Nanoscalar Modifications to Polymeric Tissue Engineering Scaffolds: Effect on Cellular Behavior

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NANOSCALAR MODIFICATIONS TO POLYMERIC TISSUE ENGINEERING SCAFFOLDS: EFFECT ON CELLULAR BEHAVIOR

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the

Graduate School of

The Ohio State University

Heather M. Powell, M.S.

*****

The Ohio State University 2004

Dissertation Committee:

Professor John J. Lannutti, Adviser Approved by

Professor Douglas A. Kniss

Professor Derek Hansford

Professor David Rigney ______Adviser Professor William Brantley Graduate Program in Materials Science and Engineering ABSTRACT

Polymeric scaffolds provide a surface that can facilitate cell growth and tissue morphogenesis. Of particular interest is the role of nanoscalar features on cell behavior.

Nanoscale topographies can be generated on two-dimensional polymeric substrates via reactive ion etching. The magnitude and morphology of the resultant surfaces can be tailored by varying the gas media, etching time and power used. Nanofibrillar surfaces were produced on polyethylene terephthalate films via oxygen-plasma etching. These nanofibrils were dimensionally similar to collagen fibers. Cells cultured on nanofibrillar surfaces were shown to have a disrupted cytoskeleton, lower levels of cell-substrate signaling, reduced strength of adhesion and an inhibition of lipid droplet coalescence.

The results suggest that cells can detect nanoscalar surface topographies and alter their function in response to these environmental stimuli.

While nanofibrillar surfaces can be considered pseudo-three dimensional, they cannot produce 3-D cell structures. Thus truly three dimensional scaffolds must be fabricated to determine the role of nanoscalar fibers on cell organization and function.

Electrospinning was employed to generate 3-D meshes of polycaprolactone, a common biodegradable polymer. These nonwoven meshes were comprised of 500 nm fibers with an average pore size of 5 µm. In addition to forming mats of nonwoven fibers,

ii electrospinning technology can also produce tubular scaffolds. These tubular scaffolds were seeded with human vascular smooth muscle cells and cultured for two days. After 2 days in culture, cells assumed a helical orientation around the lumen of the tube, an architecture which closely mimics natural blood vessels. Thus electrospun scaffolds facilitate the growth and organization of cell populations in a manner which imitates the natural tissue.

iii

Dedicated to Jon, Bill and Sharon for all their support.

iv ACKNOWLEDGMENTS

I wish to thank my adviser, Dr. John Lannutti and co-adviser, Dr. Douglas Kniss, for their advice on my research. Dr. Lannutti has been indispensable during the course of my study, providing the fundamental engineering and polymer principles. Dr. Douglas

Kniss has provided an environment in which I have had the opportunity to increase my knowledge on cell biology immensely. The combination of their expertise has allowed me to research tissue engineering problems from initial scaffold fabrication to final cell seeded product.

I also greatly acknowledge the members of the Perinatal Lab Group in Dr. Kniss’ lab for their technical support throughout the years. I specifically would like to thank Dr.

Yubing Xie for sharing her knowledge of cell culture, Dr. William Ackerman for discussions on immunocytochemistry technique and lipid bodies, and Taryn Summerfield for help with the initial reactive ion etching. I am very grateful to have had to chance to work with all the members of the Lannutti group and the Perinatal Lab.

Financial support from the National Defense Science and Engineering Graduate

Fellowship program and the University Fellowship are acknowledged.

v VITA

August 23, 1978………………………….Born in Warren, Ohio

August 1999……………………………...B.S. Geology/Paleobiology Emphasis Bowling Green State University

August 2000-present……………………. Graduate Research Associate The Ohio State University

September 2000 – September 2001……...University Fellow The Ohio State University

September 2001-present…………………National Defense Science and Engineering Graduate Fellow (NDSEG) The Ohio State University

January 2003……………………………..M.S. Materials Science and Engineering The Ohio State University

PUBLICATIONS

Research Publications

1. Kohm, A., H. Powell, I. Wood, J. Dyce, and J. Lannutti “Apparent Skid Damage Controls Implantation Time” Journal of Biomedical Materials Research: Applied Biomaterials, 2004.

2. Wang, H., JK Lee, A. Moursi, D. Anderson, P. Winnard, H. Powell, and J. Lannutti. “Microstructural disassembly of calcium phosphates” Journal of Biomedical Materials Research 2004; 68A(1): 61-70.

3. Powell, H. and J. Lannutti “Nanofibrillar Surfaces Via Reactive Ion Etching” Langmuir 2003, 19: 9071-78.

4. Wood, I., H. Powell, A. Kohm, J.J. Lannutti and J. Dyce. “Evaluation of femoral head damage during canine total hip replacement: a comparison of four reduction techniques”. Veterinary Comparative Orthopaedics and Traumatology. 2003, 16: 184- 90.

vi 5. Xie, Y., T. Sproule, Y. Li, H. Powell, JJ Lannutti, DA Kniss “Nanoscale modifications of PET polymer surfaces via oxygen-plasma discharge yield minimal changes in attachment and growth of mammalian epithelial and mesenchymal cells in vitro” Journal of Biomedical Materials Research 2002, 61(2):234-245.

FIELD OF STUDY

Major Field: Materials Science and Engineering

Minor Field: Tissue Engineering

vii

TABLE OF CONTENTS

Page

Abstract…………………………………………………………………………………....ii

Dedication……………………………………………………………………………...... iv

Acknowledgements………………………………………………………………...... ……v

Vita………...……………………………………………………………………………..vi

List of Tables…………………………………………………………………………....xiii

List of Figures…………………………………………………………………………...xiv

Chapters:

1. LITERATURE REVIEW

1.1 Introduction...... 1 1.2 The body's natural scaffold……………………………………………………………3 1.3 Replacement Scaffolds: Material Selection...... 7 1.3.1 Degradable Polymers...... 10 1.3.2 Non-degradable Polymers...... 12 1.3.2 Non-degradable Polymers...... 13 1.3.2 Non-degradable Polymers...... 14 1.4 Replacement Scaffolds: Scaffold Processing...... 15 1.4.2 Solvent Casting and Particulate Leaching ...... 19 1.4.3 Gas Foaming ...... 21 1.4.4 Membrane Lamination...... 22 1.4.5 Melt Molding ...... 22 1.5 Modification of Scaffolds ...... 24 1.5 1 Surface Topography...... 25 1.5.2 Drug Delivery ...... 34 1.6 The Future of Tissue Engineering Scaffolds ...... 38 References...... 40

viii 2. CELLULAR RESPONSE TO NANOSCALE TOPOGRAPHY; EFFECT OF 20 NM HIGH PROTRUSIONS ...... 50

2.1 Abstract...... 50 2.2 Introduction...... 51 2.3 Materials and Methods...... 53 2.3.1 Polymer discs...... 53 2.3.2 Oxygen plasma treatment ...... 54 2.3.3 Scanning electron microscopy (SEM) ...... 54 2.2.4 Atomic force microscopy (AFM) ...... 55 2.3.5 Culture and medium...... 55 2.3.6 Cell culture experiments ...... 56 2.3.7 Cell number and hormone secretion ...... 56 2.3.8 Cell morphology ...... 57 2.3.9 Immunocytochemistry ...... 57 2.3.10 3T3-L1 cell differentiation...... 58 2.3.11 Cell apoptosis...... 58 2.3.12 Statistical analysis...... 59 2.4 Results...... 59 2.4.1 Surface Characteristics...... 59 2.4.2 Cell Morphology...... 60 2.4.3 Cytoskeletal Protein Expression ...... 64 2.4.4 Cell Proliferation...... 66 2.4.5. Hormone Secretion ...... 67 2.4.6 Cell Differentiation ...... 67 2.4.7 Cell Apoptosis...... 70 2.5 Discussion...... 70 2.6 Conclusions...... 75

References...... 77

3. NANOFIBRILLAR SURFACES VIA REACTIVE ION ETCHING...... 81

3.1 Abstract...... 81 3.2 Introduction...... 82 3.3 Materials and Methods...... 84 3.3.1 Polymer Films...... 84 3.3.2 Reactive Ion Etching...... 84 3.3.3 Scanning Electron Microscopy (SEM) ...... 85 3.3.4 Atomic Force Microscopy ...... 85

ix 3.4 Results...... 87 3.4.1 Scanning Electron Microscopy...... 87 3.4.2. Atomic Force Microscopy ...... 89 3.5 Discussion...... 93 3.5 1 Nanofibril Formation by Dewetting...... 94 3.5.2 Thermal Effects and Surface Instability ...... 98 3.5.3 Effects of Prior Surface Deformation ...... 101 3.6 Conclusions...... 103

References...... 105

4. TOPOGRAPHICAL CONTROL OF CYTOSKELETAL ORGANIZATION...... 109

4.1 Introduction...... 109 4.2 Materials and Methods...... 111 4.2.1 Cell culture...... 111 4.2.2 Surface Modification via Reactive Ion Etching...... 111 4.2.3 Scanning Electron Microscopy (SEM) ...... 112 4.2.4 Atomic Force Microscopy ...... 112 4.2.5 Cell Proliferation and Morphology...... 112 4.2.6 Immunocytochemistry ...... 113 4.2.7 Western Blotting ...... 114 4.2.8 Adhesion Studies ...... 115 4.2.9 Statistical analysis...... 117 4.2.10 X-ray Photoelectron Spectroscopy (XPS) ...... 117 4.3 Results...... 117 4.3.1 Oxygen plasma etching generates nanofibrillar surfaces ...... 117 4.3.2 Proliferation ...... 118 4.3.3 Nanofeatures alter cell morphology and the actin cytoskeleton ...... 122 4.3.4 Substrate nanofibrils alter FAK expression and activation...... 127 4.3.5 Substrate etching alters relative strength of adhesion...... 129 4.4 Discussion...... 131 4.5 Conclusions...... 139

References...... 140

5. NANOTOPOGRAPHY ALTERS PREADIPOCYTE DIFFERENTIATION...... 144

5.1 Introduction...... 144 5.2 Materials and Methods...... 146 5.2.1 Cell Culture and Differentiation ...... 146 x 5.2.2 Surface Modification via Reactive Ion Etching...... 147 5.2.3 Immunocytochemistry ...... 147 5.2.4 Image Analysis...... 149 5.2.5 Western Blotting ...... 149 5.2.6 Cyclic AMP (cAMP) Production...... 150 5.2.7. Statistical analysis...... 151 5.3 Results...... 151 5.3.1 Cytoskeletal Organization...... 151 5.3.2 Lipid droplet formation and coalescence...... 156 5.3.3 PPAR-γ Expression...... 159 5.3.4 cAMP Production...... 159 5.3.6 .Protein Coatings on Lipid Bodies...... 160 5.4 Discussion...... 160 5.5 Conclusions...... 164

References...... 166

6. ELECTROSPINNING REVIEW ...... 168

6.1 Introduction...... 168 6.1.1 Mimicking the ECM ...... 168 6.2 Electrospinning ...... 171 6.2.1 Theory of Electrospinning ...... 171 6.2.2 Variability in Fiber Diameter...... 176 6.2.3 Materials for Electrospinning ...... 180

References...... 191

7. FABRICATION AND CHARACTERIZATION OF 3D POLYCAPROLACTONE ELECTROSPUN SCAFFOLDS...... 194

7.1 Introduction...... 194 7.2 Materials and Methods...... 195 7.2.1 PCL solutions...... 195 7.3 Results and Discussion ...... 198 7.3.1 Scaffold Architecture...... 198 7.3.2 Scaffold Degradation ...... 203 7.4 Conclusions...... 206

References...... 207

8. ELECTROSPUN TUBULAR SCAFFOLDS FOR VASCULAR TISSUE ENGINEERING ...... 208

8.1 Introduction...... 208 8.2 Materials and Methods...... 211 8.2.1 Tubular Scaffolds...... 211 8.2.2 Cell culture experiments ...... 211 8.2.3 Cellular morphology ...... 214 8.2.4 Immunocytochemistry ...... 214 8.3 Results...... 215 8.4 Discussion...... 221

References...... 223

9. CONCLUSIONS AND RECCOMENDATIONS……………………………………..227

9.1 Effect of 20 nm-high protrusion ...... 227 9.2 Control of Surface Roughness with Reactive Ion Etching ...... 227 9.3 Disruption of Cytoskeletal Organization By Nanofibrillar Surfaces...... 228 9.4 Alteration of Adipogenic Differentiation by Nanotopographies ...... 228 9.5 Extending these Principles to Three-Dimensions...... 229 9.6 Recommendations...... 230

