Growth of Mesenchymal Stem Cells on Electrospun Type I Collagen Nanofibers - 《stemcells》干细胞学杂志干细胞之家



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发表于 2009-3-5 00:00 |只看该作者 |倒序浏览 |打印

作者：Yu-Ru V. Shiha,b, Chung-Nan Chenc, Shiao-Wen Tsaid, Yng Jiin Wange, Oscar K. Leea,b,f,g作者单位：aDepartment of Orthopaedics and Traumatology, Taipei Veterans General Hospital, Taipei, Taiwan;bInstitute of Biopharmaceutical Sciences, National Yang-Ming University, Taipei, Taiwan;cBiomedical Engineering Research Laboratory, Industrial Technology Research Institute, Hsinchu, Taiwan;dInstitute of  ) b( H( I& ^* u# H


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      【摘要】
   We reconstituted type I collagen nanofibers prepared by electrospin technology and examined the morphology, growth, adhesion, cell motility, and osteogenic differentiation of human bone marrow-derived mesenchymal stem cells (MSCs) on three nano-sized diameters (50¨C200, 200¨C500, and 500¨C1,000 nm). Results from scanning electron microscopy showed that cells on the nanofibers had a more polygonal and flattened cell morphology. MTS (3--2H-tetrazolium compound) assay demonstrated that the MSCs grown on 500¨C1,000-nm nanofibers had significantly higher cell viability than the tissue culture polystyrene control. A decreased amount of focal adhesion formation was apparent in which quantifiable staining area of the cytoplasmic protein vinculin for the 200¨C500-nm nanofibers was 39% less compared with control, whereas the area of quantifiable vinculin staining was 45% less for both the 200¨C500-nm and 500¨C1,000-nm nanofibers. The distances of cell migration were quantified on green fluorescent protein-nucleofected cells and was 56.7%, 37.3%, and 46.3% for 50¨C200, 200¨C500, and 500¨C1,000 nm, respectively, compared with those on the control. Alkaline phosphatase activity demonstrated no differences after 12 days of osteogenic differentiation, and reverse transcription-polymerase chain reaction (RT-PCR) analysis showed comparable osteogenic gene expression of osteocalcin, osteonectin, and ostepontin between cells differentiated on polystyrene and nanofiber surfaces. Moreover, single-cell RT-PCR of type I collagen gene expression demonstrated higher expression on cells seeded on the nanofibers. Therefore, type I collagen nanofibers support the growth of MSCs without compromising their osteogenic differentiation capability and can be used as a scaffold for bone tissue engineering to facilitate intramembranous bone formation. Further efforts are necessary to enhance their biomimetic properties.
      【关键词】 Mesenchymal stem cells Electrospin Nanofibers Type I collagen Focal adhesion7 r  W: y8 g) \; w% [; `0 K
   INTRODUCTION

Tissue engineering emerges as an interdisciplinary field that employs the doctrines of engineering and life sciences toward the development of biological substitutes that restore, maintain, or improve tissue function . Electrospinning is a process that can produce polymer fibers with diameters ranging several orders of magnitude, from the micrometer range typical of conventional fibers down to the nanometer range, while providing pores for cell in-growth. Owing to their small diameters, electrospun fibers possess very high surface-to-area ratios and are expected to display morphologies and material properties very different from their conventional counterparts.' L6 ~' Y4 f) e: J8 y3 I" [" F
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Adult human mesenchymal stem cells (hMSCs) isolated from the bone marrow have been shown to differentiate into a variety of mesodermal cell lineages in vitro, including adipocytes, chondrocytes, myocytes, and osteoblasts .6 t- W3 n; z) |( T

The purpose of this study is to evaluate MSC adhesion, growth, and differentiation on an electrospun, three-dimensional (3D) nanofibrous type I collagen scaffold that shares closer resemblance to the native ECM than do two-dimensional (2D) culture systems.
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MATERIALS AND METHODS5 x) q3 u5 J" h
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Isolation and Culture of Bone Marrow hMSCs

