SURFACE MODIFICATION OF PLGA ELECTROSPUN SCAFFOLDS FOR WOUND HEALING AND DRUG DELIVERY APPLICATIONS. A Thesis. Presented to

Transcription

1 SURFACE MODIFICATION OF PLGA ELECTROSPUN SCAFFOLDS FOR WOUND HEALING AND DRUG DELIVERY APPLICATIONS A Thesis Presented to The Graduate Faculty of The University of Akron In Partial Fulfillment of the Requirements for the Degree Master of Science Jacob A. Iselin December, 2008

2 SURFACE MODIFICATION OF PLGA ELECTROSPUN SCAFFOLDS FOR WOUND HEALING AND DRUG DELIVERY APPLICATIONS Jacob A. Iselin Thesis Approved: Accepted: Advisor Dr. Yang H. Yun Dean of the College Dr. George K. Haritos Committee Member Dr. Stanley E. Rittgers Dean of the Graduate School Dr. George R. Newkome Committee Member Dr. Darrell H. Reneker Date Department Chair Dr. Daniel B Sheffer ii

3 ABSTRACT PLGA scaffolds used for wound healing and drug delivery can be surface modified to control cellular attachment to the fibers. An in situ technique has been developed whereupon surface modifying polymers migrate to PLGA microfiber surfaces during electrospinning processes by thermally induced phase separation. Biodegradable scaffolds capable of blocking or encouraging cellular adhesions have been fabricated. An electrospun scaffold is proposed in which a primary scaffold facilitates enhanced cellular attachment, while a secondary scaffold, that blocks cellular interactions, is intended to carry a drug payload to aid in tissue development. Thus, PLGA is surface modified by blending with collagen (I), fibronectin, PEG-grafted-chitosan, and PEG-PLGA block copolymer. Physical characterization of the surface-modified PLGA scaffolds is performed to define the shape, fiber diameter, and surface texture of the electrospun fibers. Additionally, assessment of contact angle and mechanical strength are examined to distinguish wettability and tensile characteristics, respectively. Biological characterization is performed to assess cellular adhesion characteristics, to examine the cellular preference between two different surface-modified blends in a single scaffold, and for the probing of vinculin proteins constituting focal adhesion sites to determine cellular adhesion mechanisms. Results indicate that electrospinning produces textured surface-modified microfibers not significantly different from pure PLGA fiber scaffolds. Contact angle results indicate that surface modification alters surface hydrophilicity. iii

4 Tensile testing determines that surface modifications do not alter yield or tensile strength, but have an effect on the elastic modulus. Results of cellular adhesion studies indicate that the addition of collagen promotes cellular binding, but is not significantly greater than the control. Cellular adhesion among competitive pairings of the modified blends shows no significant differences. Probing of vinculin proteins is inconclusive. Conclusions based upon the physical and biological characterizations indicate that the TIPS technique used for surface modification of PLGA is an ineffective technique, although collagen (I) and PEG-grafted-chitosan may represent the most efficacious biomaterials for use as surface modifiers of PLGA scaffolds. iv

5 DEDICATION I would like to dedicate this thesis to my mother, Mary Anne Ricci, for her passion and zest for life, which have shaped this author s character and personality. Her willingness to support my quest of higher education and the understanding of the hardships associated with the journey are deserving of the highest honors and gratitude. I love you Mom. Thank you. Additionally, I would also like to dedicate this work to my former mentor, Dr. Charles G. Orosz ( ), who was an extraordinary man, as well as the foremost immunologist in his field. Known simply as CGO, he succumbed to a rare immunological disease at just 56 years young, which left his family, friends and colleagues in a state of shock and loss. CGO is sorely missed today, but will always be remembered by this author for the impact he had on my life as an ambitious researcher. He commanded an infectious admiration, and I cherish the opportunity to have been his friend. He is survived by his wife Nancy, and their three children Kate, Matt, and Molly. v

6 ACKNOWLEDGEMENTS Foremost, I want to thank my advisor, Dr. Yang H. Yun, Assistant Professor of the Department of Biomedical Engineering at the University of Akron, for his mentorship and guidance in completing this Master s thesis. He provided a level of detail, precision, and righteousness in helping me design and execute this project that was unmatched by the lot of my previous experiences. His commitment to excellence in scientific research is a virtue I will carry with me in all my future endeavors. I would also like to extend my utmost appreciation to Dr. Stanley E. Rittgers, Professor of the Department of Biomedical Engineering at the University of Akron, for his patience, vision, advice, and encouragement. The impact of his values as an instructor, researcher, and friend upon this author are deserving of the highest of recognitions. I would like to extend my gratitude to Dr. Darrell H. Reneker, Professor of the Department of Polymer Science at the University of Akron for his dedication to my thesis committee and his counsel on the topics concerning polymer synthesis and electrospinning. Additionally, I would like to extend my utmost thanks to Dr. Glen O. Njus, Research Associate Professor of the Department of Biomedical Engineering at the University of Akron for his contribution of intellect, expertise, and time with respect to the material testing aspects of this project. His benevolent approach to mentorship was a special gift that this author will not soon forget. Special acknowledgement is due to Dr. Daniel B. Sheffer, Associate Professor and Chair of the Department of Biomedical Engineering at the University of Akron, for his advice, vi

7 guidance, funding of this project. Additionally, I would like to thank Dr. Sheffer for his consultation on matters of statistics and his kind friendship during difficult times. I would like to extend my appreciation to Ms. Jeanette Killius, Director of Microscopy at NEOUCOM for her assistance with confocal microscopy, Dr. Tao Han, former graduate assistant of Dr. Darrell Reneker, for his instruction in the art of electrospinning, and to Anand Parikh, Ph.D. candidate, and graduate assistant of Dr. Glen Njus for his expertise and assistance with material testing. Finally, I would like to acknowledge my colleagues in the Department of Biomedical Engineering. Great admiration is bestowed upon Mr. Andrew Ditto, Mr. Parth N. Shah and Ms. Tasha Williams for their constant intellectual support and amicable fellowship. vii

8 TABLE OF CONTENTS Page LIST OF TABLES... xi LIST OF FIGURES... xii CHAPTER I. INTRODUCTION & SPECIFIC AIMS Introduction Specific Aims...3 II. BACKGROUND Polymers: PLGA, Copolymer Component of a Biodegradable Scaffold Electrospinning Hydrophobic, Hydrophilic, and Amphiphilic Interactions Surface Modification of PLGA Electrospun Fibers Focal Adhesion Sites & Cellular Attachment Specific Adhesion Non-Specific Adhesion...23 III. METHODS & MATERIALS Electrospinning Physical Characterization: Scanning Electron Microscopy...26 viii

9 3.3 Physical Characterization: Contact Angle Physical Characterization: Tensile Testing Biological Characterization: Cell Culture Biological Characterization: Fixation and Staining Protocols Biological Characterization: Immunofluorescence Biological Characterization: Quantification of Cellular Adhesion...33 on Fiber Scaffolds 3.9 Biological Characterization: Quantification of Cellular Preference...34 Between Two Surface-Modified Microfiber Matrices Interwoven in One Scaffold 3.10 Qualitative Comparison of Vinculin Deposition on Surface-Modified...35 Microfiber Scaffolds 3.11 Data Analysis & Statistics...36 IV. RESULTS Fabrication of Surface-Modified PLGA Electrospun Fibers Physical Characterization: Scanning Electron Microscopy Physical Characterization: Contact Angle Physical Characterization: Tensile Testing Biological Characterization: Antibody Titrations Biological Characterization: Cellular Adhesion on Fiber Scaffolds Biological Characterization: Cellular Adhesion Between...64 Competitive Fiber Scaffolds 4.8 Biological Characterization:Qualitative Assessment of Vinculin...67 Deposition and Confocal Microscopy ix

10 V. DISCUSSION Fabrication of PLGA Surface-Modified Microfiber Scaffolds Physical Characterization: The Effect of Surface Modification...77 on Fiber Diameter 5.3 Physical Characterization: The Effect of Surface Modification...78 on Contact Angle 5.4 Physical Characterization: Material Characteristics Biological Characterization: Analysis of Cellular Adhesion Biological Characterization: Competitive Cellular Adhesion Biological Characterization: Qualitative Analysis of Specific vs...88 Non-Specific Adhesion VI. CONCLUSIONS & FUTURE EXPERIMENTATION Conclusions Future Experimentation Recommendations for Patterning Fibers...93 REFERENCES...94 APPENDICES APPENDIX A: CELLULAR ADHESION AT INTERSECTING FIBERS APPENDIX B: FABRICATION OF NANO-SCALE FIBERS APPENDIX C: STATISTICS EXAMPLE: FIBER DIAMETER x

11 LIST OF TABLES Table Page 1. Polymer microfiber blends Competitive pairings of PLGA surface-modified blends Mean fiber diameters of PLGA polymer microfiber blends Means of the advancing and receding angles of contact...47 angle measurements 5. Dimensions of PLGA surface-modified electrospun mats...49 for tensile testing 6. Failure modes of PLGA surface-modified electrospun samples Material characteristics of surface-modified PLGA electrospun...55 microfiber films 8. Cellular adhesion of human fibroblasts on PLGA surface modified microfibers 9. Statistical values of Student s t-test on competitive microfiber scaffolds...66 xi

12 LIST OF FIGURES Figure Page 1. Chemical composition of PEG-grafted-chitosan Chemical composition of PLGA-PEG-PLGA triblock copolymer Depiction of a cellular focal adhesion binding through integrin proteins Protein configuration of a focal adhesion site (specific adhesion) Fluorescent microscopic images of cellular (a) actin filaments stained...21 by rhodamine phalloidin, and (b) associated vinculin proteins stained by anti-vinculin Ab. 6. Fluorescent images of actin filament staining of human fibroblasts...38 seeded on PLGA microfibers + titrated w/w % PEG-PLGA block copolymer (a) 1% BCP, (b) 2% BCP, (c) 5% BCP, (d) 10% BCP mounted on culture plate cover slips (63x-obj). 7. Electrospun surface-modified PLGA fibers: (a) PLGA alone,...40 (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer (300x-mag). 8. SEM image of pure PLGA fibers showing surface texture (2000x-mag) Axiovision software image of fiber diameter measurements (1500x-mag) Fiber diameter distribution of electrospun microfiber groups (a) pure...44 PLGA, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer. 11. Measurements using Axiovision software of (a) sample length/width...50 and (b) sample thickness (20x-obj). xii