REFERENCES…....……..……………………..………………………………………..222

xii LIST OF TABLES

TABLE PAGE

1.1 Summary of surface roughness techniques………….…………………………. 33 2.1 Quantification of PET surface roughness……………………………………….62 4.1 Chemical analysius of untreated and treated PET substrates via XPS…………123 6.1 Comparison of mechanical properties of electrospun material to tissue……….183 7.1 PCL scaffold fabrication parameters…………………………………………...196

xiii LIST OF FIGURES

FIGURE PAGE

1.1 Fibroblasts in their natural scaffolding…………………………………………....2 1.2 Structure of collagen………………………………………………………………4 1.3 Model of cell adhesion to the extracellular matrix………………………………..5 1.4 Signal transduction from the extracellular matrix to the nucleus…………………8 1.5 Images of scaffolds constructed by various fabrication techniques……………….9 1.6 SEM of a hybrid synthetic and biopolymer scaffold…………………………….12 1.7 FDA approved tissue engineered skin replacement……………………………...13 1.8 Interaction of Biobrane with endigenous tissue………………………………….13 1.9 PLGA natrix degradation………………………………………………………...16 1.10 Fiber bonded PLGA scaffold…………………………………………………….18 1.11 Solvent cast-particulate leached PLGA scaffold………………………………...20 1.12 Tubular PLLA scaffold…………………………………………………………..23 1.13 Surface pillars and wells…………………………………………………………27 1.14 Compilation reflectance and fluorescence image ofr cells on rough surface……28 1.15 Surface topography via reactive ion etching………………………………….….30 1.16 Release kinetivs of DNA from microspheres……………………………………37 2.1 Surface topography of plasma etched PET coverslips……………………….…..61 2.2 SEM micrograph of cells grown on untreated and treated PET coverslips….…..63 2.3 Cytoskeletal expression of cells on PET disks……………………………….….65 2.4 Cell growth kinetics of 3T3-L1, JEG-3 and MCF-7 cells…………………….…68 2.5 Hormone production of JEG-3 cells on PET substrates…………………………69 2.6 Differentiation of preadipocytes on control and treated surfaces………………..71 2.7 TUNEL assay for apopotosis of cells on PET…………………………………...73 3.1 SEM of as-received PET film……………………………………………………86 3.2 High resolution SEM of PET plasma etched with various gases………………...88 3.3 SEM of surface of PET etched with oxygen-nitrogen mixture…………………..90 3.4 AFM of PET etched with Ar for 30 min…………………………………………91 3.5 Average roughness and maximum peak heights for all films……………………92 3.6 Schematic of morphological development of PET nanofibrils…………………..95 3.7 SEM of thermally bonded PET…………………………………………………..97 3.8 SEM of 1 min oxygen etched PET………………………………………………99 3.9 SEM iomages of mechanically deformed then etched PET…………………….102 4.1 Parallel plate device used for adhesion studies…………………………………116 4.2 SEM images of etched PET after exposure to water…………………………...119 4.3 Wet mode AFM of thermally bonded and etched PET…………………………120

xiv 4.4 MTS assay of 3T3-L1 cells grown on as-received and etched PET……………121 4.5 SEM micrographs of fibroblasts grown on smooth and nanofibril PET……….124 4.6 F-actin staining of 3T3-L1 cells………………………………………………...125 4.7 Confocal images showing FAK, phopho-FAK and F-actin……………………126 4.8 Densitometry of FAK and phosphor-FAK at Tyr 397 and 925………………...128 4.9 Relative strength of adhesion of cells…………………………………………..130 4.10 Model for integrin binding on smooth and nanofibrillar surfaces……………...137 4.11 Migratory ruffles seen in cells grown on nanofibrillar surfaces………………..138 5.1 SEM of thermally bonded and etched PET……………………………………..148 5.2 F-actin stain of cells on control and treated disks………………………………152 5.3 F-actin stain of differentiated cells……………………………………………..153 5.4 F-actin stain for 15-30 day differentiated cells…………………………………154 5.5 PPAR-γ and neutral lipid staining of differentiated cells………………………155 5.6 Distribution of lipid droplet size……………………………………………….157 5.7 cAMP expression in response to norepinephrine, isoproteranol and control….158 5.8 Western blot for ADRP………………………………………………………...161 6.1 Schematic for conventional fiber making processes…………………………...170 6.2 Generation of a polymer jet from an electrified solution……………………….172 6.3 Schematic of electrospinning apparatus………………………………………..173 6.4 Photograph of spraying phenomenon…………………………………………..175 6.5 Evolution of bending/whipping instability……………………………………..177 6.6 Fiber morphology of a function of solution viscosity………………………….178 6.7 Low viscosity solution spinning………………………………………………..179 6.8 Electrospun PEO……………………………………………………………….181 6.9 Electrospun collagen……………………………………………………………185 6.10 TEM of electrospun collagen………………………………………………...…186 6.11 Smooth muscle cells grown on PE-co-PVA……………………………………188 6.12 Smooth muscle cells grown on electrospun collagen…………………………..189 7.1 PCL scaffolds electrospun with varying solution concentrations………………197 7.2 Fiber diameter reduction with decreased solution concentration………………199 7.3 Beaded fibers…………………………………………………………………...200 7.4 Balance between flow rate and jet ejection…………………………………….201 7.5 Effect of reducing jet travel distance…………………………………………...202 7.6 Degradation of PCL: pH versus time…………………………………………..204 7.7 Morphologial signs of degradation……………………………………………..205 8.1 Electrospinning apparatus for tube construction……………………………….212 8.2 Macroscopic image of electrospun tubes……………………………………....213 8.3 Morphology of electrospun PCL tubes………………………………………...216 8.4 SEM of cells lining the lumen of the tube……………………………………...217

xv 8.5 Gallery of LSM images from lumen to mid thicknedd of scaffold……….……218 8.6 Confocal images from the mid-thickness to outer tube………………………..219 8.7 SEM of cells “migrating” through scaffold……………………………………220

xvi

CHAPTER 1

LITERATURE REVIEW

1.1 Introduction

Transplantation of organs such as the liver, kidney and heart has been a successful therapy for otherwise incurable diseases. Unfortunately, recipients far out number the available donors and many patients die while waiting for an organ. Thus alternate means of replacing or repairing diseased and damaged tissues are needed. The field of tissue engineering hopes to meet these clinical needs by producing functional three-dimensional tissues and organs from cells obtained either from the patient or from foreign tissue. In the most ideal case, tissue from the patent is broken down into individual cells and coaxed into forming tissue of the appropriate size and shape by using a physical scaffold

(Figure 1.1). The scaffold organizes the cells on a microscopic level and provides cues to stimulate growth, migration and differentiation. Tissue growth would then take place either in vivo or in vitro until a functional tissue is formed and can be transplanted into the patient. Although this process appears to be somewhat straightforward, complex fully functional organs have not yet been engineered. For example, skin replacements have been made which mimic the natural architecture of skin but cannot produce secondary differentiated structures such as sweat gland and hair follicles [1]. Consequently, tissue

Figure 1.1. Scanning electron micrograph of fibroblasts in connective tissue. The extracellular matrix surrounding these fibroblasts (red arrow) is comprised largely of collagen fibrils (yellow arrow) [2]

engineering still presents many challenges to materials scientists and biologists alike.

Materials scientists have the responsibility to create a scaffold, which can direct the arrangement of cells into the appropriate 3-D structure and provide the necessary molecular signals to make the cells form the desired tissue. In order to meet the goal of engineering organs and improving organ function further investigations into cell-material interactions are needed to determine what scaffold properties induce certain cellular behaviors.

1.2 The body’s natural scaffold

Tissues are not comprised solely of cells, they are complex structures made of cells, extracellular matrix and intracellular signaling systems. The extracellular matrix

(ECM) is an intricate network of proteins and polysaccharides that are secreted locally and organized into a meshwork by the cells that are in close association with it. The extracellular matrix provides the substrate upon which cells attach and determines the tissue’s physical properties (Figure 1.1)[2]. The relative amounts and types of matrix macromolecules and their organization gives rise to many forms of the matrix, each adapted to the functional requirements of the tissue.

The ECM is comprised of two main classes of macromolecules: 1) polysaccharide chains of the class called glycosaminoglycans (GAGs) and 2) fibrous proteins of two functional types; structural (ex. collagen) and adhesive (ex. fibronectin).

The polysaccharide chains are quite inflexible and tend to adopt a highly extended conformation which occupy a very large volume relative to their mass and form

Figure 1.2. Structure of a typical collagen molecule A) Single collagen alpha chain. Each amino acid is represented by a sphere. The molecules are arranged in a left handed helix with three amino acids per turn with a glycine at every third residue. B) A model of a triple alpha strand collagen fiber. [2]

Figure 1.3. A model showing how integrins (adhesive proteins) in the plasma membrane connect intercellular actin filaments (part of the cellular cytoskeleton) to the extracellular matrix. [3]

hydrogels at very low concentrations. Although the relative amount of polysaccharides is low compared to the amount of fibrous proteins, it occupies most of the extracellular space and provides the compressive mechanical support to the tissue.

In addition to the structural support the GAGs provide, the ECM contains a significant amount of fibrous proteins that either provide more structural support or play an adhesive role. The majority of fibrous proteins in the ECM are collagens (type I, II,

III, IV, and V). The relative amount and type of collagen is determined by tissue demands. For example, collagen type I is the main fibrous protein in the ECM of skin.

The rope-like structure of this collagen provides tensile strength to the tissue (Figure

1.2)[2]. Along with the structural fibrous proteins, adhesive proteins are also present in the

ECM. Adhesive proteins not only provide a structural framework upon which the cells can organize, they offer a mode in which the cells can interact with the matrix. The cells are ultimately linked to the ECM via these transmembrane proteins which tie the matrix the cells cytoskeleton. Integrins are a key adhesive proteins that function as transmembrane linkers mediating communication between the ECM and the cytoskeleton as well as functioning as signal transducers (Figure 1.3)[3]

The ECM was originally thought to play a relatively inert role in tissue function and development; it was just to stabilize the structure. The ECM has been shown to have a much more complex, active role in regulating the behavior of the cells in contact with it

[2]. Hausehka and Konigsberg demonstrated that collagen promoted conversion of myoblasts to myotubes and that collagen and glycosaminoglycans are crucial to salivary

6 gland morphogenesis [2]. ECM molecules interact with cell surface receptors which transmit signals across the cell membrane to molecules in the cytoplasm. A cascade of signaling events transfer the signal through the cytoskeleton into the cell nucleus. As a result the expression of specific genes is altered and various proteins are produced which can affect the ECM (Figure 1.4) [4-6]. Cells are bound to and migrate across the matrix by forming and breaking contacts with the surface via transmembrane proteins, integrins, and can receive mechanical signals from the ECM to proliferate and differentiate. (Figure

1.4). It is the extracellular matrix that provides the mechanical stability and signaling to create an environment in which cells can develop into functional tissues.

1.3 Replacement Scaffolds: Material Selection

Tissue engineering scaffolds are designed to be a surrogate ECM onto which cells can be seeded to grow and organize into tissue. The common theme of all tissue engineering scaffolds is that they are designed to provide a 3-D environment onto which the cells can attach and migrate. The scaffold should ultimately guide the cells into organizing themselves like the natural tissue and provide the cells with the necessary cues to proliferate and differentiate on the correct time frame. Material selection for scaffolds is extremely important. Scaffolds materials as well as their degradation products must be biocompatible, in that it does not elicit an inflammatory response or act as a toxin to the cells. Furthermore, scaffold materials should permit cell adhesion, promote cell growth, and allow retention of differentiated cell function. The material should be reproducibly processable and mechanically strong. A number of different materials have been used as scaffolds from bio-polymers like collagen to poly (lactide-co-glycolide) to

Figure 1.4. Schematic showing various cell signaling routes. The cell can receive physical/mechanical signals from the extracellular matrix and adhesion receptors. Soluble signals in the form of growth factors can also bind with their corresponding receptor to initiate complex signaling cascades within the cell to the nucleus ultimately resulting in alteration of cell behavior. [4]

Figure 1.5. Images of different types of scaffolds a) porous calcium phosphate scaffold [BD] b) porous poly (lactic acid) scaffold [13] c) woven poly (lactide- co-glycolide) scaffold [13]

9 hydroxyapatite and come in a range of architectures from fiber matrices to foamed structures (Figure 1.5) [7-11]. While metals and ceramics have been quite successful, in the medical field especially in orthopedics their processabilty is limited. As a result, polymeric materials have received the most attention in soft tissue engineering applications due to the ease of control over their processing.

1.3.1 Degradable Polymers

Many biocompatible materials can be used to construct scaffolds however a degradable one is preferable because the role of the tissue scaffold is only temporary.

Many degradable materials exist including natural bio-polymers which the body’s intrinsic mechanisms are able to degrade and clear the degradation products. Collagen is undoubtedly the most frequently utilized degradable bio-polymer for scaffolds. Collagen based matrices have been used for blood vessels and for tendon repair [12,8]. Urothelial cells have been grow on collagen scaffolds. The authors conclude that the tissue formed on these scaffolds is suitable for grafting [13]. Swope et al have produced skin equivalents by seeding collagen foams with dermal fibroblasts. Collagen scaffolds seeded with preadipocytes developed layers of adipose tissue upon implantation in immunodeficient mice, demonstrating that the pre-adipocyte cells differentiated within the scaffold. After 3 and 8 weeks in vivo new vessels were present supplying nutrients to the system [14]. Other biopolymers such as chitosan have been used as scaffolds. Chitosan, an antigenic natural polysaccharide whose structure is similar to glycosaminoglycans has been used in drug

10 delivery and in artificial skin [15]. Although these materials have been successful in some studies, their poor control of enzymatic degradation is a major drawback to using biopolymers as scaffold materials.

Synthetic degradable scaffolds were engineered to avoid the uncontrolled degradation commonly associated with biopolymer scaffolds. Polymers commonly used are polyesters, polyanhydrides, polyorthoesters, polycapralactone and polycarbonates[16].

Polyesters such as poly(glycolic acid) (PGA), poly(lactic acid) (PLA) and their copolymers poly [lactic-co-(glycolic acid)] (PLGA) are some of the most commonly used degradable polymers. PGA had its initial medical success as a totally degradable suture [17], however the extremely hydrophilic nature of the polymer allows for quick hydrolysis and complete degradation within 2 to 4 weeks [18]. Thus copolymerization with the more hydrophobic PLA resists water uptake and decreases degradation rates.

PLGA materials have been used extensively as scaffolds for engineering bone, cartilage and the meniscus.