Isolation of bone marrow hMSCs was achieved from a method described previously . Briefly, human bone marrow was aspirated from the iliac crest of healthy donors with informed consent. Mononuclear cells were obtained with a commercially available kit (RosetteSep; StemCell Technologies, Vancouver, BC, Canada, http://www.stemcell.com) according to the manufacturer's instructions, and nonadherent cells were washed away. Single cell-derived, clonally expanded cells were subsequently obtained by limiting dilution and maintained in an expansion medium consisting of Iscove's modified Dulbecco's medium (IMDM) (Invitrogen, Carlsbad, CA, http://www.invitrogen.com) and 10% fetal bovine serum (FBS) (HyClone, Logan, UT, http://www.hyclone.com), supplemented with 10 ng/ml basic fibroblast growth factor and 100 units of penicillin, 1,000 units of streptomycin, and 2 mmol/l L-glutamine (Invitrogen). The cultures were maintained in an incubator at 37¡ãC with 5% CO2. These isolated bone marrow-derived cells were characterized and reported previously. Their surface phenotype was negative for CD7, CD14, CD16, CD19, CD33, CD34, CD38, CD45, CD127, CD133, CD135, and HLA-DR. On the other hand, they were positive for CD29, CD44, CD73, CD90, CD105, CD166, HLA-ABC, and MSC-specific antigen SH-2 (data not shown). The ability to differentiate into mesenchymal lineages, including osteoblasts, chondrocytes, and adipocytes, was demonstrated before the MSCs were used for further experiments (data not shown).
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Reconstitution and Fabrication of Type I Collagen Nanofibers8 _5 E) c1 e+ n% x
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Reconstitution of type I collagen was accomplished with slight modifications to a method in the literature . Briefly, type I collagen was prepared from bovine skin that was lipid-depleted by a mixture of chloroform and methanol, then soaked with 0.5 M acetic acid, and added with pepsin. The insoluble material was removed by centrifugation, and NaCl was added to the remaining supernatant; the material was incubated overnight, centrifuged again to obtain a precipitate that was dissolved, and removed of salt with 0.5 M acetic acid. The purified collagen was stored in acetic acid at 4¡ãC.

Type I collagen was constructed into nanofibers by an electrospin process described in previous literature, with some modifications -carbodiimide) and eventually formed fiber diameters in the range of 50¨C200, 200¨C500, and 500¨C1,000 nm, depending on the concentration of type I collagen.

Cell Seeding and In Vitro Differentiation( `6 U  ~8 X! c/ S% A( n+ k- |/ M

Prior to cell seeding, the nanofibers were sterilized by ultraviolet irradiation overnight for both top and bottom surfaces in a laminar flow hood. The nanofibers were then placed in dishes or plates (Becton, Dickinson and Company, Franklin Lakes, NJ, http://www.bd.com) when cells were seeded for culture. Initial cell seeding density was at 3,000 cells per cm2, and only an adequate volume of culture medium was used initially to suspend cells so that when dropped onto the nanofibers it would not overflow the edges of the surface of the coated nanofibers due to the effects of surface tension. Once the seeded cells had adhered onto the nanofiber surface, inert metal rings were settled on the nanofibers to prevent it from detaching from its glass base.( W# Z- z8 M  `8 c' T8 p6 y/ M
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To induce osteogenic differentiation, we treated cells with osteogenic medium for 2 or 3 weeks and changed the medium twice each week. Osteogenic medium consists of IMDM supplemented with 0.1 mM dexamethasone (Sigma-Aldrich), 10 mM ß-glycerol phosphate (Sigma-Aldrich), and 0.2 mM ascorbic acid (Sigma-Aldrich).% z# w3 b0 |" M3 @" \. T/ U! D$ f5 I
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Scanning Electron Microscopy
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Samples of different diameters with or without cells were fixed in 10% formalin, dehydrated through increasing concentrations of ethanol, and dried overnight in a dessicator. Dehydrated samples were mounted on aluminum stubs, sputter-coated with gold-palladium, and examined in a scanning electron microscope (model S-800; Hitachi Software Engineering, Yokohama, Japan, http://www.hitachi-soft.com) at an accelerating voltage of 15 kV.
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Cell Viability Assay' s/ y; p& w- j  j
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Cell viability was evaluated using an MTS (3--2H-tetrazolium compound) assay (Promega Corporation, Madison, WI, http://www.promega.com). Cells were seeded onto each collagen nanofiber with diameters in the range of 200¨C500 or 500¨C1,000 nm and cultured in 24-well plates. After 7 days, the culture medium was washed away, and 60 ml of MTS solution was added to each well containing 300 ml of IMDM. Cells were then incubated in the dark at 37¡ãC for 4 hours. One-hundred twenty milliliters of each sample solution was transferred to a well of 96-well plates to measure the absorbance of the formazan product at an optical density (OD) of 490 nm.
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Fluorescence Staining

