Fabrication and Optimization of Methylphenoxy Substituted

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Fabrication and Optimization of Methylphenoxy Substituted Polyphosphazene Nanofibers for Biomedical Applications Lakshmi S. Nair,† Subhabrata Bhattacharyya,| Jared D. Bender,+ Yaser E. Greish,@ Paul W. Brown,@ Harry R. Allcock,+ and Cato T. Laurencin*,†,‡,§ Department of Orthopaedic Surgery, University of Virginia, Charlottesville, Virginia 22903, Department of Biomedical Engineering, University of Virginia, Charlottesville, Virginia 22908, Department of Chemical Engineering, University of Virginia, Charlottesville, Virginia 22904, Department of Chemistry, University of Virginia, Charlottesville, Virginia 22903, Department of Chemistry, The Pennsylvania State University, University Park, Pennsylvania 16802, and Intercollege Materials Research Laboratory, The Pennsylvania State University, University Park, Pennsylvania 16802 Received April 21, 2004; Revised Manuscript Received July 21, 2004

Electrospinning has developed as a unique and versatile process to fabricate ultrathin fibers in the form of nonwoven meshes or as oriented arrays from a variety of polymers. The very small dimension of these fibers can generate a high surface area, which makes them potential candidates for various biomedical and industrial applications. The objective of the present study was to develop nanofibers from polyphosphazenes, a class of inorganic-organic polymers known for high biocompatibility, high-temperature stability, and low-temperature flexibility. Specifically, we evaluated the feasibility of developing bead-free nonwoven nanofiber mesh from poly[bis(p-methylphenoxy)phosphazene] (PNmPh) by electrospinning. The effect of process parameters such as nature of solvent, concentration of the polymer solution, effect of needle diameter, and applied potential on the diameter and morphology (beaded or bead-free) of resulting nanofibers were investigated. It was found that solution of PNmPh in chloroform at a concentration range of 7% (wt/v) to 9% (wt/v) can be readily electrospun to form bead-free fibers at room temperature. The mean diameter of the fibers obtained under optimized spinning condition was found to be approximately 1.2 µm. The beadfree, cylindrical nanofibers formed under the optimized condition showed a slightly irregular surface topography with indentations of a few nanometer scale. Further, the electrospun nanofiber mats supported the adhesion of bovine coronary artery endothelial cells (BCAEC) as well as promoted the adhesion and proliferation of osteoblast like MC3T3-E1 cells. Introduction Nanosized materials or structures are receiving great interest due to the unusual properties these materials possess as compared to conventional macromaterials. Several nanosized materials such as nanotubes, nanowires, nanocrystals, nanorods, nanospheres, and nanofibers are currently under investigation for various high technology applications.1 Among these, polymeric nanofibers are fibers having diameters ranging from the submicron to nanometer scale.2 Polymeric nanofibers exhibit unusual properties due to their very small diameter, which leads to high surface area, high aspect ratio, and better mechanical properties. The nonwoven mats formed from these fibers have additional advantages of controllable pore size, very high porosity, and permeability. These remarkable properties underlie the substantial * To whom correspondence should be addressed. Phone: (434) 2430250. Fax: (434) 243-0252. E-mail: [email protected]. † Department of Orthopaedic Surgery, University of Virginia. ‡ Department of Biomedical Engineering, University of Virginia. § Department of Chemical Engineering, University of Virginia. | Department of Chemistry, University of Virginia. + Department of Chemistry, The Pennsylvania State University. @ Intercollege Materials Research Laboratory, The Pennsylvania State University.

interests in these materials for industrial, biomedical, and electronic applications. Extensive research has been undertaken to develop nanofibers from a wide range of polymer solutions and melts.3,4 Nondegradable polymeric nanofiber matrixes are currently being used for developing high performance filters and catalyst systems.3,4 Composites reinforced with nanofibers have unique physical and mechanical properties and have been identified as potential candidates for high technology applications.3 Nondegradable nanofiber fabrics due to their lightweight, high permeability, and ability to absorb toxic materials are potential candidates for novel military fabrics.4 They can also serve as templating materials for the development of nanotubes and nanorods, nanoelectronic devices, and nanosensors.3,4 Furthermore, electrospun scaffolds from biostable and biocompatible polymers such as polyesters, polyurethanes, silicones, and poly(ethylene-co-vinyl acetate) have been fabricated as drug delivery devices, wound dressings, and prosthetic devices such as vascular grafts.5-7 Biodegradable polymeric nanofiber matrices that combine the unusual nanoscale properties with the biodegradability of the matrix polymer have been identified as potential candidates for medical applications, which require the transient existence of the materials in the

10.1021/bm049759j CCC: $27.50 © 2004 American Chemical Society Published on Web 10/22/2004

