Methods of making three dimensional structures having aligned nanofibers and the resulting structures produced by such methods

Abstract

Apparatus for producing a three dimensional nanofiber structure includes (1) at least two spaced electrodes; (2) a spinner adapted to rotate the at least two spaced electrodes; (3) a syringe assembly adapted to eject a polymer solution from a syringe of the syringe assembly towards the at least two spaced electrodes while the at least two spaced electrodes are rotated by the spinner; and (4) a power supply assembly for providing the two spaced electrodes at a first electric potential, and for providing the syringe at a second electric potential which is different from the first electric potential. A composition of matter may include (1) a least one layer of nanofibers in which a distribution of angles of fibers is aligned; and (2) at least one gel layer, wherein the at least one layer of microfibers and the at least one gel layer alternate to form a laminate.

Claims

1. A method for producing a three dimensional nanofiber structure, the method comprising: a) rotating, around an axis of rotation, at least two spaced electrodes provided at a first electric potential, wherein a spacing of the at least two spaced electrodes with respect to each other is orthogonal to the axis of rotation; and b) ejecting a polymer solution from a syringe, provided at a second electric potential which is different from the first electric potential, towards the at least two spaced electrodes while the at least two spaced electrodes are rotating.

2. The method of claim 1 wherein the syringe is spaced between 3-10 cm from the at least two spaced electrodes.

3. The method of claim 1 wherein the at least two spaced electrodes rotate at a speed of 1000-1500 rpm.

4. The method of claim 1 wherein a difference between the first electric potential and the second electric potential is at least 10,000 V DC.

5. The method of claim 1 wherein the polymer solution is ejected from the syringe at a rate of at least 1 mL per hour.

6. The method of claim 1 wherein the syringe has a 20 ga needle.

7. The method of claim 1 further comprising: moving the syringe relative to the at least two spaced electrodes while the at least two spaced electrodes are rotated and the polymer solution is ejected.

8. The method of claim 1 wherein at least one of the first and second electric potentials is ground.

9. A method comprising: a) rotating at least two spaced electrodes provided at a first electric potential; and b) ejecting a polymer solution from a syringe, provided at a second electric potential which is different from the first electric potential, towards the at least two spaced electrodes while the at least two spaced electrodes are rotating; and c) providing a substrate within a space defined by the rotation of the at least two spaced electrodes, wherein the polymer solution ejected from the syringe is also directed towards the substrate, thereby providing nanofibers on the substrate.

10. The method of claim 9 further comprising: providing, after the substrate has been provided with nanofibers, a gel layer onto the nanofibers provided on the substrate.

11. The method of claim 10 wherein the gel layer is provided onto the nanofibers provided onto the substrate by (1) dipping the nanofiber-provided substrate into an alginate solution, and (2) then dipping the resulting nanofiber-provided substrate into CaCl2) solution to crosslink the alginate.

12. The method of claim 1 wherein the at least two spaced electrodes rotate at a speed of 1050-1350 rpm.

13. The method of claim 1 wherein a difference between the first electric potential and the second electric potential is at least 15,000 V DC.

14. The method of claim 1 wherein the polymer solution is ejected from the syringe at a rate of at least 1.5 mL per hour.

15. The method of claim 1 wherein the two spaced electrodes are spaced from the axis of rotation.

16. The method of claim 1 wherein a void is defined between the two spaced electrodes.

17. The method of claim 1 wherein the at least two spaced electrodes are spaced from one another at a distance of D, and wherein each of the at least two spaced electrodes are spaced from the axis of rotation at a distance of D/2.

18. The method of claim 1 wherein each of the at least two spaced electrodes is spaced from the axis of rotation by an equal distance.

19. The method of claim 1 wherein each of the at least two spaced electrodes is shaped as a prong having one open end and one attached end.

Description

3. BRIEF DESCRIPTION OF THE DRAWINGS

(1) FIG. 1 is a diagram illustrating a sample horizontal electrospinning apparatus setup.

(2) FIG. 2 is a microphotograph illustrating electrospinning with bending instability at the end of the mesh, 0.25 ms exposure. (From: Reneker et al., 2000; Journal of Applied Physics: 87 (9).)

(3) FIG. 3 is a diagram illustrating an electrospinning setup used for rotating mandrel collector. (From: Matthews et al., 2002; Biomacromolecules (3).)

(4) FIG. 4 illustrates the elongation of an electrospun nanofiber during deposition on two anode blades. (From: Teo et al., 2005; Nanotechnology: 16(9).)

(5) FIG. 5 illustrates the impact of ECM attachment points on cell culture growth.

(6) FIG. 6A illustrates Lymphocyte alignment in parallel to matrix, and FIG. 6B illustrates change of angle of migration according to the matrix. (From: Woolf et al, 2003; Blood: 102 (9).)