13 12. Examples of the tensile failure modes: (a) mid-substance, (b) grip Stress versus strain curves of each replicate for (a) pure PLGA,...52 (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer. 14. Fluorescent images of the titration of vinculin protein staining on human...60 fibroblasts (a) 1:100, (b) 1:1000 dilution (63x & 20x-obj). 15. Fluorescent image of human fibroblasts on a glass substrate showing actin...61 filaments (red), nuclei (blue), and vinculin staining (green) (20x-obj). 16. Example of Axiovision software images of fiber measurement...64 for surface area and cell density determination (20x-obj). 17. Example of Axiovision software image of fiber measurement from a sample...67 of PLGA+ 1% Collagen + crystal violet dye (red stained microfibers) versus _PLGA+ 1% PEG-g-chitosan (10x-obj). 18. Fluorescent images of vinculin staining of human dermal fibroblasts seeded...69 on PLGA microfibers mounted on culture plate cover slips (63x mag obj). 19. Layered confocal image cascades of vinculin staining of human dermal...70 fibroblasts seeded on PLGA microfibers mounted on culture plate cover slips (a) trial 1, (b) trial 2 (40x-obj). 20. Positive control sample of vinculin protein staining (green) of human...72 fibroblasts mounted on German glass cover slips (20x-obj). 21. Fluorescent images of surface-modified PLGA microfiber samples probed...72 for vinculin staining (63x-obj); (a & b) PLGA+ 1% PEG-g-chitosan, (c) PLGA+ 1% collagen. xiii

14 CHAPTER I INTRODUCTION & SPECIFIC AIMS 1.1 Introduction Novel biodegradable tissue-engineered scaffolds for use in biomedical engineering applications, such as drug delivery, wound healing, tissue regeneration, and organ replacement, have garnered considerable interest in the last decade. Common technologies involve electrospinning of synthetic biodegradable polymers such as PLA, PGA, and copolymers such as PLGA for fabricating matrix scaffolds used to grow seeded cells for tissue development [1-4]. Additionally, scaffolds composed of native human proteins of the collagen and elastin families have been investigated as useful biomaterials for constructing tissue scaffolds [5]. All of these biomaterials individually have shown promise as viable tissue scaffold materials, as well they have shown flexibility in fabrication, post-processing, and cellular adhesion characteristics that have prompted investigators to look further into their applications [1-6]. An intriguing method to enhance fibrous scaffolds is blending of these synthetic polymers with natural polymers, molecules, and proteins in order to surface modify electrospun nano- and microfibers. Surface modification is a process by which a secondary molecule is blended with a biodegradable polymer in order to enhance the protein adsorption characteristics, modify surface polarity, or vary the hydrophobicity of the fiber surface. The tissue engineering goal of surface modification is to selectively 1

15 promote or inhibit cellular adhesion in a novel tissue scaffold. Human cells have an affinity for surfaces through which they can bind using transmembrane integrin receptor proteins [7-8]. This method of cell-to-substrate adhesion is a highly conserved trait of most eukaryotic cells and can be used to promote cellular adhesion to microfiber surfaces. Having the ability to selectively modify tissue scaffold surfaces may have major future implications for drug delivery applications primarily. Co-blending scaffolds that promote cellular adhesion and tissue growth on one microfiber, while simultaneously inhibiting cellular adhesion for the purpose of delivering drugs through standard biodegradative kinetics of the biopolymer in an adjacent secondary microfiber, displays an intriguing possibility. These surface-modified scaffolds would afford the medical community the opportunity to create improved wound dressings with specific drug delivery opportunities. Of further significance, the opportunity to seed a pluripotent cell type to a synthetically fabricated polymer scaffold containing genetic material, incorporated to induce cell differentiation and proliferation poses greater merit. By taking advantage of the cell s ability to adhere to material surfaces, we will investigate the cellto-substrate adhesion of primary human dermal fibroblasts and identify preferential cellular adhesion abilities. It is hypothesized that PLGA electrospun fibers can be surface-modified by the incorporation of a secondary polymer, such as collagen (I), fibronectin, PEG-graftedchitosan, or PLGA-PEG-PLGA triblock copolymer. Collagen and fibronectin will be employed to promote cell adhesion to microfiber surfaces, while PEG-grafted-chitosan and PLGA-PEG-PLGA triblock copolymer will be incorporated to inhibit cellular adhesion to microfiber surfaces. The objective of altering the adhesion characteristics of 2

16 biodegradable microfiber scaffolds is to create a bimodal scaffold in which one fiber type harbors cellular attachment and proliferation while the other fiber type facilitates drug delivery. The feasibility of developing these novel electrospun scaffolds with surfacemodified characteristics will be investigated in this experiment by performing the following specific aims. 1.2 Specific Aims Specific Aim #1: Fabricate PLGA microfiber scaffolds that are surface-modified with collagen (I), fibronectin, PEG-grafted-chitosan, and PLGA-PEG-PLGA triblock copolymer. Rationale Surface modification of electrospun PLGA microfibers may preferentially alter cell adhesion characteristics. PLGA microfibers are surface modified by blending alternative secondary polymers with the PLGA polymer solutions. The primary control polymer solution is a 10% w/v solution of PLGA (1.15 i.v.) in DCM or HFIP. Natural polymers such as collagen and fibronectin may promote specific adhesion by increasing the incidence of focal adhesion contacts on the microfiber surface. On the contrary, the synthetic polymers such as PEG-grafted-chitosan and PLGA-PEG-PLGA triblock copolymer are believed to inhibit cellular adhesion at the microfiber surface, because of the presence of PEG groups on the polymer fiber surface. 3

17 Specific Aim #2: Evaluate the physical characteristics, specifically size, shape, surface morphology, contact angle, and material strength of the surface-modified fiber scaffolds. Rationale Surface modification of biodegradable PLGA microfibers is hypothesized to create preferential cellular attachment to electrospun matrix scaffolds by either promoting or inhibiting integrin binding. Characterizing the fiber diameter and shape of the experimental electrospun fibers by scanning electron microscopy is a necessary analysis in order to ensure that surface modification does not alter these parameters or the fiber s morphology. Furthermore, it is prudent to show that the secondary polymers used to surface modify the fibers are being expressed on the material surface, and that these molecules are the effectors of surface wettability and ultimately differential integrin binding. Additionally, analysis of material characteristics by tensile testing is intended to ensure that the surface modification procedure does not diminish material strength. Null Hypotheses Null Hypothesis 1 (N 1 ) H 0 : A statistical difference does not exist in the mean fiber diameter among untreated PLGA microfibers, PLGA +1% collagen(i) microfibers, PLGA +0.1% fibronectin microfibers, PLGA +1% PEG-grafted-chitosan microfibers, and PLGA +10% PLGA-PEG-PLGA triblock copolymer microfibers. 4

18 Null Hypothesis 2 (N 2 ) H 0 : A statistical difference does not exist in the mean contact angle measurements among untreated PLGA microfibers, PLGA +1% collagen(i) microfibers, PLGA +0.1% fibronectin microfibers, PLGA +1% PEG-graftedchitosan microfibers, and PLGA +10% PLGA-PEG-PLGA triblock copolymer microfibers. Null Hypothesis 3 (N 3 ) H 0 : A statistical difference does not exist in the mean yield strength among untreated PLGA microfibers, PLGA +1% collagen(i) microfibers, PLGA +0.1% fibronectin microfibers, PLGA +1% PEG-grafted-chitosan microfibers, and PLGA +10% PLGA-PEG-PLGA triblock copolymer microfibers. Null Hypothesis 4 (N 4 ) H 0 : A statistical difference does not exist in the mean ultimate failure among untreated PLGA microfibers, PLGA +1% collagen(i) microfibers, PLGA +0.1% fibronectin microfibers, PLGA +1% PEG-grafted-chitosan microfibers, and PLGA +10% PLGA-PEG-PLGA triblock copolymer microfibers. 5

19 Null Hypothesis 5 (N 5 ) H 0 : A statistical difference does not exist in the mean elastic modulus among untreated PLGA microfibers, PLGA +1% collagen(i) microfibers, PLGA +0.1% fibronectin microfibers, PLGA +1% PEG-grafted-chitosan microfibers, and PLGA +10% PLGA-PEG-PLGA triblock copolymer microfibers. Null Hypothesis 6 (N 6 ) H 0 : A statistical difference does not exist in the mean plastic modulus among untreated PLGA microfibers, PLGA +1% collagen(i) microfibers, PLGA +0.1% fibronectin microfibers, PLGA +1% PEG-grafted-chitosan microfibers, and PLGA +10% PLGA-PEG-PLGA triblock copolymer microfibers. 6

20 Specific Aim #3: Evaluate the biological characteristics, specifically the cellular adhesion and cellular preference, of the surface-modified fiber scaffolds. Rationale The biological characterization of cellular adhesion to surface-modified PLGA electrospun fibers is evaluated. Quantification of cellular attachment on straight fibers and intersecting fibers is examined in order to make conclusions about the efficacy of secondary polymers present on PLGA fiber surfaces. Collagen and fibronectin may promote cellular adhesion to PLGA fibers by utilizing the integrin binding cascade involved in specific adhesion. PEG-grafted-chitosan and PLGA-PEG-PLGA- triblock copolymer may inhibit cellular adhesion, because cellular binding proteins on the cell plasma membrane have a low affinity for the hydrophilic PEG functional groups. Additionally, cellular preference, also known as competitive affinity, for one fiber substrate over a differing set of fibers is investigated. Cellular substrate selectivity is of interest, and it is believed that cells migrate to fiber surfaces in their locale that offer enhanced surface characteristics for binding and cell-to-substrate stabilization. Null Hypotheses Null Hypothesis 7A (N 7A ) H 0 : A statistical difference does not exist among untreated PLGA microfibers, PLGA +1% collagen(i) microfibers, and PLGA +0.1% fibronectin microfibers with respect to increasing the quantity of mean cellular adhesion on electrospun fibers. 7

21 Null Hypothesis 7B (N 7B ) H 0 : A statistical difference does not exist among untreated PLGA microfibers, PLGA +1% PEG-grafted-chitosan microfibers, and PLGA +10% PLGA-PEG- PLGA triblock copolymer microfibers with respect to decreasing the quantity of mean cellular adhesion on electrospun fibers. Null Hypothesis 8 (N 8 ) H 0 : A statistical difference does not exist in the quantities of mean cellular adhesion among the competitive pairings of PLGA +1% collagen(i) versus PLGA +1% PEG-grafted-chitosan, PLGA +0.1% fibronectin versus PLGA +1% PEGgrafted -chitosan, PLGA +1% collagen(i) versus PLGA +10% PLGA-PEG-PLGA triblock copolymer, PLGA +0.1% fibronectin versus PLGA +10% PLGA-PEG- PLGA triblock copolymer. 8

