Journal of Materials Science

, Volume 48, Issue 15, pp 5113–5124

Biocompatibility evaluation of protein-incorporated electrospun polyurethane-based scaffolds with smooth muscle cells for vascular tissue engineering

Authors

  • Lin Jia
    • Key Laboratory of Textile Science & Technology, Ministry of Education, College of TextilesDonghua University
    • Faculty of Engineering, Center for Nanofibers and Nanotechnology, E3-05-14, Nanoscience and Nanotechnology Initiative, National University of Singapore
    • Faculty of Engineering, Center for Nanofibers and Nanotechnology, E3-05-14, Nanoscience and Nanotechnology Initiative, National University of Singapore
    • Key Laboratory of Textile Science & Technology, Ministry of Education, College of TextilesDonghua University
  • Dan Kai
    • NUS Graduate School for Integrative Sciences and Engineering, National University of Singapore
  • Seeram Ramakrishna
    • Faculty of Engineering, Center for Nanofibers and Nanotechnology, E3-05-14, Nanoscience and Nanotechnology Initiative, National University of Singapore
Article

DOI: 10.1007/s10853-013-7359-9

Cite this article as:
Jia, L., Prabhakaran, M.P., Qin, X. et al. J Mater Sci (2013) 48: 5113. doi:10.1007/s10853-013-7359-9

Abstract

Nanotechnology has enabled the engineering of a variety of materials to meet the current challenges and needs in vascular tissue regeneration. In this study, four different kinds of native proteins namely collagen, gelatin, fibrinogen, and bovine serum albumin were incorporated with polyurethane (PU) and electropsun to obtain composite PU/protein nanofibers. SEM studies showed that the fiber diameters of PU/protein scaffolds ranged from 245 to 273 nm, mimicking the nanoscale dimensions of native ECM. Human aortic smooth muscle cells (SMCs) were cultured on the electrospun nanofibers, and the ability of the cells to proliferate on different scaffolds was evaluated via a cell proliferation assay. Cell proliferation on PU/Coll nanofibers was found the highest compared to other electrospun scaffolds and it was 42 % higher than the proliferation on PU/Fib nanofibers after 12 days of cell culture. The cell–biomaterial interaction studies by SEM confirmed that SMCs adhered to PU/Coll and PU/Gel nanofibers, with high cell substrate coverage, and both the scaffolds promoted cell alignment. The functionality of the cells was further demonstrated by immunocytochemical analysis, where the SMCs on PU/Coll and PU/Gel nanofibers expressed higher density of SMC proteins such as alpha smooth muscle actin and smooth muscle myosin heavy chain. Cells expressed biological markers of SMCs including aligned spindle-like morphology on both PU/Coll and PU/Gel with actin filament organizations, better than PU/Fib and PU/BSA scaffolds. Our studies demonstrate the potential of randomly oriented elastomeric composite scaffolds for engineering of vascular tissues causing cell alignment.

Introduction

Cardiovascular diseases, such as atherosclerosis and coronary arterial restenosis, are the leading cause of morbidity and mortality in the developed world [1]. Small-diameter blood vessel substitutes are a requisite for patients requiring replacement of coronary blood vessels [2]. Tissue-engineered vascular grafts are a promising way to meet the replacement requirements. However, a bioengineered graft needs careful selection, such that the problems of thrombogenicity and poor vasoactivity are avoided and the graft requires enough mechanical and biochemical properties to be fully successful in human body system [3]. Smooth muscle cells (SMCs) are the major cells in the vascular tissue, and during vascular development, SMCs play a key role in morphogenesis of the blood vessels and exhibit high rates of proliferation, migration, and production of extracellular matrix (ECM) components such as collagen, elastin, and proteoglycans that make up the major portion of the blood vessel wall [4].

Numerous efforts are ongoing to develop functional scaffolds for the repair and regeneration of tissues and internal organs, and for being functional, the scaffold should be biologically and structurally active. The biological function of the scaffold is to support attachment, growth, and proliferation of cells. The structural functions are to provide a biocompatible template allowing for the ingrowth of cells, blood vessel formation, new tissue development, and maintenance of ECM. For these purposes, the scaffold should have sufficient porosity with interconnected pores and preferably bioresorbable in order to allow for its gradual replacement by new tissues [5]. Nanofibrous scaffolds produced by electrospinning possess several advantages which are critical for tissue regeneration, including high surface area to volume ratios for better cell incorporation and nutrition perfusion and controlled fiber diameters to mimic the fibrous architecture of ECM. The process also has the advantage of incorporating multiple polymers and bioactive ingredients to prepare scaffolds with tunable mechanical and biodegradation properties. Because of these reasons, electrospun nanofibers are widely used in tissue engineering [68]. Various biodegradable and biocompatible polymers are available and a broad spectrum of nanofiber-based scaffolds with different mechanical and biochemical properties are being used in tissue engineering. Thermoplastic polyurethane (PU) is a renowned class of polymers with excellent mechanical properties and good biocompatibility, and it has been evaluated as a coating material for breast implants, catheters, and prosthetic heart valve leaflets [9]. PU is an aliphatic biodegradable polymer and it is known to have a biostability better than poly(ester urethane). Despite good mechanical properties and predictable biodegradation kinetics, PU lacks cell-recognition moieties and is not favorable as a pure matrix for tissue regeneration [10]. However, native proteins can be incorporated into PU through blending and further electrospun to obtain PU composite fibers to improve the biocompatibility of PU, while preserving its mechanical strength.