(7) FIG. 7 are microphotographs illustrating the effect of scaffold nanotopography on GTS complex polarization (in red with white arrowheads pointing) in hESCs. It should be noted that flat substrate resulted in radial distribution of -tubulin, longitudinally. Aligned nanotopography resulted in more polarized GTS. (From: Gerecht et al, 2007; Biomaterials: 28 (28).)

(8) FIGS. 8A-8F are fluorescent microphotographs of ANSCs on planar surface (FIGS. 8A and 8D), aligned fibrous electrospun 480 nm scaffold (FIGS. 8B and 8E) and random fibrous scaffold (FIGS. 8C and 8F). Neural differentiation patterns were investigated after 5 days, labeling with anti-Tuj 1 antibodies. Note increased neuron proliferation patterns on aligned scaffold (FIG. 8E). (From: Lim et al, 2010; Biomaterials: 34 (32).)

(9) FIG. 9 is a diagram illustrating a single piece two-electrode collector electrospinning apparatus setup and folding demonstration.

(10) FIG. 10 is a diagram illustrating a rotating spindle electrospinning system.

(11) FIG. 11 is a diagram illustrating a system for coating an object during electrospinning weaving.

(12) FIG. 12 illustrates traces of every visible line over the individual nanofibers on the presented SEM microscopy image, which was used with ImageJ software to analyze the nanofiber.

(13) FIG. 13 is a table including data related to an angular output of the traced lines.

(14) FIGS. 14A and 14B are bright-field microscopy photographs of electrospun PLA nanofiber (7.5% w/w), with solvent DCM:DMF. FIG. 14A is the result when using a control static screen collector at 400. FIG. 14B is the result when using a rotating dual electrode collector at 1050 rpm and 1000.

(15) FIGS. 15A and 15B are SEM microscopy photographs depicting the alignment of PLA nanofibers during deposition onto a rotating dual electrode collector at 1050 rpm (at 2000) and a static screen negative control (at 10,000), respectively.

(16) FIG. 16 is a plot depicting an angular distribution of nanofiber deposited on rotating collector, 1050 rpm.

(17) FIG. 17 is a bright-field microscopy photograph at 1000.

(18) FIG. 18 is a SEM microscopy photograph depicting the severance of nanofibers during their deposition on a fast rotating (1350 rpm) collector.

(19) FIG. 19 is a plot depicting an angular distribution of nanofiber deposited on rotating collector, 1050 rpm.

(20) FIG. 20 is a bright-field microscopy photograph of the PLA nanofiber at 400, when DCM:pyr is used as a solvent and the collector rotation is 1050 rpm.

(21) FIG. 21 is a SEM microscopy photograph of the PLA nanofiber at 1000, when DCM:pyr is used as a solvent and the collector rotation is 1050 rpm.

(22) FIG. 22 is a plot depicting an angular distribution of nanofiber deposited on rotating collector, 1050 rpm.

(23) FIG. 23 summarizes is a table summarizing diametric measurement data corresponding to nanofibers made with different electrospinning methods and/or parameters.

(24) FIGS. 24A and 24B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 3 cm distance.

(25) FIGS. 25A and 25B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 4 cm distance.

(26) FIGS. 26A and 26B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 5 cm distance.

(27) FIGS. 27A and 27B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 6 cm distance.

(28) FIGS. 28A and 28B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 10 cm distance.

(29) FIGS. 29A and 29B illustrate a process of fabricating a nanofiber-gel laminated article of manufacture.

(30) FIG. 30 is a photograph of a glass tube that has been inserted into the center of electrodes of electrospinning apparatus and coated for 5 minutes with a nanofiber.

(31) FIG. 31 is a photograph of a nanofiber/hydrogel laminated glass tube.

(32) FIG. 32 is a photograph of nanofibers formed on aluminum wire electrodes.

4. DETAILED DESCRIPTION

4.1 Definitions

4.1.1 Fiber Alignment

(33) The meaning of alignment of fibers, such as nanofibers for example, is provided here. Deposited nanofiber is usually defined by the random bending instability, a property of electrospinning still not very well understood, but known to be responsible for fiber thinning and elongation. To address this problem, example embodiments consistent with the present invention try to override bending instability by using a rotating dual collector design. While our design increases the alignment of nanofibers, bending instability still occurs and some fibers will not be aligned perfectly around the arbitrary axis of alignment. The following describes how one may evaluate empirically the alignment of nanofibers.