While synthetic polymers provide tighter control of degradation rates and result in a scaffold with increased mechanical stability, they lack cell recognition and are commonly hydrophobic which detracts from efficient cell seeding. In contrast, biopolymer scaffolds have good cell recognition and are hydrophilic, but have poor mechanical strength. Thus creating hybrid synthetic-natural degradable scaffolds combines the advantageous properties of both. Dunn et al. prepared a collagen-PLA hybrid scaffold by dipping collagen fibers into a solution of PLA in chloroform and drying overnight [19]. Characterization of the resultant scaffold reveals that it has a tensile

Figure 1.6. Scanning electron micrograph of a hybrid synthetic collagen polymer made of a PLA woven mesh (white arrow) and collagen mesh (red arrow) [13]

12 Figure 1.7. Schematic figure of the components of Biobrane before implantations. [21]

Figure 1.8. Schematic of the interaction of Biobrane with existing dermal layers. The silicone outer membrane protects the wound from further infection from bacteria. The nylon mesh aids in the clotting of the wound to create good adherence of the dressing to the wound. Allowing for swift re-epithelialization of the wound. [21]

13 strength two times as great as the collagen alone [19]. Coating PLGA sponges with collagen has also been shown to increase its hydrophilicity facilitating faster cell adherence. Synthetic biodegradable polymer meshes have also been hybridized with collagen sponge to combine their advantages [20] (Figure 1.6).

1.3.2 Non-degradable Polymers

In the most ideal case, a scaffold would degrade on the same time scale that the cells can produce the natural ECM, leaving only native ECM and cells behind. As a result the immune response to the polymer would cease. Although many authors note that the breakdown products of PLA, PGA, and PLGA are water and CO2 and are well accepted by the body, one must take into consideration the complete degradation process whereby the polymer is first broken down into its repeat units: lactic and glycolic acid. The release of lactic and glycolic acid causes a local decrease in pH which many speculate creates a harsh environment for the cells. In addition, many degradable polymers used for tissue engineering scaffolds do not have a linear rate of degradation and often lose mechanical integrity long before natural ECM can be formed. Thus many non-degradable polymer scaffolds have been employed. For example a skin substitutes like Biobrane™, utilize a non-degradable nylon fabric embedded partially into a silicone film (Figure 1.7) [21]. The fabric provides an intricate 3-dimensional structure of threads to which collagen has been chemically bound. Blood and sera clot in the nylon fabric, which firmly attaches the dressing to the wound until re-epithelialization occurs (Figure 1.8). Other wound healing products such as Transcyte™ utilize ultrathin membranes attached to nylon mesh which provides a 3-D scaffold for growth of dermal tissue [22]. Fibroblasts residing in the mesh

14 proliferate and secrete structural proteins and growth factors that help to construct a 3-D human dermal matrix. Non-degradable materials have also been extensively utilized in research where properties of the surface such as surface chemistry, surface energy and surface topography are to be studied [23-46]. If the material were degrading while the experiment was taking place one could not rule out the possibility that the biological response was due to the instability of the surface not the specific surface properties. For example, Semler et al. investigated the change in surface character upon degradation and found that in porous PLGA scaffolds the surface morphology changes significantly while degrading (Figure 1.9) [47]. While non-degrading scaffolds are not as clinically ideal as degradable scaffold they provide a stable surface upon which cells can expand and can add additional strength to the tissue.

1.4 Replacement Scaffolds: Scaffold Processing

Not only must replacement scaffolds be constructed from materials which are biocompatible, the scaffold also must have an architecture that promotes tissue growth.

The scaffold must be porous and have high surface area to volume ratio. Pore size plays a critical role in tissue development; it allows for the infiltration of cells into the construct as well as provides internal surface area for cell attachment. The scaffold must have significant structural strength, which is most important in load-bearing replacements. The properties of the solid polymer and well as the processing have a profound effect on the scaffolds physical properties. All of these characteristics can be altered by the processing technique. Methods used to manufacture scaffolds are dependent on the polymer used and the application of the scaffold. For example a process which produces high

Figure 1.9. Reflective light micrograph of PLAGA matrix showing the effects of degradation of surface morphology. A,B,C, and D indicate observations take at 0, 48, 144, and 240 hrs of incubation in phosphate buffer solution (pH 5). Scale bar is 5 microns. [47]

16 compressive strength would not be needed in a non-load bearing scaffold and a scaffold that would have biological molecules incorporated into its structure could not be processed at high temperatures. Many novel fabrication techniques have been created which produce scaffolds with certain characteristics, the most popular of which are discussed in the following sections.

1.4.1 Fibers and Fiber Bonding

Fibers are an advantageous scaffold material because they have a very high surface area to volume ratio. Fibers are also commercially available and are already used in medicine as degradable suture materials. To create a scaffold the long fibers are made into felts or tassels [48]. Unfortunately researchers found that the non-woven fiber meshes and tassels are not dense enough to support large populations of cells and often do not possess the necessary structural stability [48].

To increase the stability of the fiber meshes a process called fiber bonding was created [49]. The process uses two polymers, one of which was a non-woven mesh

(polymer A) the other is dissolved in a solvent (polymer B) which does not dissolve polymer A. The polymer solution is poured over the mesh and allowed to evaporate. The composite is then heated to the melting temperature of the polymer B. The fibers join at their cross points. Polymer B is required to prevent the collapse of the fibers which would occur if they were heated alone. The entire composite does not melt together due to the immiscibility of polymer A and B. The composite is then cooled and polymer B is dissolved away by the solvent. Fiber bonding produces a scaffold with structural

Figure 1.10. SEM image of a fiber bonded PLGA scaffold. [49]

18 integrity, high porosity and high surface area to volume ratio (Figure 1.10)[49].

Unfortunately, this technique does not lend itself to easy independent control of porosity and pore size and uses toxic organic solvents.

1.4.2 Solvent Casting and Particulate Leaching

Solvent casting and particulate leaching overcome some of the drawbacks of fibers bonding specifically it can produce constructs with specific porosity, pore size, surface are to volume ratio and crystallinity [50]. This process is widely used for PLLA and PLGA, and can be applied to any other polymer soluble in a solvent like chloroform or methylene chloride. Solvent casting and particulate leaching begins by adding a known quantity of sized salt into a polymer solvent (often chloroform) solution. This salt- polymer solution is then cast into a glass container. Because salt is insoluble in chloroform, chloroform evaporates and leaves behind a polymer-salt composite which is highly crystalline. The crystallinity can be altered by a final processing step. If the salt- polymer composite is then immersed in water to leach out the salt the resultant scaffold will be highly crystalline. If the salt-polymer composite is heated above the melting temperature of the polymer, the composite can be cooled at varying rates to produce different levels of crystallinity (slow cool high crystallinity fast cool low crystallinity).

The salt will then be leached out by soaking the composite in water. This creates a highly porous (up to 93%) scaffold with interconnected pores (Figure 1.11) [51]. The porosity can be altered by the amount of salt added to the solution. Pore size can be changed by using different sizes of salt, pores up to 500 µm have been created. Surface area-to-volume ratio is also dependent on the size and quantity of salt particles used. This technique can

Figure 1.11. SEM image of a solvent cast-particulate leached PLGA scaffold with 90% porosity using 300-500 um diameter sodium chloride particles. [51]

20 up to 2 mm thick. Consequently 3-D scaffolds can be constructed only if the produce highly tailorable scaffolds, however it can only produce membranes membranes are laminated. In addition the organic residues left behind can have toxic effects in vitro and can produce an inflammatory response in vivo.

1.4.3 Gas Foaming

To overcome the problems associated with the use of organic solvents a method, called gas foaming, was created which uses no organic solvents [52]. In gas foaming, polymer pellets are compression molded into disks and then exposed to CO2 at high pressure. The high pressure allows the CO2 to saturate the disks and upon rapid release of the pressure pores are nucleated in the polymer from the CO2 gas. Unfortunately, the pore structure formed is a closed pore structure therefore this process must be combined with a particulate leaching technique to produce a scaffold that can form a viable tissue which requires the influx of nutrients through pores to survive [53]. Salt particles and polymer pellets are mixed together and compressed into disks. The disks are exposed to the high pressure CO2 which is consequently released. The scaffold is then submerged in water leaching out the salt and leaving behind an interconnected porous scaffold. This process eliminates the use of organic solvents, but still adds additional ions to the body, and does not allow for scaffold of specific geometries to be created.

21 1.4.4 Membrane Lamination

The need for specific three–dimensional scaffold geometries for regeneration of tissue whose function is partially dependent on its shape has made membrane lamination a popular process for creating highly porous scaffold with anatomical shapes [54]. Porous membranes made by solvent casting and particulate leaching are bound together by applying chloroform to their contacting surfaces. The chloroform solubilizes the surface of the polymer, the surface chains intermingle leaving a bond between the two surfaces when the chloroform if evaporated. Membrane lamination is only useful if the lamination process does not alter the bulk and leaves no boundary between layers. This process can also be used to create tubular scaffolds. Mooney et al. wrapped solvent casted and particulate leached membranes around a Teflon™ tube and laminated its overlapping edged creating a tubular stent for intestinal applications (Figure 1.12) [55]. This process has been used with PLLA and PLGA, and the product shows no boundary layer.

Unfortunately, this process cannot be used with polymers that are only soluble in highly toxic organic solvents because the solvent residues harm the body.

1.4.5 Melt Molding

One disadvantage to membrane lamination is that it does not make smooth 3-D geometrical shapes, the scaffolds are fairly jagged in nature [54]. Melt molding allows very intricate shaped to be formed. To produce a scaffold via melt molding a polymer containing sieved gelatin microspheres is poured into a Teflon mold and raised to a temperature above the polymers glass transition temperature allowing the polymer to take the shape of the mold. The polymer is cooled, removed from the mold and soaked in

Figure 1.12. Tubular PLLA scaffold prepared by a solvent casting and particulate leaching technique. Scaffold is suitable for peripheral nerve regeneration.[55]

23 deionized water. Gelatin is soluble in water and leaches out leaving a porous scaffold identical to the mold. The porosity and pore size can be controlled by the size and quantity much like salt leaching. The benefits to this process are no toxic organic solvents are used and it can be done at a lower temperature depending on the polymer, which can allow for incorporation of biomolecules. However, the temperature required for this process increases as crystallinity increases up to a point where gelatin is no longer soluble in water. There are many additional scaffold processing techniques each with their own advantages and disadvantages. The key in scaffold processing is to balance the properties of the polymer and the scaffolds application with the processing technique.

1.5 Modification of Scaffolds

Although scaffolds can be constructed utilizing wide range of materials in various architectures, it is apparent that there is a missing link to the equation. Many scaffolds are unable to support viable cell population for tissue growth. Others are not capable of organizing cells into the necessary organization and all scaffolds currently lack the ability to provide all of the necessary environmental cues to evoke the wide array of cellular processes required to produce fully functional tissues. Because of this, immense research has been conducted on modifying the surface of tissue engineering scaffolds [24-33, 37-39, 41-

44, 46-47, 57-71].

Interests in scaffold modification derive from the need to optimize substrates prior to expansion of cell populations in tissue engineering operations. For example, the length of time it would take to engineer a functional tissue or organ in vitro is much longer than desired if to be used in a clinical settings, thus increased proliferation and organization

24 could be beneficial in these applications. In disease therapy applications, an increase in expression of a given protein may be desired. Investigations of cell-substrate interactions have taken place to determine what surface properties can elicit specific cell responses in order to optimize the scaffold for its particular application.

1.5 1 Surface Topography

Surface topography has been extensively investigated and is identified as one of the classic factors which can control cell behavior on a surface. In the early 1930’s, the work of Weiss demonstrated that cells grown on an oriented substratum assume a corresponding orientation and migrate in that direction [72]. At that time the ability of various roughened surfaces to guide the cell was under debate due to discrepancy of results from different labs, thus a means of creating reproducible surface roughness was needed. Micro-machining, the process used to produce silicon microelectronic components, was one of the first techniques employed to produce surfaces with tightly controlled surface parameters with high reproducibility and is still used frequently today.

The majority of early surfaces topography studies investigated the effect of micromachined grooves 1 to 100 microns wide and 1 to 5 microns deep [25-26, 30, 59] in silicon or other materials cast into a micro-machined silicon mold. Although the materials used in these early studies are not commonly used as tissue scaffolds, they were easily machined and thus were a good model material for subsequent studies. The information gained from these studies can be extrapolated to more appropriate systems. Surface topography in the form of microgrooves was found to have the ability to orient cells in the direction

25 of the grooves, a process termed “contact guidance” [72]. For example, rat dermal fibroblasts cultured on silicone rubber cast from micro-machined silicon templates were found to be highly oriented on grooves that were less than 4 microns in width and randomly oriented on grooves wider than 4 microns with no effect of groove depth [30]. In contrast Wojciak-Stothard et al. found that macrophage initial orientation is dependent on groove depth with cells orientating faster on deeper grooves [69]. The cells spread faster on shallow grooves but elongate faster on deeper grooves. On titanium coated epoxy replicas of grooved photoresists, periodontal epithelial cells were directed more effectively on grooves deeper than 0.5 um [25-26]. Other authors have noted that fibroblasts extend their cellular processes into 1 micron deep grooves to produce intensive contact and mechanical interlocking with the surface to improve adhesion [43-44]. VonRecum and

VanKoten also noted that micron sized surface topography modified cell response in culture and in vivo [67]. They suggested that a surface with 1-3 micron sized grooves provides the best biocompatibility. Turner et al. have also utilized photolithography techniques to produce silicon surfaces with various geometries ranging from wells to circular and square shaped pillars (Figure 1.13) [65]. Their study indicated that astroglial cells prefer surfaces with pillar containing topographies to smooth surfaces (Figure 1.14).