Nanofibers for Biomedical Applications

body such as scaffolds for tissue engineering, controlled drug delivery devices, adhesion prevention barriers, and wound dressings.3,4 Several fabrication techniques are currently being investigated to develop nanofiber matrices from polymers. These include template synthesis, phase separation, drawing, self-assembly, and electrospinning.3 Among these, electrospinning is an efficient method to develop polymeric fibers with diameters in the nanometer range, and this method has attracted a great deal of attention recently. In an electrospinning process, an electric potential is applied to a pendent droplet of polymer solution or melt from a syringe or capillary tube. At this point, a couple of mutually opposite forces come into play. Surface tension and viscoelastic forces of the polymer solution tend to retain the hemispherical shape of the droplet, whereas the charge induced by the electric field tends to deform the droplet from a hemispherical shape to a conical shape named a Taylor cone.8,9 When the voltage is increased beyond a threshold value, the electric forces in the droplet overcome the opposing surface tension forces, and a narrow charged jet is ejected from the tip of the Taylor cone. This emerging jet may break up into droplets, a process termed electrospraying, which is typical of low viscosity solutions due to surface tension effects. In the case of polymer solutions or melts with higher viscosities, the emerging polymer jet does not break up into droplets. Instead, the polymer jet starts traveling through the air in a nearly straight line due to the stabilization imparted by the longitudinal stress of the external electrical field on the charge carried by the jet. However, the polymer jet rapidly reaches a point of instability due to the repulsive forces arising from the opposite charges present in the polymer jet. The jet then passes through a series of electrically driven bending instabilities when the unstable jet bends back and forth followed by a bending, winding, spiraling, and looping path in three dimensions as demonstrated by Reneker et al. using high-speed videography.10 In the loops of the bending instability, the polymer jet get continuously stretched. This bending and stretching of the fiber is accompanied by a significant reduction in fiber diameter and rapid evaporation of the solvent resulting in the formation of ultrathin fibers. These fibers then are randomly deposited on the grounded collector resulting in the formation of nonwoven nanofiber mats. However, some recent studies indicate that the resulting fibers can also be collected in the form of highly oriented arrays, by employing various external mechanical or electrostatic forces.3,11,12 Recent studies show that highly oriented nanofibers can elicit favorable biological responses due to its ability to better mimic the biological microarchitecture. Thus, Xu et al. demonstrated the significant increase in adhesion, proliferation, and phenotype expression of smooth muscle cells on an aligned nanofiber matrix as compared to polymer films showing the favorable biological response to an aligned matrix.13 The uniqueness of the electrospinning process is that it provides ample flexibility to control the polymer fiber diameter and morphology by varying several governing parameters.14-17 These include solution (nature of the solvent molecular weight of the polymer and concentration of the polymer solution), process (electric potential, working dis-

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tance, and hydrostatic pressure), and environmental (temperature, humidity, and air velocity) parameters. The various properties of electrospun nanofibers such as mechanical strength, fiber diameter, porosity, and biocompatibility depend significantly on the properties of the specific polymer. Therefore, studies are currently ongoing to explore the feasibility of developing nanofibers from a wide range of polymers for variety of applications. Polyphosphazenes form a novel class of polymer with an inorganic backbone comprised of alternating phosphorus and nitrogen atoms with each phosphorus atom bearing two organic or organometallic side groups.18 The versatility of polyphosphazenes lies in the fact that the physical, chemical, and biological properties of a specific polymer depend on the nature and ratio of the side groups on phosphorus atoms. This, combined with the synthetic tailorability of the polyphosphazenes, wherein different polymers can be synthesized from a single intermediate, poly(dichlorophosphazene) via the macromolecular substitution route, makes polyphosphazenes a promising class of future materials. More than 700 different polyphosphazenes have been developed so far, and depending on the nature of the side groups, these polymers exhibit a wide range of thermal, electrical, optical, and biological properties.18 Biodegradable polyphosphazenes are prepared by replacing the chlorine atoms of polydichlorophosphazene by groups such as imidazolyl, amino acid esters, glyceryl, glycosyl, and lactic or glycolic acid esters, which sensitize the polymer backbone to hydrolytic degradation.19 Biodegradable polyphosphazenes have been identified as potential candidates for various biomedical applications due to their nontoxic and neutral degradation products. Furthermore, the synthetic flexibility of the polyphosphazene system allows the development of polymers with well-controlled degradation rates. This is achieved by incorporating hydrolytically less sensitive aromatic groups together with the groups that promote polyphosphazene degradation. Laurencin et al. have evaluated co-substituted polyphosphazenes with hydrophobic p-methyl phenoxy groups in conjunction with the hydrolytically labile amino acid esters or imidazolyl groups, as polymeric systems with wide ranges of degradation.20 These polymers were found to exhibit high osteocompatibility and have been suggested as potential candidates for bone tissue engineering. Our current research involves the development of novel biodegradable amino acid ester polyphosphazenes that bear p-methyl phenoxy side groups to modulate the degradation of the polymers for bone tissue engineering application. The corresponding homopolymer poly[bis(p-methylphenoxy)phosphazene] (PNmPh) (Structure 1) is hydrolytically stable. We are also exploring various scaffold fabrication techniques to develop three-dimensional (3-D) polymer matrices with an interconnected porous structure. Nanofiber based matrices are particularly interesting as tissue engineering scaffolds and drug delivery applications due to their very high surface area and narrow fiber diameter, which closely resembles the extracellular matrix present in native tissue.21 In addition to various industrial and high technology applications, nanofiber matrices of hydrolytically stable polymers can find many applications in biomedical fields such as wound dressing, controlled drug delivery and prosthetic device development.

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Kataphinan et al. reported an attempt to develop nanofiber matrices of poly[(diphenoxy)phosphazene] (PDPP) by electrospinning.22 However, electrospinning of PDPP from THF solution resulted in the formation of non-uniform fibers with very high fiber diameters. The objective of the present study was to evaluate the feasibility of developing ultrafine fibers from PNmPh by electrospinning and to optimize the governing parameters that affect electrospinning to develop bead-free uniform cylindrical fibers as candidates for various biomedical applications. Figure 1. Shear viscosity vs shear rate of 8% (wt/v) PNmPh solution in chloroform.

