Skip to main content
NCBI home page
Tissue Engineering. Part A logoLink to Tissue Engineering. Part A
. 2010 Jun 23;16(10):3219–3230. doi: 10.1089/ten.tea.2010.0004

Colonization and Osteogenic Differentiation of Different Stem Cell Sources on Electrospun Nanofiber Meshes

Yash M Kolambkar 1, Alexandra Peister 2, Andrew K Ekaputra 3, Dietmar W Hutmacher 4,,5, Robert E Guldberg 5,✉
PMCID: PMC2947421  PMID: 20504075

Abstract

Numerous challenges remain in the successful clinical translation of cell-based therapies for musculoskeletal tissue repair, including the identification of an appropriate cell source and a viable cell delivery system. The aim of this study was to investigate the attachment, colonization, and osteogenic differentiation of two stem cell types, human mesenchymal stem cells (hMSCs) and human amniotic fluid stem (hAFS) cells, on electrospun nanofiber meshes. We demonstrate that nanofiber meshes are able to support these cell functions robustly, with both cell types demonstrating strong osteogenic potential. Differences in the kinetics of osteogenic differentiation were observed between hMSCs and hAFS cells, with the hAFS cells displaying a delayed alkaline phosphatase peak, but elevated mineral deposition, compared to hMSCs. We also compared the cell behavior on nanofiber meshes to that on tissue culture plastic, and observed that there is delayed initial attachment and proliferation on meshes, but enhanced mineralization at a later time point. Finally, cell-seeded nanofiber meshes were found to be effective in colonizing three-dimensional scaffolds in an in vitro system. This study provides support for the use of the nanofiber mesh as a model surface for cell culture in vitro, and a cell delivery vehicle for the repair of bone defects in vivo.

Introduction

Bone is one of the few adult tissues with the capacity to regenerate. However, large, unstable, or infected bone defects remain a challenging clinical problem.1 Tissue engineering strategies that deliver cells, growth factors, and genetic material on scaffolds have demonstrated considerable potential in developing bone graft substitutes.2,3 Delivery of exogenous cells capable of forming bony tissue may be especially important to repair bone defects in patients with a limited endogenous progenitor cell supply, such as older patients, smokers, or patients with certain diseases.4 The success of cell-based therapies for bone regeneration has been limited, in part, by the inadequate availability of large quantities of osteogenic cells and an effective cell delivery system.

The identification of a cell source that may be easily harvested, expanded to large numbers, and controllably differentiated may be tremendously beneficial clinically for the reconstruction of damaged tissues. Bone-marrow-derived mesenchymal stem cells (MSCs) have demonstrated a strong potential for differentiation into bone-forming cells, and have been shown to promote repair of critically sized bone defects in preclinical animal studies.5–7 These cells are well suited for autologous transplantation, making them a feasible cell source for clinical deployment due to the lack of immunogenic issues associated with this transplantation modality. However, MSCs are associated with reduced mineralization capacity in older donors and following expansion to achieve therapeutic cell numbers.8,9

Amniotic fluid stem (AFS) cells are c-Kit expressing cells isolated from amniotic fluid that have demonstrated a high self-renewal capacity and the ability to differentiate into a diverse range of cell types, including those from the adipose, muscle, neuronal, cartilage, and bone lineages.10–13 Recently, our lab has demonstrated that these cells can produce robust mineralization in three-dimensional (3D) constructs in vitro and in vivo.14,15 Importantly, AFS cells have also shown a lack of senescence through 250 population doublings and display an absence of tumorigenicity in vivo.12 However, not much is known of their osteogenic potential compared to MSCs. A critical step toward clinical translation is the quantitative comparison of the proliferation and bone-forming capacity of different cell sources.

The delivery of stem cells to the site of injury, through either systemic introduction or local delivery, is another critical consideration for the success of cell-based therapies. Site-specific delivery has the advantage of being able to deliver large numbers of cells directly to the required area. In tissue engineering strategies, this typically involves placing cells on a 3D scaffold, followed by implantation at the injury site. However, the lack of initial vascularity at the center of a 3D scaffold limits the transport of nutrients to, and waste products from, the cells. This presents a very harsh environment that makes cell survival extremely difficult.16,17 An alternative is to deliver cells to the periphery of the defect via a thin membrane or scaffold. This delivery strategy may enhance cell survival by positioning the cells in proximity to the surrounding highly vascularized tissues, and thereby providing for nourishment and clearance of waste products.

Electrospinning has recently emerged as a technique to fabricate scaffolds for tissue engineering, with fiber diameters ranging from tens of nanometers to as large as 10 μm.18–20 The nanofiber mesh obtained by this process is a unique scaffold membrane that possesses structural features with a size scale similar to extracellular matrix (ECM) components, high porosity, and large surface-area-to-volume ratios. These properties allow for enhanced cellular attachment and spreading,21,22 and therefore nanofiber meshes may serve as an effective delivery vehicle for cells to a defect site in vivo. However, it is important to first evaluate their efficacy in supporting cell function and as a cell delivery vehicle in vitro. Although a few studies have investigated the osteogenic differentiation of progenitor cells on nanofiber meshes, more thorough analyses are needed to characterize the differentiation and mineralization process, and to quantify mineral deposition.23–25 In addition, nanofiber meshes may be utilized as an ECM-mimetic surface for evaluating cell behavior, and therefore serve as an improved in vitro cell culture system, compared to flat tissue culture plates.26

The aim of this study was to investigate the attachment, colonization, and osteogenic differentiation of human MSCs (hMSCs) and human AFS (hAFS) cells on electrospun nanofiber meshes. We demonstrate that electrospun meshes are able to robustly support these functions for both cell types. Compared to tissue culture plastic, there is delayed initial attachment and proliferation, but enhanced mineralization at a later time point. Differences in the kinetics of osteogenic differentiation were observed between hMSCs and hAFS cells. Cell-seeded nanofiber meshes were also effective in colonizing 3D scaffolds in an in vitro model. These results provide support to further evaluate the nanofiber mesh as a cell delivery vehicle for the repair of bone defects in vivo.

Experimental Procedures

Fabrication of nanofiber meshes

A polymer solution was made by dissolving 13% (w/v) poly (ɛ-caprolactone) (PCL; Sigma-Aldrich, St. Louis, MO) in a 40:60 volume ratio of dichloromethane:dimethylformamide (Sigma-Aldrich). PCL pellets were added to the solvent mixture, and gently stirred for 16–24 h. The polymer solution was loaded in a 3 mL syringe (Becton-Dickinson, Franklin Lakes, NJ), and a 22-gauge blunt stainless steel needle (Jensen Global, Inc., Santa Barbara, CA) was attached to the syringe end. The syringe was mounted on a syringe pump (Harvard Apparatus, Holliston, MA), and the pump was set to infuse at a rate of 0.75 mL/h. A flat, 6 × 6 inch copper plate (McMaster-Carr, Atlanta, GA) covered with aluminum foil was used to collect the fibers, and placed at a distance of 20 cm from the needle end. Fibers were electrospun for 50 min at a voltage of 14 kV, supplied by a high-voltage power supply (Gamma High Voltage Research, Ormond Beach, FL), to obtain a thin sheet of nanofiber mesh. To remove any residual solvent, the meshes were placed in a desiccators for at least 1 day before further use.

