Design and Fabrication of Tubular Scaffolds via Direct Writing in a Melt Electrospinning Mode
© The Author(s) 2012
Received: 5 November 2011
Accepted: 9 December 2011
Published: 9 February 2012
Flexible tubular structures fabricated from solution electrospun fibers are finding increasing use in tissue engineering applications. However it is difficult to control the deposition of fibers due to the chaotic nature of the solution electrospinning jet. By using non-conductive polymer melts instead of polymer solutions the path and collection of the fiber becomes predictable. In this work we demonstrate the melt electrospinning of polycaprolactone in a direct writing mode onto a rotating cylinder. This allows the design and fabrication of tubes using 20 μm diameter fibers with controllable micropatterns and mechanical properties. A key design parameter is the fiber winding angle, where it allows control over scaffold pore morphology (e.g. size, shape, number and porosity). Furthermore, the establishment of a finite element model as a predictive design tool is validated against mechanical testing results of melt electrospun tubes to show that a lesser winding angle provides improved mechanical response to uniaxial tension and compression. In addition, we show that melt electrospun tubes support the growth of three different cell types in vitro and are therefore promising scaffolds for tissue engineering applications.
Tubular scaffolds fabricated from electrospun fibers are finding increasing use in tissue engineering (TE) applications, including vascular [1–11] (reviewed in detail by Naito et al.), neural [12–14] (reviewed in detail by Bell et al.) and more recently growth factor delivery [15–19]. Typically, solution electrospun fibers are collected as thin non-woven meshes , which has limited their application because the small pore sizes and compromised interconnectivity associated with the random layering of sub-micron diameter fibers acts as a barrier to rather than promotes cell infiltration and subsequent vascularization .
For the design and fabrication of tubular scaffolds, control over both the microscopic and macroscopic structure should be reflected in cellular and mechanical responses at varied anatomic locations and biological environments . For example, differences in pressure-dependent mechanical properties between native arteries and artificial grafts induce hydrodynamic flow disturbances and stress concentrations, thereby causing tissue damage as well as impairing cellular function, illustrating the need to match compliance of a designed artificial graft with a small-diameter artery . Whereas rigid nerve guides used in neural TE may cause inflammation due to the stresses exerted by movement, electrospinning allows the fabrication of flexible microfiber and nanofiber guides from the same materials . In bone regeneration, the presence of a solution electrospun tubular scaffold promotes mineralized matrix synthesis, prevents extra-anatomical mineralization, and guides an integrated pattern of bone formation during osteogenic protein delivery in the functional repair of large bone defects in rats . Scaffold morphology which has been studied in terms of tissue growth includes pore size and, more recently pore geometry. Pore size affects cellular activities such as pore spanning (with this ability varying depending on cell type), and the spatial organization of cells within pores. Cell number can also be affected, where a larger pore size often leads to a reduced surface area to volume ratio which then effects cell attachment and proliferation. The effect of pore geometry on tissue growth, such as the influence on tissue orientation and the pattern of tissue growth within pores has been investigated to a lesser extent .
What makes this area a particularly promising one is that electrospinning makes it possible to fabricate flexible microfiber and nanofiber tubular scaffolds from polymeric materials, using a broad range of controlled morphologies. In the majority of cases rotating devices are used to fabricate electrospun fibrous tubes. However it is difficult to accurately control fiber arrangements due to the inherent chaotic nature of the electrified jet in solution electrospinning. Although there are approaches for the organized deposition of fibers reported in the literature [22, 24–27], there still remains considerable difficulties in fabricating fibrous tubes with controllable micropatterns . Added to this, charge accumulation effects on solution electrospun fibers tend to restrict the amount of layers which can be collected and remain bound as one coherent structure . As an alternative to solution electrospinning, melt electrospinning permits improved control over the location of fiber deposition and avoids technical difficulties and cytotoxicity concerns associated with the removal and the presence of residual of solvent [24, 30].
In contrast, our group has shown most recently that scaffolds with a thickness of up to 1 cm can be designed and manufactured with controlled architectures in a direct writing mode, demonstrating that melt electrospun fibers can be readily placed on top of each other, as the deposited fibers have minimal residual charge . In this work, we demonstrate melt electrospinning in a direct writing mode onto a rotating cylindrical collector. This allows the design and fabrication of tubular scaffolds of any size using micron diameter fibers, with controllable microscopic and macroscopic architectures and mechanical properties. A key design parameter is the fiber winding angle, which allows control over scaffold pore morphology (e.g. size, shape, number and porosity) and influences the mechanical properties of such a tube. Furthermore, we establish a finite element (FE) model which can be used as a predictive design tool, where in this case the simulated response to uniaxial loading of simplified tubular structures varies when the fiber angle is altered. This response is validated against mechanical testing results of fabricated electrospun tubes. We further show that the electrospun tubes support the growth of three different cell types in vitro and are therefore promising scaffolds for TE applications.