Experimental Procedures Materials. Hexachlorocyclotriphosphazene (Nippon Fine Chemical Co. Tokyo, Japan) was purified by recrystallization from hexane followed by vacuum sublimation (55 °C and at 0.05 mm Hg). All the solvents were predried before use. Tetrahydrofuran (THF, EM Science, Gibbstown, NJ) was distilled from sodium/benzophenone immediately before use in the reactions. 4-Methyl phenol (Aldrich) was sublimed and stored in a desiccator. N,N-Dimethylformamide (DMF) (Sigma, St. Louis, MO) was vacuum-distilled and stored over molecular sieves (4°A). Metallic sodium (Sigma) was used as received. Synthesis of Poly[bis(methylphenoxy)phosphazene] (PNmPh). The synthesis of poly[bis(methylphenoxy)phosphazene] was carried out using a two-step process as reported earlier.23 Polydichlorophosphazene was first prepared by the thermal ring opening polymerization of hexachlorocyclotriphosphazene at 250 °C according to a reported procedure.17 In a second step, the complete replacement of the chlorine atoms in polydichlorophosphazenes with p-methylphenoxy groups was performed as follows. Briefly, 10.0 g (8.62 × 10-2 mol) of poly(dichlorophosphazene) was dissolved in warm THF (500 mL). In a separate flask, p-methyl phenol (55.2 g, 0.587 mol) was dissolved in THF and was added to an ice bath cooled suspension of sodium (11.1 g, 0.483 mol) in THF (200 mL). The salt solution was warmed for 24 h until all the sodium had reacted. At that time, the polymer and salt solutions were added separately to an autoclave reactor. The reaction mixture was stirred and heated in the autoclave at 150 °C for 24 h at 3.5 bar. The mixture was then concentrated and precipitated into deionized water. The polymer (Structure 1) was reprecipitated from THF into deionized water (3×), hexane (3×), and ethanol (1×) to obtain a tough white polymer. The molecular weight of the polymer was determined by GPC and was found to be 642,000 (Mn) and 1,909,000 (Mw) with a polydispersity index of 3.0. The characterization data include 1H NMR δ 7.0-7.5 (m, 4H), 2.3 (s, 3H); 13C NMR δ 149.6, 132.1, 129.1, 121.0, and 20.6 ppm; 31P NMR δ 19.6 ppm (s). The Tg of the polymer was found to be 2.1 °C.

Table 1. Viscosity of Polymer Solutions Containing 7, 8, and 9 % (wt/v) of PNmPh in Chloroform concentration (%)

shear stress (Pa)

viscosity (Pa S)

7 8 9

1.0 1.0 1.0

1.136 2.423 2.753

Characterization. NMR spectra were obtained at 298 K using a Bruker AMX-360 NMR spectrometer resonating at 360.23 MHz for 1H, 145.81 MHz for 31P, and 90.56 MHz for 13C. All 1H and 13C NMR samples were prepared with deuterated THF (Isotec, 99.5%), unless otherwise noted, and referenced to tetramethylsilane (TMS). 31P NMR shifts are relative to 85% phosphoric acid as an external reference, with positive shift values downfield from the reference. The molecular weight of the polymer was estimated using a Hewlett-Packard HP 1090 gel permeation chromatograph equipped with an HP 1047A refractive index detector and calibrated against polystyrene standards (Polysciences). The samples were eluted at a flow rate of 1 mL/min at 40 °C with solution of 0.1% tetrabutylammonium bromide (Aldrich) in THF (OmniSolv). A 1% (wt/v) solution of the polymer in THF was used for molecular weight determination. The viscosities of the polymer solutions in chloroform were determined by AR 2000 Advanced Rheometer (TA instruments, New Castle, DE) at 25 °C. The geometry used was a cone and plate arrangement. As the cone and the plate subjects the polymer solution to controlled shear stress, the resulting strain or shear rate is measured. The shear stress was swept from 0 to 10 Pa in a programmed manner. A computer interfaced to the machine recorded the resulting shear stress/viscosity versus shear rate data. Figure 1 shows the shear viscosity versus shear rate of 8% solution of PNmPh in chloroform under a shear sweep of 0-10 Pa. Single point viscosity measurement (peak hold -1 Pa) was used to determine the viscosities of different concentrations of polymer solutions. Table 1 shows the viscosities of 7, 8, and 9% (wt/v) of PNmPh solutions. Glass transition temperatures were determined by differential scanning calorimetry (DSC) using a Perkin-Elmer-7 thermal analysis system equipped with a computer. Calibration of the Perkin-Elmer-7 thermal analysis system was achieved through use of an indium standard. Polymer samples were heated from -100 to 100 °C under an atmosphere of dry nitrogen. Approximately 30 mg of sample were hermetically sealed in aluminum pans and heated at rates of 10, 20, and 40 °C/