Collagen is the most abundant proteins in the human body, a key element of the native ECM, and it imparts structural integrity and tensile strength to tissues [11]. In native blood vessels, along with smooth muscle cells (SMCs), collagen confers wall strength and resistance to vessel rupture in a harsh in situ environment [12]. Additionally, collagen is resorbable, has high water affinity, low antigenicity, good cell compatibility, and the ability to promote tissue regeneration [13]. Collagen has been used in a variety of tissue-engineering applications because of its predominance in the ECM, non-immunogenicity, and available methods of isolation from a variety of sources. He W et al. [14] utilized collagen-coated poly(L-lactic acid)-co-poly(ε-caprolactone) (PLCL) to create a suitable scaffold for vascular tissue engineering. Gelatin is a derivative of collagen, acquired by denaturing the triple-helix structure [15]. Due to the similarities between gelatin and collagen and its natural origin, it has become an attractive polymer for tissue-engineering applications. In addition to the similar mechanical properties to collagen, gelatin exhibits excellent biodegradability, non-antigenicity, and cost efficiency [16, 17]. Electrospun gelatin scaffolds have been used for applications such as nerve, cartilage, bone, and dermal tissue-engineering applications [1821]. For vascular applications, much of the work with gelatin has been centered on its use as a sealant or coating to enhance the biological properties of synthetic scaffolds. Ma et al. [22] demonstrated that the grafting of gelatin on electrospun poly(caprolactone) (PCL) nanofibers can improve endothelial cell spreading, proliferation, and even control the cell orientation. Fibrinogen is a 340-kDa plasma glycoprotein that plays a key role in blood clotting and wound healing [23]. It is a major protein with capacity to bind a wide array of molecules and it could be beneficial from a vascular prosthetic engineering point of view. Fibrinogen has been shown to contain RGD integrin-binding sites that can attach to fibroblasts (FBs) and endothelial cells (ECs). Additionally, fibrinogen has demonstrated high affinity to growth factors such as the vascular endothelial growth factor (VEGF), fibroblast growth factor (FGF), and a number of other cytokines [2427]. Fibrinogen-based materials have been widely used in the tissue engineering of cartilage, bone, skin, and blood vessel [2831]. He CL et al. [32] fabricated hybrid PLCL/fibrinogen nanofibrous scaffolds and they found that L929 cells on hybrid scaffolds achieved a relatively higher level of cell attachment. Bovine serum albumin (BSA) is a large globular protein with the molecular weight of 66000 Daltons and it contain a single polypeptide chain of about 583 amino acid residues devoid of carbohydrate residues. Bovine serum albumin (BSA) is used as coating agent or as a nutrient for cells. It has been used to protect a variety of growth factors during electrospinning of growth factor-containing nanofibers [33]. In addition, BSA is the first biomolecule that has been used in biomedical applications, because it is a relatively large biomolecule and usually has a tendency to accumulate at the interface of solid surfaces, and its adsorption on the surface of solid composite nanoparticles can act as adsorption models for other biomacromolecules [34, 35]. Studies by Parizek et al. [36] showed that the polyethylene (PE) scaffold activated with plasma and subsequently grafted with BSA can promote the adhesion, proliferation, and phenotypic maturation of SMCs.

Polymer blending is one of the most effective methods for preparing biocomposite scaffolds with tailored properties for particular tissue-engineering applications [37]. During this study, we utilized four different types of proteins, namely collagen, gelatin, fibrinogen, and bovine serum albumin, to individually blend with PU and fabricate composite PU/protein nanofibers via electrospinning. The chemical and mechanical characteristics of the different electrospun nanofibers together with its cell biocompatibility evaluations were carried out during this study in order to identify the most suitable and biofunctional scaffold for vascular tissue regeneration.