For focal adhesion studies, cells grown on nanofibers for 48 hours were washed with PBS and fixed in 3.7% formaldehyde (Sigma-Aldrich) for 15 minutes at room temperature and then permeabilized with 1% Triton X-100 (Sigma-Aldrich) for 5 minutes. After several washes, cells were blocked with 5% FBS for 1 hour at 37¡ãC and then incubated with mouse primary antibodies against human vinculin (1:400; Sigma-Aldrich) for 40 minutes at 37¡ãC. After washing, cells were incubated with cyanine (Cy)-2-labeled goat anti-mouse immunoglobulin G secondary antibody for 1 hour (1:100; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, http://www.scbt.com). F-actin was incubated simultaneously with Cy-2 antibody using Alexa Fluor 488 phalloidin (1:300; Invitrogen). Image quantification of vinculin area and cell area was accomplished by ImageJ software.: S2 }$ P, B* k
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Nucleofection of Cells with Green Fluorescent Protein

Cells were nucleofected with green fluorescent protein (GFP) according to the manufacturer's instructions (Amaxa Biosystems, Cologne, Germany, http://www.amaxa.com). Briefly, 4.5 x 105 cells were suspended in 100 µl of hMSC nucleofector solution, mixed with 2 mg of pmaxGFP plasmid DNA provided by the manufacturer, and transfected using program U-23 (high-transfection efficiency) of the nuclefector device.5 _0 B6 h" q6 s1 x" H8 d9 L

Time-Lapse Microscopy* ~6 t) s3 [  \/ O

Cells were placed under a time-lapse microscope (model IX71; Olympus, Tokyo, http://www.olympus-global.com) in a chamber with 5% CO2, and a fluorescent photo with wideband filters capturing wavelengths of 510 nm and higher was taken at 10-minute intervals for a total of 4 hours with Olympus DP Controller software. The distance of linear cell migration was quantified with ImageJ software (NIH, Bethesda, MD, http://www.rsb.info.nih.gov/ij).
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Alkaline Phosphatase Activity Assay, G# \2 F3 z% s) U

Osteogenic media were washed away, and cells on collagen nanofibers placed in 48-well plate were lysed with 0.2 ml of 0.05% SDS lysis buffer and incubated at 37¡ãC for 10 minutes. Without washing, 0.6 ml of a substrate made with equal volumes of 16 mM 4-nitrophenyl phosphate disodium salt hexahydrate (Sigma-Aldrich) and 2-amino-2-methyl-1-propanol (pH 10) mixed with 0.2% 1 M MgCl2 was added into the lysed cells to react for 30 minutes in the dark at 37¡ãC. The reaction was stopped with 0.6 ml of 0.02 M NaOH. One-hundred microliters of solution per well of 48-well was transferred into a 96-well plate, and absorbance was measured at an OD of 405 nm. For the normalization of alkaline phosphatase (ALP), another set of cells was cultured under the same conditions but was measured by MTS assay at an OD of 490 nm.

