyang, G.; McCann, J. T.; Xia, Y. Collecting electrospun nanofibers with patterned electrodes. Nano Lett. 2005, 5 (5), 913916. (7) Zong, X.; Ran, S.; Kim, K. S.; Fang, D.; Hsiao, B. S.; Chu, B. Structure and morphology changes during in vitro degradation of electrospun poly(glycolide-co-lactide) nanofiber membrane. Biomacromolecules 2003, 4 (2), 416-423. (8) Li, D.; Xia, Y. Electrospinning of nanofibers: reinventing the wheel? AdV. Mater. 2004, 16 (4), 1151-1170. (9) Boland, E. D.; Matthews, J. A.; Pawlowski, K. J.; Simpson, D. G.; Wnek, G. E.; Bowlin, G. L. Electrospinning collagen and elastin: preliminary vascular tissue engineering. Front. Biosci. 2004, 9, 14221432.
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