Materials and methods

Materials

Human aortic smooth muscle cells, smooth muscle cell medium (SMCM), fetal bovine serum (FBS), smooth muscle growth supplement (SMCGS), and penicillin/streptomycin solution (P/S) were obtained from ScienceCell, USA. 4,6-diamidino-2-phenylindole, dihydrochloride (DAPI), Alexa Fluor 488, and Alexa Fluor 594 were all purchased from Invitrogen Corporation, USA. Collagen type I, gelatin type A, bovine serum albumin, fibrinogen, and 1,1,1,3,3,3-hexafluor-2-propanol (HFP) were purchased from Sigma-Aldrich Pte Ltd., Singapore. Polyurethane (PU) was generously provided by the Lubrizol Corporation. Mouse anti-human alpha smooth muscle actin (SMA) and anti-smooth muscle myosin heavy chain antibody (MHC) were obtained from Abcam (Cambridge, UK).

Electrospinning of nanofibers

PU and powder of collagen I(PU/Coll), gelatin A(PU/Gel), fibrinogen(PU/Fib), and bovine serum albumin(PU/BSA) were dissolved in HFP at a ratio of 75:25(wt%) to obtain 6 %(w/v) solution each. The polymer solution was individually placed in a 3-ml plastic syringe attached to a 27-G blunted stainless steel needle using a syringe pump (KDS 100, KD Scientific, Hollistion, MA) and fed at a flow rate of 1.0 ml/h. A high voltage of 15 kV (Gamma High Voltage Research, USA) was applied when the polymer solution was drawn into fibers and collected on an aluminum foil-wrapped collector kept at a distance of 12 cm from the needle tip. Nanofibers collected on aluminum foil and 15-mm cover slips were dried overnight under vacuum and used for characterization and cell culture experiments, respectively. In addition, pure PU nanofibers were fabricated from 6 %(w/v) solution and utilized as a control.

Morphology, chemical, and mechanical properties of nanofibers

The electrospun nanofibers were sputter coated with gold (JEOL JFC-1200 Auto Fine coater, Japan) and visualized by scanning electron microscopy (SEM, FEI- QUANTA 200 F, the Netherlands) at an accelerating voltage of 10 kV. The average fiber diameters were measured based on the SEM micrographs by measuring about 100 random fibers using image analysis software (ImageJ, National Institutes of Health, Bethesda, MD). The porosity of the electrospun scaffolds was studied by a capillary flow porometer (CFP-1200-A). Applied with low pressure, capillary flow porometry provides a simple and non-destructive technique that allows for the rapid and accurate measurement of pore size and distribution. Attenuated total reflectance Fourier transform infrared (ATR-FTIR) spectroscopic analysis of electrospun nanofibrous scaffolds was performed on an Avatar 380 FTIR spectrometer (Thermo Nicolet, Waltham, MA) over a range of 400–4000 cm−1 at a resolution of 2 cm−1. The hydrophilic/hydrophobic nature of the electrospun nanofibrous scaffolds was measured by water contact angle measurement using VCA Optima Surface Analysis System 213 (AST products, Billerica, MA). During the measurements, electrospun nanofibrous membranes on the coverslips were directly positioned on a testing plate. Subsequently, deionized water was dropped onto the samples. Three different positions of the same sample were measured and recorded. The tensile properties of electrospun nanofibers were evaluated using a tabletop tensile Tester (Instron 5345, USA) using a load cell of 10-N capacity at ambient conditions. Rectangular specimens of dimensions 10 × 20 mm were cut (using a specimen frame) and peeled off from the nanofiber sheet spun on aluminum foil and tested at a cross-head speed of 10 mm min−1. Six samples were tested for each type of electrospun membrane during this study. Tensile strength and elongation at break were calculated based on the generated stress–strain curves generated for each scaffold.

In vitro cell culture and seeding

SMCs were cultured in SMCM supplemented with 2 % FBS, 1 %SMCGS, and 1 % P/S in a 75-cm2 cell culture flask. Cells were incubated at 37 °C in humidified atmosphere containing 5 % CO2. Cells were fed with fresh media after every 3 days and trypsinized using trypsin–EDTA for cell seeding experiments. The 15-mm coverslips with electrospun nanofibers were placed in 24-well plates and pressed with a stainless steel ring to insure complete contact of the scaffolds with the wells. The specimens were sterilized under UV light, washed thrice with phosphate-buffered saline (PBS), and subsequently incubated in SMCM overnight before cell seeding. SMCs were seeded at a density of 10,000 cells per well on PU, PU/Coll, PU/Gel, PU/Fib, PU/BSA nanofibrous scaffolds, and tissue culture polystyrene (TCP as control) placed in 24-well plates.