Nanofiber mesh morphology

The morphology of the nanofiber meshes was examined using a scanning electron microscope. A small piece of the dry nanofiber mesh was cut and mounted on a metal stub using double-sided adhesive tape. A thin layer of gold was then deposited on the mesh sample for 80 s using a sputter coater (Quorum Technologies, East Granby, CT). The gold-coated sample was then viewed under a Hitachi S-800 Field Emission scanning electron microscope (Hitachi HTA, Pleasanton, CA) with 10 kV accelerating voltage. The diameters of the fibers were quantified by analyzing the scanning electron microscopy images (at 7000 × magnification) using a custom MATLAB® (MATLAB® 7.0 R14; The MathWorks, Inc., Natick, MA) program. A total of at least 75 distinct fibers were measured from four randomly chosen locations.

Culture of AFS cells and MSCs

hAFS cells were kindly provided by Dr. Anthony Atala and Dr. Shay Soker at the Wake Forest Institute for Regenerative Medicine (Winston-Salem, NC). The isolation method and culture protocols have been described previously.10,12 Briefly, back-up human amniocentesis cultures were harvested by trypsinization, and subjected to c-Kit immunoselection. hAFS cells were subcultured routinely at a dilution of 1:4 to 1:8, and not permitted to expand beyond 70% confluence. The hAFS cells were passaged in alpha-minimum essential medium supplemented with 16% fetal bovine serum (embryonic stem cell [ESC]-qualified FBS), 100 U/mL penicillin, 100 μg/mL streptomycin, 2 mM L-glutamine (Invitrogen, Carlsbad, CA), 18% Chang B, and 2% Chang C (Irvine Scientific, Santa Ana, CA). In all experiments, cells were used at passages 16–17.

hMSCs derived from the bone marrow were obtained from the Tulane University Center for Gene Therapy (New Orleans, LA) at passage 1. Cells were isolated using bone marrow aspirates from the iliac crest of normal adult donors as previously described.27 For expansion, these cells were plated at a density of 50 cells/cm2, and cultured in the hMSC growth medium. The hMSC growth medium consisted of alpha-minimum essential medium (Invitrogen) supplemented with 16% FBS (Atlanta Biologicals, Atlanta, GA), 100 U/mL penicillin, 100 μg/mL streptomycin, and 2 mM L-glutamine (Invitrogen). The cells were subcultured once they reached a confluency of ∼70%. Passage 2–3 hMSCs were then used for all experiments.

Cell culture on nanofiber fiber meshes

Square (15 mm) samples were cut from nanofiber mesh sheets using scissors. Samples were placed in 24-well culture plates, submerged in 200 proof ethanol (Sigma-Aldrich), and sterilized by allowing the ethanol to evaporate overnight. After the samples had dried completely, they were prewetted with sterile 70% ethanol for 30 min. The 70% ethanol was then aspirated, and sterile dead weights were placed around the samples to prevent them from floating. The mesh samples were next rinsed three times with excess sterile phosphate-buffered saline (PBS; Mediatech, Inc., Manassas, VA). An 800 μL volume of the medium was placed in each well containing the samples. The control groups received the hMSC growth medium, whereas the osteogenic groups were further supplemented with 10 nM dexamethasone, 6 mM β-glycerol phosphate, 50 μg/mL ascorbic acid 2-phosphate, and 50 ng/mL L-thyroxine (Sigma-Aldrich). hMSCs and hAFS cells were then seeded onto nanofiber meshes in ∼200 μL of the hMSC medium such that the density of cells was 20,000 cells/cm2. Cells were also cultured in tissue culture treated 24-well plates at the same density for comparison. The medium was changed every 3–4 days, and the constructs were cultured for up to 4 weeks.

Cell viability

On days 1, 7, 14, and 28, the viability of the cells in the constructs was assessed using the Live/Dead® staining kit (Molecular Probes, Eugene, OR, Invitrogen). Harvested constructs were rinsed in PBS and incubated in 4 μM calcein-AM and 4 μM ethidium homodimer-1 for 45 min at room temperature. The samples were again rinsed in PBS, and images were obtained on a Zeiss LSM 510 confocal microscope (Carl Zeiss, Thornwood, NY). Green fluorescence of calcein-AM was detected by using a 488-nm Argon ion laser and a band pass 505–550 filter. Red fluorescence of ethidium homodimer-1 was detected by using a 543-nm helium–neon laser and a long pass 560 filter.

DNA content

Samples were harvested after 1, 7, 14, and 28 days to evaluate the construct cellularity, which was assessed by determining the DNA content. The cells were first lysed by freeze-thawing the constructs three times in 0.05% Triton X-100 (Sigma-Aldrich) with vigorous vortexing. To freeze the samples, they were placed in dry-ice-cooled methanol (Sigma-Aldrich) for 5 min. Samples were then thawed in a room-temperature water bath. The DNA amount in the lysate was quantified using the PicoGreen® dsDNA Quantitation Kit (Molecular Probes), and standardized using Lambda DNA solutions of known concentrations. A working solution of the PicoGreen reagent was made following the manufacturer's protocol, and incubated with experimental samples in the dark for 5 min at room temperature. The fluorescence was measured on a plate reader (HTS 7000; Perkins-Elmer, Waltham, MA) at an excitation of 485 nm and emission of 535 nm. All samples were run in triplicate, and the DNA content was normalized to the culture surface area of the samples.

Alkaline phosphatase activity

To determine the osteogenic differentiation of the cells on nanofiber meshes, the alkaline phosphatase (ALP) activity assay was performed. In this assay, the release of p-nitrophenol from p-nitrophenyl phosphate by the ALP enzyme is measured.28 The same lysate solution that was used to determine DNA content was used for this purpose. The ALP substrate working solution was made by mixing equal parts of 20 mM p-nitrophenyl phosphate, 1.5 M 2-amino-2-methyl-1-propanol (pH 10.25), and 10 mM MgCl2. The experimental samples were mixed with the freshly made substrate working solution, and incubated for 1 h at 37°C. The reaction was stopped by adding 1 N NaOH, and the absorbance was measured at 405 nm on a plate reader (PowerWave XS; Biotek, Winooski, VT). All samples were run in triplicate and compared to p-nitrophenol standards. The ALP activity was normalized by the incubation time and the amount of DNA obtained from the PicoGreen assay.