Nanofibers for Biomedical Applications Scheme 1. Schematic Diagram of the Electrospinning Apparatus

min. The final Tg values were determined through extrapolation to 0 °C/min heating rate. Electrospinning. Scheme 1 shows the schematic diagram of the electrospinning apparatus used in the present study. The apparatus consists of a 20 mL glass syringe fitted with a blunt end needle and a ground electrode. The grounded electrode consists of a copper plate covered with aluminum foil/interface fabric/Teflon coated substrate placed at a predetermined distance (working distance) from the needle tip. The syringe was fixed parallel to the collection screen, and the polymer solution was allowed to flow under gravity and electrical pressure. A Gamma High Voltage Supply ES40P-20W (0-40 kV, 20 W, Gamma High Voltage Research) with a low current output was used as the power source. A positive voltage was applied to the polymer solution in the glass syringe by attaching an alligator clip to the needle from the positive lead. The electrospinning was carried out at ambient temperature and pressure. The spun nanofiber mats were dried under vacuum at room temperature for 24 h. In the present study, electrospinning was optimized to obtain bead-free ultrafine fibers from PNmPh by varying parameters such as nature of the solvent, needle diameter, concentration of the solution, and applied electric field. The applied electric field was varied from 27 to 36 kV, the concentration of the solution was varied from 1 to 9% (wt/ v) in chloroform, and the needle diameter was varied using 25- and 18-gauge needles. The effect of the nature of solvents on electrospinning was investigated using solvents such as chloroform, THF, and a co-solvent system consisting of DMF and THF in a 1:1 ratio. The diameter, morphology (beaded/non-beaded), shape (circular/irregular cross-section), and surface topography of the formed fibers (rough/smooth) of the fibers were examined using scanning electron microsocpy. The scanning electron micrographs (SEM) of the fibers were obtained after coating the fibers with gold for 4 min at 12-15 mA at 5 V under 100 mTorr pressure using a Hummer V sputtering system (Technics Inc., Baltimore, MD). The samples were viewed under JSM-6400 scanning electron microscope (JEOL, Boston, MA) operated at an accelerating voltage of 20 kV at various magnifications. Average fiber diameters were determined by measuring 100 fibers selected randomly from each electrospun mats. Cell Culture. The electrospun nanofibers were sterilized by ethanol followed by irradiation with UV light for 15 min on each side. The mouse immortalized calvarial cells


(MC3T3-E1) used in the present study were a gift from Dr. H. Sudo, Tohoky Dental University, Tomitamachi, Koriyma, Japan. Bovine Coronary Artery Endothelial cells (BCAEC) were procured from Cell Applications, Inc., San Diego, CA. Cell Seeding. MC3T3-E1 cells were plated in a monolayer in tissue culture flasks (75 cm2) and cultured to confluence in alpha-modification of minimum essential media supplemented with 10% fetal bovine serum and 1% penicillin and streptomycin. The media was replaced every other day, and culture was maintained in a tissue culture incubator at 37 °C and 5% carbon dioxide. Once the cells reached 80% confluency, they were trypsinized and seeded onto the surface of nanofiber matrixes fixed on the bottom of well plates at a seeding density of 50,000 cells/well. Cellular constructs were harvested at 4 and 7 days, washed with PBS, fixed with 4% glutaraldehyde for 24 h at 4 °C, dehydrated through a series of graded alcohol treatments, and then air-dried overnight. The cellular constructs were sputter coated and observed by SEM at an accelerated voltage of 20 kV. BCAEC were plated in monolayer in a tissue culture flask (75 cm2) and cultured to confluence in Bovine endothelial growth medium (Cell Applications, Inc.). At 80% confluency, the cells were trypsinized and seeded on nanofiber matrixes and cultured for 24 h. After 24 h, in culture, the cellular constructs were harvested, washed with PBS, fixed in glutaraldehyde, and observed by SEM as in the case of MC3T3-E1 cell constructs. Results Effect of Electrospinning Parameters on PNmPh Nanofibers. Various electrospinning parameters can affect the formation of nanofibers from a specific polymer.3 Therefore, the effect of some of these parameters has been investigated to develop bead-free uniform fibers from PNmPh by electrospinning Nature of the Solvent. Studies have shown that the nature of the solvent can significantly affect the electrospinning process and thereby the diameter and morphology of the resulting nanofibers.24 Hence, the effect of different solvents on electrospinning of PNmPh was investigated. Three different solvent systems were used to evaluate the effect of solvent system on PNmPh, fiber formation, THF, chloroform, and a 1:1 THF/DMF mixture. The use of THF/DMF cosolvent did not facilitate PNmPh electrospinning. Figure 2a,b shows the SEMs of electrospun nonwoven mats formed from 8% (w/v) polymer solution in THF and chloroform, respectively, under an applied potential of 33 kV, working distance of 30 cm, and using an 18-gauge needle. Effect of Needle Diameter. Different gauge needles were used to evaluate the effect of needle diameter on the diameter and morphology of electrospun fibers. The fibers were electrospun from 1 and 3% (wt/v) solution of PNmPh in chloroform at a constant voltage of 33 kV and working distance of 30 cm using 25-gauge (diameter 0.5 mm) and 18-gauge (diameter 1.2 mm) needles under gravity. In the case of the 1% (wt/v) solution, the rate of fluid flow was found to be very high with the 18-gauge needle and thereby resulted in no fiber formation. Figure 3a shows the SEM of

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Figure 2. (a) SEM of electrospun PNmPh fibers from THF at a concentration of 8% (wt/v) of the polymer at 33 kV using 18-gauge needle showing the formation of highly nonuniform distorted fibers. (b) SEM of electrospun PNmPh fibers from chloroform at a concentration of 8% (wt/v) of the polymer at 33 kV using 18 gauge showing the formation of distinct uniform fibers.