Cell proliferation

The adhesion and proliferation of SMCs on different electrospun nanofibers and TCP (control) were studied using 3-(4,5-dimethylthiazol-2-yl)-5-(3-car-boxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) colorimetric assay. The kit (CellTiter 96 AQueous, One Solution cell proliferation assay; Promega, Madison, WI) was used according to the manufacturer’s directions. After culturing the cells for a period of 3, 6, 9, and 12 days, the cell-scaffold constructs were washed once with PBS and incubated with 1 ml of DMEM containing 20 % of CellTiter 96 AQueous One Solution reagent. After incubating for a period of 3 h, the deeply colored culture medium was aliquoted to a 96-well plate (100 μl/well), and the absorbance was recorded at 490 nm using a microplate reader. The sample with culture medium, but without cells, was set as the control to determine the background absorbance to be subtracted.

Morphological evaluation of SMCs

Morphological study of SMCs grown on different electrospun nanofibers was performed after 6 days of cell culture by SEM. The cell-scaffold constructs were rinsed twice with PBS and fixed in 3 % glutaraldehyde for 3 h. The scaffolds were further rinsed in deionized water and dehydrated with increasing concentrations of ethanol (50, 70, 90, and 100 %) for 10 min each. Finally, the cell-scaffold constructs were treated with hexamethyldisilazane and air dried in a fume hood overnight. The cell-scaffold construct was observed under SEM after gold coating.

Immunostaining studies

SMCs grown on different electrospun nanofibers and TCP for 9 days were processed for immunocytochemistry. After 9 days of cell culture, the cells were fixed in formalin at room temperature for 30 min. The samples were then washed three times with PBS and incubated in 0.1 % TritonX 100 solution for 5 min to permeabilize the cell membrane. Non-specific sites were blocked by incubating the cells in 3 % BSA for 90 min. The samples were then incubated with either one of the SMC- specific marker proteins, SMA or MHC, at a dilution 1:100 for 120 min at room temperature. Further, the hybrid solution of secondary antibody goat anti-mouse AF 488 (green) (1:400) for SMA or AF 594 (red) (1:400) for MHC and DAPI (1:500) was added for 90 min. The samples were washed with PBS repeatedly and further mounted over a glass slide using Vectashield mounting medium and examined under the fluorescent microscope (Olympus FV 1000).

Statistical analysis

All the data presented are expressed as mean ± standard deviation (SD) of the mean. Single-factor analysis of variance was carried out to compare the means of different datasets, and a value of p ≤ 0.05 was considered statistically significant.

Results and discussion

An important challenge in vascular tissue engineering is the development of a suitable vascular graft substitute that is “smart” enough to instruct the in vivo environment to form vascular tissue. Polyurethanes (PU) have low toxicity and good mechanical properties, but lack chemical functionalities, such as carboxyl or amine groups for cell adhesion, and hence pure PU is not always suitable for tissue engineering. Due to these reasons, we incorporated a natural protein (collagen, gelatin, fibrinogen, or bovine serum albumin) to produce PU composite scaffolds to enhance the cell adhesion on them. The goal of this study was to evaluate the ability of electrospun composite PU/protein materials, which mimic both the physical structural features and the bioactive character of native vascular tissue to promote SMC attachment, proliferation, and to assess its functionality of expression of SMC-specific proteins.

Morphology, chemical, and mechanical properties of nanofibers

Electrospinning was used as a method to fabricate nanofibrous scaffolds which mimic the native morphology of ECM in tissues, where the size of most features is in the nanometer scale. The topography of nanofibers plays an important role in regulating the initial cell behavior such as cell adhesion and proliferation. Blending PU with native protein including collagen, gelatin, fibrinogen, and bovine serum albumin at a weight ratio of 75:25 was carried out to obtain continuous, uniform, and smooth bead-free fibers. The morphology of the electrospun PU/Coll, PU/Gel, PU/Fib, and PU/BSA nanofibers was observed by SEM (Fig. 1) and the fiber diameters were in the range of 273 ± 76, 256 ± 70, 248 ± 65, and 245 ± 107 nm, respectively. Compared with pure PU nanofibers (result are not shown in this study), the incorporation of native protein reduced the fiber diameter of the PU/protein scaffolds. One possible explanation for this is that the conductivity of the electrospun solutions increased with the addition of protein into the PU solution[38]. It was reported that cells attach to and organize well around fibers with diameters smaller than the diameter of the cells [39]. Furthermore, the fibrillar structure of the scaffolds was found to promote cell attachment, proliferation, and the colony-forming capacity of cells in vitro [40].
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Fig. 1

SEM images of electrospun nanofibers with in situ figures of their pore size distributions; a PU/Coll, b PU/Gel, c PU/Fib, d PU/BSA