2.1 Experimental Materials
Poly(ε-caprolactone) (PCL) with MW = 50 kDa was kindly supplied by Perstorp, UK Ltd. (United Kingdom) and used as received. The superior rheological and viscoelastic properties within the group of aliphatic polymers makes PCL the polymer of choice to be applied during the development of scaffolds based on melt extrusion processes. Coupled with relatively inexpensive production routes and FDA approval, this provides a promising platform for the production of longer-term degradable and biocompatible implants which may be manipulated physically, chemically and biologically to possess tailorable degradation kinetics to suit a specific anatomic site as reviewed in detail by Woodruff and Hutmacher .
2.2 Fabrication of Tubes
2.2.1 Melt Electrospinning
PCL pellets were loaded into a plastic Luer-lock 3 mL syringe (B-Braun, Australia). The syringe was placed in a custom glass water jacket housing (Labglass, Australia) through which heated water at 78°C was circulated using a recirculating water tank (Ratek, Australia). The syringe was heated for 1 h to provide a homogenous polymer melt. A 21 G hypodermic needle (Becton–Dickinson, Australia) with the tip flattened was attached to the syringe to be used as the spinneret. The feeding rate of the polymer melt in the syringe was controlled at 50 μL/h using a programmable syringe pump (World Precision Instruments, USA). A high voltage (Emco High Voltage Corporation, USA) of 12 kV was applied to the needle and the distance between the tip of the needle and the rotating collector was 40 mm.
2.2.2 Single Fiber Production
To produce single melt electrospun fibers, the electrospinning settings described above resulted in an average fiber diameter of 60 μm. The fibers were collected on a flat collector translating at 250 mm/min to draw them into a straight line.
2.2.3 Tube Winding
Parameters used to fabricate melt electrospun tubes with different winding angles
Translational speed (mm/min)
Effective translational speed (mm/min)
Number of collector rotations/translation
Tangential speed (mm/min)
Resultant vector speed (mm/min)
Effective resultant vector speed (mm/min)
Actual winding angle (°)
Effective winding angle (°)
Measured winding angle (°)
29.3 ± 2.3
45.4 ± 2.2
58.3 ± 3.0
29.1 ± 2.8
41.1 ± 4.8
59.0 ± 2.8
2.3 Mechanical Testing
Mechanical testing of single fibers in tension, as well as tubes in tension and compression was undertaken. An Instron 5848 Micro Tester with a 5 N load cell (Instron, Norwood, MA) was used to apply static forces. The rigid load frame was configured vertically to suit each testing case, with the set up consisting of mounted grips or plates to fix the specimens between the base beam and the movable actuator . In each tube testing case, displacement was applied and the reaction force measured by the 5 N load cell recorded using Instron WaveMaker Runtime 32 software (Instron, Norwood, MA). Single melt electrospun fibers were placed between air pressure grips separated by 10 mm and extended up to 225% at a 1%/s rate at room temperature (n = 10) . Compression and tension tube tests were performed for cases 1a–1c, by applying a 10% deformation with a strain rate of 1%/s (n = 5) . For tensile testing of tubes, small plastic 6 mm diameter cylinders (2 mm thick) were glued to the top of modified syringe plungers for fixation to air pressure grips in the Instron Micro Tester. Each end of a tubular sample was bonded using epoxy glue to a fixation cylinder and then mounted in the Micro Tester using compressed air clamps. For compression, tubular samples were placed between a flat plate and a 10 mm diameter cylinder mounted on the moving actuator, so that the axis of the tube was in line with the central axis of the moving actuator.
For morphology observation, scaffold surfaces were sputter coated with gold (a 10 nm thick layer) using a Leica Microsystems EM SCD005 (Germany) and scanning electron microscopic (SEM) examination carried out using an FEI Quanta 200 Environmental SEM (Netherlands) at an accelerating voltage of 10 kV. Images were prepared and processed using Corel DRAW X4 and PHOTO-PAINT X4 (Corel Corporation, Australia) and Adobe Photoshop and Illustrator CS4 (Adobe Systems Incorporated, Australia) using image auto-adjust functions. For fiber diameter measurements, five images were taken at randomly chosen locations for each winding angle case. Images were imported into ImageJ 1.41 software (NIH, USA) and ten fiber diameter measurements taken for each fiber orientation. Similarly on these images, for pore size measurements a polygon was drawn along the edges of four intersecting fibers to define the outer bounds of a void (or pore) created by the fibers . For porosity measurements, a middle 3 mm section of each tubular sample was analyzed using micro computed tomography (μCT). Specimens were scanned on a Scanco μCT40 (Scanco Medical AG, Switzerland) at 6 μm resolution, employing 55 kV and 145 μA with 500 ms integration time. After segmentation, a bone morphometric analysis algorithm was used to calculate porosity as the ratio of the material volume to the total volume scanned. The mean of measured values, standard error of the means, and differences between the means (using one-way ANOVA with the assumption of normally distributed data) were then calculated using PASW Statistics 18 software (SPSS Inc., USA). Optical images of tubes and experimental conditions were taken using a Canon EOS 450D digital single lens reflex (DSLR) camera with a Canon EF-S 17-85 mm USM lens (Canon.Inc, Australia).