fibers formed from 1% (wt/v) PNmPh solution using a smaller diameter needle (25 gauge). Figure 3b shows the SEMs of fibers obtained from a 3% polymer solution using the 25-gauge needle. The gravity induced flow rate of the 3% (wt/v) solution through the 25-gauge needle was found to be low as compared to the 1% (wt/v) solution. The use of an 18-gauge needle significantly increased the spinning rate of the 3% (wt/v) PNmPh solution as compared to the 25-gauge needle and resulted in the formation of uniform fibers (Figure 3c). With polymer solutions having a concentration higher than 3% (wt/v), the 25-gauge needle was found to be inefficient for gravity induced electrospinning. Solution Concentration. Previous studies have shown that one of the major factors in determining nanofiber diameter and morphology formed by electrospinning is the solution concentration.14,15,33 In the present study, it was found that the concentration of PNmPh solution below 3% (wt/v) in chloroform resulted in the formation of composite fibers with large number of beads along the fibers. Similarly, above 9% (wt/v), the high viscosity of the solution adversely affected the electrospinning process. Therefore, the effect of polymer concentration in the range of 3-9% (wt/v) on fiber diameter was evaluated. Figure 4 shows the SEMs of PNmPh fibers

Figure 3. (a) SEM of electrospun PNmPh fibers from chloroform at a concentration of 1% (wt/v) of the polymer at 33 kV using 25-gauge needle showing the formation of composite morphology. (b) SEM of electrospun PNmPh fibers from chloroform at a concentration of 3% (wt/v) of the polymer at 33 kV using 25-gauge needle showing the formation of composite fibers with spindle shaped beads. (c) SEM of electrospun PNmPh fibers from chloroform at a concentration of 3% (wt/v) of the polymer at 33 kV using 18-gauge needle showing the formation of nonuniform fibers.

fabricated from different polymer concentrations at a constant electrospinning voltage of 33 kV using the 18-gauge needle at a working distance of 30 cm. Figure 4a shows the SEM of fibers obtained by electrospinning 5% (wt/v) PNmPh solution in chloroform, and Figure 4b shows the corresponding SEM from 6% (wt/v) polymer solution.



Figure 4. (a) SEM of electrospun PNmPh fibers from chloroform at a concentration of 5% (wt/v) of the polymer at 33kV using 18-gauge needle showing the formation of almost bead free fibers with larger diameter distribution. (b) SEM of electrospun PNmPh fibers from chloroform at a concentration of 6% (wt/v) of the polymer at 33kV using 18-gauge needle showing the formation of bead free fibers.

Figure 5. Frequency distributions of electrospun fibers from 7, 8, and 9% (wt/v) solutions at a applied potential of 33 kV, working distance of 30 cm, and using 18-gauge needle.

Figure 5 shows the frequency distribution of fiber diameters obtained from 7, 8, and 9% (wt/v) polymer solution when spun at 33 kV and at a working distance of 30 cm. The most uniform fiber diameter distribution was obtained from 8% (wt/v) polymer solution; hence, that concentration was used to optimize the applied potential. Effect of Applied Voltage. Figure 6 shows SEM micrographs of electrospun nanofibers formed at various applied potentials from a constant polymer concentration of 8% (wt/ v) and at a working distance of 30 cm using an 18-gauge needle. Electrospun fibers obtained from an 8% (wt/v) solution in chloroform spun at an applied potential of 27 kV resulted in the formation of bead-free fibers. The fibers, however, displayed a highly irregular shape or cross-section

Figure 6. (a) SEM of electrospun PNmPh fibers from chloroform at a concentration of 8% (wt/v) of the polymer at 27 kV using 18-gauge needle showing the formation of fibers with distorted structure. (b) SEM of electrospun PNmPh fibers from chloroform at a concentration of 8% (wt/v) of the polymer at 33 kV using 18-gauge needle showing the formation of more cylindrical fibers with junctions. (c) SEM of electrospun PNmPh fibers from chloroform at a concentration of 8% (wt/v) of the polymer at 36 kV using 18-gauge needle showing the formation of uniform cylindrical fibers.

(Figure 6a). Increasing the applied potential from 27 to 30 kV did not seem to improve the shape of the electrospun fibers. In comparison, with the fibers spun at 27 kV (Figure 6a) and 30 kV, more uniform and cylindrical fibers were obtained at an applied potential of 33 kV (Figure 6b). Further increase in applied potential to 36 kV (Figure 6c) resulted in the formation of almost cylindrical fibers with circular cross-section. Figure 7 shows the mean diameters of the fibers formed from 8% (wt/v) of PNmPh solution in chloroform electrospun at a working distance of 30 cm, using an 18-gauge needle and under various electric potentials (27-36 kV). The values were reported as means ( standard deviations, and the statistical analysis was performed using

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Figure 7. Relationship between average fiber diameter and applied voltage of 27-36 kV of fibers electrospun from 8% (wt/v) solution in chloroform at a working distance of 30 cm and using 18-gauge needle.