The porosity of the electrospun scaffolds was studied by a capillary flow porometer (CFP-1200-A) and the mean pore size of PU/Coll, PU/Gel, PU/Fib, and PU/BSA nanofibers was 0.357, 0.328, 0.301, and 0.286 μm, respectively. Applied with low pressure, capillary flow porometry provides a simple and non-destructive technique that allows for the rapid and accurate measurement of pore size and distribution [41]. The fiber diameter and fiber mass play a dominant role in controlling the pore diameter of the networks [42]. In our study, the thickness of all PU/protein nanofibers was kept identical (approximately 25 mm). The mean pore sizes obtained for the nanofibers were inversely proportional to the diameter of the fibers and our results were also in accordance with the results reported by other researchers [41, 43]. Uniform distribution of pore sizes was observed for PU/Coll, PU/Gel, and PU/Fib nanofibers (Fig. 1a, b, c). However, uneven distribution of pore sizes was observed for PU/BSA nanofibers(Fig. 1d), which was also clear from a bigger standard deviation of the PU/BSA fiber diameters and could be due to the uneven distribution of fiber diameters.

Surface characterization of the electrospun nanofibers was carried out by ATR-FTIR. Figure 2 shows the ATR-FTIR spectra of electrospun PU/Coll, PU/Gel, PU/Fib, and PU/BSA compared to those of the PU nanofibers. The spectroscopy of electrospun PU has a characteristic absorption band at 2960, 1710, 1530, 1220, 1110, and 777 cm−1, which represents υ(C–H), υ (C=O), υ (C=C), υ (C–C), υ (C–O), and υ (C–H), respectively, on substituted benzene [20]. In addition to the characteristic absorption band of PU, the electrospun PU/Coll, PU/Gel, PU/BSA, and PU/Fib nanofibers also showed the presence of N–H stretching peak at 3310 cm−1 for amide A and C–H stretching at 3062 cm−1 for amide B, suggesting the presence of amino functional groups on their surfaces. Additionally, the C=O stretching and N–H deformation peaks were also observed on electrospun PU/Coll, PU/Gel, PU/BSA, and PU/Fib nanofibers at 1651 cm−1(amide I) and 1535 cm−1 (amide II), respectively. Because the υ (C–C) absorption peak at 1530 cm−1 of PU is very close to the N–H deformation peaks at 1535 cm−1, the two peaks broadened to produce a bigger and more prominent peak (compared to the respective peak of pure PU scaffold) at 1535 cm−1 on the surface of PU/Coll, PU/Gel, PU/BSA, and PU/Fib nanofibers. Comparing the four spectroscopies of electrospun PU/Coll, PU/Gel, PU/Fib, and PU/BSA nanofibers in Fig. 2, we found that the intensity of the N–H stretching peak at 3310 cm−1 for amide A on the PU/Coll nanofibers was stronger and the intensity of N–H deformation peaks at 1535 cm−1 for amide II on PU/Fib was less intensive compared to the other scaffolds. Although the electrospun PU/Coll, PU/Gel, PU/BSA ,and PU/Fib nanofibers express the presence of amino functional groups on their surfaces, the intensity of these groups is different and this might further affect the cell behavior when they are utilized as scaffolds for cell culture. The spectra also illustrate no other reactions between PU and the incorporated protein and hence it was clear that the individual polymers kept their structure independent in the composite scaffolds.
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Fig. 2

FTIR spectra of electrospun PU, PU/Coll, PU/Gel, PU/Fib, and PU/BSA nanofibers

The surface hydrophobic–hydrophilicity of electrospun nanofibers has a major influence on cell adhesion and proliferation behavior. Pure PU nanofibers are hydrophilic, but the nanofibers got wrinkled, start shrinking due to the extremely high elastic properties of pure PU, and this is not a desirable aspect required of a vascular graft, which might also influence the cell growth and behaviors. Hence, we incorporated a native protein into the PU polymer to enhance the cell adhesion and proliferation on composite PU/protein scaffolds. The inset of Fig. 3 shows the surface hydrophobic–hydrophilicity of electrospun PU/Coll, PU/Gel, PU/Fib, and PU/BSA nanofibers with a contact angle of 32.7 ± 3.2°, 28.1 ± 2.1°, 40.7 ± 4.5°, and 43.4 ± 4.2°, respectively. Results showed that the incorporation of proteins helped to retain the hydrophilicity of PU nanofibers and assisted in maintaining the integrity of the scaffolds. However, the hydrophilicity of PU/Fib and PU/BSA nanofibers was lesser than PU/Coll and PU/Gel scaffolds. This is because both collagen and gelatin contain hydroxyl groups which can form hydrogen bonds with water molecules, thereby providing higher hydrophilicity to PU/Coll and PU/Gel scaffolds.
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Fig. 3