2.5 Finite Element Modeling
2.5.1 Preparation of Model for Mechanical Testing Simulation
Tube geometry was prepared in ANSYS® Mechanical APDL, Release 12.1 (ANSYS Inc., USA). To simulate the winding process, helical lines were created in pairs representing one back and forth translation of the rotating mandrel on which electrospun fibers were collected. Each fiber pair originated at the same location, with one helix line directed clockwise and the other counter clockwise. For computational efficiency, ten fiber pairs were distributed equally around the circumference of a 6 mm diameter circular base (assuming equally distributed fibers). Multiple geometries were created where the winding angle was varied from 15° to 85° , however in each case the total tube height was 10 mm.
Three noded quadratic BEAM188 elements with a circular cross section of 60 μm in diameter (resultant diameter of single fiber fabrication) were used to mesh the geometry. Enabling shear stress calculations through the elements by using Thimoshenko beam theory, the BEAM188 element is the most accurate beam element provided by ANSYS 12.1 and suits the fiber geometry in this model . In order to accurately represent the circular structure, the elements were used in their quadratic form. Following refinement analysis, the division arc spanned by one element was defined to be not bigger than 3° to be able to approximate the circular structure using straight elements . The helical lines were defined for two cases: independent of each other (not connected) to obtain information about influence of winding angle on the stability of the geometry in terms of single fibers; and alternatively, contact between the elements where the helical lines crossed each other was defined as bonded to represent fusion between the fibers during melt electrospinning.
2.5.3 Material Model
In each simulation case, all nodes atz = 0 (bottom of the tube) were constrained in all degrees of freedom (fixed supports). All nodes located atz = 10 mm were loaded with a deformation of ±1 mm parallel to the axis of symmetry to simulate 10% uniaxial tension and compression (Fig. 1b). For model validation, each geometry variant was rotated 45° about the central axis of the tube (torsion) (Fig. 1c).
The reaction forces at the end of each fiber (z = 10 mm) were added, giving a total reaction force in the tensile and compressive loading cases. The total torsion reaction was calculated by transforming the resultant x and y directed forces into radial and tangential forces, then summing and dividing them by the associated arc length.
2.6 In Vitro Biocompatibility Assessment
Primary human osteoblasts (hOBs) and mesothelial cells as well as mouse osteoblasts (mOBs) were utilized in this study in order to demonstrate that the electrospun fibers comprising the tubes are non-cytotoxic and support cell colonization.
2.6.1 Biomimetic Coating of PCL Tubular Scaffolds for In Vitro Studies with Osteoblasts
To enhance osteo-induction, PCL tubes were coated with a layer of calcium phosphate (CaP). The coating process consisted of three steps: surface activation with alkaline treatment [Sodium hydroxide (NaOH)]; treatment with simulated Body Fluid 10× (SBF10×) to deposit the CaP; and post-treatment with NaOH. The preparation of SBF10× was adapted from Yang et al. . For 1 L of solution, reagents were dissolved in ddH20 in the following order: 58.430 g NaCl, 0.373 g KCl, 3.675 g CaCl2·2H2O and 1.016 g MgCl2·6H2O. The next reagent (1.420 g of Na2HPO4) was dissolved separately in 20 mL of ddH20 and added drop by drop into the main solution while maintaining the pH level at 4 by adding hydrochloric acid (HCl) 32% in order to avoid precipitation of calcium cations and phosphate anions. The tubes were first cleaned by immersion in 70% ethanol solution under vacuum for 15 min for the purpose of removing entrapped air bubbles, then the structures were immersed into pre-heated (37°C) NaOH 2 M and a 5 min vacuum treatment was performed at room temperature. For the rest of the activation step, the scaffolds were placed at 37°C for 30 min to accelerate the etching process. The tubes were then rinsed with ddH2O until the pH level dropped to approximately 7. Meanwhile, NaHCO3 was added to the SBF10× solution until a pH of 6 was reached. This activated SBF solution was filtered (0.2 μm filter) and another 5 min vacuum treatment at room temperature was performed to ensure that the solution fully penetrated the tubes. The samples were thereafter placed at 37°C for another 30 min. The solution was replaced with freshly activated and filtered SBF and placed again at 37°C for 30 min. The tubes were rinsed in ddH2O and then immersed in pre-heated NaOH 0.5 M for 30 min at 37°C. Finally the tubes were rinsed with ddH2O until the pH level dropped to approximately 7 and then dried overnight in a dessicator.