While the ability of varying groove magnitudes and periodicities to elicit a specific cellular response varied from lab to lab and cell type to cell type, most researcher have

a b

c d

Figure 1.13. Scanning electron micrograph of different surface topographies. a) wells 1µm deep, 0.5µm wide and

0.5µm apart b) pillars 1µm tall, 0.5µm in diameter and 1µm apart c) surface with same size pillars 5µm spacing d) square plateaus 1µm tall 1µm apart. Scale bar in a,b, and c is 2µm. Scale bar in d is 20µm. [65]

Figure 1.14. Compilation reflectance and fluorescence microscopy images illustrating that astroglial cells prefer rough areas (light gray region) over smooth surfaces. Scale bar is 50 µm. [65]

28 come to the conclusion that cells behave differently on textured surfaces than smooth surfaces but have not come to an agreement on what scale of grooves are best for each type of cell.

It is very promising that cells can be guided by their substrate. This ability is extremely important in applications such as spinal cord injury and nerve regeneration [73-

74]. For example, to recover function in a spinal cord injury the implanted scaffold must guide the axons along the spinal cord [73]. While contact guidance is important in these applications, a large number of cell populations sit on a more random surface in the body

[73, 75].

Abrams et al. characterized the basement membrane, the substrate upon which most epithelial cells attach, as an apparently random, highly porous, three–dimensional structure; thus a more random topography may be ideal for applications such as skin regeneration and other types of organ replacement [75]. Several authors have investigated the effects of less regular surface topographies by using acid etching, sand-blasting and various polishing techniques. For example, chondrocytes grown on polished Ti have a reduction in alkaline phosphatase activity with increasing surface roughness (Ra 0.22 um to 4.24 um) [24]. In contrast, osteoblasts show decreased proliferation, DNA synthesis and increased alkaline phosphatase activity as surface roughness increased in titanium surfaces [41]. Sandblasted poly(methyl methacrylate) (PMMA) increased migration of cells 3 times for corneal cells on rough surfaces (peak to valley height 3.34 microns)

Figure 1.15 Rough surface created by reactive ion etching. Peaks are 230 nm tall and 57nm in diameter 65]

30 compared with smooth surfaces (peak to valley height 70 nm) [60]. Adhesion was also higher on rough cellular behavior however the disparities in production method and cell type do not surfaces. These studies demonstrate that random surface roughness does affect allow good comparisons from experiment to experiment.

In order to produce a random, reproducible surface topography many researchers turned to reactive ion etching (RIE) to create surface roughness in their materials.

Reactive ion etching has been used for many years in the plastic industry to roughen the surface prior to printing [76]. RIE provides a reproducible method to alter surface characteristics of a substrate without changing the bulk characteristics. Turner et al. reactive ion etched silicon wafers with Cl2, CF2, and O2 gasses to produce 57 nm diameter peaks that were 230 nm in height (Figure 1.15). Cell behavior on the RIE surfaces was compared to wet etched silicon surfaces (100-250 nm in width, 115 nm in height). Primary astrocytes preferred the RIE etched areas over the smooth areas and in contrast transformed astrocytes showed a preference for the smooth areas [65]. Xie et al investigated the effect of nanometer sized random surface topography by etching

[10] polyethylene terephthalate disks with a 25 % O2 -75% Ar gas mixture . JEG-3, 3T3-

L1, and MCF-7 cells has the same adhesion, proliferation and cytoskeleton expression on

[10] rough surface, Ra 16 nm, and smooth surfaces, Ra 4 nm . Random surface roughness has been shown to increase cell adhesion in most cases however long term cell behaviors such as apoptosis and differentiation (processes which are crucial to a developing tissue) have not been extensively studied on these surfaces.

31 Surface roughness does effect cellular behavior. The literature suggests that roughened surfaces in the dimensions of 1~50 µm showed better early adhesion of cells, increased osteoblastic function and functional gene expression, reduced fibrous encapsulation, and enhanced integration of subcutaneous implants, compared with smooth surface and grooved surfaces have the ability to orient cells along the groove [33,

68, 58, 67]. However a consensus on what geometry and magnitude of surface roughness is ideal for different cell types has not been determined (Table 1). One potential cause of this is the lack of quantified surface topographies. Within a subset of the literature, only qualitative descriptions of the surface such as, scanning electron and optical micrographs, are reported, therefore it is not possible to compare surfaces from study to study. In those that do provide quantitative surface roughness parameters, the total surface area examined is not large enough to be deemed representative of the entire surface. Secondly, in most investigations there is no range of roughness; only comparison between smooth and rough which can be orders of magnitude different. Thus a more systematic approach to surface roughness investigations would provide scientists with more definitive insight into the effect of surface roughness on cell behavior. In addition, many researchers only investigate initial effects such as cell adhesion, proliferation up to 3 weeks, orientation and cell shape.

Type of Surface Material Roughness Process Used Cell Type Biological Effect Ref Titanium parallel grooves Micro- gingival human fibroblasts oriented themselves with Brunette coated silicon, (3um deep with machining fibroblasts the orientation of the grooves, 1986 silicon and spacing of 4.9, fibroblasts on rough surfaces appear titanium 6.6, 8.3, and more round and have more filapodia coated 80um) than on flat surfaces photoresist

Silicone and grooves (depth Micro- in vivo soft thick fibrous capsule around the Parker et poly-lactic 1.0um, width machining tissue response implant but not significantly different al 2002 acid 10.0um) in goat model than smooth implant

Silicone grooves (depth Micro- rat dermal grooves of 2.0 and 5.0um spacing denBraber 0.5 um, spacing machined fibroblasts were able to orient cells better than a et al 1995 2.0, 5.0, 10.0 um) silicon wafers 10um spacing were used a mold for silicone Fused silica grooves (30- Micro- murine cells on grooves surfaces oriented Wojciak- 282nm deep, machining macrophages themselves along the grooves, had a Stothard spacing 25um) greater number of protrusions of the et al 1996 cell membrane, and an increase in the amount of F-actin when compared to cells cultured on smooth surfaces

Silicone square pillars or Micro- human cells on 2 and 5um pillars had Green et wells 2,5, or 10 machining abdomen increased proliferation and cell density al 1994 um in height or fibroblasts in comparison to 2 and 5 um wells. 10 depth and smooth um wells and pillars were not silicone statistically different in biological response than smooth surfaces

Polystyrene grooves (depth Micro- bone cells cells on 5.0um grooved substrates had Chesmel 0.5 or 5.0 um) machined isolated from significantly higher velocities than et al 1995 silicon wafers neonatal rat with smooth or 0.5um grooves were used a calvaria substrates, cell colonies were highly mold for oriented with the substrate whereas solvent cast isolated cells were found to ignore polystyrene surface topography Titanium grooves (vertival micromachining epithelial cells grooves directed cell locomotion, Brunette coated silicon, walled grooved with a however 0.5 um deep grooves were 1986 titanium and v-shaped evaporation less directive than deeper grooves, coated grooves 0.5-60um coating of Ti cells did not flatten well on grooved photoresist, deep) substrates and epon- araldite

Table 1.1. Summary of surface roughness modifications to cell growth substrates.

33 Studies looking at long-term cell cultures, differentiation, gene production and apoptosis would provide information on the cellular processes required to produce a functioning tissues.

1.5.2 Drug Delivery

Alternate methods to control cell behavior have been investigated, primarily the incorporation of drugs or growth factors into the scaffold for delivery upon degradation or via diffusion [55, 77-91]. In natural tissue generation, a complex and orchestrated delivery

of soluble signals to the cells occurs. Thus to optimize scaffold performance those soluble signals have been integrated into the polymer.

An early attempt at delivering growth factors via scaffolds used adsorbed basic fibroblast growth factor (bFGF) on Dacron. Greater than 40% of the bFGF was released within the first 24 hours and all of the adsorbed molecules were gone within two weeks

[87]. The bFGF did increase the proliferation and migration of the fibroblasts which illustrates that the growth factor retained its biological activity when adsorbed to the surface, however this method did not allow for a substantial amount of bFGF to be released from the scaffold. A double emulsion method has been used to incorporate a larger amount of growth factors into microspheres. The microspheres are then either formed into scaffolds or mixed into the scaffold material prior to forming. Nof and Shea used the double emulsion process to incorporate DNA into microspheres that were then pressed into a non-porous disks or gas-foamed into interconnected open pore scaffolds

34 [84]. The disks and scaffolds exhibited sustained plasmid release for 21 days and had a minimal initial release burst when compared with non-formed microspheres (Figure

16)[84].

Concerns regarding the harsh conditions of the double emulsion technique, namely the use of organic solvents, have been postulated to have deleterious effects on the integrity of the growth factor and may result in deactivation [90]. Therefore other means of drug delivery via polymer scaffolds have been investigated. Growth factors have been entrapped in scaffolds by adding the molecules to the polymer solution prior to compression molding. Murphy et al. mixed vascular endothelial growth factor (VEGF) into a poly(lactide-co-glocolide)-salt solution before compressing molding into a 4.2 mm disk [45]. The disks were then gas foamed causing the VEGF and NaCL particles to become entrapped in the polymer. Soaking in CaCl2 for 24 hours then leached the NaCl particles out. Endothelial cells grown on these porous degradable scaffolds had a significant increase in proliferation for all time intervals when compared with scaffold prepared by the same method without the VEGF. Although sustained release of the

VEGF from the scaffold was achieved, 30% of the VEGF was released within 1 day and as much as 65% of the VEGF was released by day three. This type of release profile is common for many scaffolds. Emulsion freeze-dried PLGA scaffolds released an initial burst of bovine serum albumin within the first two days followed by slow sustained release over 12 weeks (Figure 1.16) [89]. Although these methods of incorporation have been shown to readily entrap growth factors, retain their bioactivity and reduce the initial release burst, they cannot incorporate large quantities of the growth factor into their

35 structure. On average, the maximum drug that can be incorporated into the scaffold is about 0.05 wt. % of the scaffold (about 0.5ug of drug for every 1mg of polymer). To increase the loading capability, supercritical carbon dioxide (SCCO2) processing methods have been used.

Supercritical fluid technology has traditionally been used for chemical extraction and synthesis. In the mid-1980’s SCCO2 was used for bio-catalytic reactions and by the mid-90’s it was used to develop sized protein powders for drug delivery without the use of solvents [90]. Research over the past several years has demonstrated that simple swelling, foaming, and extraction can be achieved using supercritical fluids. When judiciously applied, these modification techniques do not degrade the mechanical properties of the native polymer. Furthermore, the use of carbon dioxide as a supercritical fluid provides complete sterilization of the material in addition to its often- cited benefits as an environmentally benign and non-toxic solvent. SCCO2 is currently used to create porous and non-porous polymer scaffolds that entrap growth factors [52, 77].

[77] The SCCO2 process allows for much greater loading of the drug up to 50% w/w . The work by Howdle et al indicates that a scaffold with controlled porosity and pore size can be impregnated with large amounts of bioactive molecules without losing their activity.

Most recent research has shown that ibuprofen can be impregnated into PVP from

SCCO2 solution and PLA composites encapsulating enzymes were formed using SCCO2,

[91, 87] with retained enzyme activity . SCCO2 has also been used to impregnate

Figure 1.16 Release kinetics of DNA from microspheres, compressed disks and foamed disks. Formation of microspheres into either foamed or solid scaffolds dramatically decreases the initial release burst seen in non-formed microspheres, [84]

37 polymethyl methcrylate with a fluorescently labeled protein [86]. The cross-section of the protein-impregnated disk indicated that protein impregnation had occurred to a depth of

30 µm [86]. These studies indicate that proteins can be embedded into a polymer via

SCCO2 and retain their activity. While SCCO2 provides a method of infusing polymers with proteins and drugs without the use of organic solvents and at higher loading capabilities, there are problems with the ability of the technique to provide a uniform penetration front.

Drug delivery promises to be a useful tool in tissue engineering, however the ability to embed accurate quantities of growth factors or drugs into the polymer and more control over the release profile is needed before they can be put into any clinical situation.

1.6 The Future of Tissue Engineering Scaffolds

Research in tissue engineering can be dated back to 1933 when Bisceglie implanted mouse tumor cells that were encased in a polymer membrane into the abdomen of chick embryos and showed the survival of these cells. Tissue engineering has since evolved into a highly interdisciplinary field in which almost every tissue and organ structure is actively being studied. However, few products have entered clinical trials and even fewer have been approved be the FDA for clinical use in humans. A major obstacle in the engineering of organs is the ability to organize cells in the proper manner and deliver nutrients to and from the system. Strategies for increased vascularization of the engineered tissue include release of growth factors such as vascular endothelial growth

38 factor from the scaffold. Cell organization has also been enhanced by new scaffold processing techniques which allow for a much more controlled fabrication of the scaffold architecture and morphology to acquire the appropriate tissue architecture. Scaffolds in the future will not only provide the correct microenvironment for the cells to reside in, but also supply the appropriate physiological stresses and soluble signals for functional tissue development. Advances in scaffold design will ultimately lead us closer to the goal of engineering fully functional tissues to restore, maintain or enhance tissues and organs.

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CHAPTER 2

CELLULAR RESPONSE TO NANOSCALE TOPOGRAPHY; EFFECT OF 20 NM

HIGH PROTRUSIONS

2.1 Abstract

Surface topography is believed to be a potent factor affecting cell morphology, proliferation and differentiation. The effect of surface roughness in the micron to multi- micron range has been investigated for a variety of cell types. In this study, the influence of nanoscale surface roughness was examined in terms of its effects on morphology, cytoskeleton expression, proliferation, differentiation, function and apoptosis.

Polyethylene terephthalate (PET) discs were etched using an oxygen plasma to produce uniform and decidedly nanoscale values of surface roughness. Three distinct types of cell lines − mouse 3T3-L1 preadipocytes, human JEG-3 choriocarcinoma cells and human

MCF-7 breast adenocarcinoma cells − were cultured on the plasma-treated discs.