22 CHAPTER II BACKGROUND 2.1 Polymers: PLGA, Copolymer Component of a Biodegradable Scaffold An ideal tissue engineered polymer scaffold is one that is mechanically and thermodynamically stable and has the ability to function biologically at an implant site [9]. In order to create a biocompatible, non-toxic, and biodegradable scaffold that facilitates cellular attachment and uninhibited tissue growth, a polymer should be chosen that has a track record for its biodegradation kinetics and biocompatibility. Croll et al. submit that, For many soft tissue applications, scaffolds must be large, very highly porous, and soft, yet have enough strength to resist the contractile forces generated by growing tissue [10]. In other soft tissue applications it may be beneficial to employ a polymer that has the ability to contract with the growing tissue without relinquishing the scaffold stability and integrity. For the reasons listed above, the FDA-approved copolymer poly(lactic-co-glycolic acid), also known as PLGA, is the primary biodegradable polymer chosen for this experiment. PLGA is the copolymer product of PLA, poly(lactic acid), and PGA, poly(glycolic acid), combined in a ring-opening polymerization reaction. PLGA is an amorphous polymer with degradation kinetics in the range of days, but as high as one year in some formulations [11-12]. PLGA is soluble in organic solvents and can be electrospun into nano- and microfibrous mats that resemble a randomly oriented, non-woven matrix 9

23 scaffold. PLGA is relatively hydrophobic as compared to the natural extracellular matrix (ECM), is unable to interact specifically with cells, and does not possess the necessary functional groups for the attachment of biologically active molecules [10]. Additionally, PLGA is a hydrophobic species because of the presence of glycolide subunits which contain elevated levels of methyl functional groups [10-11]. A primary advantage to using PLGA fibrous scaffolds is that they can be complexed with hydrophilic molecules to surface modify the microfiber, as well as to modify the internal morphology so that the microfiber can be used to carry a payload of either hydrophilic or hydrophobic drugs [1,11]. This flexibility is critical for incorporating a diverse array of drugs into the polymer microstructure. The degradation kinetics of the polymer microfiber matrix can be controlled by the modulation of the scaffold s morphology, porosity, and composition [1]. 2.2 Electrospinning With respect to tissue engineering, electrospinning is the process that is applied to fabricate matrix scaffolds composed of natural, synthetic, biodegradable, or nonbiodegradable polymers. Electrospinning is a process that uses an electric field to control the formation and deposition of polymers [5-6,14]. Electrospinning is a rapid, efficient, and cost-friendly process. The procedure of electrospinning begins by creating a polymer solution, which is the mixture of solid polymer with an organic solvent. The next step is to eject the polymer solution across a charged electrical field where a charge imbalance in the solution occurs. At a critical voltage the charge imbalance overcomes the surface 10

24 tension of the polymer solution and a polymer jet is formed at the ejection tip. The solvent in the solution evaporates as the polymer jet is drawn through the electric field toward the grounded target, which may be a spinning mandrel or a flat platform. The continuous flow of the solution creates an uninterrupted filament that collects as a nonwoven scaffold [5]. Adjustments of the solution concentration, voltage, nozzle-tocollection plate (gap) distance, ejection flow rate, and nozzle diameter, collectively will produce variant forms of polymer nanofibers and microfibers. For polymers such as PLGA, the resulting scaffold still may have organic solvent residues, and thus it is necessary to vacuum dry scaffolds that will come into contact with live cells and tissues. Vacuum drying is advised for 1-7 days dependent upon the fiber size, morphologic nature, and porosity [1,14]. 2.3 Hydrophobic, Hydrophilic, and Amphiphilic Interactions The technique of surface modification in this study is thermodynamically induced phase separation (TIPS). To understand the mechanism by which phase separation occurs in a polymer fiber requires defining the principles of hydrophobic, hydrophilic and amphiphilic interactions. Hydrophobic (water-fearing) molecules are typically nonpolar compounds such as fatty acids that have long hydrocarbon chains [15]. Hydrophilic (water-loving) compounds are polar (charged) molecules that possess a permanent dipole moment. A good example of a hydrophilic molecule is water. A hydrophobic interaction is best described as the association of nonpolar molecules in an aqueous solvent. Nonpolar compounds cannot engage in hydrogen bonding with water and will aggregate 11

25 together via van der Waals interactions when mixed in an aqueous solvent in order to minimize their exposure to the polar surroundings. An example of this is an oil and water mixture in which oil droplets rapidly form at the surface after the mixture is stirred. The oil molecules aggregate into droplets and separate into a layer, or a phase, that is thermodynamically favorable for the nonpolar molecules. Hydrophobic interactions are driven by increased entropy (disorder). David Chandler, in a discussion in Nature, describes this process by asserting that, When a hydrophobic group projects into an aqueous solvent, the water molecules become ordered in a cage around the hydrophobic group. These solvent molecules become disordered when the hydrophobic group withdraws from the surrounding solvent [16]. Hydrophilic interactions are different than hydrophobic interactions. Hydrophilic interactions describe the process by which hydrophilic (polar) molecules participate in hydrogen bonding in an aqueous environment [15]. The ability to form hydrogen bonds with water makes a hydrophilic compound soluble in the solvent. Molecules containing both hydrophobic (nonpolar) and hydrophilic (polar) regions are known as amphiphilic or amphipathic molecules [15]. Amphiphilic interactions are a combination of the models described above taking place in separate locales on one molecule. Common amphiphilic substances are soaps, detergents, cholesterols, and phospholipids. Proteins and block copolymers can be amphiphilic as well and are known to often have several hydrophobic and hydrophilic portions. Amphiphilic interactions occur typically at interfaces of polar and nonpolar solvents [15]. Introducing an amphiphilic molecule (amphiphile) into such an environment causes the amphiphile to orient itself in such a manner so that its polar region projects toward the hydrophilic 12

26 solvent and its nonpolar region projects toward the hydrophobic solvent. The result is the amphiphile creating an energetically balanced partition between the immiscible biphasic solvents [15]. Amphiphilic interactions form the foundation of the phase separation technique employed in this study. Once mixed with PLGA, proteins such as collagen and fibronectin, as well as synthetic polymers such as PEG-grafted-chitosan and PLGA-PEG- PLGA triblock copolymer are believed to phase separate and migrate to the interface [10-11,19]. The proteins and polymers could orient their hydrophobic segments inward toward the PLGA-rich core and extend their hydrophilic segments outward radially at the surface. Thus, a modified surface can be created that presents hydrophilic species to the surrounding environment. 2.4 Surface Modification of PLGA Electrospun Fibers The purpose of modifying a biomaterial surface is to redefine the interaction that the material has with the body while maintaining the bulk material characteristics. For instance, coating a hydrophobic polymer species with a hydrophilic protein may cause enhanced cellular interactions uncharacteristic of just the hydrophobic polymer alone. Surface biocompatibility of a biomaterial is directly associated with the chemical composition of the materials surface [17]. The chemical composition of a biomaterial surface will mediate the material s wettability and surface charge [17], as well as mediate the adsorptive characteristics of the associated proteins that regulate cellular adhesion and migration [18]. Administering a diverse array of proteins or variant polymers to the 13

27 microfiber surface can modify the surface composition of pure electrospun polymer fibers. Surface modification of PLGA microfibers is achievable by introducing secondary molecules that are incorporated onto the surface of the polymer fibers by techniques such as phase separation, plasma surface treatment, aminolysis, and etc [17]. Thermodynamically induced phase separation (TIPS) is the primary method for surface modifying PLGA in this application [10]. Phase separation of hydrophobic PLGA molecules occurs when amphiphilic molecules with highly hydrophilic regions, such as grafted poly(ethylene oxide) (PEG), are introduced during electrospinning fabrication [10,15,19]. The nonpolar, hydrophobic properties of the PLGA polymer is thought to create an energetically favorable situation for the migration of the amphiphilic species, i.e. PEG-grafted-chitosan, to the surface of the fiber as it is spun in order to avoid thermodynamic (charge) instability in the final solid fiber structure. In theory, the phase separation of the two biomaterials results in a material surface interface comprised of a decreased amount of hydrophobic methyl groups and an increased amount hydrophilic functional groups, such as the terminal carboxy groups of collagen or the hydrocarbon chains of PEG [10,19-21]. Furthermore, multiple hydrocarbon chains extending from PEG molecules on PLGA fiber surfaces are flexible and are known to decrease surface interactions with human cells by steric repulsion. The phenomenon is highly dependent on the chain length and saturation density of PEG [19]. Surface modification of PLGA microfibers allows for the opportunity to create divergent biodegradable matrix scaffolds. An attractive characteristic of PLGA is that it can be easily surface-modified with the addition of natural molecules and synthetic 14

28 polymers [2,10,19,22]. For example, the additions of purified bovine collagen (type I), and purified bovine fibronectin, have been shown to enhance the degree of cellular attachment to surface modified PLGA nanofibers [2,5,22-23]. Molecules of collagen and fibronectin, both natural hydrophilic polypeptide molecules, are believed to favorably orient themselves thermodynamically at the surface of the hydrophobic PLGA fiber. The terminal carboxy functional groups present on collagen and fibronectin proteins have an affinity for binding serum proteins, which may allow for promoted cellular attachment at the material surface [2,5,10,19,22-23]. Although it is unknown exactly how these molecules conform on the fiber surface or in what orientation they reside, it is evident that their presence affects the wettability, hydrophilicity, and surface polarity of modified PLGA fiber surfaces [2,22-23]. On the contrary, poly(ethylene glycol) (PEG) grafted to the biomaterial chitosan (PEG-grafted-chitosan; PEG-g-CHN) can integrate onto the surface of PLGA fibers and presumably inhibit cellular attachment because of the hydrophilic properties of the PEG group [20-21,24]. The chemical composition of the synthetic, amphiphilic, PEG-graftedchitosan molecule is shown in Figure 1. PEG is a hydrophilic species that is considered a Stealth molecule within the body [10]. Mammalian proteins have general difficulty adsorbing to a PEG-saturated surface, because PEG molecules have saturated hydrocarbon side chains that manifest at the material surface and induce steric hindrance [10,19]. The polar steric hindrance created inhibits common serum proteins from binding to the material surface, which consequently minimizes cellular attachment. As stated previously, the thermodynamic instability and subsequent phase separation of PEG molecules within hydrophobic, PLGA-rich structures creates a surface with PEG 15