Total RNA Extraction and Reverse Transcription¨CPolymerase Chain Reaction+ O9 N  u" Y# w$ [+ I3 e
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Total RNA was extracted from 3 x 105 differentiated cells using RNeasy Kit (Qiagen Inc., Valencia, CA, http://www1.qiagen.com) according to the manufacturer's instructions. The concentration of RNA samples was quantified using a spectrophotometer (Eppendorf, Hamburg, Germany, http://www.eppendorf.com) at OD 260/280, and RNA samples were reverse-transcribed using reagents (Genemark Technology, Taiwan, http://www.genemark.com.tw) according to the manufacturer's instructions. cDNA was amplified after initial denaturation at 94¡ãC for 2 minutes using a Nucleic Acid Purification/Amplification Kit (Genemark Technology) at 94¡ãC for 1 minute, 61.9¡ãC for 1 minute, and 72¡ãC for 2 minutes for 32 cycles. Primers used for amplification were osteocalcin: ACATCTATCCGGGAGGAAATC (sense), CTGGCGGTCTCCTCACTC (antisense); osteonection: AGGTATCTGTGGGAGCTAATC (sense), ATTGCTGCACACCTTCTC (antisense); osteopontin: GACCTGACATCCAGTACCC (sense), GTTTCAGCACTCTGGTCATC (antisense); and glyceraldehyde-3-phosphate dehydrogenase (GAPDH): GAGTCCACTGGCGTCTTC (sense), GACTGTGGTCATGAGTCCTTC (antisense). For single-cell reverse transcription-polymerase chain reaction (RT-PCR), 1,000 cells were suspended in 1 ml of phosphate-buffered solution and single cells were isolated under a light microscope (Nikon Corporation, Tokyo, Japan, http://www.nikon.com). Cells were immediately lysed at two freeze-thaw cycles with liquid nitrogen. RT-PCR was accomplished with a one-step RT-PCR kit RobusT II (Finnzymes, Espoo, Finland, http://www.finnzymes.fi), and cDNA was amplified for 41 cycles under the same amplification conditions mentioned above. The primer sequences used for type I collagen were GTGATGCTGGTGCTAAAGG (sense) and GGTCCAGCATTTCCAGAG (antisense).
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Statistical Analysis- m2 y) n5 Z& J+ X; I# V$ g( v

All values were expressed as mean ¡À SD and compared using Student's two-tailed t test or analysis of variance (Tukey's post hoc test) with p
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RESULTS1 k! d" W% R+ n/ q3 d

Characterization of Type I Collagen Nanofibers

Scanning electron photomicrographs (Fig. 1) revealed a 3D, nonwoven nanofibrous type I collagen that had interconnected pores and was fabricated into three sets of diameters ranging from 50¨C200 nm (Fig. 1A) and 200¨C500 nm (Fig. 1B) to 500¨C1,000 nm (Fig. 1C). Although the nanofiber diameters for each type I collagen concentration had a range of thickness distribution for different individual fibers, each individual fiber still possessed a uniform thickness along its entire length.

Figure 1. Scanning electron microscopy micrographs of type I collagen nanofibers electrospun at 30 kV with fiber diameters of 50¨C200 nm (A), 200¨C500 nm (B), and 500¨C1,000 nm (C).

Cell Morphology on Type I Collagen Nanofibers
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After 3 days of culture, cells displayed a more polygonal and flattened cell morphology on the 500¨C1,000-nm nanofiber (Fig. 2A, 2B) compared with those on tissue culture polystyrene (TCPS) (Fig. 2C, 2D). Interestingly, cells on the nanofibers also displayed higher numbers of filopodia-like thin protrusions, and cell contours in proximity to the nanofibers were extended toward and along the lengths of the nanofibers. Figures 1D¨C1F show cells on 50¨C200-, 200¨C500-, and 500¨C1,000-nm nanofibers, respectively, viewed with immunofluorescent staining at x630 magnification using a laser confocal microscope (Leica Microsystems, Wetzlar, Germany, www.leica-microsystems.com).