Untreated PET discs were used as a control. Cytoskeletal proteins (F-actin and cytokeratin peptide-8) exhibited similar patterns of expression. Cell morphology was also similar on either surface. Cell growth kinetics for the three types of cells and hormone secretion from the JEG-3 cells was not significantly different from that of the controls

(P>0.05). However, the differentiation of preadipocyte 3T3-L1 cells into lipid-laden fat

50 cells was affected by nanoscale surface topography. In addition, the 15dPGJ2-induced apoptosis of the JEG-3 and MCF-7 cells revealed distinct differences between the two surfaces. Plasma-treated surfaces showed more differentiated and apoptotic cells, respectively, compared to the controls. These results indicate that nanoscale roughness has little effect on cellular adhesion, proliferation and hormone secretion in the cell lines tested in this work. However, we observed a synergistic effect of topography and nanoscale roughness in vitro when these surface modifications were present.

2.2 Introduction

Surface topography upon which cells grow is one of the classic factors known to influence cellular behavior and has been extensively investigated. Roughened surfaces having tissue-like dimensions of 1-50 µm have shown better early adhesion of cells,1-3 increased osteoblastic function and functional gene expression,4-6 reduced fibrous encapsulation,7 and enhanced integration of percutaneous implants compared to corresponding smooth surfaces.8 The majority of research on surface topography involves substrata having roughness features greater than 0.5 µm.9 However, surface roughness in the 10 nm range may also influence biological activity.10 Topography as small as 10 nm may preferentially adsorb small biomolecules and ions.7 Nanoscale three- dimensional fibrous scaffolds containing fibers with diameter ranging from 50-500 nm have been investigated to optimize cell attachment and function.11 Such nanoscale

51 surface engineering efforts have recently generated a flurry of activity based on the premise that biological response to a non-cytotoxic substrate may be controlled as much by topography as chemical composition.9, 12-19

Our interests in this issue derive from the need to optimize the surface of fiber, whose diameters are already in the micron range, prior to expansion of cell populations for tissue engineering applications. In examining the issue of nanoscale topography, we sought to firmly establish the possible growth dependence (or complete lack thereof) of cells. We compared the growth characteristics in vitro of three distinct cell populations mesenchyme-derived, fibroblast-like cells, epithelial breast cancer cells and placental trophoblast-like choriocarcinoma cells in response to nanoscale topography. These cell types were employed to determine whether surface topography on the nanometer scale could influence a broad range of cell types.

The optimal substrate for cell function should possess many similarities to the topography of the natural cell-substrate interface. Throughout the vertebrate body, cells lie on a basement membrane a porous, fibrous meshwork of extracellular matrix (ECM) proteins such as fibronectin, laminin, type I and IV collagens, hyaluronic acid, chondroitin sulfate or dermatan sulfate and proteoglycans. The biochemical and mechanical properties of plasma membrane–ECM interactions modulate cell processes such as adhesion, proliferation, locomotion, differentiation and cell shape which then influence gene expression and control cell cycle activity.20 For example, hyaluronic acid inhibits cell-cell adhesion, while the rigidity of the cell membrane influences the strength of integrin-cytoskeleton linkages.21,22 The basement membrane also possesses a complex

52 three-dimensional topography with nanometer-sized features. Recent studies show that the basement membrane consists of fibers ranging from 5-77 nm in width and pores 3-72 nm in diameter, supporting the need for substrata roughness of the same dimensions as the ideal for in vitro cell culture. 12-13, 23-24

To achieve this level of roughness, we used polyethylene terephthalate (PET), a pertinent choice, as it is utilized extensively in vivo, in vitro and ex vivo. Oxygen plasma discharge was used to modify the polymer and produce suitable values of surface topography. It has been demonstrated previously that plasma-treated surfaces improved cell adhesion on PET for mice fibroblasts, myoblast cells, and endothelial cells without observable cytotoxic effects.25-27 In this study, oxygen plasma etching alternated with rinsing in 70% ethanol was used to produce a relatively uniform distribution of roughness across the surface of PET coverslip discs. Atomic force microscopy (AFM) was used to determine that the resulting Ra was ~16 nm. The results show that, in agreement in much of the literature, proliferative behavior was unaffected. However, apoptosis and differentiation on these surfaces provide clear indications of a previously unsuspected role of nanoscale topography in influencing cellular responses.

2.3 Materials and Methods

2.3.1 Polymer discs

® Polyethylene terephthalate (PET), Thermonox coverslip discs (Nalgene Nunc

International) were used in this study. The thickness and diameter of the PET discs were approximately 200 µm and 13 mm, respectively.

53 2.3.2 Oxygen plasma treatment

The plasma reactor (EA Fiscione, Model 1400) used in this study consisted of a plasma chamber with an oil-free vacuum system comprised of a molecular drag pump backed by a diaphragm pump. A high frequency oscillating power supply located outside of the plasma chamber was coupled to the chamber with a HF antenna creating low energy plasma within the chamber. The gas supplied to the system was a mixture of 25% oxygen and 75% argon. An internal pressure regulator was present to provide the proper pressure to the process gas valve set at 10 mTorr. Discs were placed horizontally in pairs on an aluminum sample holder and inserted through a single port into the plasma chamber. The surface modification process consisted of exposing the disc surface to oxygen plasma for 7.5 minutes followed by a rinse using 70% ethanol. This cycle was repeated three times to produce the abundant nanoscale roughness. We conducted several experiments showing that this topography was both highly reproducible and stable in the presence of 70% ethanol. This was necessary due to the usage of 70% ethanol for sterilization of the discs prior to cell seeding. Untreated PET discs rinsed with 70% ethanol were used as a control.

2.3.3 Scanning electron microscopy (SEM)

The surface topography of the treated and untreated surfaces was qualitatively examined using scanning electron microscopy (Philips model XL-30). Treated and untreated PET discs were coated with 10 nm of gold using a sputter coater and analyzed under SEM with 6 keV accelerating voltage.

54 2.2.4 Atomic force microscopy (AFM)

The surface topography of the discs was quantified using atomic force microscopy

(Digital Instruments model Nanoscope IIIa). Several areas, 100 µm2, on both the untreated and treated discs were analyzed in tapping mode with a 2 volt oscillating amplitude using a silicon tip and cantilever. The surface characteristics were evaluated both qualitatively and quantitatively using Digital Instruments software version 4.22r2.

Topographic roughness parameters Ra (average deviation from arithmetic centerline),

Rmax (maximum peak height), Rq (root mean square deviation of the surface) and Np

(average peak to peak distance) were determined for each sample surface.

2.3.5 Culture and medium

Mouse 3T3-L1 preadipocytes, human JEG-3 choriocarcinoma cells and human MCF-

7 breast adenocarcinoma cells (a generous gift from Dr. Lisa Yee, The Ohio State

University), obtained from the American Type Culture Collection (ATCC) were used in this study. Mouse 3T3-L1 cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, Grand Island, NY) supplemented with 10% fetal bovine serum

(FBS, Gibco). Human JEG-3 cells and MCF-7 cells were maintained in Ham’s

F12/Dulbecco’s modified Eagle’s medium (F12/DMEM, 1:1) (Gibco, Grand Island, NY) with 10% FBS, 2 mM L-glutamine and 1 mM sodium pyruvate. All cells were cultured at 37°C in humidified 5% CO2 incubator.

55 2.3.6 Cell culture experiments

The plasma-treated and control PET discs were placed in 24-well cell culture plate and sterilized by immersion in 70% alcohol for 30 min followed by washing with phosphate-buffered saline (PBS) three times and the medium once. Seeding cultures were prepared by trypsinizing mouse 3T3-L1 cells, human JEG-3 or MCF-7 cells grown in a 75 cm2 flask to make a cell suspension containing 0.2-1 x 107 cells/ml. Each well containing the PET disc was inoculated with a known amount of cells (5 x 104) and then incubated in a humidified 5% CO2−95% air incubator (37 °C). For SEM, immunocytochemistry and apoptosis studies, cells were incubated for 48 h. For cell growth and hormone production experiments cells were incubated for 3-6 days and sampled at different time point. The spent media were replaced with fresh media every 2 days. Each experiment was done in duplicate and repeated at least once.

2.3.7 Cell number and hormone secretion

Cell-PET disc samples were taken every day for cell number determination. Cell number was determined by dye exclusion using trypan blue (0.4% in 0.85% saline)

(Sigma, St. Louis, MO). A hemocytometer and an inverted phase contrast microscope were used for cell counting. Liquid samples (150 µl each) were taken from each well of human JEG-3 cell culture at various time intervals for 17β-estradiol and progesterone assays. 17β-Estradiol and progesterone were assayed by specific EIA kits (Cayman

Chemical Co., Ann Arbor, MI) as outlined by the manufacturer. Each assay was run in duplicate wih a standard curve and measured at 405 nm.

56 2.3.8 Cell morphology

Cell morphology on the two different surfaces was studied using SEM. Cells on PET discs were fixed with 3% glutaraldehyde in a phosphate buffer (pH 7.4) for 2 h. They were then washed with the phosphate buffer for 30 min. The samples were osmicated in

1% OsO4 for 1 h and washed again. After dehydration in a graded ethanol series, the samples were dried in a critical point dryer. After sputter coating with gold/palladium, the samples were examined using a scanning electron microscope (Philips model XL 30).

2.3.9 Immunocytochemistry

The expression of cytoskeletal proteins including cytokeratin peptide-8 and F-actin were examined by immunocytochemistry or phalloidin-NBD-FITC staining using an epi- fluorescence microscope. Briefly, the control or treated PET disc with cells was fixed in

4% paraformaldehyde in PBS for 15 min. After fixation, the samples were rinsed gently with PBS and the cells were permeabilized with 0.5% Triton X-100 in PBS for 10 min.

Samples were then pretreated for 15 min with 5% horse serum (HS) in PBS followed by incubation for 60 min with the primary antibody of cytokeratin peptide-8 (1:100,

Chemicon, Temecula, CA) or FITC-conjugated phalloidin (to stain F-actin) (1:50,

Molecular Probes, Eugene, Oregon). After rinsing with PBS, F-actin stained sample was examined under a fluorescence microscope (Nikon ECLIPSE TE 200). For cytokeratin staining, the sample was rinsed with 5% HS in PBS and then incubated with anti-mouse

IgG conjugated to FITC (1:100) for 30-45 min before examination under the microscope.

57 Negative controls for cytokeratin were prepared following the same procedures and stained with secondary antibody only. The complete negative pictures were obtained and showed high specificity of the antibody.

2.3.10 3T3-L1 cell differentiation

Mouse 3T3-L1 cells were grown on treated and untreated PET discs to confluence for

2 days. Differentiation was induced after 2 days of incubation with 10 µg/ml insulin

(Life Technologies, Grand Islands, NY), 1 µM dexamethasone (Sigma, St. Louis, MO) and 500 µM isobutyl-methylxanthine (IBMX) (Sigma, St. Louis, MO) in DMEM containing 10% FBS (IDX mixture). After two days of differentiation, cells were incubated in DMEM containing 10% FBS and replaced every other day. At day 6 after differentiation, cells were stained with oil red O for 15 min and then observed under microscopy. The oil red O stock solution (300 mg/l in 99% isopropanol) was diluted in

6:4 with distilled water and filtered immediately before each use.

2.3.11 Cell apoptosis

After 24 h of incubation, human JEG-3 and MCF-7 cells on plasma-treated and

12,14 untreated discs were treated with 10 µM 15-deoxy ∆ prostaglandin J2 (15dPGJ2)

(BIOMOL Research Laboratories, Inc, Plymouth Meeting, PA) for 5 h to induce cell apoptosis. Cells on the discs were fixed with 4% paraformaldehyde (Sigma, St. Louis,

MO) for 25 min at 4°C. After twice washing with PBS for 5 min, the cells were permeabilized by 0.2% Triton X-100 solution (USB, Cleveland, OH) in PBS for 5 min, followed again by twice washing with PBS for 5 min. An Apoptosis Detection System,

Fluorescein kit (Promega) was used for apoptosis analysis based on the TdT-mediated

58 dUTP Nicked-End Labeling (TUNEL) assay as outlined by the manufacturer. The positive control was prepared by DNase treatment followed by the same washing and fixing procedure. The negative control was incubated in the buffer without TdT Enzyme.

Cells were visualized using a Nikon ECLIPSE TE 200 inverted microscope after exposure to 1 µm/ml propidium iodide (PI) (CALBIOCHEM, La Jolla, CA) for 15 min.

Natural apoptosis of human JEG-3 and MCF-7 was also performed as above, following 7 days of cultivation without change of medium.

2.3.12 Statistical analysis

For cell growth and hormone secretion experiments, a one-way analysis of variance

(ANOVA), followed by the Turkey-Kramer multiple comparison analysis, was performed. The data were presented as the mean and standard deviation, and p<0.05 was considered significant.

2.4 Results

2.4.1 Surface Characteristics

At relatively high magnifications, SEM revealed that the plasma-treated surface showed a relatively rougher surface than untreated PET discs (Fig. 2.1). AFM revealed that the treated surface contained many peaks roughly 50 nm in height. In contrast, theuntreated surface was relatively smooth with minor sinuous hills (Fig. 2.1C, D). The topographic parameters indicate a marked difference between the two surfaces with the average roughness (Ra) of the treated surface being four times that of the control (Table

1).