29 molecules [10,19-21,25-29]. Thus, the hydrocarbon functional groups of PEG molecules presumably increase the material surface s hydrophilicity. Chitosan, a hydrophobic molecule, is thermodynamically favored over PEG with respect to PLGA, and thus it is thought that chitosan becomes entrenched within the PLGA fiber. Figure 1. Chemical composition of PEG-grafted-chitosan [30] Similarly, the synthetically fabricated PLGA-PEG-PLGA triblock copolymer (referred to as PEG-PLGA block copolymer in this text) has amphiphilic properties much like that of PEG-g-chitosan [31-32]. It is believed that the block copolymer orients itself in much the same way as PEG-g-chitosan [25-29]. Specifically, thermodynamic instability of PEG molecules within a PLGA rich environment is the overriding cause for migration of PEG molecules to the fiber surface interface [26]. As is the case with PEGg-chitosan, PGA and PLA components of PEG-PLGA block copolymer are believed to orient their position in order to interact with the hydrophobic PLGA, while the hydrophilic PEG segment is thought to orient itself radially outward or outside of the PLGA-dense core. Again, the PEGylated surface may induce steric hindrance and consequently reduce cellular adhesion. PEG-PLGA block copolymer is also a candidate 16

30 for surface modification of PLGA microfibers in this experiment, and its molecular configuration is shown in Figure 2, below. Figure 2. Chemical composition of PLGA-PEG-PLGA triblock copolymer [33] Through the surface modification of PLGA microfibers, by way of collagen (type I), fibronectin, PEG-g-CHN, and PEG-PLGA block copolymer, the ability to create a simultaneous bimodal scaffold system is plausible. Surface modifying with collagen or fibronectin is intended to promote cellular adhesion, while the incorporation of PEG-g- CHN or PEG-PLGA block copolymer should theoretically inhibit adhesion. 2.5 Focal Adhesion Sites & Cellular Attachment Cells have a specific procedure by which they attach themselves to substrates such as surface-modified polymer microfibers. To understand the mode by which eukaryotic cells adhere to non-living surfaces, it is necessary to understand the molecular biology of cellular adhesion. Focal adhesion sites are cellular attachment points along a cell s 17

31 plasma membrane that extend across the plasma membrane [7-8]. They form through a cascade of intracellular linker proteins, known as integrins and adherens, and conjugate with cytoskeletal actin filaments. Focal adhesion sites facilitate cell-to-substrate binding, unlike cell-to-cell contacts, which are typically referred to as zonula adherens or adherens junctions. Focal adhesions also function as major sites for transduction of signals that control cell shape and movement [34]. When observed microscopically, focal adhesions between a cell and a substrate are typically 1-2 µm in length, 0.5 µm in width, and form a gap between the cell membrane and the substrate of approximately nm [8]. Integrin proteins, a class of heterodimeric transmembrane proteins and the primary adhesion receptor of extracellular matrix, have receptor sites on their extracellular domain that bind extracellular matrix-associated proteins, such as fibronectin [7]. Integrins are widely known as promiscuous receptors because of the binding sequence motifs that bind to a wide range of extracellular components. For example, the matrix ligand sequence motif arginine-glycine-aspartic acid (RGD) is common to 33% of all integrin family proteins [8]. Conversely, integrins are not the primary transmembrane proteins that assist in cell-to-cell binding. Cell-to-cell binding is carried out by cadherin proteins, which form the most simplistic protein links between neighboring cells [8]. 18

32 Figure 3. Depiction of a cellular focal adhesion binding through integrin proteins [35]. The intracellular domain of an integrin molecule conjugates with a set of intracellular proteins known as adherens, which include talin, paxillin, and vinculin [7]. The adheren proteins form an indirect junction complex of integrin proteins to α-actinin by their association with focal adhesion kinase (FAK). Subsequently, α-actinin binds to actin filaments of the cytoskeleton, and a focal adhesion site is formed [8]. In addition to the aforementioned components of focal adhesions, cytoplasmic adheren proteins filamin and F-actin are also known to preferentially associate with focal adhesion structures [36]. An example of the specific protein configuration of vinculin with respect to integrin binding at focal adhesion sites is depicted in Figure 4. 19

33 Figure 4. Protein configuration of a focal adhesion site (specific adhesion). [37] Focal adhesion sites are of paramount interest in this study, because they offer a glimpse into how a eukaryotic cell, a primary human fibroblast in this case, is interacting with surface-modified PLGA microfibers. Focal adhesion sites are considered the strongest of all cell-to-substrate interactions [17]. The adheren protein vinculin, a large cytoplasmic protein, plays an integral role in linking actin to junctional membrane proteins. Every focal adhesion site has associated vinculin proteins [8,15]. Thus, molecular probing for vinculin is one avenue by which we may garner an indication as to the cellular adhesion mode, locality, distribution, and quantity of focal adhesion sites present in a cell s plasma membrane along a substrate. Vinculin is a 117 kd (1066 amino acids) adheren protein that conjugates with transmembrane integrin molecules to form focal adhesion contact points [7]. Vinculin 20

34 contains multiple binding domains for other cytoskeletal proteins and focal adhesion components as well as phosphorylation sites [34]. The 3D structure of the vinculin protein consists of a globular head domain fused to a proline-rich domain that connects an elongated tail domain. The head domain contains binding sites for adheren proteins talin and α-actinin as well as a self-binding site (vinculin) on the tail region for a folded configuration. The tail region also contains bindings sites for actin and paxillin. Histological immunofluorescent staining of cellular vinculin proteins is depicted in Figure 3. (a) (b) Figure 5. Fluorescent microscopic images of cellular (a) actin filaments stained by rhodamine phalloidin, and (b) associated vinculin proteins stained by antivinculin Ab. [38] 21

35 2.5.1 Specific Adhesion In order to understand the mode by which cells apply their internal machinery to create a focal adhesion site, it is necessary to understand how cell-to-substrate binding occurs. Cellular binding classifications by known biochemical pathways include: tight junctions, gap junctions, zonula adherens, desmosomes, hemidesmosomes, or focal adhesions [7]. Specific adhesion is a reference to cellular integrin binding (focal adhesion) to biological substrates that are known and homogenously pure. For instance, coating a culture disc with the extracellular matrix protein fibronectin determines a known surface and induces specific integrin binding mechanisms [17]. In this configuration, cells bind to the fibronectin substrate through a known biochemical pathway, focal adhesion integrin binding. Specifically, in connection with this experiment, the surface modification of PLGA fibers with collagen and fibrinogen forms protein binding domains that associate with integrin proteins on the plasma membrane of a cell, in this case a human fibroblast. The mode of integrin binding with respect to the surface modification of PLGA fibers in this case is termed specific adhesion because there is certainty as to the modifying component on the substrate surface. 22

36 2.5.2 Non-Specific Adhesion In contrast to specific adhesion, cell-substrate focal adhesion sites exist in which the cell utilizes integrin binding, but the composition of surface-adsorbed proteins of the substrate is heterogeneous and consequently uncertain. This mode of cellular adhesion is termed non-specific adhesion [17]. Here integrin binding is employed, but the proteinspecific composition of the substrate is not specific to a single homogenous protein. Nonspecific adhesion is a common theme in the use of biomaterials within the body, because when biomaterials are implanted, they become coated with serum proteins. Serum proteins vary widely in their phenotype and terms of their ability to induce integrin binding. Thus, non-specific adhesion is the only justifiable classification that can be made for the mode of adhesion based upon the heterogeneity of the protein-adsorbed substrate surface. A common example of non-specific adhesion is with cell culture plates, which are coated with the extracellular matrix proteins fibrinogen, fibronectin, and vibronectin in order to take advantage of focal adhesion site binding. The mixed protein components lead to a cell-to-substrate binding arrangement that is varied and uncertain among the three proteins types. 23

37 CHAPTER III METHODS & MATERIALS A tissue-engineered biodegradable polymer matrix scaffold with variant surfacemodified characteristics was investigated for cell adhesion properties. These scaffolds were prepared by electrospinning PLGA microfibers with added bovine collagen type I, human fibronectin, PEG-grafted-chitosan, and PEG-PLGA block copolymer in order to surface modify the polymer fiber surface of the scaffold. The surface-modified microfiber matrix scaffolds were evaluated on their ability to either promote cellular adhesion through enhanced integrin binding or inhibit cellular adhesion through the addition of PEG molecules to the polymer microfiber surface. Pure, unmodified, electrospun PLGA microfibers represented the primary control for all experiments henceforth. Table 1 depicts the formulations of each polymer fiber that was fabricated in this investigation. Table 1. Polymer microfiber blends 1 o Polymer 1 o Solvent w/v 2 o Surface Mod. Mol. 2 o Solvent w/w PLGA (1.15 i.v.) DCM 10% n/a n/a n/a PLGA (1.15 i.v.) DCM 10% Bovine Collagen (I) HFIP 1% PLGA (1.15 i.v.) DCM 10% Human Fibronectin HFIP 0.1% PLGA (1.15 i.v.) DCM 10% PEG-g-Chitosan DMSO 1% PLGA (1.15 i.v.) DCM 10% PEG-PLGA BCP HFIP 10% 24

38 3.1 Electrospinning Electrospinning procedures were performed using 50/50 poly(dl-lactide-coglycolide) (PLGA), inherent viscosity: 1.15 dl/g in 30 o C, molecular weight: kda (Absorbable Polymers, Pelham, AL). The solid PLGA was dissolved in dichloromethane (DCM) or (VWR International, Batavia, IL) 1,1,1,3,3,3-hexafluoro-2- propanol (99+%) (HFIP, Sigma Chemical Company, St. Louis, MO) in a 10% w/v polymer solution. The solution was loaded in a 5-mL sterile luer-lock syringe (nonrubber plunger) (National Scientific, Rockwood, TN) and was ejected through an 18- gauge stainless steel surgical needle (BD Biosciences, Franklin Lakes, NJ). The solution was ejected at 0.19 ml/min by a steady flow syringe pump (Harvard Apparatus, Model 22/4, Holliston, MA). An electrical gradient was created by setting a high-voltage DC voltmeter to 20 kv DC (Del Electronics, Mount Vernon, NY) across a cm gap distance. Microfibers were spun and collected upon a grounded 15x15 cm 2 brass platform, covered with aluminum foil. Samples of microfibers collected on the foil were used for SEM imaging, contact angle measurements, and tensile testing. Additionally, ultra-low adhesion polymer culture plates (Corning 3262, Corning, NY) were used to fabricate 1.25 cm diameter sample discs. These sample discs were used to collect microfibers for cell culture experiments. The discs were placed on top of the grounded foil platform during the electrospinning process. The ultra-low adhesion coating of the Corning 3262 culture plate does not allow eukaryotic cellular adhesion, and creates a counter-current substrate for cellular adhesion to the local PLGA microfibers. The variant surface-modified PLGA microfiber solutions were mixed in the format as described above in Table 1. Acid soluble bovine collagen (I) was purchased from 25