Calcium content

To quantify matrix mineralization, the calcium deposited by cells on nanofiber meshes after 28 days was determined using the Arsenazo III dye.29 Samples were vortexed with 1 N acetic acid overnight to extract the calcium into solution. The extract was mixed with the Arsenazo III reagent (Diagnostic Chemicals Limited, Oxford, CT) and incubated for 30 s at room temperature, and the absorbance read at 650 nm on a plate reader (PowerWave XS; Biotek). The samples were assayed in triplicate and compared to calcium chloride standards.

Calcein staining and quantification

For observation of the mineral deposited by the cells, the samples were stained using calcein on day 28.30,31 Briefly, a stock solution of 100 μg/mL calcein (Sigma-Aldrich) in PBS (pH 7.4) was added to the medium on top of the samples, such that the final concentration of calcein was 10 μg/mL. The samples were incubated in the calcein solution for 4 h in the incubator. After rinsing twice with PBS and fixing with 10% neutral-buffered formalin (EMD Chemicals, Gibbstown, NJ), samples were rinsed with excess of deionized water. The fluorescence of the samples was read on a fluorescence plate reader (HTS 7000; Perkins-Elmer) at an excitation of 485 nm and emission of 535 nm. After this, the same samples were imaged using an inverted microscope (Axio Observer.Z1; Carl Zeiss) and an FITC filter.

Fourier transform infrared spectroscopy

On day 28, constructs were also harvested for analyzing the chemical composition of the mineral deposited on the nanofiber meshes. Samples were dehydrated in 100% ethanol and dried at 50°C overnight. Acellular PCL nanofiber mesh was used as a negative control. After dehydration, the samples were cut into small pieces, mixed with potassium bromide (Sigma-Aldrich), and pressed into pellets using a custom built apparatus. Samples were analyzed with a Nicolet Nexus 470 FTIR spectrometer (Thermo-Nicolet, Madison, MI). Sixty-four scans were acquired at 4 cm−1 resolution under nitrogen purge.

Cell delivery by nanofiber mesh in vitro

The ability of a cell-seeded nanofiber mesh to serve as a cell delivery vehicle was studied using an in vitro model. AFS cells were seeded on to nanofiber mesh samples (15 × 10 mm) at a density of 200,000 cells/cm2. The cells were allowed to attach to the mesh overnight. On the following day, each cell-seeded mesh was wrapped around a cylindrical collagen scaffold (dry dimensions: 4 mm diameter and 9 mm length) aseptically, such that the cells were facing the scaffold (Fig. 7A). The scaffolds were punched from a fibrous collagen sheet (average pore size 61.7 μm, 93.7% pore volume; Kensey Nash Corporation, Exton, PA). The mesh was held in position by placing two interrupted silk sutures through the mesh and scaffold at the two ends of the scaffold. For comparison, we also seeded 300,000 cells throughout collagen scaffolds by pipetting the cell suspension directly in the scaffolds. There was no nanofiber mesh in this control group. The constructs were statically cultured in the hAFS cell growth medium. After 2 weeks, the mesh was taken off, following which the mesh and scaffold were stained with the Live/Dead staining kit (Molecular Probes, Invitrogen) to observe the cell migration into the scaffold. A confocal microscope (Zeiss LSM 510; Carl Zeiss) was used to take serial images to create 3D images.

FIG. 7.

FIG. 7.

Open in a new tab

Cell-seeded nanofiber meshes for in vitro delivery. (A) To investigate the use of nanofiber meshes for cell delivery, AFS cells were seeded on nanofiber meshes and wrapped around a three-dimensional (3D) collagen scaffold for 2 weeks in vitro. For comparison, cells were seeded throughout the scaffold. (B–E) Three-dimensional confocal images of the Live/Dead-stained scaffold and mesh. The projections of the 3D images are shown. The surface and top views are views of the 3D image looking from the top. The side view is the view of the 3D image looking from the side. The green box indicates the area and view being analyzed. (B) The collagen scaffold with cells seeded throughout had more cells on the exterior with numerous dead cells in the interior. (C) When a cell-seeded mesh was wrapped around the scaffold, cells migrated on to the peripheral surface of the scaffold and displayed high viability. (D) Top and cross-sectional surfaces of scaffold wrapped with cell-seeded mesh. Cells also colonized the top surface of the scaffold and migrated more than 500 μm into the scaffold from the mesh. (E) The seeded mesh was confluent with cells. Images were taken at 10 × magnification. Scale bar indicates 200 μm and applies to all images. Color images available online at www.liebertonline.com/ten.

Data analysis

Results are presented as mean ± standard error of the mean. Analysis of variance was performed on data, with pair-wise comparisons done using the Tukey multiple comparison procedure. A p-value of < 0.05 was considered significant. Residuals were used for diagnosing the appropriateness of the model by analyzing the constancy of error variance and normality of error terms.32 Wherever required, remedial measures were taken by transforming the data according to the Box-Cox procedure,33 or by using weighted least squares to make the error variance constant and the error distribution normal.32,34 Minitab® 15 (Minitab, Inc., State College, PA) was used to perform the statistical analysis.

Results

Morphology of nanofiber meshes

PCL nanofiber meshes were electrospun on a flat collector plate. The mesh formed a circular area of ∼8 cm diameter. The thickness of the mesh was found to vary with location, with the central areas thicker than the edges. The thin mesh samples from the edges were discarded and not used for cell culture. Fibers appeared to be smooth and uniform, with minimal bead formation (Fig. 1A, B). The quantification of the fiber diameter using a custom MATLAB® program demonstrated that the mean fiber diameter was 591 nm with a standard deviation of 199 nm. The fiber diameter histogram revealed that most of the fibers were between 300 and 900 nm, with the highest frequency occurring in the 500–600 nm range (Fig. 1C).

FIG. 1.

FIG. 1.

Open in a new tab

Nanofiber mesh morphology. Nanofiber meshes were electrospun from a 13% (w/v) poly (ɛ-caprolactone) (PCL) solution made in 40:60 dichloromethane:dimethylformamide. A scanning electron microscope was used to examine the morphology of the nanofibers. (A) Scanning electron microscopy image at low (1000 × ) magnification. (B) Scanning electron microscopy image at high (7000 × ) magnification. (C) The diameter of the fibers was quantified using a custom MATLAB® program, and the diameter distribution was plotted on a histogram. The mean diameter of the fibers was found to be 591 ± 199 nm.