Figure 2. Scanning electron microscopy photomicrograph of cells on 500¨C1,000-nm nanofibers (A, B) and tissue culture polystyrene (C, D). Representative confocal microscopy of cell morphology of nanofibers with diameters of 50¨C200 nm (E), 200¨C500 nm (F), and 500¨C1,000 nm (G). Nanofibers with diameters of 500¨C1,000 nm were stained with rhodamine (red). Magnifications: x1,500 (A, D), x3,500 (B, C), x630 (E¨CG).
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Effect of Nanofibers on MSC Viability
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To determine whether the various diameters of nanofibers had a significant effect on the growth of MSCs, an MTS assay was carried out after 7 days of cell culture on nanophase collagen fibers. As shown in Figure 3, mean cell viability of MSCs grown on the 200¨C500-nm collagen was 30% greater whereas that of cells on 500¨C1,000-nm collagen was 67% greater compared with TCPS surfaces. Results are from triplicate assays measured at OD 490 (p " s* s% Z0 b* v+ M; @

Figure 3. 3-(4.5-Dimethythiazol-2-yl)-5-(3-carboxy-methoxyphenyl)-2-(4-sul-fophenyl)-2H-tetrazolium compound assay of MSC viability on different diameters (200¨C500 and 500¨C1,000 nm) of type I collagen nanofibers. Statistical analysis revealed a difference between the 500¨C1,000-nm nanofibers and the polystyrene control on the viability of MSCs (*p

Focal Adhesion Quantification

Relative focal adhesion formation was performed by quantifying the fluorescent signals of vinculin and F-actin obtained from immunofluorescent microscopy (Fig. 4). Cells cultured on the 200¨C500-nm, 500¨C1,000-nm, and polystyrene surfaces all displayed bright green vinculin spots (white arrows, Fig. 4A, 4C, 4E), corresponded with its red F-actin stains (Fig. 4B, D, 4F) on the same image under a different excitation wavelength. All quantified units are displayed as pixels drafted from the image. The number of individual vinculin stains was counted and quantified by normalizing to relative cell area from five random replicates (Fig. 4G). Both cells grown on 200¨C500-nm and 500¨C1,000-nm collagen displayed a reduced number of vinculin stains, which was 39% less compared with cells on TCPS, in which the 200¨C500-nm group was statistically significant (p
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Figure 4. Immunofluorescent microscopy for staining of vinculin and F-actin. Vinculin (A, C, E) is stained as bright green spots (white arrows) corresponding to red F-actin stain (B, D, F) of the same cells on 200¨C500-nm (A, B), 500¨C1,000 nm (C, D), and tissue culture polystyrene (E, F). Fewer vinculin stains are visually identified by cells grown on the nanofibers compared with control. (G): The 200¨C500-nm and the 500¨C1,000-nm groups had a significantly lower amount of vinculin (*p
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Distance of Cell Migration

Time-lapse microscopy of cells on TCPS (Fig. 5A), 50¨C200-nm (Fig. 5B), 200¨C500-nm (Fig. 5C), and 500¨C1,000-nm (Fig. 5D) nanofibers during a 4-hour period was taken, and linear distance of cell migration was quantified. Cell migration was 56.7% (p = .035), 37.3% (p = .001), and 46.3% (p = .006) higher on the 50¨C200-nm, 200¨C500-nm, and 500¨C1,000-nm nanofibers than on TCPS, respectively (Fig. 5E).