39 Elastin Products Co. (No. C857, MW: 120 kda; MO, USA) while human fibronectin from plasma was purchased from Sigma-Aldrich (F2006, MW: 450 kda; St. Louis, MO, USA). PEG-grafted-chitosan was synthesized in our lab Dr. Yang H. Yun from methoxy poly(ethylene glycol) (mpeg-oh, MW 2000) and chitosan (CHN, degree of deacetylation 85%, medium viscosity) purchased directly from Sigma-Aldrich (St. Louis, MO, USA). The PEG-grafted-chitosan is in the form of a brush polymer containing one covalently linked PEG group for every chitosan molecule [24. PLGA- PEG-PLGA triblock copolymer (MW: kda) was synthesized in our lab by Dr. Parth Shah. The final electrospinning volume for each polymer solutions was 1.0 ml. Organic solvents such as dichloromethane (DCM) and hexafluoro- isopropanol (HFIP) (Sigma-Aldrich, USA) are toxic to live cells, and therefore the fiber scaffolds were vacuum dried at room temperature for hours to evaporate residual organic solvents. 3.2 Physical Characterization: Scanning Electron Microscopy Scanning electron microscopy was employed to qualitatively assess the morphology of electrospun PLGA microfibers (Hitachi S2150, Japan). An electrospun sample was deposited to a foil substrate, of which a 5 mm 2 sample was cut, and then mounted to a stainless steel slug. The experimental surface was sputter coated (Emitech K575x Turbo Sputter Coater, Ashford, KY) with silver/palladium. More specifically with respect to fiber morphology, assessments of size, shape, texture, smoothness, and continuity were evaluated. Images were taken at 2000x 26

40 magnification to visualize the fiber characteristics. Individual fiber diameter was determined using image software provided by Axiovision (Carl Zeiss Inc, Maple Grove, MN) that correlates the pixels to a scale measurement. Six random images containing a minimum of fibers/image were used to evaluate fiber diameter for each polymer blend. Mean fiber diameter was calculated for each polymer microfiber blend. 1% collagen in PLGA, 01% Fn in PLGA, 1% PEG-g-CHN in PLGA, and 10% PEG-PLGA block copolymer in PLGA were the experimental group, while PLGA alone was used as the control. The number of data points was variable among groups. Six random photographs were taken for each of 3 trials for a total of 18 pictures per group. However, each picture encompasses a varying number of fibers. Thus, a varying number of fibers within the pictures led to variation in the total data points for each experimental group. 3.3 Physical Characterization: Contact Angle The measurement of contact angle on a material surface offers a very convenient means to determine the relative hydrophilicity or hydrophobicity of a surface and is indicative of the associated surface energy. Evaluating the relative wettability amongst experimental surface-modified microfibers was conducted by contact angle studies. Surface wettability of the polymer microfiber surfaces in this experiment were evaluated using a Rame-Hart Goniometer (Akron University, Akron, Ohio). A dense electrospun mat of microfibers was spun onto a glass cover slip and used for testing [11]. The electrospun-coated cover slips were applied with a ~20 µl water droplet from a retractable pipette. Snapshots from a Hitachi model VK-C150 camera (Japan) captured 27

41 still images of the advancing and receding edges of the water droplet on the polymercoated surface. The advancing and receding angles were analyzed using a computer software tool, Image J (NIH, Bethesda, Maryland), which aids in still-photograph analysis. 3.4 Physical Characterization: Tensile Testing In order to investigate if the process of surface modification of PLGA microfibers affects the material characteristics of the experimental surface-modified scaffolds as compared to the control group, pure PLGA microfiber scaffolds, tensile testing was performed. The tensile testing was performed in order to determine values of the material characteristics including the yield, failure, modulus of elasticity, and toughness. ASTM standard D638-82a for tensile testing of plastics was utilized and the testing procedure was performed under the direction of Dr. Glen Njus, Department of Biomedical Engineering, at the University of Akron [39]. A Shore Western (SC-300C Servo Controller, CS-601 HPS Controller, Load Frame, Monrovia, CA) tensile testing apparatus was employed to investigate the material strength of each microfiber type. Values for the yield point, failure point, elastic modulus, and the energy absorbed, also known as toughness, were evaluated. Polymer scaffolds were fabricated as described above in Section 3.1; however, for tensile testing sample preparations, the final volume of each formulation was increased to 5 ml. Microfibers were electrospun into thick mats on non-stick aluminum foil. Microfiber mats were vacuum dried at room temperature for hours. The dense 28

42 mats were peeled away from the foil and cut, using a scalpel, into small rectangular samples (5mm x 20mm) for tensile testing. Photographs of each sample were taken in order to make fine measurements using Axiovision software, which was used to determine the sample dimensions. In order to determine the thickness, each sample was placed on its side under phase contrast microscopy, and still photographs were taken depicting the profile. Again, Axiovision software was used to determine the measurement of sample thickness by utilizing a scaling tool that measures pixels. The samples were loaded into plastic clamps used for tensile testing and secured in place vertically between the load cell (Lebow Model lb, Eaton Corp., Carol Stream, IL) and the ram of the Shore Western load frame. The 25lb load-cell was used to capture measurements of applied tensile force to the sample. Data was collected at a rate of 10 samples per second under displacement control. One thousand data points were collected for each trial. The strain rate of the experimental setup was 0.25-mm per second. Each sample was carried out to failure. The gain of the system was calculated to be 20,000. Experimental parameters that were measured included time, linear displacement (LVDT), and force. 3.5 Biological Characterization: Cell Culture Human dermal fibroblasts were procured from discarded foreskin samples of male newborns from Akron General Medical Center. Foreskins were enzymatically digested to isolate fibroblasts and graciously received from the laboratories at the Kenneth Calhoun Research Lab. Culturing of human dermal fibroblasts was performed by seeding 29

43 approximately one million fibroblasts into a Falcon culture dish ( Polystyrene Tissue Culture Disc, 100x20 mm, BD Biosciences, Franklin Lakes, NJ). Cells were suspended in 10 ml of human dermal fibroblast feeding media which includes: Dulbecco s Modified Feeding Media (DMEM), (Mediatech Inc, Derndon, VA), 10% Fetal Bovine Serum (Cambrex Bio Sciences, Walkersville, MD), 1% Antibiotic/ Antimycotic (Penicillin, Streptomycin, Amphotericin; Mediatech Inc, Derndon, VA), and 1% GlutaMAX L-glutamine 100x (Invitrogen, Carlsbad, CA). Fibroblasts were cultured in a CO 2 incubator (Napco Model 6300) at 37 o C, 5% CO 2. Passage of fibroblasts was performed using 0.25% Trypsin-0.53mM EDTA preparation. Sample discs with electrospun PLGA microfibers were consequently seeded with 2000 fibroblasts/cm 2, loaded in Falcon 12-well tissue culture plates (353043, BD Biosciences, Franklin Lakes, NJ). Seeded sample discs were incubated in 2 ml of fibroblast feeding media for 72 hours at 37 o C, 5% CO Biological Characterization: Fixation and Staining Protocols Fixation is the histological process by which tissues are preserved, and arrested from all biochemical processes, and provided mechanical stability and strength from decay [24]. Fixatives used for biological tissues are chemicals that preserve a biological material in a frozen state for storage and histological analysis. In this experiment setup many common organic solvents used for fixation also dissolved the polymers used to construct the matrix scaffolds or degraded the culture dishes. For that reason, methanol and acetone were ruled out immediately, because they degraded polystyrene culture 30

44 plates and/or PLGA [40]. Formaldehyde (Fisher Scientific, USA) was a consensus candidate for this histological approach in that it does not react adversely with the aforementioned polymers. A titration of formaldehyde was performed to determine a prudent concentration for fixation that corresponded with viable antibody staining. The fixative was titrated at concentrations of 0.125%, 0.25%, 0.5%, and 1.0% in PBS and applied to PLGA scaffold samples. The final determination for the fixation protocol was performed by immersing the samples for 10 minutes, at 4 o C, in 0.2% formaldehyde solution. Antibody staining, as described in the next section of this text, was performed and the results were evaluated under fluorescent light microscopy. Immunofluorescent studies were performed on the seeded microfiber sample discs. As noted previously, fixation of the samples was performed for 10 minutes at 4 o C using 0.2% formaldehyde solution (Fisher Scientific, USA). Samples were then permeabilized using 0.2% Triton X solution for 5 minutes at 4 o C. Sample discs were then washed/blocked 3 times for 10 minutes each with a 1% fetal bovine serum (FBS) solution. Next a 1:100 mouse anti-vinculin, clone V284 (Chemicon, Billerica, MA), monoclonal primary antibody solution (50 µl) was incubated at 4 o C overnight upon each sample disc. Next, samples were washed 3 times for 10 minutes each with 1% FBS solution. Samples were then incubated for 45 minutes at 4 o C with a 1:250 dilution of FITC-labeled sheep anti-mouse Immunoglobulin G (IgG) secondary antibody (Ab) F6257 (Sigma-Aldrich, St. Louis, MO). Also included in this secondary Ab solution were a 1:40 dilution of rhodamin phalloidin R415 (Molecular Probes, Eugene, OR), and a 1:500 dilution of Hoechst trihydrochloride (33342 Fluoro pure grade H21492; Molecular Probes, Eugene, OR). Finally, samples were washed 4 times in 1% FBS solution. 31

45 Mounting the fluorescent-labeled sample discs involved placing the disc fiber side down on a glass coverslip (Corning Cover Glass No.1, 24x40 mm) with a droplet of Fluoromount G (Southern Biothech, Bingham, AL) to reduce fluorchrome bleaching. The coverslips sealed using acrylic varnish and were mounted to glass microscope slides using double-sided tape. Finally, fluorescent microscopy studies were performed using a Zeiss Axiovert 200 microscope (Carl Zeiss Inc, Thornwood, NY). Objectives used with the Axiovert microscope include: 10x PNF, 20x Apo, and an oil-immersion 63x PNF. A 10x-magnified eyepiece objective was employed. The camera used to capture fluorescent images was a Zeiss AxioCam HRm (Carl Zeiss Inc) camera with a camera mount adapter (60 C 1 1.0x, Carl Zeiss Inc). 3.7 Biological Characterization: Immunofluorescence A fluorescent microscope was employed to visualize fluorescently labeled samples. Fixed and slide-mounted samples were viewed under phase contrast light, rhodamine red filtered (ex. 540 nm (green); em. 565 nm (red)), fluorescien iso-thiocyanate (FITC) filtered (ex. 488 nm (blue); em. 520 nm (green)), and Hoechst filtered light (ex. 352 nm (violet); em. 461 nm (blue)). A confocal microscope was employed to visualize layers of cell-seeded microfiber samples (Olympus Fluoroview). The laser used to probe the layers of the fluorescently labeled samples was an argon green laser (488nm). An objective of 40x with a 10x eyepiece objective was used. 32