59 2.4.2 Cell Morphology

In this study, mouse 3T3-L1 cells were used as a model cell line for mesenchymal cell types; human JEG-3 and MCF-7 cells were used as model cell lines for epithelial cell types. The morphology of 3T3-L1, JEG-3 and MCF-7 cells grown on treated and untreated surfaces (as the control) at day 2 were examined using SEM. On both surfaces, undifferentiated 3T3-L1 cells exhibited a flat fusion morphology, formed dense layers with more diffuse borders and reached confluence. There was no significant morphological difference between the cells cultured on the two different surfaces. The control surface not covered by 3T3-L1 cells appeared to be clean and smooth, while the unobscured treated surface was rough and covered with a layer of dense matrix, probably extracellular matrix (ECM) protein deposition (Fig. 2.2A, B). Some of the 3T3-L1 cells on the O2 plasma-treated surface exhibited a stellate shape having numerous filamentous extensions (data not shown). The JEG-3 cells spread out and grew as a monolayer on both surfaces. On the the control surface, cells flattened with numerous microvilli, while on plasma-treated surface the cell membrane appeared more ruffled having many global nodules (Fig. 2.2C, D). MCF-7 cells exhibited the same tendency for spreading with some cytoplasmic expansions firmly adhered on either surface. The only difference was

A B

C D

® Figure 2.1. Surface morphology of Thermanox PET discs. SEM micrographs of the untreated (A) and plasma-treated (B) surfaces, and AFM micrograghs of the untreated (C) and plasma-treated (D) surface. A distinctive, uniform surface roughness is produced (B and D). The untreated surfaces are used as the control

ROUGHNESS CONTROL TREATED

Ra 4.8 16

Rmax 10 58

Rq 6 21

Np 2220 690

TABLE 2.1. Surface roughness (nm) of treated and untreated PET discs.

A Control 3T3-L1 B 3T3-L1

C Control JEG-3 D JEG-3

E Control MCF-7 F MCF-7

Figure 2.2. SEM micrographs of cells grown on PET discs. Mouse 3T3-L1 cells on the untreated (as control) (A) and plasma-treated (B) surfaces; human JEG-3 cells on the untreated (C) and plasma-treated (D) surfaces; human MCF-7 cells on the untreated (E) and plasma-treated (F) surfaces. The untreated surfaces are used as the control.

63 that cells on the control surface appeared to be polygonal with few pseudopodia while some cells on the treated surface took on a spherical shape with some filopods extending from the cell body to the surface (Fig. 2.2E, F).

2.4.3 Cytoskeletal Protein Expression

Cytoskeletal proteins are important for maintaining cell shape. They are intimately involved in cell adhesion, spreading and migration along the substratum. Cytoskeletal components include actin filaments, polymerized microtubules and cell type-specific intermediate filaments. In this work, F-actin in 3T3-L1, JEG-3 and MCF-7 cells was labeled with FITC-phalloidin and visualized using fluorescence microscopy. Cell orientation on the surfaces and the organization of the actin cytoskeleton was revealed by

F-actin expression. On both untreated and treated surfaces, JEG-3 cells fused into multinucleated cells and the F-actin microfilaments were expressed at every mononucleated cell periphery (Fig. 2.3A, B). For MCF-7 cells, F-actin was concentrated at the spreading edge of the cells (Fig. 2.3C). There was no significant difference in the expression pattern for JEG-3 and MCF-7 cells grown on the two types of surfaces. The actin filaments of 3T3-L1 cells on the O2 plasma-treated surfaces oriented in the direction of the long axis of the cells, while the cells themselves did not display any particular orientation (Fig. 2.3D). The filaments contained many thick stress fibers and spread out well on the substratum. The F-actin pattern of 3T3-L1 cells was similar on both surfaces

(data not shown).

64 A Control JEG-3 B JEG-3

C MCF-7 D 3T3-L1

E JEG-3 F MCF-7

Figure 2.3. Cytoskeletal expression on PET discs (200X). F-actin expression in human JEG-3 cells on the untreated (control) (A) and plasma-treated (B) surfaces; F-actin expression of human MCF-7 cells (C) and mouse 3T3-L1 cells (D) on PET surfaces (red staining showed the cell nuclei); cytokeratin peptide-8 expression of human JEG-3 cells (E) and MCF-7 cells on PET surfaces.

65 Cytokeratin peptide-8, one of the intermediate filaments, was used as a general epithelial cell marker for MCF-7 and JEG-3 cells. High levels of cytokeratin expression were visualized in JEG-3 and MCF-7 cells grown on untreated or treated surfaces. The thick intermediate filaments formed a circle around the cell nucleus (Fig. 2.3E, F).

Cytokeratin expression in both MCF-7 and JEG-3 cells on the treated surfaces was similar to that of the control (data not shown). The similar cytoskeletal expression indicated that the plasma-treated surface had little effect on cell shape, spreading and reorganization.

2.4.4 Cell Proliferation

Cell proliferation was monitored by determining the increase in cell number with time in culture. The growth kinetics on plasma-treated discs and untreated control surfaces was compared for 3T3-L1, JEG-3 and MCF-7 cells. On untreated and treated surfaces, the number of 3T3-L1 cells increased until 3 days and reached confluence (Fig. 2.4A). There was no significant difference in cell number between treated and untreated surfaces.

Similarly, no difference in JEG-3 cell growth between O2 plasma-treated surfaces or untreated surfaces was observed during the period of cultivation (Fig. 2.4B). For MCF-7 cells, there was no difference in cell growth between two surfaces before 48 h. After 48 h, the cell number on plasma-treated surfaces was slightly higher than control (Fig. 2.4C).

However, this difference was not statistically significant (P>0.05).

66 2.4.5. Hormone Secretion

The production of 17β-estradiol and progesterone by JEG-3 cells was an important indicator of differentiated placental trophoblast function. JEG-3 cells are malignant choriocarcinoma cells, which secrete high levels of 17β-estradiol and progesterone when provided with appropriate steroid substrates. The 17β-estradiol secretion of JEG-3 cells grown on untreated and O2 plasma-treated surfaces was compared and similar kinetics were found during the 4 days of cultivation (Fig. 2.5A).

Progesterone secretion by JEG-3 cells grown on plasma-treated discs was slightly higher than secretion by cells grown on the untreated surface. However, the difference was not statistically significant (P>0.05) (Fig. 2.5B). Since the cell growth kinetics on both surfaces were similar, the specific production of 17β-estradiol and progesterone were also similar, indicating that plasma treatment did not change demonstrably cell physiology and biochemical function under the parameter tested here.

2.4.6 Cell Differentiation

Mouse 3T3-L1 cells are preadipocytes that can differentiate into triglyceride-rich adipocytes in vitro by the appropriate treatment.[28] These cells appear morphologically similar to fibroblasts in the undifferentiated state (Fig. 2.6A). At confluence, induction of differentiation by the IDX mixture led to dramatic cell shape changes. Adipocytes were spherical in shape and accumulated neutral lipid droplets as revealed by oil red O staining

(Fig. 2.6B). Adipose cell formation involved an increase in adipocyte size and

3.0 3T3-L1 Control 2.5 Treated

2.0

1.5

1.0

0.5

0.0 0 20406080100120 A Time (h) 10 9 Control JEG-3 8 Treated ) 5 7

6 5 4 3 Cell Number (X10 2 1 0 0 20406080100120140 B Time (h) 3.0 MCF- 7 Control 2.5 Treated

2.0

1. 5

1. 0

0.5

0.0 0 20406080100120140

Time ( h) C

Figure 2.4. Cell growth kinetics on the untreated and plasma-treated PET discs. (A) Mouse 3T3-L1 cells; (B) human JEG-3 cells and (C) human MCF-7 cells.

1800

1600 Control

1400 Treated

1200

1000

800 β

600

400

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0 A 0 102030405060708090100 Time (h)

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6000 Progesterone (pg/ml) Progesterone 4000

2000

0 0 102030405060708090100 B Time (h)

Figure 2.5. Hormone production of JEG-3 cells on the

untreated and plasma-treated PET discs. (A) 17β-Estradiol and (B) progesterone. 69 the formation of new adipocytes from preadipocytes. There was no significant difference in adipocyte size, but adipocyte number on the plasma-treated surface was slightly higher than that on the control surface (Fig. 2.6C, D).

2.4.7 Cell Apoptosis

JEG-3 and MCF-7 cells are human carcinoma cells. They can be induced to

12,14 29 undergo apoptosis by treatment with 15-deoxy∆ prostaglandin J2 (15dPGJ2). The cells can also undergo natural apoptosis during long-term culture. The nucleus of the apoptotic cell is highly condensed and fragmented. The fragmented DNA of apoptotic cells was visualized by fluorescence microscopy using the TUNEL assay. The total cell number in the culture was detected using PI staining. In the study of basal apoptosis, there was no significant difference in the percentage of apoptosis for JEG-3 cells grown on the two surfaces (Fig. 2.7A-D). Similar results were seen in MCF-7 cells (data not shown). For both the MCF-7 and JEG-3 cells, however, 15dPGJ2-induced apoptosis on the O2 plasma-treated surface was somewhat stronger than that of cells grown on the untreated surface (Fig2.7E-H).

2.5 Discussion

® Oxygen plasma modification of Thermanox discs clearly does not produce any apparent cytotoxic effects. Mouse 3T3-L1 cells, human JEG-3 cells and MCF-7 cells behaved similarly on plasma-treated and untreated surfaces in terms of adhesion,

A B

C D

Figure 2.6. Images of mouse 3T3-L1 cells (200X). Preadipocytes (A); differentiated adipocytes (B); adipocytes on the untreated (C) and plasma- treated (D) PET discs.

71 proliferation, differentiation, hormone secretion and apoptosis. There were statistically indistinguishable differences in cell morphology, MCF-7 cell proliferation and progesterone secretion. The cytoskeleton appeared similar in mouse 3T3-L1 cells, human JEG-3 cells and MCF-7 cells on both type of surfaces.

The cellular effects of substrate topography have been observed to be highly dependent on degree of roughness and the specific cell type under examination. On a surface composed of 0.5 µm grooves, bone cells showed minimal responses to surface topography.30 Human osteoblast-like cells (MG 63) behaved similarly on titanium discs having an average surface roughness < 3 µm.31-34 There was no statistically significant

4 difference in cell proliferation on three surfaces with Ra’s of 3, 6 and 9 µm. Hatano et al. specified a roughness of about 810 nm as an optimum in the culture of rat calvarial osteoblasts.6 However, their work also showed that roughness other than this value produced indistinguishable results. In addition, osteoblast adhesion was reported to be unaffected by topography during the first week of culture, but became sensitive to this parameter after 14 days in vitro.35 From the literature it is clear that one ideal value of surface roughness cannot be defined for all cells.The present experiments indicate that nanoscale topography has negligible effects during the early stages of cell culture and may not influence overall cellular behavior. Serum protein adsorption is a significant factor for cell adhesion. Cell adhesion to a substrate follows a sequence involving serum protein adsorption to the substratum, cell contact, initial attachment and then spreading.36

Protein adsorption is almost spontaneous, depositing within the first minute of contact,

A A Control JEG-3 B Control JEG-3

C JEG-3 D JEG-3

E Control MCF-7 F Control MCF-7

G MCF-7 H MCF-7

Figure 2.7 Images of cell apoptosis on PET discs (200X). Apoptotic human JEG- 3 cells revealed by TUNEL assay on the untreated (A) and plasma-treated (C) surfaces. Total human JEG-3 cells were revealed by PI staining on the untreated (B) and treated (D) surfaces. 15PGJ2-induced apoptotic human MCF-7 cells revealed by TUNEL assay on untreated surface (E) and plasma-treated surface (G) and total human MCF-7 cells revealed by PI staining on untreated surface (F) and treated surface (H). The untreated surfaces are used as the control.

73 and cells in suspension then gravitate to this coated substrate. Significant amounts of vitronectin and fibronectin were adsorbed onto polyester PGA and PLGA from serum- containing medium; smooth muscle cell adhesion was regulated by the type and amount of protein adsorbed.37 As our data suggest, however, similar amounts and types of proteins adsorbed on two surfaces having nanoscale roughness differences ofapproximately 12 nm (data not shown). This protein adsorption from serum may have overridden the differences in surface topography to normalize the adhesion portion of the cellular growth curve by coating the landscape of the culture surface.

In contrast, the enhanced biomolecule-cell interactions suggest a synergistic relationship with nanoscale surface topography. Significant differences between plasma- treated and untreated PET discs were observed during hormone-induced 3T3-L1 differentiation and 15dPGJ2-induced apoptosis of JEG-3 and MCF-7 cells. Mouse 3T3-

L1 differentiation on the plasma-treated surfaces was enhanced when biochemical stimuli were present. Increased cell signaling via integrins38 and the preferential polymerization of actin10 are two examples of intracellular processes affected by external surface topography. Wojciak-Stothard et al. noted a concentration of αV integrins along groove/ridge boundaries suggesting that integrins tend to cluster in areas of topographical change.10 The integrin's role in linking the extracellualr matrix and the cytoskeleton of cells has been widely reported.39,40 An additional role of integrins as signal transducers has also been entertained.40 If the apoptotic signals induced by an extracellular

74 biomolecule originates at a receptor on the cell membrane and is transduced through a signaling cascade to the nucleus, a concentration of integrins in response to the nanoscale topography could enhance signal transduction.41 Clustering of β1 integrins has also been shown to induce tyrosine phosphorylation in several kinds of cells.38 Drug-induced apoptosis in leukemia cells by p53 depends on tyrosine phosphorylation.40 Therefore, it can be speculated that a nanoscale-triggered increase in tyrosine phosphorylation will increase the rate of apoptosis. More recently, 15dPGJ2 has been proposed to function via non-apoptotic mitochondrial membrane depolarization;41 the interaction of microtubules and mitochondria suggests that enhancement of microtubule density and/or organization in the cell membrane in response to nanoscale topography may contribute to increased depolarization of the mitochondrial membranes (relative to the ‘smooth’ substrate) promoting cell death.10

2.6 Conclusions

Nanoscale PET surface topography was produced using oxygen plasma etching and was examined by SEM and quantified by atomic force microscopy. Cellular adhesion, proliferation, differentiation, function and apoptosis were evaluated in both mesenchymal and epithelial cells. This surface modification was biocompatible with all three cell types studied herein. Statistically indistinguishable differences in morphology, growth and function were observed in cells cultured on O2 plasma-treated versus untreated surfaces. We demonstrate that this level of nanoscale surface topography has only limited effects on these adhesion, growth and proliferation. This may be due to a

75 diluting effect of protein adsorption that could override the 12 nm difference in surface roughness. However, 15dPGJ2-induced apoptosis and differentiation show clear differences suggesting that subtle intracellular influences do exist and can control cell responses to a given surface in the appropriate environment. This has logical consequences for long-term tissue engineering applications incorporating cellular modifications.