(34) To analyze the nanofiber, the present inventors have used ImageJ software, version 1.48. ImageJ is an open source analytical program that allows for evaluation of visual data in an obtained image. Using the software, the present inventors were able to trace every visible line over the individual nanofibers on the presented SEM microscopy image. (See, e.g., FIG. 12.) ImageJ generates an angular output of the traced lines. FIG. 13 is a table including this data, as processed manually, and includes the observed angular counts within a certain degree range. On an ideally fiber, all fibers would be counted as angles around 90. Again, due to bending whipping instability this is not the case. So an aligned nanofiber scaffold would have most angles around the central axis (90), with other fibers normally distributed around it. This approach ensures that most fibers are aligned around the same axis and the least fibers are perpendicular to that axis. To test for normal distribution, normal distribution goodness of fit statistical analysis was used. This allows the data to be decomposed, and rearranged into how it would look like under ideal normal distribution. Observed data can then be compared to the ideal expected data. The output of such analysis is the Chi-square value, which, if fits into the critical value range, allows one to rule out the null hypothesis that the obtained nanofibers are not angularly normally distributed around the alignment axis.

(35) Referring to FIG. 13, the table provides a sample Excel calculation table of statistical analysis of the angular distribution, based on ImageJ output, of PLA nanofibers formed on a rotating collector (1050 rpm, DCM:DMF 70:30 solvent). Individual z scores were calculated using:

(36) $z=\frac{x_i-\stackrel{\_}{x}}{s}$

(37) Expected frequency values were established using a normal distribution frequency table with matching z scores. Chi-square values were calculated as follows:

(38) $X^2=\underset{i=1}{\overset{k}{.Math.}}\left[\frac{{\left(O_i-E_i\right)}^2}{E_i}\right],$

(39) where Oi and Ei are observed and expected angular counts, respectively. Here, probability of obtaining X.sup.2 as large as 11.037 if the null hypothesis (that data is not normally distributed) is true is higher than 0.10, indicating that the angles might, in fact, be normally distributed around 90.

(40) Now, the obtained X.sup.2 value can be compared to the established percentiles of the Chi-Square Distribution. For eight (8) degrees of freedom, we have X.sup.2.sub.90 of 13.362 and X.sup.2.sub.95 of 15.507. Our found value is lower than X.sup.2.sub.90,8DoF so we can again conclude that angles might, in fact, be normally distributed around 90. For comparison, X.sup.2 value for the negative control static screen is 45.87, which is larger than X.sup.2.sub.90,8DoF so the null hypothesis for that cannot be rejected (note, here we also compare to the same X.sup.2.sub.90,8DoF because we have the same sample size hence same DoF in negative control, which is not necessarily the case for the whole experiment).

(41) Thus, in the following, aligned may be defined in terms of p, given that the lower the p is, the closer is the match. Since, however, an ideal normal distribution count is recalculated for every trial, only how much the obtained results match the expected (under normal distribution) results can be evaluated. Hence, it is possible to determine, statistically, whether the fibers are aligned or not aligned.

4.1.2 Three Dimensional

(42) In some instances, three dimension is meant to convey that fibers that are formed in multiple layers. A standard setup with a static screen forms random and interconnected (see SEM images for static control) fibers as a mat, which is different from a layered (3D) structure formed on a rotating collector consistent with the present invention.

4.2 Example Apparatus for Producing 3D Nanofiber Structure

4.2.1 Linear, Single Piece Static Double Electrode Collector

(43) FIG. 9 is a diagram illustrating a single piece two-electrode collector electrospinning apparatus setup and folding demonstration. As shown, the apparatus setup includes polymer solution 1, a spinneret needle (highlighted in green) 2, electrodes (Red positive and blue negative) 3, a folded single piece aluminum foil (e.g., grounding) counter-electrode collector 4. Note that the distance between electrodes, d, can easily be set during folding. In FIG. 9, the distance between the electrodes is 3 cm. The distance h here represents the distance between the spinning needle and collector electrodes.

(44) To ensure an even charge distribution, a collector was prepared from a single piece aluminum foil (Reynolds Wrap Aluminum Foil). (In this research no cell work was performed, thus a relatively toxic but conductive and malleable metal was chosen.) Rolling the foil as shown in FIG. 9 and bending along the measured lines allowed for a formation of a single collector with two exposed parallel electrodes of equal length with distance din between, which is preset during folding.

(45) As FIG. 9 shows, after folding, the collector (D) was attached to a (e.g., grounding) counter-electrode (C). After the polymer solution (A) was loaded into the syringe (Norm-Ject 10 mL) and mounted onto the syringe pump (Harvard Apparatus 11 Plus Syringe Pump), electrode (C) was attached to the needle (20 gauge diameter) (B). Controlled distance between the needle and the collector h (chosen to be 10 cm for shown sample) and distance between two collecting electrodes, d (in FIG. 9, a sample 3 cm was chosen), was fixed during the entire spinning process. An electrode, as well as the counter-electrode, were attached to the voltage source (Ultravolt's HV Rack) and a voltage regulation mode was set to provide a constant voltage throughout the entire electrospinning process.