46 3.8 Biological Characterization: Quantification of Cellular Adhesion on Fiber Scaffolds Primary human dermal fibroblasts were seeded to the polymer fiber scaffolds and examined among each experimental scaffold type. Cell culture of primary human dermal fibroblasts was performed by the protocol set forth in Section 3.5 of this report. Cells adhered to polymer microfiber segments were quantified by counting cell nuclei that were fluorescently stained using the nuclear dye Hoechst. The microfiber surface area composed within each replicate was determined by measuring the length of each fiber in an image magnified using a 20x-objective and phase contrast microscopy. Surface area was calculated by using the mean fiber diameter of each fiber type investigated and determined from Section 3.2. Cell number was normalized to the mean surface area of the microfibers from each image, and the values were pooled among replicates. Six images containing a minimum of 5-10 fibers were examined for each fiber blend to determine the magnitude of cellular density. Fixation, actin staining, and nuclear staining were performed by the protocols set forth in Section 3.6, above. Statistical significance of the experimental surface-modified polymers outlined in Table 1 were compared to the control group, PLGA fibers alone. 33

47 3.9 Biological Characterization: Quantification of Cellular Preference Between Two Surface-Modified Microfiber Matrices Interwoven in One Scaffold Cell preference for an individual surface-modified polymer scaffold over a differing surface-modified scaffold was investigated. PLGA surface-modified scaffolds were electrospun concurrently into one scaffold. By incorporating the crystal violet dye (Sigma-Aldrich, St. Louis, MO) at a w/v concentration of 0.10% 0.20% into one of the polymer solutions during scaffold fabrication, the ability to differentiate between polymer types under fluorescent microscopy was achieved. Crystal violet is hydrophobic and does not leach out of PLGA fibers. Collagen-modified PLGA microfibers were spun concurrently with PEG-g-CHN-modified PLGA microfibers, into one matrix scaffold. Crystal violet dye was added at a low concentration (~0.1% w/v) to one of the surfacemodified polymer solutions to distinguish the randomly oriented fibers from the nondyed matrix that was spun concurrently. Additionally, crystal violet was added to individual polymer solutions evenly among the six replicates. Swapping of the dye among the experimental groups was performed to protect against the possible interference of the cell-to-substrate binding caused by crystal violet. Conclusions of cellular adherence preferences were determined between blended polymer types in the same matrix scaffold. Cellular density was determined for each microfiber type within each pairing. Six images (10x-objective) both color and fluorescent containing a minimum of 5-10 fibers each were examined for each fiber pairing. Fixation, actin staining and nuclear staining were performed by the protocols set forth in Section 3.6, above. 34

48 Table 2. Competitive pairings of PLGA surface-modified blends Polymer 1 Polymer 2 PLGA+ 1% Collagen vs PLGA+ 1% PEG-g-CHN PLGA+ 1% Collagen vs PLGA+ 10% PEG-PLGA BCP PLGA+ 0.1% Fibronectin vs PLGA+ 1% PEG-g-CHN PLGA+ 0.1% Fibronectin vs PLGA+ 10% PEG-PLGA BCP 3.10 Qualitative Comparison of Vinculin Deposition on Surface-Modified Microfiber Scaffolds In order to understand the method of cellular attachment to surface-modified PLGA fibers, fluorescent imaging techniques were employed to examine vinculin protein staining. Vinculin protein staining was executed on all the samples of seeded PLGA matrix scaffolds. Six images (63x-objective) containing a minimum of 5-10 fibers each were examined for each fiber blend. Fixation, actin staining and nuclear staining were performed by the protocols set forth in Section 3.6, above. Comparison of the relative fluorescent intensity of vinculin staining among the experimental polymer blends was investigated qualitatively. Aggregation of vinculin deposits and their relative intensity with respect to the natural-modified polymer as compared to the PEG-modified polymers was the focus of the experiment. 35

49 3.11 Data Analysis & Statistics All experiments and data collected within Sections were completed with replicates of n=3. Data collected were first tested for normality by performing a Shapiro & Wilk test. If a normal distribution was found, a single factor ANOVA with a post-hoc test (Student-Newman-Kuehl) was employed in order to make determinations of statistical significance. For tensile testing, a single factor ANOVA and post-hoc test was performed; however, an additional statistic was performed. Dunnett s t-test was utilized in order to compare all experimental values to the control. A priori limits of significance were set to p=0.05. All experiments and data collected within Sections were completed with replicates of n=6. Specifically, data collected from experiments involved in Section 3.8 was analyzed in the same manner as described in the paragraph above (ANOVA, SNK, & Dunnett s). However, data collected from experiments in Section 3.9 was tested for normality by performing a Shapiro & Wilk test. Subsequently, in order to make determinations of statistical significance between competing polymer types, a Student s t- Test was utilized, because only two means were being compared. A priori limits of significance were set to p=

50 CHAPTER IV RESULTS 4.1 Fabrication of Surface-Modified PLGA Electrospun Fibers Bovine collagen (type I) and human fibronectin are each used in trial experimentations of cell adhesion studies to determine if the manufacturer specifications of 1% w/v (1mg) and 0.1% w/v (100ng), respectively, are suitable for fabrication procedures. PEG-grafted-chitosan is used at a concentration of 1% w/v in DMSO. The PEG-PLGA block copolymer is titrated in cellular adhesion studies to determine the ideal w/w concentration for surface modification addition to PLGA electrospun microfiber scaffolds. The block copolymer is titrated at concentrations of 1%, 2%, 5%, and 10% w/w Figure 6 depicts the titration s key findings. Figures 6a-c show cellular adhesion that is similar to control PLGA fiber scaffolds. However, the results displayed in Figure 6d shows decreased cellular adhesion along the fiber scaffold. The reduced cellular adherence when 10% block copolymer is incorporated in the microfiber indicates that PEG molecules may be present at the surface of the microfiber at a substantial saturation density to attenuate cellular attachment. The 10% w/v concentration was chosen as the ideal concentration for further testing. 37

51 (a) (b) Figure 6. Fluorescent images of actin filament staining of human fibroblasts seeded on PLGA microfibers + titrated w/w % PEG-PLGA block copolymer (a) 1% BCP, (b) 2% BCP, (c) 5% BCP, (d) 10% BCP mounted on culture plate cover slips (63x-obj). 38

52 (c) (d) Figure 6. Fluorescent images of actin filament staining of human fibroblasts seeded on PLGA microfibers + titrated w/w % PEG-PLGA block copolymer (a) 1% BCP, (b) 2% BCP, (c) 5% BCP, (d) 10% BCP mounted on culture plate cover slips (63x-obj). (continued) 39

53 4.2 Physical Characterization: Scanning Electron Microscopy PLGA fibers and the experimental surface-modified blends described in Table 2 are all successfully electrospun. Scanning electron microscopy (SEM) is employed to assess the size, shape, surface texture, and fiber morphology. The following images in Figure 7 show randomly oriented, micro- and nano-scale microfibers. Morphologically, the images exhibit fibers randomly oriented in a matrix scaffold that has both straight and curved segments. (a) (b) Figure 7. Electrospun surface-modified PLGA fibers: (a) PLGA alone, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer (300x-mag). 40

54 (c) (d) (e) Figure 7. Electrospun surface-modified PLGA fibers: (a) PLGA alone, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer (300x-mag). (continued) Upon review under higher magnification (Figure 8), fibers are cylindrical in shape, or at least semi-cylindrical. In general, the surface topography is macroscopically smooth, but high magnification reveals a textured appearance. Fiber diameters for pure PLGA fibers and the four experimental surface-modified groups are displayed in Table 3. Fiber diameters among all groups range from 0.3 µm 41

55 20.1 µm. Mean fiber diameter among all groups is 2.6 +/- 0.1 µm. An example of the fiber diameter measurements using Axiovision software is depicted in Figure 9, while graphics of the individual fiber distributions are shown in Figure 10. The p-value determined using a single factor ANOVA statistical test yields a value of p = Thus, the null hypothesis that states that surface modification of PLGA fibers does not alter the fiber diameter is not rejected based upon these findings. Although values of the mean fiber diameter display a difference of almost twice the fiber diameter size between the control group (mean fiber diameter = 3.6 µm) and the experimental group PLGA + 1% collagen (mean fiber diameter = 1.8 µm), a statistical significance is not present. Figure 8. SEM image of pure PLGA fibers showing surface texture (2000x-mag). 42

56 Table 3. Mean fiber diameters of PLGA polymer microfiber blends. Polymer Trials Pictures N Mean (µm) Std. Error Mean (µm) PLGA /- 0.4 PLGA +1% Collagen /- 0.5 PLGA +0.1% Fibronectin /- 0.5 PLGA +1% PEG-g-Chitosan /- 1.0 PLGA +10% PEG-PLGA BCP /- 0.5 Figure 9. Axiovision software image of fiber diameter measurements (1500x-mag). 43

57 Frequency Frequency Fiber Diameter Distribution of PLGA alone Diameter (microns) (a) Fiber Diameter Distribution of PLGA + 1% Collagen Diameter (microns) (b) Figure 10. Fiber diameter distribution of electrospun microfiber groups (a) pure PLGA, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEGg-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer.