2.6.2 Human Primary Osteoblast Cell Culture and Seeding onto PCL Tubular Scaffolds
hOBs were isolated as previously described . Cells were maintained in culture up to passage 4 in alpha minimum essential media (α-MEM) (Invitrogen, Australia) supplemented with 10% FBS (Invitrogen, Australia), 100 IU/mL penicillin (Invitrogen, Australia), and 0.1 mg/mL streptomycin (Invitrogen, Australia). Prior to cell seeding, PCL tubes were sterilized by immersion in 70% ethanol for 30 min, followed by 20 min UV exposure. Dry scaffolds were transferred into a 24-well plate and 1.6 × 105 hOBs suspended in 40 μL of culture media were equally distributed onto the scaffolds. After leaving the moistened scaffold for 2 h at 37°C, during which initial cell attachment to the scaffold occurred, 1 mL of culture medium was carefully added. Cells were grown over a total culture period of 4 weeks in 5% CO2 and at 37°C. Medium was changed every 3 days. After 2 weeks of culture, cells were cultured under osteogenic conditions (i.e. in cell culture medium supplemented with 50 μg/mL ascorbate-2-phosphate (Sigma-Aldrich, Australia), 10 mM β-glycerophosphate (Sigma-Aldrich, Australia), and 0.1 μM dexamethasone (Sigma-Aldrich, Australia)).
2.6.3 Mouse Primary Osteoblast Cell Culture and Seeding onto PCL Tubular Scaffolds
MOBs were received as a gift from Dr. Jean-Pierre Levesque (Mater Medical Research Institute, Brisbane, Australia). Cells were maintained in culture in α-MEM (Invitrogen, Australia) supplemented with 10% FBS (Invitrogen, Australia), 100 IU/mL penicillin (Invitrogen, Australia), 0.1 mg/mL streptomycin (Invitrogen, Australia) and 50 μg/mL ascorbate-2-phosphate (Sigma-Aldrich, Australia). Prior to cell seeding, PCL tubes were sterilized by immersion in 70% ethanol for 30 min, followed by 20 min UV exposure. Dry scaffolds were transferred into a 24-well plate and 3 × 105 mOBs suspended in 37.5 μL culture media were equally distributed onto the scaffolds. After leaving the moistened scaffold for 2 h at 37°C, during which initial cell attachment to the scaffold occurred, 1 mL of culture medium was carefully added. Cells were grown over a total culture period of 4 weeks in 5% CO2 and at 37°C. Medium was changed every 3 days. After 2 weeks of culture, cells were cultured under osteogenic conditions, i.e. in cell culture medium supplemented with 50 μg/mL ascorbate-2-phosphate (Sigma-Aldrich), 10 mM β-glycerophosphate (Sigma-Aldrich), and 0.1 μM dexamethasone (Sigma-Aldrich).
2.6.4 Culture and Seeding of Human Mesothelial Cells onto PCL Meshes
Human mesothelial Met-5A cells were obtained from the American Type Culture Collection (ATCC) and maintained in the recommended media199 . Briefly, 2 × 105 cells were seeded on top of sterilized PCL meshes, grown for up to 28 days and processed using confocal laser scanning microscopy (CLSM) as described previously .
2.6.5 Analysis of Cell Viability
After 28 days of 3D culture, cell viability was assessed by fluorescein diacetate (FDA) (Invitrogen, Australia) and propidium iodide (PI) (Invitrogen, Australia) staining. Samples were rinsed three times with pre-warmed phenol red free α-MEM (Invitrogen, Australia). Samples were then incubated in 2 μg/mL FDA and 10 μg/mL PI in phenol red-free medium at 37°C for 15 min and washed using phosphate buffered saline (PBS). Thereafter, the hydrated specimens were immediately imaged using a confocal microscope (TCS SP5 II, Leica, Australia) with a 10/20× immersion oil objective. 2 μm thick Z–stacks were acquired over a total height of 400–600 μm and from these, 3D projections were generated using LAS AF software (v.1.8.2 build 1465, Leica Microsystems, Australia). Then, the scientific image processing software Imaris (x64; v.7.3.0, Bitplane, Switzerland) was used to prepare 3D reconstructions.
2.6.6 Phalloidin/DAPI Staining of Cells Growing on PCL Scaffolds
Cells were fixed with 4% paraformaldehyde for 20 min at room temperature (RT) and permeabilized using 0.2% Triton X-100 in PBS for 5 min at RT. Thereafter, samples were washed twice with PBS and blocked for 10 min in 2% bovine serum albumin (BSA) (Sigma, Australia) in PBS. Samples were then incubated with 5 μg/mL 4′,6-diamidino-2-phenylindole (DAPI) (Invitrogen, Australia) and 200 U/mL rhodamine-conjugated phalloidin in 2% BSA/PBS for 1 h at RT. After washing with PBS, hydrated samples were imaged using a laser scanning confocal microscope (TCS SP5 II, Leica). Z-stacks and 3D projections were prepared as described above.