Figure 5. Time-lapse microscopy of green fluorescent protein-nucleofected cells on tissue culture polystyrene (TCPS) (A), 50¨C200-nm (B), 200¨C500-nm (C), and 500¨C1,000-nm (D) nanofibers represented by overlap of images from the start (green cells) to the end (red cells) of cell migration during the 4-hour observation period. (E): Linear distance of cell migration was quantified and was 56.7%, 37.3%, and 46.3% for cells on 50¨C200-nm, 200¨C500-nm, and 500¨C1,000-nm nanofibers, respectively, compared with those on TCPS (n = 9). Same letters correspond to same level of significance.1 @/ L, K2 H0 a( g) K' H- v/ b: T
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ALP Activity

Cells grown on nanofibers demonstrated no difference in ALP production after 4 days of differentiation. Differences were seen after 8 days between ALP production by cells on nanofibers and those cultured on polystyrene (Fig. 6; n = 3). Cells coated on 50¨C200-nm collagen showed lower ALP activity (only 56%) when compared with cells on polystyrene. Similar results could be observed for the 200¨C500-nm collagen with a reduced production in ALP (only 63%). However, ALP production between the nanofiber groups and polystyrene had no difference by the 12th day of differentiation.+ ~) Z( Z( I; J2 V% [. V
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Figure 6. Alkaline phosphatase (ALP) assay after 12 days of osteogenic induction on type I collagen nanofibers with diameters of 50¨C200 and 500¨C1,000 nm. No noticeable differences in ALP activity are observed for both the 50¨C200-nm and 500¨C1,000-nm nanofibers (*p ( a5 @; B  V" Z* E
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Gene Expression of MSCs Differentiated into the Osteoblast Lineage% b7 {6 g1 W% N8 A0 M
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Osteoblast-lineage gene expression was detected using RT-PCR analysis for osteogenic genes after differentiation (Fig. 7). After induction for 2, 3, and 4 weeks, cells seeded on all diameters of collagen nanofibers revealed extremely similar expression of osteocalcin, osteopontin, and osteonectin compared with cells induced on polystyrene (Fig. 7A). After 2 weeks of induction, single-cell gene expressions of type I collagen of six individual cells seeded on the nanofibers demonstrated various levels of expression but were overall higher than those on TCPS (Fig. 7B). GAPDH gene expression of another set of six individual cells was shown as a sensitivity control between individual cell samples (Fig. 7C). The relative GAPDH gene expression was similar among the six individual cells, which demonstrated low errors in experimental techniques among different samples.4 k. H3 J; P8 c5 _3 a6 a, e
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Figure 7. Reverse transcription-polymerase chain reaction (RT-PCR) analysis of osteogenic gene expression. (A): Comparison among cells seeded on tissue culture polystyrene, 50¨C200-nm 200¨C500-nm, and 500¨C1,000-nm nanofibers after 2, 3, and 4 weeks of induction reveals similar effects on osteoblast gene expression. (B): Single-cell RT-PCR of six individual cells on each nanofiber diameter revealed a higher level of type I collagen gene expression on the nanofibers than on tissue culture polystyrene. (C): Sensitivity control of six different individual cells demonstrated by GAPDH gene expression. Abbreviation: GAPDH, glyceraldehyde-3-phosphate dehydrogenase.
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DISCUSSION
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Extensive efforts have been made to develop scaffolds able to support and promote tissue growth for clinical applications. In this study, we have evaluated the adhesion, growth, motility, and differentiation of hMSCs on electrospun type I collagen nanofibers of different diameters. It was found that cell morphology pictured by scanning electron microscopy substantiates the presumption that the structural properties of the random, nanofibrous ECM alter the cytoskeleton and architecture of the cells and may have exerted an effect on cellular physiology. Higher cell viability suggests that under our fabrication standards, the type I collagen nanofibers provide favorable growth conditions and survival for MSCs. This appears to be advantageous if low numbers of cells are initially available for cell therapy in which the scaffold may aid in the viability and expansion of cells and enhance the rate of tissue formation.
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Studies on cell adhesion were performed by immunofluorescent staining for the formation of vinculin-associated focal adhesion and focal complexes. Quantifying the amount and area . Whether and how the altered mechanical properties resulting from the nanofibers affect focal adhesion and cellular response need to be investigated further.
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The nanofibers also supported osteogenic differentiation in which cellular ALP production was apparent and similar to that on TCPS after 12 days. Gene expression analysis of MSCs after osteogenic induction demonstrated similar or higher levels of osteoblast-lineage RNA transcript production between the nanofiber and TCPS at a single-cell or multiple-cell level, indicating that the nanofibers support intrinsic properties of MSC differentiation. The results from cytocompatible evaluations of the fabricated electrospun type I collagen nanofibers¡ªcombined with multipotent cells such as MSCs as a source for cell therapy and tissue engineering¡ªprovide substantial potential in the treatment of patients with intramembranous bone injuries such as craniofacial reconstructions. A recent study reported that nanofibrous scaffold fabricated from PCL was able to support MSC differentiation into osteoblastic, chondrogenic, and adipogenic lineages . It would therefore be interesting to compare the growth and differentiation properties of MSCs on nanofibrous scaffolds made from type I collagen and PCL.