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CHAPTER 3

NANOFIBRILLAR SURFACES VIA REACTIVE ION ETCHING

3.1 Abstract

The etching behavior of polyethylene terephthalate (PET) versus reactive ion etching under various gas compositions was examined. An RF power supply operating at

30Khz was used to produce plasmas from Ar. N2, O2, Ar-N2, Ar-O2, and N2-O2. Exposure times were typically 10, 20 and 30 minutes. Exposure to the relatively inert gases and gas mixtures (Ar, N2, Ar-N2) produced a polygonal pattern of protrusions surrounding shallow cavities. In contrast, the presence of oxygen or oxygen-containing plasmas added fibrils to these polygons typically originating at their triple points. Fibrillar dimensions ranged with exposure time but were observed to be up to 300 nm in length and approximately 20 nm in diameter. Atomic force microscopy (AFM) was used to quantify the surface roughness and show that the inert gas compositions (Ar, N2, Ar-N2) produced statistically indistinguishable values of Ra (14.1 + 1.7 nm, 15 + 2.5 nm, 14.2 + 2.9) significantly larger than the as-received film. The pure oxygen etched films had Ra values at least twice as large as those of inert gas compositions. Prior work by Nie et al suggests that etching resistant low molecular weight fragments (LMWF) could develop on the

81 surface during exposure and lead to the observed surface morphologies. Prior surface deformation was shown to have a strong influence on the spacing and density of surface fibrils.

3.2 Introduction

Polymer surface topography is one of the classic factors controlling cell behavior on surfaces. In the early 1930’s, the work of Weiss demonstrated that cells grown on an oriented substratum assume a corresponding orientation and migrate in that direction [1].

Since then a number of investigations have taken place that examine the effect of surface roughness on cell behavior [2-8]. A variety of surface modification techniques, such as mechanical abrasion treatment with solvents, caustic solutions or acids, graft polymerization, and micro-machining, have been employed to alter cellular behaviors [9-

14]. However these techniques either lack reproducibility, utilize toxic compounds, or cannot be used on a three-dimensional surface. Therefore, many researchers have turned to reactive ion etching (RIE) to produce a random, reproducible surface topography.

Reactive ion etching has been used for many years in the plastics industry to enable printing operations [15, 16] and other modifications such as adhesives, coatings, composites, and electronics. Reactive ion etching has been extensively used due to its ability to modify large areas easily without changing bulk properties[15]. Plasma etching has been used to provide enhanced adhesion, hydrophilicity and biocompatability [15,17,18].

Reactive plasmas contain a collection of charged particles (ions and electrons), excited neutrals, radicals, and UV radiation [15,19,20] which react both physically and chemically with the surface to remove contamination, introduce chemical functionalities,

82 and induce chain scission and cross-linking [15,21-24]. The predominant reactive species in radio frequency plasmas are positive ions and photons capable of breaking intrapolymer bonds to form reactive radicals [21,23,25,26]. These reactive radicals usually interact with the surrounding gas to produce new surface functionalities such as carbonyl, hydroxyl, or carboxylic acid groups created from radical-gas interactions. Both the chemical structure of the polymer and the type of plasma determine the extent and character of degradation

[22]. Polymeric structures containing oxygen are more susceptible to oxygen plasmas whereas structures containing benzene rings are more resistant to oxidation and thus etching [22]. On the other hand, the ability of argon plasma to etch the oxygen-containing polymer polyethylene terephthalate (PET) is controversial. Fischer et al. reported that argon plasma alone does not etch PET [27] while Beake et al. reported that it does [21]. The reported effects of oxygen plasma etching are, in contrast, quite consistent. Etching rates

[28,29] typically increase with oxygen atom concentration in the plasma . The addition of N2

[15] and O2to the gas amplifies the oxygen atom concentration and etch rates . Rates of material removal appear to be faster at lower gas pressures [21]. Etching rates have been reported to be linear in some gases such as argon and nitrogen but parabolic in the presence of oxygen or air [23]. Bourceanu et al reported that etch rates varied with gas with highest etch rates seen with oxygen followed by air, nitrogen, and argon [23].

Exposure to the plasma also results in loss of C-H bonds as a result of hydrogen abstraction [30]. Chain scission results in a surface rich in low molecular weight fragments

83 which are then either removed from the surface via the vacuum system or remain on the surface [31-33] Etching and chemical modification often occur simultaneously and are competitive processes [15,21].

In this study, the ability of these different plasma ‘media’ (nitrogen, argon, oxygen, and 50-50 vol. % mixtures) to etch PET is investigated. High resolution SEM and atomic force microscopy are used to analyze the resultant surface as a means of providing an estimate of their biomimetic potential.

3.3 Materials and Methods

3.3.1 Polymer Films

Polyethylene terephthalate (PET), Thermanox™ plastic coverslip rectangles

(Nalgene Nunc International, Rochester, NY) were used in this study. The thickness, width and length of the PET film were 200 µm, 22 mm, and 66 mm, respectively.

3.3.2 Reactive Ion Etching

In the reactive ion etcher (Technics Micro-RIE Series 800-II, Concord, CA) we used, the platen (cathode) is connected to an RF power supply operating at 30 KHz. The chamber itself acts as the anode causing the gas entering the chamber to be ionized when an electric field is applied. Gasses were supplied to the system at a rate of 25 sccm and under 50 Watts of power with a pressure of 135 mTorr in the chamber.

To investigate the effect of gas media and time on etching ability, a test matrix was designed which utilized three exposure periods in combination with the different gasses selected on the basis of previous literature. Beake et al suggest that the most significant change in surface topography occurs after 20 minutes of exposure to oxygen

84 plasma; therefore 10, 20 and 30 min time intervals were investigated [21]. Different gases are known to possess varying etching ability and the presence of multiple gas species can have an effect. Thus, high purity gasses and 1:1 mixtures of each gas (high purity argon, oxygen or nitrogen or a 1:1 mixture of argon-oxygen, argon-nitrogen or nitrogen-oxygen) were examined. The PET films were placed flat on the center or the plasma chamber floor. The surface modification process consisted of a continuous exposure of the films to the various plasmas for 10, 20 or 30 minutes. The films were then placed upright in a microslide container to prevent subsequent contact with any other surface. Each experiment was conducted four times to ensure reproducibility.

3.3.3 Scanning Electron Microscopy (SEM)

The surface topography of the etched and non-etched films was qualitatively examined using scanning electron microscopy (FEI Sirion). Both etched and untreated films were sputter coated with a 10 nm layer of gold-palladium and then analyzed in ultra high resolution mode with a 5 keV accelerating voltage.

3.3.4 Atomic Force Microscopy

The surface topography of the films was quantified using atomic force microscopy (Digital Instruments Nanoscope IIIa, Veeco Metrology, Tucson, AZ) in tapping mode with a frequency of 225 kHz, an amplitude of 1.225 V and a scan rate of

0.500 Hz. Several areas ranging from 1 to 5 um2 on each surface were imaged and analyzed with Digital Instruments software version 4.22r2. Topographical roughness parameters Ra (average deviation from arithmetic centerline), Rmax (maximum peak height), and Rq (root mean square deviation) were determined for each surface.

Figure 3.1. High-resolution SEM of the as received PET film. The cracks seen on the surface are an artifact of the Au-Pd coating. Scale bar = 200 nm

86 3.4 Results

3.4.1 Scanning Electron Microscopy

Scanning electron microscopy revealed that each etched surface has a markedly different surface topography than the unetched PET film (Figure 1). Films etched with pure gasses have a distinct surface morphology characteristic of the gas used. PET films etched with argon exhibit the smallest amount of topographic change and are characterized by a polygonally patterned surface made up of many small protrusions.

After 10 minutes, the surface displays small (10-20 nm in diameter) protrusions in a pattern alternating with shallow divots (Figure 3.2a). After 20 minutes, the protuberances become larger (15-30 nm in diameter) and the polygonal pattern becomes less dense. The protrusions increase in size (40-50 nm in diameter) and decrease in density following the final 30 minute interval (Figure 3.2b).

Plasma etching with nitrogen is classified by larger protrusions arranged in a less regular polygonal pattern. At 10 minutes the surface has 20-40 nm diameter protrusions with shallow depressions between the raised protrusions at the edges of the polygons

(Figure 3.2c). These protuberances decrease in number and increase in length until they form short (60-80 nm in length) fibrils emerging from the triple points of the adjoining polygons (Figure 3.2d).

In contrast to the nitrogen and argon results, oxygen etching produces a distinctive fibrillar structure even at shortest time interval; only 10 minutes of etching

a. b.

c. d.

e. f.

Figure 3.2. High resolution SEM of the PET surfaces after etching for a) 10 min in Ar; b) 30 min in Ar; c) 10 min in N2; d) 30 min in N2; e) 10 min in O2; f) 30 min in O2.

88 produces fibrils 65-100 nm long and 20 nm in diameter (Figure 3.2e). Increasing exposure increases the fiber length (up to 300 nm long at 30 minutes) and decreases fibril density (Figure 3.2f, complete set of images in Appendix A)

Gas mixtures produce topologies intermediate to the pure gas morphologies. In all oxygen-containing mixtures, the fibrillar structure predominates. For example, oxygen- argon mixtures contain small protrusions at the triple points of the polygonal features at the 10 minute interval; these appear to grow into thin (15-20 nm diameter), long (200-250 nm) fibrils after 30 minutes of etching. Oxygen-nitrogen etched surfaces are also characterized by tiny protrusions at their triple points following 10 minutes of exposure.

In contrast, only 100-200 nm long fibrils are visible after 30 minutes of exposure (Figure

3.3). Argon-nitrogen mixtures produce a polygonal pattern consisting of very fine protrusions (5-15 nm in diameter). As the time of etching increases, larger protrusions are produced and increase in density until the surface contains protrusions typically 35 nm in diameter.

3.4.2. Atomic Force Microscopy

To better quantify surface topography, tapping mode atomic force microscopy was used to analyze these etched surfaces following 30 minutes of exposure. Surfaces imaged by AFM closely resemble the corresponding SEM images for surfaces etched with Ar (Figure 3.4), N2 and Ar-N2. However, the surface fibrils produced by oxygen- containing mixtures appear much flatter than in SEM. This is in part due to the complex fibril geometries and the inability of the reverse tapping probe to image angles greater

Figure 3.3. High resolution SEM of the PET surface etched with an O2- N2 gas mixture for 30 minutes

Figure 3.4. AFM image of a PET film etched with Ar for 30min.

(nm) 20 a

0 O2 N2 Ar O2-Ar O2-N2 Ar-N2 As-is Plasma Media 300

200

(nm) max

R 100

0 O2 N2 Ar O2-Ar O2-N2 Ar-N2 As-is Plasma Media

Figure 3.5. a) R (average deviation from arithmetic centerline) b) a Rmax (maximum peak height) values from films exposed to 30 minutes of the indicated plasmas

92 than roughly 80 degrees. Thus, the surface parameters calculated for surfaces etched with oxygen and oxygen-containing mixtures are clearly less than their actual values. In spite of this deficiency, we can conclude that only the oxygen surface has significantly different average roughness and maximum peak height values (Figures 3.5a & b) when compared to the other surfaces. Average roughness values for the oxygen, argon, nitrogen, argon-nitrogen, argon-oxygen, and nitrogen-oxygen etched films were 31.4 +

4.6 nm, 14.1 + 1.7 nm, 15 + 2.5 nm, 14.2 + 2.9 nm, 12.4 + 2.8 nm, and 11.5 + 1.6 nm, respectively. These values are all significantly larger than the as-received film (3.5 + 0.8 nm). Maximum peak values of pure oxygen etched films were at least twice as large as maximum values for all of the other gas combinations. Films etched with gas media other than pure oxygen all had statistically indistinguishable roughness values.

3.5 Discussion

Reactive plasmas produce a broad spectrum of effects on polymer surfaces including greater surface roughness and an increased affinity for water through the addition of polar groups to macromolecules. The hydrophilicity of the surface is not stable with time [21,25,27]. Upon storage in air for one day, the wetting angle of PET returns to its original value [25]. This hydrophobic recovery is believed to result from the movement of surface macromolecules into the bulk which homogenizes the overall surface character. In contrast to these chemical changes, surface analysis using SEM and

AFM has shown that the increase in surface roughness is stable over time.

93 In agreement with previous literature, our SEM and AFM results indicate that oxygen plasmas have the greatest potential to etch PET [25,27,31]. Positive ions are thought to be responsible for the degradation of the surface and thus a higher concentration of positive ions can lead to increased etching ability. Based on differences in chamber pressure before and during plasma etching, Borceanu et al determined that the concentration of dissociated ions in nitrogen plasmas was roughly 7% while for oxygen it was approximately 50% [27]. This could lead to a greater etch rate and surface roughness for films exposed to oxygen plasma simply because there are more dissociated species

(positive and negative) present.

Films etched with oxygen exhibited the largest surface topography followed by nitrogen then argon; the surface roughness parameters for nitrogen and argon are statistically indistinguishable. This is in contrast with previous reports [31] which conclude that argon plasmas do not possess the ability to etch PET. SEM and AFM results presented here indicate that under these conditions all gas and gas mixtures used in this study can etch PET. While etch rates and surface chemistry alterations have been reported for a number of materials, few studies describe the morphological results of the process [21,23,34,39].