4.2.2 Conventional Static Screen Electrospinning Apparatus Setup, Negative Control (Optional)

(46) As explained earlier, whipping instability creates randomness in nanofiber deposition onto the collector. To design a negative control, a setup which does not intervene with the random collecting process was chosen. For this apparatus setup, following an already established method of electrospinning as shown on FIG. 9, a horizontal apparatus was assembled with square shaped aluminum foil folded three times to form a screen with dimensions 1010 cm.sup.2. The screen was attached to the counter electrode as a single static collector. For that design the needle-collector distance h was fixed and was the same as during two-electrode collecting system.

4.2.3 Rotating Two-Electrode Spindle Collector and Coating

(47) FIG. 10 is a diagram illustrating an example rotating spindle electrospinning system consistent with the present invention. As shown, a parallel two electrode aluminum screen (A) is attached to a conducting aluminum wire (B). Transmitting aluminum wire (C) is coiled around and attached to a (e.g., grounding) counter-electrode (D). Magnetic stirring bar (E) provides rotation via stirrer (F). The circuit is complete through attachment of electrode to a needle with polymer solution (G).

(48) The setup illustrated in FIG. 10 is based on the fact that a rotating conducting aluminum wire (National Hardware V2566 diameter 18 ga) (B, in black), is attached to another static transmitting aluminum wire (C, in grey), coiled around it and attached to a (e.g., grounding) counter-electrode (D). This prevents coiling of the transmitting wire C, since the conducting wire B can rotate inside of it C freely. Rotating parallel dual electrode aluminum screen (A), (folded as shown on FIG. 9, except for skipping the last step) is similar to the static setup discussed earlier. Rotation and its regulation is provided by the attached magnetic stirring bar (E) and the magnetic stirrer. Polymer loaded into a syringe (G), which is attached to an electrode, completes the circuit. The dual electrode collector was set up so that the central axis of rotation is exactly half the distance between the electrodes, d, to ensure an even distribution of centripetal force.

(49) The setup of FIG. 10 allows for deposition of nanofiber onto the rotating electrodes; a material inserted into the rotating system is not subjected to current. FIG. 11 is a diagram illustrating an example system for coating an object (e.g., a glass tube or any other substrate that can be positioned within the rotating electrodes) during electrospinning weaving. As shown in FIG. 11, a needle (1102) attached to electrode expels the nanofiber jet onto the rotating collector (1104). A glass tube (1106) fixed in between the central axis of rotation collects the deposited nanofiber without conducting current. As shown in FIG. 11, a coating experiment setup is similar to the one of FIG. 10, but further includes a glass tube (1106) which is inserted into the central axis of rotation of collecting electrodes (1104) during the electrospinning. The glass tube was not attached to electrodes and was not subjected to current. Other substrates may be used instead. Other example substrates include, for example, a food product, a body part, etc.

4.3 Example Methods for Producing 3D Nanofiber Structure

4.3.1 Polylactic Acid Polymer Preparations

(50) Commercial grade polylactate for all experiments in this work was obtained from MakerBot, Large True Orange PLA Filament. Molecular weight (MW) of the polymer was not provided by the company. An filament was cut into pieces of an appropriate mass and solubilized in dichloromethane (DCM) (Sigma, anhydrous, 99.8%; vapor pressure 352 mmHg at 20 C. as provided by Sigma) with either Pyr (Sigma, anhydrous, 99.8%; vapor pressure 20 mmHg at 25 C. as provided by Sigma) or DMF (Sigma, anhydrous, 99.8%; vapor pressure 2.7 mmHg at 20 C. as provided by Sigma) at the ratios of DCM:Pyr 60:40 and DCM:DMF 70:30 via heating and stirring overnight. Solution was visually checked for homogeneous polymer solubilization in solvent. For the purpose of the experiments presented in this work, a fixed 7.5% w/w PLA solution was prepared in all cases. Of course, other concentrations can be used. Lower polymer concentrations were reported to produce a higher quality nanofiber, but since the commercial grade polymer's MW was not provided, it was assumed to be small, and a higher PLA concentration solution was chosen for electrospinning.

4.3.2 Electrospinning Apparatus Parametric Setup

(51) For all electrospinning experiments conducted, previously described parameters were adopted with consideration of developed MATLAB model (discussed later). Parameters were chosen as in the following table.