SUMMARY" X( l) p3 ]  w! ~' O$ K2 p

This study reports a biomimetic 3D nanofibrous scaffold capable of supporting the in vitro adhesion, growth, and differentiation of MSCs into the osteoblast lineage. However, further work will be carried out to modify the nanofibers for improved cellular interactions and reconstitute a scaffold that could enhance its biomimetic properties.) k/ S. |* j. r9 p
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DISCLOSURES

The authors indicate no potential conflicts of interest.

ACKNOWLEDGMENTS

This work was supported by grants from the National Science Council, Taiwan (NSC 94-2120-M-010-001), and from Taipei Veterans General Hospital, Taiwan (VGH 94-365-13).
      【参考文献】2 b; @. x1 C% d$ x% W3 b8 D+ I

Langer R, Vacanti JP. Tissue engineering. Science 1993;260:920¨C926.
7 u+ W: A$ ~: w0 T$ k5 Y+ S/ y
Ma Z, Kotaki M, Inai R et al. Potential of nanofiber matrix as tissue-engineering scaffolds. Tissue Eng 2005;11:101¨C109.

Pittenger MF, Mackay AM, Beck SC et al. Multilineage potential of adult human mesenchymal stem cells. Science 1999;284:143¨C147.: r2 ?5 s' H- s3 [. [$ w

Prockop DJ. Marrow stromal cells as stem cells for nonhematopoietic tissues. Science 1997;276:71¨C74.
7 v# A7 N3 h* \- d
Maitra B, Szekely E, Gjini K et al. Human mesenchymal stem cells support unrelated donor hematopoietic stem cells and suppress T-cell activation. Bone Marrow Transplant 2004;33:597¨C604.$ p/ ^4 `- c# t5 f
* b6 B" |. ?  W4 Q
Le Blanc K, Ringden O. Immunobiology of human mesenchymal stem cells and future use in hematopoietic stem cell transplantation. Biol Blood Marrow Transplant 2005;11:321¨C334.

Wnek GE, Carr ME, Simpson DG et al. Electrospinning of nanofiber fibrinogen structures. Nano Lett 2003;3:213¨C216., j: o7 ^' Q# p0 d) A# s6 ~0 ^

Jin HJ, Chen JS, Karageorgiou V et al. Human bone marrow stromal cell responses on electrospun silk fibroin mats. Biomaterials 2004;25:1039¨C1047.
# ?9 H1 w% r( B! [7 q) d
Huang L, McMillan RA, Apkarian RP et al. Generation of synthetic elastin-mimetic small diameter fibers and fiber networks. Macromolecules 2000;33:2989¨C2997.

Xie JB, Hsieh YL. Ultra-high surface fibrous membranes from electrospinning of natural proteins: Casein and lipase enzyme. J Mater Sci 2003;38:2125.