Stress–strain curves for electrospun nanofibers and the insets shown are the water contact angle values

The mechanical properties of the scaffolds are very critical, especially while choosing substrates for vascular tissue engineering. The objective of this study was to imitate the vascular ECM as much as possible, which means the fabricated substrate should possess some of the intrinsic mechanical properties of the native vascular tissue. The mechanical properties of different electrospun scaffolds, including their tensile strength and ultimate strain, were evaluated during this study. Figure 3 shows the typical stress–strain curves of the electrospun scaffolds. Mechanical property evaluation of the scaffolds in Fig. 3 shows a linear segment up to the proportionality limit, followed by a non-linear curve, which was characterized by considerable elongation without a corresponding increase in loading stress. This feature is attributed to the random and interlacing arrangement of the nanofibers; under a continuous stress, the randomly orientated fibers would slide along each other, rearranging themselves in the direction of stress. The Young’s modulus was calculated using the Secant modulus method as the curve is non-linear; here, a tangent was drawn to the curve from the origin and the slope of this tangent was taken as the Young’s modulus. The Young’s modulus of the PU/protein scaffolds ranged from 5.03 to 16.67 MPa. Dacron (PET), the commonly used large-diameter vascular graft, has a tensile modulus of about 14,000 MPa, is a very stiff material, and cannot be utilized as small-diameter vessels such as the coronary arteries, mainly because the low blood flow along with high shear makes it more prone to thrombus formation and intimal hyperplasia [44]. Ideally, less stiff (more compliant) and biodegradable scaffolds, such as the PU/protein nanofibrous scaffolds prepared in this study, are competent to endure high physiological pressure and are better choices for constructing a complete biological and fully functional coronary artery. The tensile properties of native coronary artery and electrospun PU/protein scaffolds obtained from four independent experiments are summarized in Table 1. Compared to the tensile properties of native coronary artery (Table 1), we found that the electrospun composite PU/protein scaffolds had sufficient tensile strength and elastic modulus to be utilized as a vascular graft. The tensile properties of PU/protein nanofibers fabricated during our study were also found better than the tensile strength (3.23 ± 0.57) and modulus (1.2 ± 0.3) of the cell–matrix-engineered electrospun poly(lactide-co-ε-caprolactone) vascular grafts reported by Mun et al. [45]. The PU/protein nanofibrous scaffolds are much less stiff (more compliant) compared to the poly(l-lactic acid)-co-poly(ε-caprolactone) (PLCL) nanofiber mesh, which has a tensile modulus of about 44 MPa, utilized as a vascular graft by He et al. [14]. Comparing the tensile properties among the four PU/protein nanofibrous scaffolds, we found that all the electrospun PU/protein scaffolds had comparative tensile strain except the PU/Fib scaffolds. However, the PU/Coll and PU/Gel scaffolds had the highest tensile strength compared to the PU/Fib and PU/BSA scaffolds. Our results suggested that the electrospun PU/Coll and PU/Gel nanofibrous scaffolds might be more suitable than PU/Fib and PU/BSA to provide the necessary mechanical properties required for blood vessel engineering.
Table 1

Tensile properties of electrospun nanofibers and coronary artery

Mechanical property

PU/Coll nanofibers

PU/Gel nanofibers

PU/Fib nanofibers

PU/BSA nanofibers

Coronary arterya

Ultimate tensile stress (MPa)

11.49 ± 1.31

11.64 ± 1.53

9.81 ± 0.87

9.08 ± 0.75

1.4–11.14

Ultimate strain (%)

128.75 ± 11.38

141.21 ± 10.47

171.78 ± 18.35

140.83 ± 16.24

45–99

Tensile modulus (MPa)

16.67 ± 2.17

6.25 ± 0.57

5.03 ± 0.02

11.54 ± 1.39

aSee Ref. [14]