2.6.7 SEM Analysis of Cells Growing on PCL Scaffolds
Samples were fixed with 3% (v/v) glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.3) overnight at 4°C. Fixed specimens were washed in 0.1 M sodium cacodylate buffer and dehydrated through a graded series of ethanol. Samples were then incubated in hexamethyldisilazane (HMDS) (Prositech, Australia) for 30 min and air-dried. Specimens were mounted and gold coated (SC500, Bio-Rad, Australia) prior to visualization using a Quanta 200 SEM (FEI, The Netherlands).
3 Results and Discussion
3.1 Design of Porous Tubular Structures
For a single layer of wound fibers with diameter of 25 μm, Fig. 2b demonstrates a power law relationship between the number of fiber pairs (as well as the number of pores) and the number of crossover points, which increases with a larger winding angle. Associated with this increase in number of crossover points/pores with more fiber pairs is a decrease in average pore size (assuming a homogenous distribution of fibers) which becomes more apparent with increasing winding angle (Fig. 2c).
Assuming a fixed fabrication time, a constant amount of material should be collected. However as the winding angle increases, so will the length of a single fiber from one end of the tube to the other (assuming the tube height is fixed). Therefore, this will be accompanied by an associated decrease in fiber diameter to maintain a constant volume of material. As well as showing the reduction in porosity as the number of fibers increases, Fig. 2d illustrates a decrease in porosity with increasing winding angle. This can be explained by the reduced volume occupied by the smaller diameter fiber associated with increased winding angle.
The relationships between the parameters described above are presented in the supplementary information. This allows the design of tubes with control over features such as pore size, shape and number as well as total porosity, based on the following input parameters: tube height and diameter, fiber diameter, number of fiber pairs, and winding angle. Their choice and balance between them allows the tailoring of the pore architecture to suit a specific TE application. For example, for a desired pore size and using a fixed number of fibers, a large winding angle such as 60° allows for an increased number of crossover points (for tissue spanning) and surface area to volume ratio than compared to a more acute angle. The associated decrease in pore size reduces the potential for cell and tissue infiltration which may be favorable for example to neural compared to vascular applications.
3.2 Mechanical Simulation
It has been observed that melt electrospun fibers may solidify prior to deposition (Fig. 3c) or alternatively, fuse at the points of contact if insufficient cooling has taken place prior to deposition (Fig. 3d). Matsuda et al. reported an increase in stiffness of an electrospun tubular construct with an increased degree of fusion or welding at the contact points with other fibers, attributed to the restricted freedom of movement of fibers under mechanical stretching. Therefore, a physically entangled mesh should exhibit a higher degree of fiber extension upon stretching than with bonded fibers .
For a melt electrospun tube produced in a direct writing mode with the same number of fiber pairs, a larger winding angle will facilitate more fiber bonding sites. However, for the loading cases of uniaxial tension and compression, this could be accompanied by a sacrifice in the stiffness of the construct as the orientation of the fibers diverges further from the direction of loading. Figure 3 illustrates the results of simulated uniaxial tensile and compressive loading of a tubular construct for two cases: where there is no bonding between fibers (Fig. 3a), and where the fibers are fully bonded (Fig. 3b). These results show an improvement in stiffness when the fibers are bonded compared to unbonded. In the unbonded case (Fig. 3a), stiffness reduces as the winding angle is increased from 30° to 60° according to a power law. Whereas in the bonded case (Fig. 3b), stiffness reduces linearly as the winding angle is increased from 30° to 60°. However, the fact that stiffness continues to reduce with increasing winding angle for the bonded case suggests that the orientation of fibers closer to the axis of loading improves stiffness more significantly than the increased amount of fiber bonding sites provided by fibers orientated further from the axis. Furthermore, the nature of the difference between the stiffness response (power law vs. linear) with varied winding angle to such loading cases may be used to provide an indication as to the degree of fusion between fibers in the constructs.
A limitation of this model is shown by the fact that the compressive and tensile results are equal, due to the linear approximation for the material model based on tensile testing results. With further development, such as the inclusion of compressive material properties (which would account for failure of slender fiber segments in compression due to buckling as observed in Fig. 6b) as well as cyclic loading cases (to better represent in vivo loading) such a predictive design tool could be applied to more complicated combined variable loading cases, which may benefit from the design of structures with increased complexity, such as multi-angle wound tubes.