Viability and colonization of hMSCs and hAFS cells over time

hMSCs and hAFS cells were seeded on electrospun nanofiber meshes and tissue culture plates, and cultured in the osteogenic medium for up to 28 days. The viability of the cells on the meshes was assessed on days 1, 7, 14, and 28 by the Live/Dead staining kit. The live cells are stained green, whereas the dead cells appear red. At the same time points, DNA from the samples was extracted and quantified to estimate the number of cells on the meshes, as well as in the culture wells. The Live/Dead images (Fig. 2) illustrate that both cell types attached to the nanofiber meshes by day 1 and were able to spread out by day 7. During days 7–14 the number of cells increased considerably, and by day 28, the cells were confluent on the meshes. The viability of both cell types on the meshes was found to be high, as seen by the extensive green stain, though a few dead cells were detected. No differences were observed in the viability and colonization between the two cell types.

FIG. 2.

FIG. 2.

Open in a new tab

Live/Dead staining. Human mesenchymal stem cells (hMSCs) and human amniotic fluid stem (hAFS) cells were seeded on nanofiber meshes and cultured in the osteogenic medium. On days 1, 7, 14, and 28, the viability of the cells on the meshes was assessed by imaging the constructs with a confocal microscope after staining with the Live/Dead® stain. Green, live cells; red, dead cells. Images were taken at 10 × magnification. Scale bar indicates 200 μm and applies to all images. Both cell types were able to attach, proliferate, and become confluent on the mesh. Color images available online at www.liebertonline.com/ten.

The DNA quantification over the 4-week culture period was used to compare the colonization kinetics of the two cell types on tissue culture plates and nanofiber meshes, respectively (Fig. 3). There was significant increase in DNA with time for both cell types, on plates as well as meshes, indicating cellular proliferation. On plates, the number of hAFS cells increased between days 1 and 7 but did not change significantly after that, suggesting rapid initial proliferation and confluency around day 7 (Fig. 3A). On the other hand, hMSCs increased in numbers between both days 1–7 and days 14–28. However, the later increase in DNA is because the hMSCs lift off the plate after confluence around day 7 to form a pellet and then repopulate the plate. This pelleting behavior was not seen with the hAFS cells for up to 28 days, though the hAFS cells do ultimately lift off the plate. The hMSC repopulation explains the increase in hMSC number between days 14 and 28 and the differences seen between hMSCs and hAFS cells on days 14 and 28. There were also significantly more hAFS cells than hMSCs on day 1, suggesting a higher initial attachment and/or proliferation rate.

FIG. 3.

FIG. 3.

Open in a new tab

DNA content. To evaluate sample cellularity, the DNA content was determined after cell lysis using the PicoGreen® reagent. Cells were cultured on (A) tissue culture plates and (B) nanofiber meshes. To compare the cellularity on nanofiber meshes with that on tissue culture plates, the data were plotted again for (C) hMSCs and (D) hAFS cells. DNA increased with time, indicating cellular proliferation. p < 0.05 is considered significantly different (*significantly different from the other cell type at same time point; #significantly different from the previous time point for same cell type).

The pelleting phenomenon does not occur on the nanofiber meshes, even at a later time points. There was a significant increase in DNA for both cell types between days 1–7, and an even higher increase between days 7 and 14, corresponding to the Live/Dead images (Fig. 3B). The number of cells did not change significantly after that, suggesting confluency of the cells on the nanofiber meshes. There were no significant differences in the DNA between hMSCs and hAFS cells at any time point.

Figure 3C and D compare the colonization kinetics of hAFS cells on the nanofiber meshes with that on the tissue culture plates. On day 1, there was significantly less DNA on the mesh compared to plates, suggesting that not all cells attach to the nanofibers within the first 24 h. The same trend was seen on day 7. However, by day 14, there was no significant difference between meshes and plates. On day 28, there was again no significant difference, though the lines crossed over. Although the hMSC pellet in the plates and repopulate the culture surface by day 28, the amount of DNA was not significantly different than that on the mesh.

Osteogenic differentiation of hMSCs and hAFS cells: ALP activity

The osteogenic differentiation of the cells was first investigated by analyzing the ALP activity of the cells (Fig. 4). ALP is a membrane-bound enzyme that hydrolyzes phosphate esters, which results in inorganic phosphate being available for incorporation into mineral deposits.35 There was significant increase in the ALP activity of both cell types with time, on plates as well on nanofiber meshes, suggesting osteogenic differentiation. On tissue culture plates, ALP activity peaked at day 7 for MSCs, whereas for AFS cells it increased slowly but continuously up to day 28 (Fig. 4A). Interestingly, the maximum ALP activity of the hAFS cells was greater than the hMSC peak. On the nanofiber meshes, hMSCs demonstrated a similar earlier rise in ALP activity on day 14 compared to day 28 for hAFS cells (Fig. 4B). The ALP activity of the hMSCs was significantly greater than that of hAFS cells on all time points other than day 1. The ALP response on meshes is delayed compared to that on the plate, as seen by the later increase in ALP activity. Interestingly, the ALP activity of hMSCs on meshes was sustained longer than that observed on plates, with the maximum value on meshes at day 28 greater than that on plates at day 7 (p < 0.05).

FIG. 4.

FIG. 4.

Open in a new tab

Alkaline phosphatase (ALP) activity. The osteogenic differentiation of the cells was evaluated by measuring the ALP activity of cell lysates on (A) tissues culture plates and (B) nanofiber meshes. ALP activity increased for both cell types with time, suggesting osteogenic differentiation. p < 0.05 is considered significantly different (*significantly different from the other cell type at same time point; #significantly different from a previous time point for same cell type; $significantly different from hMSC peak on plate at day 7).

Osteogenic differentiation of hMSCs and hAFS cells: Matrix mineralization

The osteogenic differentiation of the cells was further investigated by quantifying and staining the calcium deposits and by analyzing the chemical nature of the deposited mineral by Fourier transform infrared (FTIR) spectroscopy. An analysis of variance on the calcium deposited by the cells revealed that both cell type and the culture surface had a significant effect on the calcium levels (Fig. 5A). Under osteogenic stimulation, all groups demonstrated increased calcium deposition, compared to the growth medium, indicating that cells are able to differentiate to an osteoblastic phenotype on the surfaces. Calcium levels in the hMSC growth medium groups were negligible. hAFS cells deposited a higher amount of calcium than hMSCs on both plates and meshes. Also, both cell types deposited more calcium on meshes compared to plates.

FIG. 5.

FIG. 5.

Open in a new tab

Calcium quantification and Fourier transform infrared analysis. (A) The mineralization of the constructs was assessed by measuring the calcium deposited by cells. Both cell types deposited calcium in the osteogenic medium, indicating an osteoblast phenotype. p < 0.05 is considered significantly different (#significantly different from the growth medium; *significantly different from other cell type on same surface; $significantly different from plate with same cell type). (B) The chemical composition of the mineral was analyzed by Fourier transform infrared spectroscopy. The annotated peaks are the signature of a carbonate-containing, poorly crystalline hydroxyapatite, indicative of physiologic mineral. The remaining peaks are due to the PCL nanofiber mesh, as seen in the acellular mesh. Color images available online at www.liebertonline.com/ten.