(52) TABLE-US-00001 TABLE Parametric setup for electrospinning Parameter Setup chosen/measured Ambient Conditions Standard temperature and pressure (~23 C. and 1 atm); no airflow Between-electrode distance 3 cm for dual screen collector, d Electrode needle to collector 10 cm distance, h Applied voltage, U 15,000 V, DC Polymer delivery rate 1.5 mL/h Total volume of electrospun 4 mL solution Collector rotation 0, 1050 rpm and 1350 rpm Needle diameter 20 ga, 0.91 mm outer, 0.60 mm inner Solvent Either DCM:Pyr or DCM:DMF* Rotation 0, 1050 rpm, 1350 rpm

(53) The versatility, in terms of polymer solution of the proposed method, was also tested. For this, two solvents, DCM:Pyr (higher conductivity) and DCM:DMF (lower conductivity) were tested. From this, the following eight setups were evaluated:

(54) TABLE-US-00002 TABLE Solvent choice for eight electrospinning experiments First experiment Second experiment Electrospinning Setup solvent chosen solvent chosen Static screen collector DCM:Pyr DCM:DMF (negative control) Static dual electrode collector DCM:Pyr DCM:DMF Rotating dual electrode DCM:Pyr DCM:DMF collector, 1050 rpm Rotating dual electrode DCM:DMF second experiment collector, 1350 rpm not conducted Coating experiment DCM:DMF second experiment no conducted

(55) Four example setups, as well as diametric measurements of nanofibers made with different electrospinning methods and/or parameters, are described in 4.3.3-4.3.7 below.

4.3.3 Example 1: Standard Setup

(56) The parameters used in a first example electrospinning method are provided in the following table:

(57) TABLE-US-00003 Parameter Setup chosen/measured Ambient Conditions Standard temperature and pressure (~23 C. and 1 atm); no airflow Between-electrode distance 3 cm for dual screen collector, d Electrode needle to collector 10 cm distance, h Applied voltage, U 15,000 V, DC Polymer delivery rate 1.5 mL/h Total volume of electrospun 4 mL solution Collector rotation 1050 rpm Needle diameter 20 ga, 0.91 mm outer, 0.60 mm inner Solvent DCM:DMF 70:30 Note that DCM is Dichloromethane, DMF is N,N-Dimethylformamide (vapor pressure 2.7 mmHg at 20 C., conductivity 0.9-1.5 S/cm at 25 C.).

(58) FIGS. 14A and 14B are bright-field microscopy photographs of electrospun PLA nanofiber (7.5% w/w), with solvent DCM:DNIF. FIG. 14A is the result when using a control static screen collector at 400. FIG. 14B is the result when using a rotating dual electrode collector at 1050 rpm and 1000.

(59) FIGS. 15A and 15B are SEM microscopy photographs depicting the alignment of PLA nanofibers during deposition onto a rotating dual electrode collector at 1050 rpm (at 2000) and a static screen negative control (at 10,000), respectively. In each case, the solvent was DCM:DMF. Note the horizontal alignment of the nanofibers in FIG. 15A as compared with those of FIG. 15B.

(60) FIG. 16 is a plot depicting an angular distribution of nanofiber deposited on rotating collector, 1050 rpm. Observed count (expected with 90 angle highlighted) vs from normal distribution around 90. The solvent is DCM:DMF. In blue is the hypothetical axis of nanofiber alignment. N=103, DoF=8. The angular distribution was measured using ImageJ software, version 1.48. H.sub.0: sample is not normally distributed around 90. Student t-test: t.sub.observed<t.sub.crit, p<0.05. Chi-Square test of goodness of fit: X.sup.2=11.037; X.sup.2.sub.90>X..sup.2 The p value of obtaining a value of X.sup.2 as large as 11.037 if H.sub.0 is true is higher than 0.10. Reject H.sub.0; angles of fiber deposition around a rotating collector of 1050 rpm, using DCM:DMF solvent, may be distributed normally around 90.

4.3.4 Example 2: Changing Collector Rotation Speed

(61) TABLE-US-00004 Parameter Setup chosen/measured Ambient Conditions Standard temperature and pressure (~23 C. and 1 atm); no airflow Between-electrode distance 3 cm for dual screen collector, d Electrode needle to collector 10 cm distance, h Applied voltage, U 15,000 V, DC Polymer delivery rate 1.5 mL/h Total volume of electrospun 4 mL solution Collector rotation 1350 rpm Needle diameter 20 ga, 0.91 mm outer, 0.60 mm inner Solvent DCM:DMF 70:30

(62) FIG. 17 is a bright-field microscopy photograph at 1000. Note the severed ends of the nanofibers. The inventors believe that the higher velocity rotation of the collector causes breaks (ROIs) and discontinuous PLA nanofiber formation.