3.3 Fabrication of Melt Electrospun Tubes in a Direct Writing Mode
Fabrication times and resultant dimensions for two different strategies to fabricate melt electrospun tubes
Fabrication time (min)
Effective resultant vector speed (mm/min)
Average fiber diameter (μm)
15.2 ± 3.3
25.7 ± 1.8
14.8 ± 2.5
22.4 ± 1.8
15.7 ± 2.9
19.9 ± 0.9
11.1 ± 1.2
20.4 ± 1.8
17.5 ± 0.7
26.3 ± 3.2
37.4 ± 1.3
27.7 ± 2.4
3.3.1 Collection Speeds
By maintaining a constant Tp in case 1, the fabrication time (to translate back and forth 700 times) was fixed. This should ensure a constant amount of material collected. Table 2 shows that for a fabrication time of 22 min the weight of tubes fabricated in this manner was relatively uniform. However, because the length of a single fiber from one end of the tube to the other will increase with an increase in winding angle, this should be accompanied by an associated decrease in fiber diameter to maintain a constant volume, shown in Table 2.
As an alternative method to control the winding angle, the tangential speed was fixed and Tp varied. The tangential speed of 637 mm/min used in case 1 for a 45° winding angle was chosen as a reference point so that the translational speed could be varied above and below the translational speed matching this speed to obtain winding angles of 30 and 60° (Table 1). However, an approximate 1.7 fold increase and decrease in the translational speeds was required to achieve the desired changes in winding angle. Because of the associated differences in fabrication time and total volume of the tubes fabricated in this manner (Table 2), this method was not preferred for subsequent characterization and comparisons of mechanical properties based on winding angle.
3.3.2 Pore Size and Porosity
Predicted and measured values for porosity and pore size for tubes fabricated with varied winding angle
Predicted porosity (unbonded) (%)
Predicted porosity (bonded) (%)
Measured porosity (%)
Predicted pore size (×10−2 mm2)
Measured pore size (×10−2 mm2)
6.9 ± 1.9
5.0 ± 1.1
4.8 ± 0.6
Also shown in Table 3 are predicted values for pore size which reduce from 7.84 × 10−2 mm2 to 5.28 × 10−2 mm2 as the winding angle is increased from 30° to 60°. The average measured values shown in Table 3 demonstrate that pore size reduces (from 6.9 ± 1.9 × 10−2 mm2 down to 4.8 ± 0.6 × 10−2 mm2) with increased winding angle. The large variation in the measured values can be attributed to the presence of field disturbances during the electrospinning process as well as repulsive coulombic interactions between charged fibers immediately prior to deposition . However, these results demonstrate that choice of design parameters such as number of fibers, fiber diameter and winding angle can be used to predict tube architecture such as pore size, number and total porosity.
3.4 Mechanical Testing
3.5 In Vitro Biocompatibility Assessment
The first step in optimizing TE scaffold design is to understand how scaffold architecture influences cellular activities. From this point of view, solution electrospinning has a number of drawbacks associated with chaotic fiber deposition and thus random assembly characteristic to the process . Therefore, our aim is to translate melt electrospinning, which offers improved control over fiber deposition, into the direct writing of tubular scaffolds based on better control over design parameters. Pore geometry plays a vital role in governing cell infiltration, with the shape of the pore dictating the distribution of cells within, and the resultant pattern of tissue growth. For example, the evaluation of diamond-shaped pores suggests that they are less sensitive to initial conditions of cell attachment than rectangular pores, and thus more effective in guiding engineered tissue cell and collagen orientation . Edwards et al.  report that tissue growth can be predicted based on aspects of pore geometry, namely, the angle between crossing fibers and the distance between adjacent fiber crossovers. Tissue lengths at fiber crossovers were found to decrease exponentially with increasing interfiber angle, as well as decrease away from the fiber crossover, with the smallest lengths toward the fiber segment mid-point. Considering this approach in the design of TE scaffolds, it is envisaged that tissue growth may, in part, be controlled by scaffold fiber orientation (dictating the interfiber angle) and packing density (controlling the distance between adjacent fibers). Therefore, the aim of the following in vitro cultures was to demonstrate the biocompatibility of these structures for future exploration into the aforementioned issues.
3.5.1 Culture of Human Primary Osteoblasts on Tubular PCL Scaffolds
3.5.2 Culture of Mouse Primary Osteoblasts on Tubular PCL Scaffolds
3.5.3 Culture of Human Mesothelial Cells on PCL Scaffolds
4 Summary and Conclusion
Over the last 10 years there have been significant advancements in the field of electrospinning. In spite of this, new breakthroughs in manufacturing are needed to maintain the momentum of the electrospinning field. For the most part, researchers have focused on electrospinning from solution with only a handful attempts to design and fabricate scaffolds from the melt. Our group has made significant progress in the design and fabrication of scaffolds by combining melt electrospinning with a direct writing process, where matching the translation speed of the collector to the speed of the melt electrospinning jet is the key factor which establishes control over the location of fiber deposition in order to write with a continuous line. In this work we were able to accurately deposit melt electrospun fibers and create tubular scaffolds with different fiber winding angles. Control over the relative speed between translation and rotation allowed control over fiber diameter (due to induced drawing) and the winding angle. Combined with the choice of fiber number, these design parameters allowed control over scaffold architecture in terms of number of pores, their size and geometry, as well as total porosity. A computational model was developed and validated against mechanical testing results to show that melt electrospun tubes provide a stiffer response to uniaxial loading when fabricated with a smaller winding angle and no fusion between fibers.