FTIR spectroscopy was used to characterize the composition of the mineral that was deposited by the cells on the nanofiber meshes under osteogenic stimulation (Fig. 5B). To distinguish the peaks associated with the mineral from the peaks associated with the PCL mesh, an acellular piece of PCL mesh was also scanned. The cellular samples displayed amide I/II peaks at 1655 and 1550 cm−1, a carbonate peak at 870 cm−1, and a doublet split phosphate peak at 560 and 605 cm−1, which were not seen in the acellular mesh. There was also a peak at 1050 cm−1 in the cellular samples, but it overlapped with a PCL mesh peak. These peaks are the signature of a carbonate-containing, poorly crystalline hydroxyapatite, the form of mineral that is found in native bone. This suggests that both hMSCs and hAFS cells deposited mineral that possessed the characteristic bands of physiological mineral.

Samples were stained with calcein to observe the presence of calcium on the nanofiber meshes. The calcein staining demonstrated the presence of extensive calcium-containing mineral nodules, which were uniformly distributed on the meshes, as seen by the green fluorescence (Fig. 6A). This was the case in both the hMSC and hAFS cell osteogenic groups, whereas the growth medium groups stained minimally. Quantification of the fluorescence revealed that more mineral was deposited by the hAFS cells compared to the hMSCs (Fig. 6B), thus supporting the calcium data in Figure 5.

FIG. 6.

FIG. 6.

Open in a new tab

Calcein staining. Constructs were stained with calcein to observe the presence of the mineral deposits. (A) The osteogenic samples stained with calcein, illustrating that cells had deposited mineral uniformly on the nanofiber meshes. Images were taken at 10 × magnification. Scale bar indicates 200 μm and applies to all images. (B) The calcein staining was quantified by measuring the fluorescence using a plate reader. The data revealed greater mineralization by AFS cells. p < 0.05 is considered significantly different (*significantly greater from the MSC osteogenic medium; #significantly greater from growth with same cell type). Color images available online at www.liebertonline.com/ten.

Nanofiber mesh as a cell delivery vehicle

As a preliminary evaluation of the nanofiber mesh for cell delivery, AFS cells were seeded on nanofiber meshes and wrapped around cylindrical collagen scaffolds in vitro. The constructs were cultured for 2 weeks and then stained with the Live/Dead staining kit. In the cell-seeded scaffold, we observed that more cells were present at the periphery of the scaffold, even though cells were seeded throughout (Fig. 7A). We also detected a large number of dead cells in the interior of the scaffold. When the cell-seeded mesh was wrapped around a collagen scaffold, we found that numerous cells had migrated off the mesh onto the collagen scaffold and had high viability (Fig. 7C, D). The majority of cells were located close to the peripheral surface. Interestingly, we noted that the top surface was completely covered with cells (Fig. 7D). This implies that the cells were able to migrate at least 2 mm from the edge, where the mesh was present, to the center of the top surface. A few cells were also seen in the interior of the scaffold, more than 500 μm away from the peripheral surface (Fig. 7D). The mesh was completely confluent with cells, indicating that only a subset of cells migrates from the mesh onto the scaffold (Fig. 7E).

Discussion

In this study, we investigated the function of two kinds of stem cells, adult hMSCs and fetal hAFS cells, on electrospun nanofiber meshes. Both cell types were able to attach, colonize, and undergo robust osteogenic differentiation on the meshes. This indicates that the nanofiber mesh is a scaffold membrane capable of supporting vital osteoprogenitor and osteoblast functions. Other groups have also reported the ability of nanofiber meshes to promote differentiation of osteoblasts36,37 and MSCs;23,38 however, a quantitative and more thorough analysis of the matrix mineralization has been missing. We utilized a sensitive calcium assay based on the Arsenazo III dye to quantify the extent of matrix mineralization.29,39 The mineral deposited on the nanofiber meshes was confirmed to be biological in nature by FTIR spectroscopy, indicating that the process was cell mediated. Finally, we used calcein staining to observe and semiquantify the mineral deposits on the mesh. Another advantage of the calcein stain is that it can be used for continuous monitoring of the in vitro matrix mineralization process.30,31

Although acellular approaches to bone reconstruction using scaffolds and osteogenic growth factors have shown moderate clinical success, the delivery of osteogenic cells may be required for patients with a reduced local supply of responsive osteoprogenitor cells. For successful clinical translation of cell-based bone defect repair, a cell source needs to be identified that is readily available, propagated easily, has high osteogenic potential, and will be accepted by the recipient immune system. Both hMSCs and hAFS cells possess a number of these characteristics. MSCs have been studied extensively, especially for bone regeneration, and preclinical studies have shown their ability to repair bone defects in vivo.5,6 A number of human clinical trials have shown variable, but encouraging results of hMSC therapy, including the treatment of graft-versus-host disease, myocardial infarction, osteogenesis imperfecta, and large bone defects.40,41 However, hMSCs are known to progressively lose their stem cell properties during expansion, limiting the total number of cells available for therapy, and limited viability following transplantation remains a significant challenge.42 One of the objectives of this study was to compare the osteogenic capacity of this widely used adult stem cell with a more novel fetal stem cell source, the hAFS cells. AFS cells have been demonstrated to be capable of extensive self-renewal, and therefore can be expanded to large numbers and still maintain their multipotency.10–13

The colonization kinetics of hMSCs and hAFS cells on nanofiber meshes was found to be similar, suggesting comparable proliferation rates when seeded at a high density of 20,000 cells/cm2. On tissue culture plates, there were more hMSCs than hAFS cells on day 28, but this was due to the pelleting and recolonization by the hMSCs. hAFS cells demonstrated a later rise in ALP activity than the hMSCs on both plates and meshes, perhaps due to their primitive nature. ALP is a membrane-bound enzyme that plays an important role in matrix mineralization.35 ALP activity is one of the earliest markers of osteogenic differentiation and rises as the osteoprogenitors commit to the osteoblast lineage. It peaks in the matrix maturation phase in preparation of mineralization and decreases as mineralization progresses.43,44 However, ALP is expressed by other differentiated cells as well,45,46 and therefore it is important to simultaneously analyze other osteogenic measures, such as matrix mineralization. At 4 weeks, hAFS cells deposited significantly more mineral than hMSCs on both plates and meshes, as demonstrated by calcium quantification and calcein staining. Thus, we observed that, while the rise in ALP activity of the hAFS cells occurs later than in hMSCs, the hAFS cells mineralize more robustly after 4 weeks. This indicates that the kinetics of ALP activity and matrix mineralization are differentially regulated for these two different cell types. Our results demonstrate that hAFS cells have high osteogenic potential, even at the late passage numbers we have used. In addition, unlike human ESCs, hAFS cells have shown an absence of tumor formation in vivo.12 This suggests that the hAFS cells may be a feasible cell source for the repair of bone defects. They may be especially useful in the case of patients whose cells are not amenable for autologous transplantation due to disease or advanced age. Another advantage of the AFS cells is that they are suitable for convenient off-the-shelf allogeneic cell delivery, as long as the major histocompatibility complex of donor and recipient are matched. This would reduce the time and cost of delivering the cell therapy and may result in improved clinical acceptance.