Cell proliferation on electrospun nanofibers

The proliferation capacity of SMCs on electrospun nanofibers after 3, 6, 9, and 12 days was determined by MTS assay (Fig. 4). It was observed that the optical density of SMCs on all the electrospun substrates increased during the 12 days of cell culture. Results suggested that the SMCs were metabolically active on protein-containing nanofibers and demonstrated the lack of cytotoxic effects of these scaffolds. After 3 days of cell culture, the proliferation of SMCs on PU/Coll and PU/Gel nanofibers was found significantly higher (p ≤ 0.05) than the cell proliferation on PU/Fib and PU/BSA. After 12 days of cell culture, the cell proliferation on PU/Coll was the highest among all the electrospun nanofibrous scaffolds. We observed 6.6, 42.3, and 24.8 % higher cell proliferation on PU/Coll compared to the cell proliferation on PU/Gel, PU/Fib, and PU/BSA nanofibers, respectively. This is mainly because of the presence of collagen within the scaffold, whereby collagen serves a major protein of the native artery, providing biological signals to adjacent cells, has high water affinity, low antigenicity, and good cell compatibility [13]. While, gelatin is a derivative of collagen and it also has merits including high water affinity and good cell compatibility. Hence, the incorporation of either collagen or gelatin in PU enhanced the cell proliferation on both PU/Coll and PU/Gel nanofibers significantly. On the other hand, although a researcher found that the incorporation of fibrinogen can also increase the cell growth [29], compared to collagen- and gelatin-containing PU scaffolds, SMCs cultured on PU/Fib scaffolds showed the least proliferation, and this is because fibrinogen is a protein critical toward the coagulation of blood; during this study, we observed the tendency of SMCs to aggregate together on the PU/Fib scaffolds, influencing cell proliferation. Results of the cell proliferation assay suggested that the electrospun PU/Coll and PU/Gel nanofibers are more cell compatible, less cytotoxic, and could be more suitable as a vascular graft.
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Fig. 4

SMC proliferation on electrospun nanofibers and TCP as determined by MTS assay. *Significant compared with cell proliferation on PU/Fib at P ≤ 0.05

Morphological evaluation of SMCs on electrospun nanofibers

Evidence suggesting cell morphology and cell–biomaterial interaction was observed via SEM and the results are shown in Fig. 5. SMCs attached on the scaffolds and stretched across the nanofibrous substrates upon proliferation, and on the nanofibrous scaffolds (Fig. 5a–d), the cell filopodia came in direct contact with the neighboring cells as well as with the nanofibrous architecture compared to those observed on TCP (Fig. 5e). After 6 days of cell culture, the number of SMCs attached on PU/Coll and PU/Gel (Fig. 5a, b) nanofibers was much higher than the cells attached on PU/Fib and PU/BSA nanofibers (Fig. 5c, d). The SMCs attached on PU/Coll and PU/Gel nanofibers showed a characteristic spindle shape, while the SMCs attached on PU/Fib and PU/BSA nanofiber showed irregular morphology. In addition, the SMCs attached on PU/Coll and PU/Gel nanofibers showed a highly aligned phenotype (Fig. 5a, b). The key feature of arterial microarchitecture is the alignment of smooth muscle cells (SMCs) with their long axis extension toward the circumferential direction in the medial layer [46]. Vasoactivity, the constriction or dilation of blood vessels, is controlled by the contractile force produced by aligned SMCs, and the durable mechanical properties of arteries can be attributed to the alignment of SMCs and their fibrous extracellular matrix (ECM) [47]. Therefore, the alignment of SMCs is necessary for tissue-engineered blood vessels to perform their biologic functions in vivo. It has been reported that scaffolds with aligned fiber orientations may promote cells alignment through contact guidance with the nanofibers [4850]. Alternatively, mechanical forces are also applied in various studies to induce aligned orientation of cells through mechanical stimuli [3]. However, in this study, we used random nanofibers and we were able to promote cell alignment without any external stimuli. SMCs cultured on random PU/Coll and PU/Gel nanofibers showed aligned orientation of cells, suggesting that the collagen- and gelatin-containing PU scaffolds can create a residual stress environment similar to that found in native arteries, which encouraged the orientation of SMCs along the principal stress directions of the artery. Instead of using aligned fibers, with appropriate selection of polymers, a composite elastomeric polymeric scaffold could be highly preferred for vascular tissue engineering. Our results suggest that PU/Coll and PU/Gel nanofibrous scaffolds have the ability to direct cellular alignment without the use of external stimuli or compaction, giving them a distinct advantage for vascular tissue regeneration.
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Fig. 5

SEM images of SMCs on electrospun a PU/Coll, b PU/Gel, c PU/Fib, d PU/BSA nanofibers, and e TCP, after 6 days of cell proliferation