Figure 8. High SEM magnification of fibers obtained at optimized conditions [8% (wt/v), 36 kV, 18-gauge needle] showing the irregular surface topography with nanoindentations.

one way analysis of variance to determine the variance of the data (pe 0.05). Surface Topography of Nanofibers. Figure 8 shows the high magnification SEM of nanofibers obtained at the optimized condition of 8% w/v polymer solution in chloroform spun at an applied potential of 36 kV using an 18gauge needle. Cell Adhesion and Proliferation. Figure 9a shows the SEM of BCAE cells on PNmPh nanofiber matrices after 24 h post-cell seeding. Figure 9b,c shows SEMs of MC3T3-E1 cells on PNmPh nanofiber matrices after 4 and 7 days postcell seeding. Discussion Poly[bis(methyl phenoxy)phosphazene] forms an interesting member of polyphosphazene family due to the ease of functionalization of p-methyl phenoxy groups. This, coupled with the high chemical stability of a phosphorus-nitrogen backbone, makes PNmPh an ideal substrate for developing novel materials. PNmPh has been previously studied as an intermediate platform for the installation of reactive functional groups as well as a substrate for immobilizing biologically relevant molecules.23,25 In one study, protein-A was immobilized on a PNmPh polymer substrate. The immobilized protein-A binds specifically to immunoglobulins (IgG) and could be used to develop blood dialysis systems that selectively remove IgG from the blood.26 Similarly, heparinized poly(organophophazenes) have been developed

Figure 9. (a) SEM showing the adhesion of BCAEC on electrospun PNmPh fiber matrices after 24 h in culture. (b) SEM showing MC3T3E1 cells on electrospun PNmPh fiber matrices after 4 days in culture. (c) Matrices after 7 days in culture.

by complexing heparin to chemically modified PNmPh in an attempt to develop blood compatible polymers.27 Therefore, the development of PNmPh nanofiber matrixes with a highly porous architecture and high surface area can give a new dimension to the applicability of the polymer. In the present study, electrospinning of PNmPh has been optimized to develop bead-free uniform fibers. The effect of different solvent systems on PNmPh fiber formation was investigated. The use of the THF-DMF cosolvent system did not facilitate fiber formation presumably due to the low volatility of the solvent system, which caused incomplete drying of the fibers prior to deposition on the collector. Electrospinning of the THF solution of PNmPh resulted in the formation of highly non-uniform fibers and bundles of fibers with distorted shapes (Figure 2a). On the other hand, fibers formed from the chloroform solution of PNmPh showed uniform cylindrical morphology (Figure 2b). Thus, among the solvent systems investigated (THF, chloroform, and 1:1 mixture of THF/DMF), the PNmPh solution in chloroform resulted in the formation of the most uniform fibers. Previous studies show that the needle or orifice diameter has a profound effect on the diameter of polymeric nanofibers obtained by electrospinning.28,29 A decrease in needle


diameter usually results in a decrease in fiber diameter.28,29 Electrospinning of 1% (wt/v) of PNmPh in chloroform using a 25-gauge needle resulted in the formation of ultrafine fibers (Figure 3a). However, the formed fibers were found to have a composite morphology with a large number of beads along the fibers. The use of an 18-gauge needle did not facilitate the electrospinning of 1% (wt/v) polymer solution due to the very high rate of fluid flow. Electrospinning of 3% (wt/ v) solution using a 25-gauge needle resulted in the formation of composite fibers with large number of spindle shaped beads along the fibers (Figure 3b). However, the use of an 18-gauge needle for 3% (wt/v) polymer solution resulted in the formation of fibers with significantly decreased bead formation even though the fibers showed a highly irregular morphology with large variations in diameter along a single fiber (Figure 3c). Thus, in the case of PNmPh, the use of the lower gauge needle (25 gauge) resulted in the formation of lower diameter fibers as qualitatively evidenced from SEM (Figure 3b,c); however, the use of the lower gauge needle significantly increased the formation of beads at the polymer concentrations studied when they were electrospun under gravity. The concentration of the polymer solution is known to play a significant role in determining fiber diameter and morphology during electrospinning.13,14,33 In the present study, fibers were found to form from solutions having concentrations ranging from 1 to 9% (wt/v). Increase in the concentration of the polymer solution usually results in the formation of fibers having larger diameters.33 In the case of the PNmPh solution in chloroform, SEM showed an increase in fiber diameter of beaded fibers formed from a 1% (wt/v) solution (Figure 3a) to a 3% (wt/v) solution (Figure 3b) when the electron was spun at an applied field of 33 kV, working distance of 30 cm, and using a 25-gauge needle. The increase in solution concentration also significantly influenced the shape of the beads on the fibers formed from 1 and 3% (wt/ v) solution. The beads formed along the fibers became more spindle shaped as the concentration of the solution was increased to 3% (wt/v) (Figure 3b). Increase in the polymer concentration significantly affected the morphology of fibers obtained when electrospun at an applied potential of 33 kV, working distance of 30 cm, and using an 18-gauge needle. As evidenced from SEM (Figure 3c), fibers formed from the 3% (wt/v) polymer solution have highly irregular undulating shapes with circular and flat ribbonlike fibers. Fewer numbers of beads were found along the fibers; however, the fibers showed large variations in diameter. The fiber matrix formed from the 5% (wt/v) polymer solution also showed the presence of circular and ribbon shaped fibers with a large variation in fiber diameter, however, with very few beads as evidenced from SEM (Figure 4a). Electrospinning of the 6% (wt/v) PNmPh solution resulted in the formation of more circular fibers (Figure 4b). This is in accordance with other studies, which showed that increase in concentration and associated electrical forces favor the formation of circular fibers.14-15 Further increase in the solution concentration of PNmPh did not, however, show a dramatic increase in fiber diameters as observed in the case of some polymers.33 Among 7, 8, and 9% (wt/v) solutions