Fang X, Reneker DH. DNA fibers by electrospinning. J Macromol Sci Phys B 1997;36:169.
" T6 }# K4 m0 K/ W* P& L- P/ O
Matthews JA, Wnek GE, Simpson DG et al. Electrospinning of collagen nanofibers. Biomacromolecules 2002;3:232¨C238.
. ~/ v" B5 U4 K5 @& G
Huang L, Nagapudi K, Apkarian RP et al. Engineered collagen¨CPEO nanofibers and fabrics. J Biomater Sci Polym Ed 2001;12:979¨C993.

Yang F, Murugan R, Wang S et al. Electrospinning of nano/micro scale poly(L-lactic acid) aligned fibers and their potential in neural tissue engineering. Biomaterials 2005;26:2603¨C2610.

Li WJ, Tuli R, Huang X et al. Multilineage differentiation of human mesenchymal stem cells in a three-dimensional nanofibrous scaffold. Biomaterials 2005;26:5158¨C5166.7 L; h8 X. E1 g  a7 U

Yoshimoto H, Shin YM, Terai H et al. A biodegradable nanofiber scaffold by electrospinning and its potential for bone tissue engineering. Biomaterials 2003;24:2077¨C2082.

Li M, Mondrinos MJ, Gandhi MR et al. Electrospun protein fibers as matrices for tissue engineering. Biomaterials 2005;26:5999¨C6008.2 l  m- K  _0 C  x0 g3 h& L  z0 w

Lee OK, Ko YC, Kuo TK et al. Fluvastatin and lovastatin but not pravastatin induce neuroglial differentiation in human mesenchymal stem cells. J Cell Biochem 2004;93:917¨C928.9 o7 C6 c6 r1 E% G! d- v5 x
/ v; L8 h6 D1 s+ H3 M
Lee KD, Kuo TK, Whang-Peng J et al. In vitro hepatic differentiation of human mesenchymal stem cells. Hepatology 2004;40:1275¨C1284.$ E$ I- W, U- v( R5 h- K3 p- D

Lan CW, Wang FF, Wang YJ et al. Osteogenic enrichment of bone-marrow stromal cells with the use of flow chamber and type I collagen-coated surface. J Biomed Mater Res A 2003;66:38¨C46.1 |2 s2 T5 f+ d% p* c" q
: L9 X1 q& q4 A; F3 s6 j+ N! l
Mo XM, Xu CY, Kotaki M et al. Electrospun P(LLA-CL) nanofiber: A biomimetic extracellular matrix for smooth muscle cell and endothelial cell proliferation. Biomaterials 2004;25:1883¨C1890.

Hunter A, Archer CW, Walker PS et al. Attachment and proliferation of osteoblasts and fibroblasts on biomaterials for orthopaedic use. Biomaterials 1995;16:287¨C295.; F& O" d" W% v

Katz BZ, Zamir E, Bershadsky A et al. Physical state of the extracellular matrix regulates the structure and molecular composition of cell-matrix adhesions. Mol Biol Cell 2000;11:1047¨C1060.

Peterson L, Burridge K. Focal adhesions and focal complexes. In: Beckerle MC, ed. Cell Adhesion.Oxford, UK: Oxford University Press,2002;282¨C323.# t  l* V' d( u& N$ F

Cukierman E, Pankov R, Stevens DR et al. Taking cell-matrix adhesions to the third dimension. Science 2001;294:1708¨C1712.

Wang YK, Wang YH, Wang CZ et al. Rigidity of collagen fibrils controls collagen gel-induced down-regulation of focal adhesion complex proteins mediated by alpha2beta1 integrin. J Biol Chem 2003;278:21886¨C21892.+ D& ]2 s1 }0 w9 o* W% q( b, d
( B, W3 V: e7 L7 [8 W$ c
Ishii I, Tomizawa A, Kawachi H et al. Histological and functional analysis of vascular smooth muscle cells in a novel culture system with honeycomb-like structure. Atherosclerosis 2001;158:377¨C384.

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