(63) FIG. 18 is a SEM microscopy photograph depicting the severance of nanofibers during their deposition on a fast rotating (1350 rpm) collector. Orange circles overlayed on the SEM microphotograph denote severed ends.

(64) FIG. 19 is a plot depicting an angular distribution of nanofiber deposited on rotating collector, 1050 rpm. Observed count (expected with 90 angle highlighted) vs from normal distribution around 90. The solvent is DCM:DMF. In blue is the hypothetical axis of nanofiber alignment. N=89, DoF=8. The angular distribution was measured using ImageJ software, version 1.48. H.sub.0: sample is not normally distributed around 90. Student t-test: t.sub.observed<t.sub.crit, p<0.05. Chi-Square test of goodness of fit: X.sup.2=13.460; X.sup.2.sub.95>X.sup.2>X.sup.2.sub.95. The p value of obtaining a value of X.sup.2 as large as 13.460 if H.sub.0 is true between 0.05 and 0.10. Reject H.sub.0; angles of fiber deposition around a rotating collector of 1350 rpm, using DCM:DMF solvent, may be distributed normally around 90.

4.3.5 Example 3: Changing Solvent

(65) TABLE-US-00005 Parameter Setup chosen/measured Ambient Conditions Standard temperature and pressure (~23 C. and 1 atm); no airflow Between-electrode distance 3 cm for dual screen collector, d Electrode needle to collector 10 cm distance, h Applied voltage, U 15,000 V, DC Polymer delivery rate 1.5 mL/h Total volume of electrospun 4 mL solution Collector rotation 1050 rpm Needle diameter 20 ga, 0.91 mm outer, 0.60 mm inner Solvent DCM:Pyr 60:40 Note that Pyr is Pyridine (v.p. 20 mmHg at 25 C., conductivity 12.7 S/cm at 22.4 C.)

(66) FIG. 20 is a bright-field microscopy photograph of the PLA nanofiber at 400, when DCM:pyr is used as a solvent and the collector rotation is 1050 rpm. Note the beads on string formations. The inventors believe that these are probably due to high vapor pressure and conductivity of Pyr associated with higher mass flow.

(67) FIG. 21 is a SEM microscopy photograph of the PLA nanofiber at 1000, when DCM:pyr is used as a solvent and the collector rotation is 1050 rpm. Again note the bead formations as well as garland (versus straight) fiber deposition. Again, the inventors believe that these are probably due to high vapor pressure and conductivity of Pyr associated with higher mass flow.

(68) FIG. 22 is a plot depicting an angular distribution of nanofiber deposited on rotating collector, 1050 rpm. Observed count (expected with 90 angle highlighted) vs from normal distribution around 90. The solvent is DCM:pyridine. In blue is the hypothetical axis of nanofiber alignment. N=74, DoF=8. The angular distribution was measured using ImageJ software, version 1.48. H.sub.0: sample is not normally distributed around 90. Student t-test: t.sub.observed<t.sub.crit, p<0.05. Chi-Square test of goodness of fit: X.sup.2=15.203; X.sup.2.sub.95>X.sup.2>X.sup.2.sub.95. The p value of obtaining a value of X.sup.2 as large as 15.203 if H.sub.0 is true between 0.05 and 0.10. Reject H.sub.0; angles of fiber deposition around a rotating collector of 0350 rpm, using DCM:Pyr solvent, may be distributed normally around 90.

4.3.6 Comparison of the Average Diameters of Example Methods

(69) The following table includes diametric measurements of nanofibers made with different electrospinning methods and/or parameters.

(70) TABLE-US-00006 Collector type Average diameter, nm Standard error screen, pyridine (control) 110.67 4.10 rot1050, pyridine 150.00 6.84 screen, DMF (control) 131.10 10.34 rot1050, DMF 94.91 3.11 rot1350, DMF 294.29 15.80

(71) FIG. 23 summarizes this data.

4.37 Example 4: Changing Needle to Collector Distance, Static with MATLAB Analysis

(72) FIGS. 24 to 28 show, side-by-side, MATLAB model and actual nanofiber distribution for different needle to collector distances. The remaining electrospinning parameters are summarized in the following table:

(73) TABLE-US-00007 Parameter Setup chosen/measured Ambient Conditions Standard temperature and pressure (~23 C. and 1 atm); no airflow Between-electrode distance 3 cm for dual screen collector, d Electrode needle to collector 3, 4, 5, 6, 10 cm distance, h Applied voltage, U 15,000 V, DC Polymer delivery rate 1.5 mL/h Total volume of electrospun 4 mL solution Collector rotation 0 Needle diameter 20 ga, 0.91 mm outer, 0.60 mm inner Solvent DCM:DMF 70:30

(74) FIGS. 24A and 24B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 3 cm distance. Note the separation of electric potential between counter-electrodes and a low value of 1*10.sup.4V. Note also a disrupted, irregular nanofiber formation with one side preference when distance between needle and collector is too small.