studied, the electrospinning of the 8% (wt/v) solution resulted in the formation of fibers with narrow diameter distribution (Figure 5). Also, it is evident from the figure that higher counts of larger diameter fibers are formed from the 9% solution as compared to the 8% solution. This can be attributed to the higher elasticity of the 9% solution as compared to 8% solution, which pulls more material from the Taylor cone, resulting in greater material flow. Previous studies show that the electrospinning voltage may also have a significant influence on the diameter and shape of the resulting nanofibers. Typically, a larger applied voltage increases the net charge experienced by the jet, thereby increasing the electrostatic stress on the jet, resulting in enhanced fiber drawing to narrower diameters. However, it has also been reported that after reaching a minimum value, the diameter of the fibers increases with an increase in voltage due to greater material flow.29 Thus, Demir et al. showed an increase in polymer jet diameter in a sigmoidal manner with increasing voltage resulting in an increase in diameter of the resulting fibers. However, an increase in applied voltage did not show an increase in fiber diameter for PNmPh in the range of voltage studied. Thus, in the case of PNmPh, the variation of applied potential from 27 to 30 kV did not show any statistically significant differences in fiber diameters (Figure 7). Further increase in electric potential to 33 kV showed a reduction in diameters of the resulting fibers. Increasing the electric potential to 36 kV did not show any statistically significant differences in the fiber diameters as compared to fibers obtained under 33 kV (Figure 7). However, it has been found that an increase in electric field strength considerably affected the shape of the resulting fibers. At lower field (27 and 30 kV), the fibers formed were found to have a curly or distorted shape with dimples or nanoindentations on the surface (Figure 6a). An increase in the electric field from 30 to 33 kV altered the shape of the fibers, which assumed a more cylindrical and rodlike structure (Figure 6b). Increasing the applied potential to 36 kV further improved the shape of the fibers, and distinct cylindrical fibers were formed (Figure 6c). The improvement in shape of the fibers can be attributed to the increased electrostatic force exerted on the jet at higher potential. Another interesting feature observed in the case of PNmPh is the surface topography of the fiber obtained. Polymeric nanofibers with rough or nanoporous morphology can give rise to very high surface areas, and hence, can become potential candidates for various applications. The occurrence of nanopores on the surface of polymeric nanofibers has been recently observed from various polymers, and different mechanisms have been suggested.30-32 Several parameters such as nature of the solvent, glass transition temperature of the polymer, solvent-polymer interactions, and environmental parameters such as humidity and temperature have been shown to affect the nanostructured surface morphologies of electrospun fibers. The high magnification SEM micrograph (Figure 8) shows the formation of uniform cylindrical fibers of PNmPh under optimized condition, however, with a rough surface topography with dimples or nanoscale indentations. The reason for the occurrence of the surface irregularity has not been elucidated at present, even though

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similar surface irregularities have been observed in the case of low glass transition temperature polymers such as poly(ethylene oxide). The electrospun nanofiber matrixes of PNmPh could be used for developing novel drug delivery and prosthetic devices with unique properties due to the inherent differences in the properties of polyphosphazenes as compared to organic macromolecules. As reported earlier, the electrospun fibers could provide a three-dimensional highly porous structure for cell attachment, migration, and proliferation; therefore, PNmPh fibers could be used as an implant coating to enhance tissue integration. A preliminary biocompatibility evaluation of PNmPh nanofiber matrixes was performed by following the adhesion and proliferation of cells on the nanofiber matrixes. Bovine coronary artery endothelial cells, which are sensitive to the changes in growth medium and substrates, were used for cytotoxicity evaluation. Figure 9a shows appreciable adhesion on the nanofiber matrix after 24 h of seeding, indicating the cytocompatibility of the nanofiber matrixes. MC3T3-E1 cells are osteoblast like cells derived from neonatal mouse calvaria and are widely used as a model to study osteoblast physiology in vitro. The non-woven fiber mat of PNmPh formed via electrospinning was found to support the adhesion and proliferation of MC3T3-E1 cells. SEMs of cell seeded nanofiber mats after 4 days in culture showed that cells were found to adhere and migrate through the porous structure of nanofiber matrix (Figure 9a). By 7 days in culture, the cells proliferated completely, covering the matrix (Figure 9b). Overall, the present study indicates that the polyphosphazene (PNmPh) nanofiber matrixes could promote cell matrix and cell-cell interactions, making them potential candidate for various biomedical applications such as wound dressings and prosthetic organs. In the case of nanofiber matrixes, in addition to diameter and morphology of the polymeric fibers, the porosity, pore diameter, orientation of the fibers, surface area of the matrix, and mechanical integrity of the nanofiber matrixes play a crucial role in eliciting favorable cell-matrix interaction as well as to design novel controlled delivery matrixes. Further studies are underway to optimize the microarchitecture of the developed polyphosphazene fibers to fabricate candidate materials for various biomedical applications. Conclusions This study demonstrated the feasibility of developing polyphosphazene fibers in the nano- and micrometer range through electrostatic spinning. The electrospinning parameters such as solution concentration, nature of solvent, needle diameter, and potential gradient have been optimized to develop ultrafine bead-free uniform fibers from PNmPh. The fiber diameters obtained at optimized conditions were found to be ∼1.2 µm. The cylindrical fibers formed under optimized conditions showed a slightly irregular surface topography with nanoindentations. The nonwoven nanofiber mats formed from PNmPh supported the adhesion of endothelial cells after 24 h in culture. Also, the nanofiber