The scaffolds that are used in TE are generally meant to provide provisional substitutes for ECM, providing a temporary structural support combined with specific biochemical signals that encourage cells to create their own ECM environment. However, the ECM is much more than a static mechanical support for tissues. It provides the physical microenvironment of a cell. Cells are not only affected by material composition, but also by the topography and mechanical properties of the scaffold. We have shown in a series of in vitro studies that tubular melt electrospun scaffolds fabricated in a direct writing mode show excellent cell compatibility. Taken together, these in vitro studies demonstrate that hOB, mOB and human mesothelial cells were able to infiltrate entirely the fibrillar scaffolds and that the specific architecture obtained by the combination of melt electrospinning and a direct writing process is favorable for cell spanning between adjacent fibers. Future work of our group is focused on the design and implementation of in vivo studies to further explore the potential of this unique scaffold design concept.
Funding for this work was provided by a grant from the CMF Clinical Priority Program of the AO Foundation (C-10-61H) and Australian Research Council. Thanks to Dr. Ferry Melchels for μCT imaging, QUT Analytical Electron Microscopy Facility for SEM sample preparation and imaging, Dr. Leo DeBoer for confocal imaging and Dr. Jean-Pierre Levesque for providing mOB cells.
- Wu H, Fan J, Chu CC, Wu J (2010) J Mater Sci Mater Med 21:3207View ArticleGoogle Scholar
- Vaz C, Van Tuijl S, Bouten C, Baaijens F (2005) Acta Biomater 1:575View ArticleGoogle Scholar
- Tillman BW, Yazdani SK, Lee SJ, Geary RL, Atala A, Yoo JJ (2009) Biomaterials 30:583View ArticleGoogle Scholar
- Lee SJ, Liu J, Oh SH, Soker S, Atala A, Yoo JJ (2008) Biomaterials 29:2891View ArticleGoogle Scholar
- Thomas V, Donahoe T, Nyairo E, Dean DR, Vohra YK (2011) Acta Biomater 7:2070View ArticleGoogle Scholar
- Stitzel J, Liu J, Lee SJ, Komura M, Berry J, Soker S, Lim G, Van Dyke M, Czerw R, Yoo JJ (2006) Biomaterials 27:1088View ArticleGoogle Scholar
- Matsuda T, Ihara M, Inoguchi H, Kwon IK, Takamizawa K, Kidoaki S (2005) J Biomed Mater Res A 73:125View ArticleGoogle Scholar
- Nottelet B, Pektok E, Mandracchia D, Tille JC, Walpoth B, Gurny R, Moeller M (2009) J Biomed Mater Res A 89:865View ArticleGoogle Scholar
- Ju YM, Choi JS, Atala A, Yoo JJ, Lee SJ (2010) Biomaterials 31:4313View ArticleGoogle Scholar
- Inoguchi H, Kwon IK, Inoue E, Takamizawa K, Maehara Y, Matsuda T (2006) Biomaterials 27:1470View ArticleGoogle Scholar
- Naito Y, Shinoka T, Duncan D, Hibino N, Solomon D, Cleary M, Rathore A, Fein C, Church S, Breuer C (2011) Adv Drug Deliv Rev 63:312View ArticleGoogle Scholar
- Panseri S, Cunha C, Lowery J, Del Carro U, Taraballi F, Amadio S, Vescovi A, Gelain F (2008) BMC Biotechnol 8:1View ArticleGoogle Scholar
- Bini T, Gao S, Tan TC, Wang S, Lim A, Hai LB, Ramakrishna S (2004) Nanotechnology 15:1459View ArticleGoogle Scholar
- Bell J, Haycock JW (2011) Tissue Eng Part B-Rev. doi:10.1089/ten.TEB.2011.0498
- Oest ME, Dupont KM, Kong HJ, Mooney DJ, Guldberg RE (2007) J Orthop Res 25:941View ArticleGoogle Scholar
- Kolambkar YM, Boerckel JD, Dupont KM, Bajin M, Huebsch N, Mooney DJ, Hutmacher DW, Guldberg RE (2011) Bone 49:485View ArticleGoogle Scholar
- Kolambkar YM, Dupont KM, Boerckel JD, Huebsch N, Mooney DJ, Hutmacher DW, Guldberg RE (2011) Biomaterials 32:65View ArticleGoogle Scholar
- Boerckel JD, Kolambkar YM, Dupont KM, Uhrig BA, Phelps EA, Stevens HY, García AJ (2011) Biomaterials 32:5241View ArticleGoogle Scholar