Woo et al. have recently reported that a nanofibrous scaffold made by a modified solvent casting method resulted in improved expression of osteoblast phenotype versus a solid-walled scaffold.47 We compared cell function on the nanofiber meshes with that on tissue culture plastic. Compared to tissue culture plastic, there is delayed initial attachment and proliferation on the meshes. However, by 2 weeks, the cells on the meshes catch up with those on the plates, and there is no significant difference between the groups. The cells, especially the hMSCs, did not lift off the nanofiber mesh surface at high cell densities, as was seen on plates. This difference in cell attachment could be due to changes in cell adherence, material, and topographic properties or ECM deposited on the nanofiber mesh surface. Despite the initial lag in colonization on meshes compared to that on plates, we observed enhanced mineralization on meshes by both the cell types at 4 weeks. This suggests that the ECM-mimetic morphology of the nanofibers provides an environment conducive for matrix maturation. In a recent article, Smith et al. demonstrated that the use of nanofibers resulted in a greater degree of ESC differentiation, compared to films and tissue culture plates.48 This study also provides support for the use of nanofiber meshes as an improved in vitro cell culture model surface that better recapitulates the in vivo environment of cells.

Cell survival after delivery is a critical issue in the development of cell-based strategies, especially for thick tissues such as bone. The lack of initial vascularity in bone defects limits the transport of nutrients to and waste products from the center of the defect. Therefore, if cells are seeded throughout a 3D scaffold and placed at the defect site, cells located at the center of the scaffold may not survive.16,17,49,50 Delivery of cells on the periphery of bone defects via a tissue-engineered periosteum may be an effective approach to enhance cell survival by the presence of a neighboring vasculature. With time, as a continuous vasculature is established at the center, the cells may migrate toward the center due to an improved transport environment. Recently, Zhang reported that engraftment of bone morphogenetic protein-2 producing MSCs using gelfoam wrapped around nonvital allografts improved allograft incorporation and repair.51 Our results indicate that the electrospun nanofiber mesh possesses characteristics suitable for supporting cell function. In addition, its design and thickness can be controlled to obtain a membrane suitable for creating a tissue-engineered periosteum.52 To begin preliminary investigations into cell delivery, we asked following the question: Will cells migrate off the mesh and populate a 3D scaffold in vitro? We observed that the cells migrated off the mesh and colonized the scaffold within 2 weeks, traveling as far as 2 mm. In addition to the migration, part of the colonization is probably due to cell proliferation. Interestingly, we noticed better viability of the cells in the scaffold when they were delivered on the mesh compared to when they were seeded uniformly in the scaffolds. These results suggest that a cell-seeded nanofiber mesh may be an effective method to deliver cells to bone defects and maintain high viability. Future work will determine whether these effects are also observed in vivo.

In conclusion, we demonstrated that two types of stem cells, hMSCs and hAFS cells, are able to attach, colonize, and undergo robust osteogenic differentiation on electrospun nanofiber meshes. hAFS cells displayed a delayed ALP increase, but deposited significantly more mineral compared to hMSCs. Cell-seeded nanofiber meshes were effective in colonizing 3D scaffolds in an in vitro model. These results indicate that the electrospun nanofiber mesh supports osteoprogenitor cell function and may be useful as a medium for cell delivery for the repair of bone defects in vivo. In addition, this study provides support for the use of nanofiber meshes as a model surface for cell culture experiments.

Acknowledgments

This work was funded by NIH Grant AR051336. Collagen sheets were generously provided by Kensey Nash Corporation. The authors thank Dr. Charles Gersbach, Dr. Jennifer Phillips, and Dr. Ge Zhao for technical assistance with the calcium assay, FTIR, and the ALP assay, respectively. The authors would also like to thank Dr. Ayona Chatterjee for assistance with statistical analysis and Vivek Mukhatyar for helpful discussions.

Disclosure Statement

The authors do not have any competing financial interest.