(75) FIGS. 25A and 25B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 4 cm distance. Note the shared potential between counter-electrodes 2.5*10.sup.4V. Note also an uneven, one-sided distribution of nanofiber between two collecting electrodes.

(76) FIGS. 26A and 26B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 5 cm distance. Note the shared potential between counter-electrodes 2.5*10.sup.4V. Note also the gradient in nanofiber distribution between two collecting electrodes.

(77) FIGS. 27A and 27B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 6 cm distance Note a high shared potential between counter-electrodes 0.5*10.sup.5V. Note also a uniform nanofiber distribution between two collecting electrodes.

(78) Finally, FIGS. 28A and 28B are, respectively, MATLAB model and experimental results of electric field during electrospinning, 10 cm distance Note a high shared potential overlap between counter-electrodes 0.5*10.sup.5V. Note also a uniform nanofiber distribution between two collecting electrode.

4.4 Example 3D Nanofiber Structure

(79) FIG. 32 is a photograph of nanofibers formed on aluminum wire electrodes. An example nanofiber product (preferably produced by the method of claim 1) has a distribution of angles of fibers with respect to an ideal (perfectly aligned) distribution that is within a predefined aligned range. An example nanofiber has an average diameter of fibers between 70 and 300 nm, for example between 94 and 295 nm. Note that the desired average diameter may depend on the intended application. For example, a desired average diameter might depend on cell type ideal interactions.

4.5 Example Nanofiber-Gel Laminated Structure

(80) As noted in section 4.3.2 above with reference to FIG. 11, a substrate (e.g., a glass tube) can be coated with spun nanofibers in a manner consistent with the present invention. The resulting nanofiber coated substrate may then be provided with a gel coating (e.g., by dipping the nanofiber coated substrate into solution of alginate (an anionic polymer), and then dipping into Ca solution (a cationic bivalent cross linker). Such electrospinning and dipping can be repeated to prepare a laminated article of manufacture having multiple, alternating layers of fibers and gels. Such laminated articles of manufacture may be very important when tissue engineering hierarchal complex structures.

(81) One example method for making such a laminated article of manufacture is described below with reference to FIGS. 29A and 29B. As the polymer solution, PLA, 7.5% w/w in Dichloromethane:N,NDimethylformamide 70:30, is used. As the alginic acid solution, potassium alginate in water, 2.5% w/w, is used. As a Calcium Chloride solution, CaCl2 in water, 2.5% w/w, is used. Referring to FIG. 29A, first, the polymer is loaded into a syringe and a electrospinning weaving (1050 rpm, 15 kV DC) is initiated. More specifically, a glass tube is inserted into the center of the electrodes and coated for 5 minutes with a nanofiber, as depicted by 2910 of FIG. 29A, as well as FIG. 30. The resulting nanofiber coated tube is then dipped into an alginate solution (as depicted by 2920) and swiftly transferred and dipped into a calcium chloride solution (as depicted by 2930) to crosslink the viscous alginate solution, thereby forming a hydrogel layer around (as depicted by 2940 of FIG. 29A, as well as FIG. 30B). The resulting gel and nanofiber coated tube is then inserted into the center of the electrospinning electrodes once again, and the whole process is repeated n times (See, e.g., FIG. 29A) to form a nanofiber/hydrogel laminated tube. (See, e.g., FIG. 31.).

(82) Referring to the gel-nanofiber coating setup of FIG. 29B, initially, an electrospinning weaving setup is constructed with a PLA nanofiber (A) forming on the collector (C). A glass tube (B) is inserted into the central axis of rotation and is coated with nanofiber (D, first layer in black). The glass tube with first layer of nanofiber is dipped into the alginate solution (E, in red). The glass tube with nanofiber and alginate (F) is then dipped into CaCl2 solution (G, yellow) to crosslink the alginate. This forms a layer of hydrogel around the nanofiber (H). 4. This procedure is repeated multiple times to form a multilayered structure. I is the glass tube layer, K are the nanofiber layers and J are the formed alginate layers.

(83) The foregoing procedure and/or materials can be modified, especially the gelation process. For example, instead of using alginate, another anionic polymer such as carrageenan, or other anioinic polysaccharides, polyacrylic acids, or other anionic synthetic polyelectrolytes. for example, may be used instead. Further, instead of using a cationic bivalent cross linker such as a Ca solution, another cationic multivalent cross linker such as polyethyleneimine, or polylysine may be used instead. Gels can also be formed with cationic polymers such as polyethylene imine with anionic multivalent crosslinkers or covalent crosslinkers such as glutaraldehyde. In addition, gel layers may be produced by photocrosslinking, other covalent crosslinkings, interpolymer complexations or other multipolymer complexes.