- Ekaputra AK, Prestwich GD, Cool SM, Hutmacher DW (2008) Biomacromolecules 9:2097View ArticleGoogle Scholar
- Cipitria A, Skelton A, Dargaville TR, Dalton PD, Hutmacher DW (2011) J Mater Chem 21:9419View ArticleGoogle Scholar
- Pham QP, Sharma U, Mikos AG (2006) Biomacromolecules 7:2796View ArticleGoogle Scholar
- Zhang D, Chang J (2007) Adv Mater 19:3664View ArticleGoogle Scholar
- Edwards SL, Church JS, Alexander DLJ, Russell SJ, Ingham E, Ramshaw JAM, Werkmeister JA (2010) Tissue Eng Part C 17:123View ArticleGoogle Scholar
- Dalton PD, Klee D, Moller M (2005) Polymer 46:611View ArticleGoogle Scholar
- Vaquette C, Cooper-White JJ (2011) Acta Biomater 7:2544View ArticleGoogle Scholar
- Teo WE, Kotaki M, Mo XM, Ramakrishna S (2005) Nanotechnology 16:918View ArticleGoogle Scholar
- Amoroso NJ, D’Amore A, Hong Y, Wagner WR, Sacks MS (2011) Adv Mater 23:106View ArticleGoogle Scholar
- Zhang DM, Chang J (2008) Nano Lett 8:3283View ArticleGoogle Scholar
- Ramakrishna S, Fujihara K, Teo WE, Lim TC, Ma Z (2005) An introduction to electrospinning and nanofibers. World Scientific Publishing, SingaporeView ArticleGoogle Scholar
- Dalton PD, Joergensen NT, Groll J, Möller M (2008) Biomed Mater 3:034109View ArticleGoogle Scholar
- Brown TD, Dalton PD Hutmacher DW (2011) Adv Mater 23:5651View ArticleGoogle Scholar
- Woodruff MA, Hutmacher DW (2010) Prog Polym Sci 35:1217View ArticleGoogle Scholar
- Instron Corporation (2001) Instron Model 5848 Micro Tester, Reference Manual—Equipment Revision DGoogle Scholar
- Tan EPS, Lim CM (2005) Biomaterials 26:1453View ArticleGoogle Scholar
- Mistry J (1992) Compos Struct 20:83View ArticleGoogle Scholar
- Wang E, Nelson T, Rauch R (2004) Back to elements-tetrahedral versus hexahedral elements. Proceedings of the 2004 International ANSYS Conference, Pittsburgh, PA, USAGoogle Scholar
- Hughes TJR, Cottrell JA, Bazilevs Y (2005) Comput Method Appl Mech Eng 194:4135View ArticleGoogle Scholar
- Daniels AU, Chang MKO, Andriano KP (1990) J Appl Biomater 1:57View ArticleGoogle Scholar
- Chen B, Evans JRG (2005) Macromolecules 39:747View ArticleGoogle Scholar
- Rudnik E (2008) Compostable Polymer Materials. Elsevier Science, OxfordGoogle Scholar
- Ayodeji O, Graham E, Kniss D, Lannutti J, Tomasko D (2007) J Supercrit Fluid 41:173View ArticleGoogle Scholar
- Yang F, Wolke JGC, Jansen JA (2008) Chem Eng J 137:154View ArticleGoogle Scholar
- Reichert JC, Quent V, Burke LJ, Stansfield SH, Clements JA, Hutmacher DW (2010) Biomaterials 31:7928View ArticleGoogle Scholar
- Ke Y, Reddel RR, Gerwin BI, Reddel HK, Somers AN, McMenamin MG, LaVeck MA, Stahel RA, Lechner JF, Harris CC (1989) Am J Pathol 134:979Google Scholar
- Loessner D, Stok KS, Lutolf MP, Hutmacher DW, Clements JA, Rizzi SC (2010) Biomaterials 31:8494View ArticleGoogle Scholar
- Mertiny P, Ellyin F, Hothan A (2004) Compos Sci Technol 64:1View ArticleGoogle Scholar
- Hutmacher DW, Dalton PD (2011) Chem-Asian J 6:44View ArticleGoogle Scholar
- Tan EPS, Lim CT (2006) Compos Sci Technol 66:1102View ArticleGoogle Scholar
- Chung SW, Ingle NP, Montero GA, Kim SH, King MW (2010) Acta Biomater 6:1958View ArticleGoogle Scholar
- Reneker DH, Yarin AL, Fong H, Koombhongse S (2000) J Appl Phys 87:4531View ArticleGoogle Scholar
- Engelmayr GC Jr, Papworth GD, Watkins SC, Mayer JE Jr, Sacks MS (2006) J Biomech 39:1819View ArticleGoogle Scholar
Open AccessThis article is distributed under the terms of the Creative Commons Attribution License which permits any use, distribution, and reproduction in any medium, provided the original author(s) and the source are credited.