Nair et al.

matrix supported the adhesion and proliferation of osteoblastlike MC3T3-E1 cells. Acknowledgment. Authors greatly acknowledge NIH #46560 for financial support. Dr. Laurencin was the recipient of a Presidential Faculty Fellow Award. References and Notes (1) Nalwa, H. S. Handbook of nanostructured materials and nanotechnology; Academic Press: San Diego, 2000. (2) Khil, M. S.; Kim, H. Y.; Kim, M. S.; Park, S. Y.; Lee, D. Polymer 2004, 45, 295-301. (3) Huang, Z.; Zhang, Y. Z.; Kotaki, M. Ramakrishna, S. Comp. Sci. Technol. 2003, 63, 2223-2253. (4) Frenot, A.; Chronakis, I. S. Curr. Opin. Colloid Interface Sci. 2003, 8, 64-75. (5) Martin, G. E.; Cockshott, I. D.; Fildes, F. J. T. U.S. patent 4878908, 1989. (6) Stenoien, M. D.; Drasler, W. J.; Scott, R. J.; Jenson, M. L. U.S. patent 5866217, 1999. (7) Kenawy, R.; Layman, J. M.; Watkins, J. R.; Bowlin, G. L.; Matthews, J. A.; Simpson, D. G.; Wnek, E G. Biomaterials 2003, 24, 907-13 (8) Reneker, J. D. J. Electrostatics 1995, 35, 151-160. (9) Taylor, G. I. Proc. Royal Soc. London 1969, A313, 453 (10) Reneker, D. H.; Yarin, A. L.; Fong, H.; Koombhongse, S. J. Appl. Phys. 2000, 87, 4531-4547. (11) Theron, A.; Zussman, E.; Yarin, A. L. Nanotechnology 2001, 12, 384-390 (12) Zussman, E.; Theron, A.; Yarin, A. L. Appl. Phys. Lett. 2003, 82, 973-975 (13) Xu, C. Y.; Inai, R.; Kotaki, M.; Ramakrishna, S. Biomaterials 2004, 25, 877-886. (14) Deitzel, J. M.; Kleinmeyer, J.; Harris, D.; BeckTan, N. C. Polymer 2001, 42, 261-272. (15) Fong, H.; Chun, I.; Reneker, D. H. Polymer 1999, 40, 4585-4592. (16) Theron, S. A, Zussman, E.; Yarin, A. L. Polymer 2004, 45, 20172030. (17) Zong, X.; Kim, K.; Fang, D.; Ran, S.; Hsiao, S. B.; Chu, B. Polymer 2002, 43, 4403-4412. (18) Allcock, H. R. Chemistry and Applications of Polyphosphazenes; Wiley-Interscience: New York, 2002. (19) Lakshmi, S.; Katti, D. S.; Laurencin, C. T. AdV. Drug DeliVery ReV. 2002, 55, 467-82. (20) Laurencin, C. T.; Norman, M. E.; Elgendy, H. M.; El-Amin, S. F.; Allcock, H. R.; Pucher, S. R.; Ambrosio, A. A. J. Biomed. Mater. Res. 1993, 27, 963-73. (21) Li, W.; Laurencin, C. T.; Caterson, E. J.; Tuan, R. S.; Ko, F. K. J. Biomed. Mater. Res. 2002, 60, 613-621. (22) Kataphinan, W.; Teye-Mensah, R.; Evans, E. A.; Ramsier, R. D.; Reneker, D. H.; Smith, D. J. J. Vac. Sci. Technol. 2003, A21, Jul/ Aug. (23) Allcock, H. R.; Fitzpatrick, R. J. Chem. Mater. 1992, 4, 769-775. (24) Lee, K. H.; Kim, H. Y.; Khil, M. S.; Ra, Y. M.; Lee, D. R. Polymer 2003, 44, 1287-1294. (25) Wisian-Neilson, P.; Bailey, L.; Bahadur, M. Macromolecules 1994, 27, 7713-7717. (26) Allcock, H. R.; Nelson, C. J.; Coggio, W. D. Chem. Mater. 1994, 6, 516. (27) Neenan, T. X.; Allcock, H. R. Biomaterials 1982, 3 78-80 (28) Mo, X. M.; Xu, C. Y.; Kotaki, M.; Ramakrishna, S. Biomaterials 2004, 25, 1883-1890. (29) Baumgarten, P. K. J. Colloid Interface Sci. 1971, 36, 71-79. (30) Bognitzki, M.; Czado, W.; Frese, T; Schaper, A.; Hellweg, M.; Steinhart, M.; Greiner, A; Wendorff, J. H. AdV. Mater. 2001, 13, 70. (31) Megelski, S.; Stephens, J. S.; Chase, D. B.; Rabolt, J. F. Macromolecules 2002, 35, 8456-8466. (32) Casper, C. L.; Stephens, J. S.; Tassi, N. G.; Chase, D. B.; Rabolt, J. F. Macromolecules 2004, 37, 573-578. (33) Demir, M. M.; Yilgor, I.; Yilgor, E.; Erman, B. Polymer 2002, 43, 3303-3309.

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