References

  • 1.Praemer A. Furner S. Rice D.P. Musculoskeletal Conditions in the United States. Rosemont, IL: American Academy of Orthopaedic Surgeons; 1999. [Google Scholar]
  • 2.Rose F.R. Oreffo R.O. Bone tissue engineering: hope vs hype. Biochem Biophys Res Commun. 2002;292:1. doi: 10.1006/bbrc.2002.6519. [DOI] [PubMed] [Google Scholar]
  • 3.Salgado A.J. Coutinho O.P. Reis R.L. Bone tissue engineering: state of the art and future trends. Macromol Biosci. 2004;4:743. doi: 10.1002/mabi.200400026. [DOI] [PubMed] [Google Scholar]
  • 4.Bruder S.P. Fox B.S. Tissue engineering of bone. Cell based strategies. Clin Orthop Relat Res. 1999;S68 doi: 10.1097/00003086-199910001-00008. [DOI] [PubMed] [Google Scholar]
  • 5.Bruder S.P. Kraus K.H. Goldberg V.M. Kadiyala S. The effect of implants loaded with autologous mesenchymal stem cells on the healing of canine segmental bone defects. J Bone Joint Surg Am. 1998;80:985. doi: 10.2106/00004623-199807000-00007. [DOI] [PubMed] [Google Scholar]
  • 6.Bruder S.P. Kurth A.A. Shea M. Hayes W.C. Jaiswal N. Kadiyala S. Bone regeneration by implantation of purified, culture-expanded human mesenchymal stem cells. J Orthop Res. 1998;16:155. doi: 10.1002/jor.1100160202. [DOI] [PubMed] [Google Scholar]
  • 7.Tsuchida H. Hashimoto J. Crawford E. Manske P. Lou J. Engineered allogeneic mesenchymal stem cells repair femoral segmental defect in rats. J Orthop Res. 2003;21:44. doi: 10.1016/S0736-0266(02)00108-0. [DOI] [PubMed] [Google Scholar]
  • 8.McCulloch C.A. Strugurescu M. Hughes F. Melcher A.H. Aubin J.E. Osteogenic progenitor cells in rat bone marrow stromal populations exhibit self-renewal in culture. Blood. 1991;77:1906. [PubMed] [Google Scholar]
  • 9.Quarto R. Thomas D. Liang C.T. Bone progenitor cell deficits and the age-associated decline in bone repair capacity. Calcif Tissue Int. 1995;56:123. doi: 10.1007/BF00296343. [DOI] [PubMed] [Google Scholar]
  • 10.Kolambkar Y.M. Peister A. Soker S. Atala A. Guldberg R.E. Chondrogenic differentiation of amniotic fluid-derived stem cells. J Mol Histol. 2007;38:405. doi: 10.1007/s10735-007-9118-1. [DOI] [PubMed] [Google Scholar]
  • 11.Siddiqui M.M. Atala A. Amniotic fluid-derived pluripotential cells. In: Lanza R., editor; Gearhart J., editor; Hogan B., editor. Handbook of Stem Cells. Philadelphia: Academic Press; 2004. p. 175. [Google Scholar]
  • 12.De Coppi P. Bartsch G., Jr. Siddiqui M.M. Xu T. Santos C.C. Perin L. Mostoslavsky G. Serre A.C. Snyder E.Y. Yoo J.J. Furth M.E. Soker S. Atala A. Isolation of amniotic stem cell lines with potential for therapy. Nat Biotechnol. 2007;25:100. doi: 10.1038/nbt1274. [DOI] [PubMed] [Google Scholar]
  • 13.Delo D.M. De Coppi P. Bartsch G., Jr. Atala A. Amniotic fluid and placental stem cells. Methods Enzymol. 2006;419:426. doi: 10.1016/S0076-6879(06)19017-5. [DOI] [PubMed] [Google Scholar]
  • 14.Peister A. Porter B.D. Kolambkar Y.M. Hutmacher D.W. Guldberg R.E. Osteogenic differentiation of amniotic fluid stem cells. Biomed Mater Eng. 2008;18:241. [PubMed] [Google Scholar]
  • 15.Peister A. Deutsch E.R. Kolambkar Y. Hutmacher D.W. Guldberg R. Amniotic fluid stem cells produce robust mineral deposits on biodegradable scaffolds. Tissue Eng Part A. 2009;15:3129. doi: 10.1089/ten.tea.2008.0536. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Thorrez L. Shansky J. Wang L. Fast L. Vandendriessche T. Chuah M. Mooney D. Vandenburgh H. Growth, differentiation, transplantation and survival of human skeletal myofibers on biodegradable scaffolds. Biomaterials. 2008;29:75. doi: 10.1016/j.biomaterials.2007.09.014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Jager M. Degistirici O. Knipper A. Fischer J. Sager M. Krauspe R. Bone healing and migration of cord blood-derived stem cells into a critical size femoral defect after xenotransplantation. J Bone Miner Res. 2007;22:1224. doi: 10.1359/jbmr.070414. [DOI] [PubMed] [Google Scholar]
  • 18.Pham Q.P. Sharma U. Mikos A.G. Electrospun poly(epsilon-caprolactone) microfiber and multilayer nanofiber/microfiber scaffolds: characterization of scaffolds and measurement of cellular infiltration. Biomacromolecules. 2006;7:2796. doi: 10.1021/bm060680j. [DOI] [PubMed] [Google Scholar]
  • 19.Barnes C.P. Sell S.A. Boland E.D. Simpson D.G. Bowlin G.L. Nanofiber technology: designing the next generation of tissue engineering scaffolds. Adv Drug Deliv Rev. 2007;59:1413. doi: 10.1016/j.addr.2007.04.022. [DOI] [PubMed] [Google Scholar]
  • 20.Tan S.H. Inai R. Kotaki M. Ramakrishna S. Systematic parameter study for ultra-fine fiber fabrication via electrospinning process. Polymer. 2005;46:6128. [Google Scholar]
  • 21.Pham Q.P. Sharma U. Mikos A.G. Electrospinning of polymeric nanofibers for tissue engineering applications: a review. Tissue Eng. 2006;12:1197. doi: 10.1089/ten.2006.12.1197. [DOI] [PubMed] [Google Scholar]
  • 22.Murugan R. Ramakrishna S. Nano-featured scaffolds for tissue engineering: a review of spinning methodologies. Tissue Eng. 2006;12:435. doi: 10.1089/ten.2006.12.435. [DOI] [PubMed] [Google Scholar]
  • 23.Li W.J. Tuli R. Huang X. Laquerriere P. Tuan R.S. Multilineage differentiation of human mesenchymal stem cells in a three-dimensional nanofibrous scaffold. Biomaterials. 2005;26:5158. doi: 10.1016/j.biomaterials.2005.01.002. [DOI] [PubMed] [Google Scholar]
  • 24.Xin X. Hussain M. Mao J.J. Continuing differentiation of human mesenchymal stem cells and induced chondrogenic and osteogenic lineages in electrospun PLGA nanofiber scaffold. Biomaterials. 2007;28:316. doi: 10.1016/j.biomaterials.2006.08.042. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Yoshimoto H. Shin Y.M. Terai H. Vacanti J.P. A biodegradable nanofiber scaffold by electrospinning and its potential for bone tissue engineering. Biomaterials. 2003;24:2077. doi: 10.1016/s0142-9612(02)00635-x. [DOI] [PubMed] [Google Scholar]
  • 26.Klinkhammer K. Seiler N. Grafahrend D. Gerardo-Nava J. Mey J. Brook G.A. Moller M. Dalton P.D. Klee D. Deposition of electrospun fibers on reactive substrates for in vitro investigations. Tissue Eng Part C Methods. 2009;15:77. doi: 10.1089/ten.tec.2008.0324. [DOI] [PubMed] [Google Scholar]