58 Frequency Frequency Fiber Diameter Distribution of PLGA +0.1% Fibronectin Diameter (microns) (c) Fiber Diameter Distribution of PLGA + 1% PEG-g-CHN Diameter (microns) (d) Figure 10. Fiber diameter distribution of electrospun microfiber groups (a) pure PLGA, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEGg-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer. (continued) 45

59 Frequency Fiber Diameter Distribution of PLGA +10% PEG-PLGA BCP Diameter (microns) (e) Figure 10. Fiber diameter distribution of electrospun microfiber groups (a) pure PLGA, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEGg-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer. (continued) 46

60 4.3 Physical Characterization: Contact Angle Results of wettability and surface energy are evaluated by measuring the material surface contact angle. Rame-Hart Goniometer analysis of electrospun fibers results in the values found in Table 4. A marked decrease is seen in the contact angle among fibers treated with PEG-grafted-chitosan indicating a shift in the hydrophilicity of the biomaterial surface. The decrease shown indicates that polar functional groups could cause a steric effect at the water-surface interface. The reduced angular measurements can be attributed to the relaxation of a sessile water droplet on a more wettable surface. Advancing contact angle values determined in this experiment are higher than reported values from the literature [10,43]. Table 4. Means of the advancing and receding angles of contact angle measurements. Polymer Trials Advancing L (deg) θ A Receding L (deg) θ B Hysteresis (Adv - Rec) PLGA (+/- 2.80) (+/- 2.40) PLGA +1% Collagen (+/- 3.04) (+/- 3.04) PLGA +0.1% Fibronectin (+/- 1.68) (+/- 2.46) PLGA +1% PEG-g-Chitosan (+/- 3.43) (+/- 1.97) PLGA +10% PEG-PLGA BCP (+/- 3.39) (+/- 4.07) * Values in parentheses are standard error of the mean Single factor ANOVA statistics of the means of the advancing contact angles reveals significant differences among the surface-modified groups. SNK s post hoc test displays no significant difference between the control group, pure PLGA microfibers, and the surface-modified groups containing collagen and PEG-PLGA block copolymer. The 47

61 surface-modified groups containing fibronectin and PEG-grafted-chitosan are significantly different than the control (p < 0.05). However, within experimental groups collagen-modified and fibronectin-modified PLGA fibers were not significantly different than one another. PEG-g-CHN-modified and PEG-PLGA block copolymer-modified PLGA fibers are statistically different. Single factor ANOVA statistics of the means of the receding contact angles uncovers greater significant differences than the advancing contact angles (p < 0.05). SNK testing exposes significant differences of all experimental groups as compared to the PLGA control. Within the experimental groups, the only surface-modified groups that show similarity are collagen-modified and PEG-PLGA block copolymer-modified. All other variants within the experimental group show significant differences in receding contact angle. 4.4 Physical Characterization: Tensile Testing Mechanical testing of electrospun PLGA microfiber scaffolds reveal interesting material characteristics about the surface modification of PLGA scaffolds. Mechanical characteristics of electrospun fibrous membranes depend on a number of factors including the fiber structure, properties of the constituent polymers, and their interactions [44]. Table 5 displays the dimensions of the all the samples tested, and Table 6 shows the mode of failure of each sample. Figure 11 exhibits the Axiovision software images for measurement of (a) thickness and (b) the length/width, while Figure 12 displays timelapse photographs of the testing procedure. Table 7 lists the findings of the mechanical 48

62 testing. Finally, Figure 13 displays the stress versus strain curves for the control samples as well as each of the experimental samples, respectively. Table 5. Dimensions of PLGA surface-modified electrospun mats for tensile testing. Polymer Trial Width (mm) Length (mm) Cross Sectional Area (mm 2 ) PLGA Avg Thickness of Samples = 0.21 mm PLGA +1% Collagen Avg Thickness of Samples = 0.13 mm PLGA +0.1% Fibronectin Avg Thickness of Samples = 0.05 mm PLGA +1% PEG-g-Chitosan Avg Thickness of Samples = 0.12 mm PLGA +10% PEG-PLGA BCP Avg Thickness of Samples = 0.08 mm

63 (a) (b) Figure 11. Measurements using Axiovision software of (a) sample length/width and (b) sample thickness (20x-obj). 50

64 (a) (b) Figure 12. Examples of the tensile failure modes: (a) mid-substance, (b) grip. Table 6. Failure modes of PLGA surface-modified electrospun samples. Polymer Trial Ultimate Failure Strength (MPa) Strain at Failure (%) Mode of Failure PLGA 1 n/a n/a Mid-substance Mid-substance Mid-substance PLGA +1% Collagen Grip Mid-substance Mid-substance PLGA +0.1% Fibronectin Mid-substance Mid-substance Grip PLGA +1% PEG-g-Chitosan Grip Mid-substance Mid-substance PLGA +10% PEG-PLGA BCP Mid-substance Mid-substance Grip 51

65 Tensile Test: PLGA Alone PLGA 2 PLGA 3 PLGA 1 Stress ( MPa ) Strain ( % ) (a) Stress ( MPa ) Tensile Test: PLGA +1% Collagen Strain ( % ) (b) COL 2 COL 3 COL 1 Figure 13. Stress versus strain curves of each replicate for (a) pure PLGA, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer. 52

66 Stress ( MPa ) Tensile Test: PLGA +0.1% Fibronectin FIB 2 FIB 3 FIB Strain ( % ) (c) Stress ( MPa ) Tensile Test: PLGA +1% PEG-g-CHN Strain ( % ) (d) CHN 2 CHN 3 CHN 1 Figure 13. Stress versus strain curves of each replicate for (a) pure PLGA, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer. (continued) 53

67 Stress ( MPa ) Tensile Test: PLGA +10% BCP Strain ( % ) (e) BCP 2 BCP 3 BCP 1 Figure 13. Stress versus strain curves of each replicate for (a) pure PLGA, (b) PLGA+ 1% Collagen, (c) PLGA+ 0.1% Fibronectin, (d) PLGA+ 1% PEG-g-chitosan, and (d) PLGA+ 10% PEG-PLGA block copolymer. (continued) 54

68 Table 7. Material characteristics of surface-modified PLGA electrospun microfiber films PLGA PLGA +1% Collagen PLGA +0.1% Fibronectin PLGA +1% PEG-g-Chitosan PLGA +10% PEG-PLGA BCP Yield: Stress (MPa) (+/ ) (+/ ) (+/ ) (+/ ) (+/ ) Strain (%) 2.83 (+/- 1.02) Failure: 1.67 (+/- 0.22) 1.52 (+/- 0.04) 2.05 (+/- 0.28) 1.91 (+/- 0.03) Stress (MPa) (+/ ) (+/ ) 1.01 (+/- 0.36) (+/ ) (+/ ) Strain (%) (+/ ) Modulus: (+/- 5.20) (+/- 6.37) (+/- 6.82) (+/- 1.36) Elastic (MPa) (+/- 5.02) (+/ ) (+/ ) (+/ ) (+/- 8.27) Plastic (MPa) (+/ ) Toughness: (+/ ) (+/ ) (+/ ) (+/ ) Energy (J) (+/ ) (+/ ) * All table values are means of n=3 trials. ** Values in parentheses are standard deviations (+/ ) (+/ ) (+/- 77.0) The mean yield strength of PLGA surface-modified microfiber scaffolds ranges from MPa. Values of the mean yield strength increase with the addition of a secondary polymer; however, the percent strain decreases in all the experimental groups as compared to the control. Single factor ANOVA statistics of the mean yield strength of surface-modified PLGA microfibers is not significantly different than the control PLGA microfibers (p = 0.19). Dunnett s t-test, a post-hoc test comparing experimental means to a single control, in this case pure PLGA microfibers, returns no statistical difference in 55

69 each comparison. Additionally, SNK s test for differences among all means also finds no significant differences among any means of yield strength. Thus, the null hypothesis is not rejected. The range of the mean ultimate failure, also known as the ultimate tensile strength, is from MPa. Failure occurs in the mid-substance or at the grip. Discrepancies in the stress versus strain curves are apparent among specific samples in Figure 13. For example, collagen sample #1, fibronectin sample #3, chitosan sample #1, and block copolymer sample #3 individually show marked differences as compared to the other respective samples within each group. When the aforementioned anomalies in the stress versus strain curves are correlated with the failure modes at the grip, instead of in the mid-substance (see Table 6), a direct relationship between the anomaly and the failure mode is uncovered. Based upon the results in Table 6 and 7, the mean ultimate failure is increased with the addition of secondary polymers. However, much like the yield strength, the mean ultimate failure of experimental surface-modified microfibers is not significantly different from the failure of the PLGA control (ANOVA: p = 0.20). Dunnett s t-test finds no difference in each mean ultimate failure value as compared to the control. Furthermore, SNK s test finds no differences among all means for failure. Therefore, the null hypothesis cannot be rejected in this case. Of further interest in this case are the values of percent strain at failure that markedly decrease in the experimental groups as compared to the control (62.70 %). Percent strain at failure decreases as much as 2-3 times in the blended experimental groups. The stress versus strain curve of each surface-modified PLGA microfiber group displays an elastic region, followed by a plastic region. Values of the mean elastic 56

70 modulus range from MPa. The elastic modulus is smallest in the control group (14.18 MPa) and increased as much as 2-4 fold in the blended experimental groups. Investigating the statistics of the elastic modulus uncovers very interesting material characteristic differences. Single factor ANOVA exposes statistical differences between surface-modified PLGA experimental groups compared to the mean (p < 0.05). Dunnett s t- Test finds statistical differences between fibronectin-modified and PEG- PLGA block copolymer-modified PLGA microfibers and the control PLGA microfibers (p < 0.05). The SNK test for differences among means of elastic modulus expands upon the statistical differences. The PLGA control is not significantly different than the grouping of collagen-modified and PEG-g-CHN-modified microfibers (p > 0.05). Secondly, the grouping of PEG-g-CHN-modified, collagen-modified and fibronectinmodified are not significantly different, but differ from the control and the PEG-PLGA block copolymer-modified microfibers as a group (p < 0.05). Finally, PEG-PLGA block copolymer-modified, fibronectin-modified, and collagen-modified microfibers are not significantly different as a group, but do differ from the control and PEG-g-CHNmodified microfibers as a group (p < 0.05). Thus, the null hypothesis that states that surface modification does not alter elastic modulus must be rejected. It can be concluded that surface modification does alter the elastic modulus of a select group of PLGA surface-modified microfiber scaffolds. Evaluation of mean plastic modulus is of particular importance because the parameter indicates the material s ability to withstand further load past the yield point, which could be an indicator of how the scaffold as a medical device performs in a failure situation in vivo. Values of the mean elastic modulus range from ( 1.01) MPa in this study. 57

71 Positive values for the parameter are seen for PLGA alone, collagen, fibronectin, and PEG-g-chitosan-modified groups, which indicates the ability to withstand further loading. However, the PEG-PLGA block copolymer group displays a negative value indicating that this modification type cannot bear an increased load past the yield point. The results of the tensile testing display significant differences for the mean plastic modulus. Single factor ANOVA statistics return a value of F < Dunnett s t-test shows significant differences between fibronectin-modified, PEG-g-chitosan-modified, and PEG-PLGA block copolymer-modified microfibers as compared to the PLGA control microfibers (p < 0.05). Collagen-modified is the lone experimental case that does not show statistical difference from the control (p > 0.05). SNK s test for differences among means does not uncover a difference between the control and collagen-modified microfibers. However, it does reveal that PEG-PLGA block copolymer-modified PLGA microfibers are significantly different than all other cases (p < 0.05). As a group, fibronectin-modified and PEG-g-chitosan-modified microfibers do not show a difference (p > 0.05), but are significantly different from the control (p < 0.05). Collagen-modified is similar to PEGg-CHN (p > 0.05), but does not share the similarity with the fibronectin-modified case (p < 0.05). Therefore, the null hypothesis, which states that surface modification of PLGA microfibers does not alter the mean plastic modulus, must be rejected. Analysis of the mean plastic modulus of PLGA microfiber scaffolds determines that surface modification does have a significant effect on the plastic modulus of a select group of surface-modified experimental blends. 58