(84) Although the laminated article of manufacture having multiple, alternating layers of fibers and gels were described as using a PLA nanofiber, any synthetic or biological material capable of forming a fiber (such as the alternatives described in 4.6 below) may be used instead.

(85) Although not shown, cells can be introduced before coating with alginate or crosslinking. For example, stem cells may be introduced, at various steps, into the gels. Thus, soft material may be introduced into the woven structures.

4.6 Refinements, Alternatives and Extensions

(86) iA mechanical stress-strain analysis may be used to evaluate strength and stiffness of the created PLA nanofiber. Cytotoxicity may also be analyzed. Understanding these parameters is import to assess the suitability of the created mesh for use in tissue engineering of various tissues.

(87) Although polylactic acid preparations were described in 4.3.1 above, other polymers can be used instead. Such alternative polymers may include, for example, any commercial thermoplastic polymer if soluble, any commercial elastomeric polymer if soluble, and any thermosetting polymer if the setting completed after spinning. Any of the above may be done in the lab with any polymerization method: Enzymatic, condensation, free radical, electrochemical, template or any other polymerization method. That is, the polymers can be prepared by different methods In some embodiments, the polymer may be mixed with another polymer. For example, a polymer blend with miscible of phase separated morphology; phase separated by nucleation and growth or spinodal phase separation mechanism. That is, in addition to single polymers, but multiples in mixtures, blends with different morphologies, etc., may be used instead.

(88) In some example embodiments consistent with the present invention, the polymer chemical composition may be copolymer made of various monomers, block copolymer similarly, dendrimer or other macromolecule, liquid crystalline and/or semicrystalline polymer.

(89) Further, in some example embodiments consistent with the present invention, the polymer may be filled with functional nanoparticles, drug, therapeutics, gels, drug delivery systems, DNA, siRNA, mRNA, proteins, viruses, bacteria or any other inorganic, organic or biological entity. In addition, a polymer composite (e.g., filled with inorganic, carbon nanotubes, fullerenes or any modifications of the ones, grapheme and grapheme modifications) may be used.

(90) In any event, the molecular weight of the polymer (or polymer blend, or polymer chemical composition, etc.) should be large enough to allow formation of a nanofiber.

(91) Although the syringe is fixed in some of the foregoing example embodiments, it can be moved up and/or down with respect to the collector. Otherwise, a gradient with fibers concentrated at the center may result. Fibers might be even more concentrated if the electrodes frame a concave shape. Thus, in some example embodiments consistent with the present invention, the syringe dispensing the polymer solution can be moved in one or more of the x,y,z directions. Alternatively, or in addition, in some example embodiments consistent with the present invention, the receiving electrode system can be moved in one or more of the x,y,z directions. Various shapes and/or thickness gradients can be obtained by controlling (such as continuously changing) one or more of x, y, z positions of the syringe and/or the receiving electrode system.

(92) In some example embodiments consistent with the present invention, the material to be coated and/or the formed fiber shapes can be surface modified so that they become cationic, anionic, hydrophobic, or hydrophilic. This may be done to increase their affinity for other materials to absorb onto them. For example, anionic or cationic polyelectrolytes, synthetic or biological, can be attached to the surfaces for various reasons, such as for improved functionalities.

(93) Referring back to section 4.5 above, in some example embodiments consistent with the present invention, one of the functionalities may be a gel formation when the attached polyelectrolytes then bind with bi (or multi)functional molecules like Ca2+ to crosslink and form gels on the surface of the material to be coated. During such an example gelation process, other functional materials (e.g., proteinuous growth factors, signaling agents like cytokines and cells such as stem cells) can be added. These functional material(s) can be added by dipping into, pouring into, dispensing into, injecting, and/or printing and forming various gradients, diffusive layers and compositions.

(94) In some example embodiments consistent with the present invention, after the nanofiber is produced, the electrodes can be removed, and perhaps replaced with a frame having a different material. (For example, a material that cells have an affinity for.)

4.7 Conclusions

(95) Known mats of nanofilaments are too dense for tissue engineering. Nanofibers fabricated using the foregoing techniques are expected to have many applications, such as tissue engineering (e.g., tissue scaffolding) for example. Other potential applications include, for example, filters, protective clothing, stain resistant fabrics, drug delivery media/encapsulation (e.g., for control of delivery rate), light-weight solar sails, aircraft wings, and bullet proof vests, fuel cell membranes, food encapsulation, soft candy, ice cream and all kinds of food coating.