Expression of smooth muscle cell markers

To validate the SMC phenotypic protein expressions, immunofluorescent staining for alpha smooth muscle actin (SMA) and smooth muscle myosin heavy chain (MHC) was carried out for cells cultured on the nanofibrous scaffolds after 9 days of culture time. Figure 6 shows the representative images of fixed and SMA immunostained SMCs on different substrates. Indeed, alpha smooth muscle actin (SMA) is a typical protein that is one of the most abundantly present earliest markers of SMCs [51]. Confocal microscopy images showed that SMCs grown on the nanofibrous scaffolds and TCP stained positively for SMC markers. However, SMCs on both PU/Coll and PU/Gel nanofibers expressed higher density of SMA and interacted well with the nanofibers and were elongated along the nanofibers, orientating with the sarcomeric structures of mature smooth muscle cells. In addition, cells on PU/Coll and PU/Gel nanofibers exhibited SMA markers with aligned microfilament bundle phenotype, suggesting that the PU/Coll and PU/Gel nanofibers could potentially improve the chemical signal transport and force exerted by SMCs, which is crucial for vascular regeneration. Nivison-Smith reported expression of SMA filament after seeding SMCs on electrospun synthetic elastic fibers [52], and our studies showed more strong, uniform, and aligned SMA expression of SMCs cultured on PU/Coll and PU/Gel. Taken together, the results from the SMCs cultured on PU/Coll and PU/Gel materials demonstrated maintenance in SMC proliferation over time, aligned spindle-like morphology, and actin filament organization.
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Fig. 6

Immunocytochemical analysis for the expression of nuclei stained with DAPI (A1E1); (A2E2) SMC marker protein SMA and merged images showing the dual expression of both DAPI and SMA (A3–E3) on PU/Coll, PU/Gel, PU/Fib, and PU/BSA nanofibers; and TCP at ×60 magnification

The functionality of SMCs on the electrospun nanofibers was further elucidated by smooth muscle myosin heavy chain (MHC) staining. Smooth muscle myosin heavy chain (MHC) is the most discriminating marker for SMCs identified to date [4]. Figure 7 shows that the SMCs expressed MHC protein on the nanofibrous scaffolds. Similar to the expression of SMA, the SMCs seeded on PU/Coll and PU/Gel nanofibers demonstrated uniform high density of MHC with alignment orientation. On the other hand, SMCs seeded on PU/Fib and PU/BSA scaffolds showed random and uneven expression of MHC in much lesser amounts. Blit et al. [53] reported that SMCs cultured on elastin-like polypeptide-modified PU nanofibers showed the expression of MHC; our studies showed stronger, more uniform, and aligned MHC expression of SMCs cultured on PU/Coll and PU/Gel. Taken together, the results from the SMCs cultured on PU/Coll and PU/Gel materials demonstrated maintenance in SMC proliferation over time, aligned spindle-like morphology, actin filament organization, and MHC cell expression. Electrospun PU/Coll and PU/Gel nanofibrous scaffolds had sufficient mechanical properties, supported SMC proliferation, and assisted in oriented morphological alignment of cells, with potential for vascular tissue engineering.
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Fig. 7

Immunocytochemical analysis of cells grown on different scaffolds and TCP showing the expression of DAPI (A1, B1, C1, D1, E1); SMC-specific marker protein MHC (A2, B2, C2, D2, E2); merged image showing the dual expression of (A3, B3, C3, D3, E3) at ×60 magnification

Conclusion

In the present study, four different kinds of natural proteins were incorporated with PU to electrospin PU/protein nanofibers, and the proliferation of SMCs on the electrospun scaffolds was evaluated by a cell proliferation assay. Our results suggested that the SMCs were metabolically active on protein-containing nanofibers, especially on PU/Coll and PU/Gel nanofibrous scaffolds. Cell morphology and cell–biomaterial interaction from the SEM observation showed high coverage of SMCs on PU/Coll and PU/Gel nanofibers over 6 days of culture and the scaffold supported cell alignment over its surfaces. SMCs grown on PU/Coll and PU/Gel nanofibrous scaffolds exhibited a spindle-like morphology, actin filament organization, and MHC expression specific of vascular SMCs. Our study suggests PU/Coll and PU/Gel nanofibrous scaffolds as promising substrates for application as functional blood vessels.

Acknowledgements

This research was supported by an NRF-Technion grant (R-398-001-065-592) and the Nanoscience and Nanotechnology Initiative, Faculty of Engineering, National University of Singapore, Singapore. This work was also partly supported by grants (50973014 and 11172064) from the National Natural Science Foundation of China and the Foundation for the Author of National Excellent Doctoral Dissertation of P.R. China(200961), as well as sponsored by the Shanghai Rising-Star Program in Chian(10QA1400100) and the Fok Ying Tong Education Foundation(121071) to Prof. Xiaohong Qin. It was also supported by the Program for New Century Excellent Talents in University (NCET-10-0322) and the Fundamental Research Funds for the Central Universities as well as the “Shu Guang” (11SG33) project supported by Shanghai Municipal Education Commission and Shanghai Education Development Foundation. Furthermore, the first author would also like to acknowledge the China Scholarship Council for granting a scholarship that enables her to pursue this work at NUS, Singapore.

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© Springer Science+Business Media New York 2013