  • 27.Sekiya I. Larson B.L. Smith J.R. Pochampally R. Cui J.G. Prockop D.J. Expansion of human adult stem cells from bone marrow stroma: conditions that maximize the yields of early progenitors and evaluate their quality. Stem Cells. 2002;20:530. doi: 10.1634/stemcells.20-6-530. [DOI] [PubMed] [Google Scholar]
  • 28.Martin J.Y. Dean D.D. Cochran D.L. Simpson J. Boyan B.D. Schwartz Z. Proliferation, differentiation, and protein synthesis of human osteoblast-like cells (MG63) cultured on previously used titanium surfaces. Clin Oral Implants Res. 1996;7:27. doi: 10.1034/j.1600-0501.1996.070104.x. [DOI] [PubMed] [Google Scholar]
  • 29.Leary N.O. Pembroke A. Duggan P.F. Single stable reagent (Arsenazo III) for optically robust measurement of calcium in serum and plasma. Clin Chem. 1992;38:904. [PubMed] [Google Scholar]
  • 30.Uchimura E. Machida H. Kotobuki N. Kihara T. Kitamura S. Ikeuchi M. Hirose M. Miyake J. Ohgushi H. In-situ visualization and quantification of mineralization of cultured osteogenetic cells. Calcif Tissue Int. 2003;73:575. doi: 10.1007/s00223-002-1052-3. [DOI] [PubMed] [Google Scholar]
  • 31.Hale L.V. Ma Y.F. Santerre R.F. Semi-quantitative fluorescence analysis of calcein binding as a measurement of in vitro mineralization. Calcif Tissue Int. 2000;67:80. doi: 10.1007/s00223001101. [DOI] [PubMed] [Google Scholar]
  • 32.Kutner M.H. Nachtsheim C.J. Neter J. Li W. Applied Linear Statistical Models. fifth. New York: McGraw-Hill; 2005. [Google Scholar]
  • 33.Box G.E.P. Cox D.R. An analysis of transformations. J R Stat Soc B. 1964;26:211. [Google Scholar]
  • 34.Wu C.F.J. Hamada M. Experiments: Planning, Analysis, and Parameter Design Optimization. first. New York: Wiley-Interscience; 2000. [Google Scholar]
  • 35.Seibel M.J. Molecular markers of bone turnover: biochemical, technical and analytical aspects. Osteoporos Int. 2000;11(Suppl 6):S18. doi: 10.1007/s001980070003. [DOI] [PubMed] [Google Scholar]
  • 36.Venugopal J. Low S. Choon A.T. Kumar A.B. Ramakrishna S. Electrospun-modified nanofibrous scaffolds for the mineralization of osteoblast cells. J Biomed Mater Res A. 2008;85:408. doi: 10.1002/jbm.a.31538. [DOI] [PubMed] [Google Scholar]
  • 37.Badami A.S. Kreke M.R. Thompson M.S. Riffle J.S. Goldstein A.S. Effect of fiber diameter on spreading, proliferation, and differentiation of osteoblastic cells on electrospun poly(lactic acid) substrates. Biomaterials. 2006;27:596. doi: 10.1016/j.biomaterials.2005.05.084. [DOI] [PubMed] [Google Scholar]
  • 38.Shin M. Yoshimoto H. Vacanti J.P. In vivo bone tissue engineering using mesenchymal stem cells on a novel electrospun nanofibrous scaffold. Tissue Eng. 2004;10:33. doi: 10.1089/107632704322791673. [DOI] [PubMed] [Google Scholar]
  • 39.Gersbach C.A. Le Doux J.M. Guldberg R.E. Garcia A.J. Inducible regulation of Runx2-stimulated osteogenesis. Gene Ther. 2006;13:873. doi: 10.1038/sj.gt.3302725. [DOI] [PubMed] [Google Scholar]
  • 40.Abdallah B.M. Kassem M. The use of mesenchymal (skeletal) stem cells for treatment of degenerative diseases: current status and future perspectives. J Cell Physiol. 2009;218:9. doi: 10.1002/jcp.21572. [DOI] [PubMed] [Google Scholar]
  • 41.Quarto R. Mastrogiacomo M. Cancedda R. Kutepov S.M. Mukhachev V. Lavroukov A. Kon E. Marcacci M. Repair of large bone defects with the use of autologous bone marrow stromal cells. N Engl J Med. 2001;344:385. doi: 10.1056/NEJM200102013440516. [DOI] [PubMed] [Google Scholar]
  • 42.Shi S. Gronthos S. Chen S. Reddi A. Counter C.M. Robey P.G. Wang C.Y. Bone formation by human postnatal bone marrow stromal stem cells is enhanced by telomerase expression. Nat Biotechnol. 2002;20:587. doi: 10.1038/nbt0602-587. [DOI] [PubMed] [Google Scholar]
  • 43.Lian J.B. Stein G.S. Development of the osteoblast phenotype: molecular mechanisms mediating osteoblast growth and differentiation. Iowa Orthop J. 1995;15:118. [PMC free article] [PubMed] [Google Scholar]
  • 44.Aubin J.E. Regulation of osteoblast formation and function. Rev Endocr Metab Disord. 2001;2:81. doi: 10.1023/a:1010011209064. [DOI] [PubMed] [Google Scholar]
  • 45.Hinnebusch B.F. Siddique A. Henderson J.W. Malo M.S. Zhang W. Athaide C.P. Abedrapo M.A. Chen X. Yang V.W. Hodin R.A. Enterocyte differentiation marker intestinal alkaline phosphatase is a target gene of the gut-enriched Kruppel-like factor. Am J Physiol Gastrointest Liver Physiol. 2004;286:G23. doi: 10.1152/ajpgi.00203.2003. [DOI] [PubMed] [Google Scholar]
  • 46.Matsumoto H. Erickson R.H. Gum J.R. Yoshioka M. Gum E. Kim Y.S. Biosynthesis of alkaline phosphatase during differentiation of the human colon cancer cell line Caco-2. Gastroenterology. 1990;98:1199. doi: 10.1016/0016-5085(90)90334-w. [DOI] [PubMed] [Google Scholar]
  • 47.Woo K.M. Jun J.H. Chen V.J. Seo J. Baek J.H. Ryoo H.M. Kim G.S. Somerman M.J. Ma P.X. Nano-fibrous scaffolding promotes osteoblast differentiation and biomineralization. Biomaterials. 2007;28:335. doi: 10.1016/j.biomaterials.2006.06.013. [DOI] [PubMed] [Google Scholar]
  • 48.Smith L.A. Liu X. Hu J. Wang P. Ma P.X. Enhancing osteogenic differentiation of mouse embryonic stem cells by nanofibers. Tissue Eng Part A. 2009;15:1855. doi: 10.1089/ten.tea.2008.0227. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Byers B.A. Guldberg R.E. Garcia A.J. Synergy between genetic and tissue engineering: Runx2 overexpression and in vitro construct development enhance in vivo mineralization. Tissue Eng. 2004;10:1757. doi: 10.1089/ten.2004.10.1757. [DOI] [PubMed] [Google Scholar]
  • 50.Cartmell S. Huynh K. Lin A. Nagaraja S. Guldberg R. Quantitative microcomputed tomography analysis of mineralization within three-dimensional scaffolds in vitro. J Biomed Mater Res. 2004;69A:97. doi: 10.1002/jbm.a.20118. [DOI] [PubMed] [Google Scholar]
  • 51.Zhang X. Xie C. Lin A.S. Ito H. Awad H. Lieberman J.R. Rubery P.T. Schwarz E.M. O'Keefe R.J. Guldberg R.E. Periosteal progenitor cell fate in segmental cortical bone graft transplantations: implications for functional tissue engineering. J Bone Miner Res. 2005;20:2124. doi: 10.1359/JBMR.050806. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Ekaputra A.K. Zhou Y. Cool S.M. Hutmacher D.W. Composite electrospun scaffolds for engineering tubular bone grafts. Tissue Eng Part A. 2009;15:3779. doi: 10.1089/ten.TEA.2009.0186. [DOI] [PubMed] [Google Scholar]

Articles from Tissue Engineering. Part A are provided here courtesy of Mary Ann Liebert, Inc.

RESOURCES