72 4.5 Biological Characterization: Antibody Titrations Fibroblast samples seeded on German glass cover slips are tested in order to determine a suitable working concentration for repeated vinculin protein staining. Antivinculin primary Ab dilutions are titrated at concentrations of 1:100 and 1:1000 in 0.5% bovine serum albumin (BSA). Figure 14 displays the results of the titration of the primary Ab. Note that the anti-vinculin Ab is vibrant and robust in the 1:100 dilution (14a), but is washed out and muted in the 1:1000 dilution (14b). Results suggest that a 1:100 dilution of anti-vinculin is suitable for quality staining. Next, a similar experiment to the previous one is performed. Yet, rhodamine phalloidin (actin cytoskeleton; dilution 1:40) and Hoechst (nuclear; dilution 1:500) stains are combined with anti-vinculin (dilution 1:100). Figure 15 shows the co-localization of the filtered images. Note the focal adhesion sites stained green aggregated at the terminal ends of the cytoskeletal actin filaments stained red. As the cell forms pseudopods, focal adhesion sites anchor the pseudopod to the substrate. The results of this experiment are consistent with those presented by McKenzie et al. in Figure 5. 59

73 (a) (b) Figure 14. Fluorescent images of the titration of vinculin protein staining on human fibroblasts (a) 1:100, (b) 1:1000 dilution (63x & 20x-obj). 60

74 Figure 15. Fluorescent image of human fibroblasts on a glass substrate showing actin _filaments (red), nuclei (blue), and vinculin staining (green) (20x-obj). 4.6 Biological Characterization: Cellular Adhesion on Fiber Scaffolds The inspection of human fibroblast attachment to PLGA microfibers demonstrates similarities among the experimental surface-modified groups and the pure PLGA control microfibers. Single factor ANOVA statistics (p = 0.07) reveal no significant differences among cellular adhesion on microfibers. Table 8 reports the values of cellular density adjusted to the microfiber surface area. Figure 16 depicts an Axiovision software image depicting the measuring technique for quantifying surface area and to count cell nuclei attached to the fibers. 61

75 Collagen-modified microfibers show an elevated level of adjusted cellular density (2.06 x10 5 cell#/sa) that is double the value of the other experimental results, as well as the control. PEG-grafted-chitosan-modified microfibers display the least quantity of the adjusted cellular density (8.43 x10 4 cell#/sa). Results of adjusted cellular density of the fibronectin-modified and PEG-PLGA block copolymer-modified in comparison to the control suggest that these two surface modifying agents are not well suited to increase or decrease cellular adhesion on PLGA microfibers, respectively. Statistical results suggest that surface modification does not create differences among cell adhesion characteristics. However, separating the treatment groups into the natural polymers and the synthetic polymers and comparing each treatment is performed. Statistics reveal no significant differences (p > 0.05) using Dunnett s t-test. Therefore, the null hypothesis that states that cellular adhesion is not affected by surface modification of PLGA microfibers cannot be rejected. Cell adhesion is not significantly affected by surface modification of PLGA electrospun microfibers with a secondary polymer. 62

76 Table 8. Cellular adhesion of human fibroblasts on PLGA surface-modified microfibers PLGA PLGA +1% Collagen PLGA +0.1% Fibronectin PLGA +1% PEG-g- Chitosan PLGA +10% PEG-PLGA BCP Trials Pictures Fibers Counted Total Cells Counted Total Fiber Length Counted (µm) Mean Fiber Diameter (µm) Total Surface Area (cm 2 ) 4.76 x x x x x10-3 Total Cells 1.01 x x x x x10 5 Counted/ Total Surface Area (cm -2 ) SEM +/ x10 4 +/ x10 4 +/ x10 4 +/ x10 4 +/ x

77 Figure 16. Example of Axiovision software image of fiber measurement for surface area and cell density determination (20x-obj). 4.7 Biological Characterization: Cellular Adhesion Between Competitive Fiber Scaffolds When the natural polymer modifications are placed in competition with the synthetic PEG-based polymer modifications of PLGA microfibers, significant differences are not seen. Table 9 displays the statistical t values of each competitive pairing that are analyzed using Student s t-test, comparing two means. Figure 17 shows an example of the Axiovision software images used to sum the microfiber surface area and count adhered fibroblast cells. The results indicate that the pairing of collagen-modified and PEG-grafted-chitosan modified microfibers show elevated quantities of the adjusted cellular density as 64

78 compared to the other paired groups (9.05 x10 4 and 5.80 x10 4 cm -2 respectively). The difference between their means (3.25 x10 4 cm -2 ) is the greatest difference among all the competitive pairings. However, with respect to the collagen-chitosan pairing, the data shows an elevated standard error, and this result indicates that data distribution is wide ranging. The large standard error suggests that the resultant values of the adjusted cellular density may not be accurate. Student s t-test values all indicate p-values greater than 0.05 for all treatments. Statistical significance is not seen in this experiment. The null hypothesis that states that surface modification does not alter the competitive balance among natural and synthetically surface-modified PLGA microfiber groups cannot be rejected based upon the results in Table 9. Based upon the results in Table 9, values of the adjust-ed cellular density of fibronectin-modified microfibers display levels (5.58 x10 4 and 4.09 x10 4 cm -2 respectively) are comparable to the results for the synthetic PEG-modified groups (4.80 x10 4 and 5.38 x10 4 respectively). The results do not show an increase in cellular adherence for fibronectin in comparison to the PEG-modified groups, and actually show a decrease in cellular adherence as compared to the results in Section 4.6 (9.78 x10 4 cm -2 ). Thus, the results suggest that the in situ model developed for surface modification in this experiment does not significantly promote nor inhibit cellular adhesion between opposing surface modified scaffolds that are electrospun concurrently in one blended scaffold. The fibroblasts appear to not differentiate between opposing surface-modified microfibers. 65

79 Table 9. Statistical values of Student s t-test on competitive microfiber scaffolds Polymer Trials Mean Cell #/ S.A. (cm -2 ) PLGA +1% Collagen 9.05 x10 4 vs 6 (+/ x10 4 ) PLGA +1% PEG-g-Chitosan 5.80 x10 4 PLGA +1% Collagen vs PLGA +10% PEG-PLGA BCP PLGA +0.1% Fibronectin vs PLGA +1% PEG-g-Chitosan PLGA +0.1% Fibronectin vs PLGA +10% PEG-PLGA BCP * Values in parentheses are standard error of the mean (+/ x10 4 ) 3.71 x10 4 (+/ x10 4 ) 2.01 x10 4 (+/ x10 3 ) 5.58 x10 4 (+/ x10 4 ) 4.80 x10 4 (+/ x10 4 ) 4.09 x10 4 (+/ x10 4 ) 5.38 x10 4 (+/ x10 4 ) p Values Student s t-test

80 Figure 17. Example of Axiovision software image of fiber measurement from a sample of PLGA+ 1% Collagen + crystal violet dye (red stained microfibers) versus PLGA+ 1% PEG-g-chitosan (10x-obj). 4.8 Biological Characterization: Qualitative Assessment of Vinculin Deposition and Confocal Microscopy The magnitude of specific cellular adhesion, via focal adhesion site expression, versus non-specific cellular adhesion is achieved by the evaluation of fluorescent vinculin deposition by human fibroblasts adhered on PLGA surface-modified electrospun microfibers. Figure 18 displays what is believed to be focal adhesion site deposition on fibers of pure PLGA. The control samples are viewed under a confocal microscope to 67

81 view the three-dimensional deposition of the vinculin staining. Figure 19 illustrates a cascade of two sets of 5 images each, taken while probing among layered depths. The confocal laser is trimmed across a total of 5 µm, up to 2 µm above and below the centerline in 1µm increments. Vinculin deposition is apparent in the each of the cascading sets as bright green fluorescent deposits along the fiber surfaces. It is noted that vinculin staining intensity increases and then again decreases as the confocal microscope probes into and then out of the layered microfiber sections (indicated by arrows). The investigation of vinculin protein staining is further developed in the experimental surface-modified groups. However, the ability to reproduce viable results was fruitless in as many as twelve trials. Positive control samples of vinculin staining on glass slides show quality fluorescent staining in every trial. An example of the control is represented in Figure 20. Despite the quality of the positive control samples, fluorescent images of vinculin staining in experimental samples show non-distinctive, ubiquitous staining, as seen in the examples of Figure 21. Focal adhesion sites are not seen in the images indicated by a lack of aggregates of green staining seen localizing along fiber surfaces as witnessed in the preliminary trials (see Figure 18). Based upon these findings, confocal microscopy was not performed on the experimental samples. With the lack of any focal adhesion staining in the experimental groups, the mode by which fibroblasts adhere to PLGA surfaces cannot be justified as integrin-associated binding. 68

82 (a) Figure 18. Fluorescent images of vinculin staining of human dermal fibroblasts seeded on PLGA microfibers mounted on culture plate cover slips (63x mag obj). Arrows indicate aggregations of focal adhesion sites on fiber surfaces. 69 (b)

83 (1a) (1b) (2a) (2b) (3a) (3b) Figure 19. Layered confocal image cascades of vinculin staining of human dermal fibroblasts seeded on PLGA microfibers mounted on culture plate cover slips (a) trial 1, (b) trial 2 (40x-obj). Arrows indicate focal adhesion deposition. 70

84 (4a) (4b) (5a) (5b) Figure 19. Layered confocal image cascades of vinculin staining of human dermal fibroblasts seeded on PLGA microfibers mounted on culture plate cover slips (a) trial 1, (b) trial 2 (40x-obj). Arrows indicate focal adhesion deposition. (continued) 71

85 Figure 20. Positive control sample of vinculin protein staining (green) of human fibroblasts mounted on German glass cover slips (20x-obj). (a) Figure 21. Fluorescent images of surface-modified PLGA microfiber samples probed for vinculin staining (63x-obj); (a & b) PLGA+ 1% PEG-g-chitosan, (c) PLGA+ 1% collagen. 72

86 (b) (c) Figure 21. Fluorescent images of surface-modified PLGA microfiber samples probed for vinculin staining (63x-obj); (a & b) PLGA+ 1% PEG-g-chitosan, (c) PLGA+ 1% collagen. (continued) 73