3.5 1 Nanofibril Formation by Dewetting

At its most basic level, the process of surface roughening occurs by breaking of primary chemical bonds [25,27-30]. Exposure to plasma will induce both chain scission in the polymer chain and reaction with the plasma resulting in relatively polar, low

PET b. O2 plasma LMWF ( l) ~nm O plasma, heating 2

c. Localized Rim-type Rayleigh instabilities instability

PET O2 plasma, heating

PET

Figure 3.6. Schematic of morphological development on oxygen-etched PET surfaces . In a) the initial PET surface is given as etching initiates, b) illustrates the formation of a thin continuous film of relatively polar low molecular weight fragments (LMWF). At a subsequent stage in the etching process c) shows that the larger scale polygonal instability has been nucleated while smaller scale instabilities continues to exist. The final morphological stage is shown in d) in which the smaller scale instabilities no longer exist and only the rim and nanofibrillar features are still evident.

95 molecular weight fragments (LMWF) [36]. We can consider the products of the etching process to be an unstable, non-wetting thin film of liquid on the PET substrate (Figure

3.6).

At the very limits of our resolution, Figure 3.2f shows that the base of each fibril in the final etched surface appears to be well connected to the substrate. Multiple rims branch out into not only the major rim defining the polygon but also into the ‘crater’ of the polygon itself. We can rationalize the development of this morphology in the context of Figure 3.6 as a special case of the dewetting phenomenon observed in the two-phase

(polystyrene films on heated silicon substrates) system of Sharma and Reiter[40]. In our case, however, we envision the surface as an initially homogeneous, single phase material

(Figure 3.6a) that becomes a two-phase system (Figure 3.6b) due to the development of this etch-resistant LWMF film. Rayleigh instability [41] then appears to develop (Figure

3.6c) at two different levels: locally (20 nm) and at a length scale defined by the observed polygons (~150 nm). Given that the initial film thickness could be less than 1 nanometer, the local instabilities at the earliest etching periods seem quite reasonable.

The subsequent development of distinctive ‘nanofibrils’ on top of the polygonal substructure (Figure 3.6d) differs from the results of Sharma and Reiter[40]. In their case a thin (~10 nm) film dewetted and the triple points in the resulting polygonal arrangement were the highest features in the overall structure. In our case it seems likely that a polygonal structure of protrusions due to localized dewetting is formed first and the fibrillar structure is generated subsequently as the temperature and scale of the dewetting phenomena increase.

Figure 3.7. SEM image of “thermally bonded” PET etched for 30 minutes in oxygen.

97 3.5.2 Thermal Effects and Surface Instability

As etching time increases an additional factor, temperature, must come into play and influence the morphology of the oxygen-containing etching atmospheres in a manner that causes them to diverge substantially from those produced by inert gases. This is likely due to increases in temperature caused by the greater reactivity of O2 plasmas with

[42] polymers and our own simple observations: O2-etched substrates are hotter to the touch. Thus the transition from the small protrusions of Figure 3.2a to the short fibrils of

Figure 3.2e and subsequently to the extended fibrils of Figure 3.2f is driven by the decreases in viscosity, increases in wetting angle and greater transport rates engendered by increases in temperature. The etching temperature clearly exerts significant effects on the evolving nanostructure. An additional illustration of this is found in Figure 3.7 in which the “thermal bonding” technique of Dems and Rodriguez [43] was used to decrease the surface temperature. Substantial changes (smaller polygon size, apparent fibril clustering) in the nanostructure occur even though the etching exposure is identical to that used to produce Figure 3.2f.

The mechanism outlined above potentially provides a common mode for the creation of these morphologies. Thus O2-treated films exposed for shorter time intervals should generate lower amounts of LMWF and should produce morphologies similar to Ar and N2 etching. To test this hypothesis, a 1 minute oxygen-etched PET surface was produced and analyzed using SEM (Figure 3.8). Although the protrusions on the surface are not strictly identical to the Ar or N2 etched films a similar pattern of polygons surrounding shallow cavities is evident. This short exposure appears to capture an early

Figure 3.8. SEM image of 1 minute oxygen-etched PET. Note the nascent polygonal pattern emerging from an otherwise uniform pattern of protrusions.

99 stage of polygon development in which the local Rayleigh instabilities between the evolving polygon rims still exert a nanoscale (20nm) influence on the local surface morphology (Figure 3.6c).

Full description of this system would require quantification of forces in these polar films on relatively apolar substrates and the relationship between the film dynamics and the equilibrium substrate wetting characteristics [44]. Reliable methods for measuring these forces are lacking for even relatively simple systems and are non-existent for our system which is continuously changing in temperature and, probably, chemical composition. We can, however, invoke the following general equation [45,41,46,47] describing the shape and dynamics of such a thin film polar film on an apolar solid:

∂ ∂ ∂3 ∂ ∂2∆ ∂ h ⎛ 3 h⎞ ⎡ 3⎛ G⎞ h⎤ 3µ + ⎜γfbh ⎟− h ⎜ ⎟ =0 ∂ ∂ ⎜ ∂ 3 ⎟ ∂ ⎢ ⎜ ∂ 2 ⎟ ∂ ⎥ t x⎝ x ⎠ x ⎣ ⎝ h ⎠ x⎦

where h(x, t) is film thickness, x describes planar motion across the substrate, t is time, µ viscosity and γfb is the interfacial surface tension of the film (f) against the plasma or “bounding phase” (b) and ∆G is the total excess free surface energy. The viscous force

∂ ∂ ∂3 ⎛ h ⎞ ⎛ 3 h ⎞ 3µ retards the growth of any instability. The interfacial tension ( ⎜γfbh ⎟ ) ⎜ ⎟ ⎜ 3 ⎟ ⎝ ∂t ⎠ ∂x ⎝ ∂x ⎠ stabilizes the film until rupture occurs. The current set of observations makes it clear that both of these terms are quickly overwhelmed in the initial stages of the etching process but likely exert transient barriers to the evolving nanostructure. The intermolecular term

100 2 ∂ ⎡ ⎛ ∂ ∆G ⎞ ∂h⎤ 2 ( h3⎜ ⎟ ) contains a variable diffusivity h3 (∂ ∆G ) in which a negative ∂ ⎢ ⎜ ∂ 2 ⎟ ∂ ⎥ ∂h2 x ⎣ ⎝ h ⎠ x ⎦

2 diffusivity ( ∂ ∆G <0) leads to interfacial instability allowing the growth/existence of ∂h2 thicker /‘capped’/ nanofibrillar regions at the expense of the thinner/trough regions

(Figure 3.6d).

3.5.3 Effects of Prior Surface Deformation

Processing history has also been shown to influence surface morphology. In their

AFM analysis of biaxially-oriented polypropylene films, Nie et al developed a relationship between the draw ratios and the configuration of the resulting nanometer- scale fiber-like network structure [48]. Following etching, however, the fiber-like network was completely obscured by completely anisotropic low molecular weight oxidized deposits. Similarly, we see no directionality in either the protrusions or fibrils on any of the etched films with apparent exception of the 10 minute N2-etched film (Figure 3.2c).

Nie et al also observed that shear deformation of the underlying polymer chains appears to have an effect on the resultant surface morphology[49]. An increased density and ordering of the polymer chains in the deformed region were observed [49]. To investigate the effect of shear deformation on etched PET films, two PET films were lightly rubbed together prior to etching to produce micron sized scratches on the surface. After a 30 minute oxygen plasma exposure, the scratched surface exhibits changes in local morphology (Figure 3.9). Areas of tension (the indented portion of the scratch) and compression (the ‘hill’ of material displaced from the scratch) appear to be created. The

101

Figure 3.9. SEM image of a scratched then subsequently etched PET surface a) low magnification of entire scratch b) high magnification image of the highlighted tensile deformed region 2) high magnification image of region under compression. The scale bar = 2 microns in a) and 200 nm in b and c.

102 area under tension is characterized by lower fibrils densities compared to the non- deformed surface (Figure 3.9b). The area under compression displays a higher density of fibrils (Figure 3.9c). Higher densities of low molecular weight oxidized deposits were also seen on deformed polypropylene films [50]. This suggests a relationship between the stress state of the polymer and subsequent etching mechanisms. Deformation reorients the polymer strands creating areas of decreased and increased local chain density. In addition, polymer chains under local tensile stress are likely to be more reactive. These differences illustrate that micropatterning techniques that result in local changes in stress state can be used to examine the effects of both the micro- and the nano-scale on the same surface.

The ability to produce such distinct, scalable surface morphologies may be key to optimizing tissue engineering. Much effort has been directed toward creating biomaterials with chemistries and architectures that closely resemble natural structures.

Such surface-based biomimesis is intended to provide cells with structural cues absent current generation of the biomaterials. The architectural similarity of oxygen-etched polymer surfaces to native tissues (e.g extracellular matrix) may bring biomedicine closer to the ex vivo generation of replacement organs.

3.6 Conclusions

The ability to “tune” nanoscalar morphology has significant potential for controlling the biological response to a biomaterial. PET films plasma etched with argon, oxygen, and nitrogen gases show that each gas produces unique surface morphologies.

103 Oxygen is clearly capable of producing surfaces having the highest levels of surface roughness and a distinctive fibrillar structure. The latter has the greatest resemblance to naturally occurring structures such as collagen fibers and other non-globular proteins. We intend to exploit this ‘biomimesis’ to determine the true influence (if any) of these nanoscalar structures on biological function in a tissue engineering setting.

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108

CHAPTER 4

TOPOGRAPHICAL CONTROL OF CYTOSKELETAL ORGANIZATION

4.1 Introduction

A unifying principle in tissue engineering is the seeding of cells onto scaffolds for the purpose of generating 3D tissue-like structures. An important challenge is the promotion of cellular adhesion, orientation, cell-matrix and cell-cell communication which will ‘encourage’ the formation of neo-tissue that adequately mimics the properties of the desired endogenous tissue. Many investigators have studied the effect of scaffold properties on cell behavior to determine which characteristics elicit desired responses [1-6].

Surface topography had been extensively studied and is identified as one of the classic factors known to control cellular organization, proliferation and morphology [7-15] , its ability to alter either strength of adhesion to a scaffold surface or cell-substrate signaling remains poorly defined [16].

A comprehensive understanding of cell-substrate interactions requires investigations of cellular adhesion and cell-surface signaling. Adhesion to a surface is the initial step in the formation of cell communities and the eventual morphogenesis of neotissue. These adhesion events are mediated via interactions of specialized cell surface

109 proteins, integrins, with extracellular matrix constituents. Engagement of integrin receptors with the ECM ligands leads to the formation of focal adhesions (FAs) that complete the linkage of the ECM to the cytoskeleton. Focal adhesions are believed to regulate adhesion and locomotion in response to environmental stimuli and act as sites of force generation and signal transmission [17-18]. The formation and turnover of FAs are under the regulation of protein tyrosine kinases, specifically focal adhesion kinase (FAK)

[19].

FAK is a cytoplasmic tyrosine kinase that localizes FA following integrin clustering and mediates FA dynamic signaling [20-21]. Autophosphorlyation of FAK at several tyrosine residues (Tyr397, Tyr407, Tyr577, Tyr861, and Tyr925) enhances the activity of FAK [17] and allows FAK to associate with the Src tyrosine kinase, triggering downstream signaling events [17-19, 22]. Tyrosine phosphorylation of FAK is crucial to the subsequent phosphorylation of other FA associated proteins such as paxillin [17]. Because

FAK is key to many cellular processes such as adhesion, migration, signaling and cell survival. The ability of a surface to influence FAK expression, localization and activation would in theory provide greater control of cell behavior.

In this study, the ability of nanoscalar surfaces to alter cellular adhesion, morphology, cytoskeletal arrangement and FAK expression, localization and activation was investigated. High resolution scanning electron microscopy and atomic force microscopy were used to characterize the untreated and nanoscalar polyethylene terephthalate (PET) surfaces prior to culture

110 Cells will be cultured on the surfaces and analyzed for general morphology via scanning electron microscope (SEM), actin organization via phalloidin staining and cell- substrate interaction through immunofluorescence and immunoblotting with FAK and phospho-FAK.

4.2 Materials and Methods

4.2.1 Cell culture

Mouse 3T3-L1 preadipocytes obtained from the American Type Culture

Collection (ATCC) were used in this study. Cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS, Gibco), 2 mM L-glutamine, 1mM sodium pyruvate, and penicillin-streptomycin. Cultures were incubated in a chamber with 5% CO2/95% air at

37°C .

4.2.2 Surface Modification via Reactive Ion Etching

Polyethylene terephthalate (PET), Thermanox™ plastic coverslips (Nalgene Nunc

International, Rochester, NY) were used in this study. The thickness and diameter of the

PET films were 200 µm and 13 mm, respectively. For the adhesion studies rectangular

(60 mm x 20 mm) Thermanox™ coverslips were used. PET substrates were modified utilizing reactive ion etching following a method described previously [23] using a

Technics Micro-RIE Series 800-II (Technics, Concord, CA) at 50 W, 160 mTorr with oxygen supplied to the system at a rate of 25 ccm. PET coverslips were etched

111 continuously for 30 min with oxygen plasma then placed upright in a multi-well cell culture plate to prevent subsequent contact of these delicate nanoscalar features with any other surface.

4.2.3 Scanning Electron Microscopy (SEM)

The topography of the etched and non-etched surfaces was qualitatively examined using scanning electron microscopy (FEI Sirion). Both etched and untreated films were sputter coated with a 10 nm layer of gold-palladium and analyzed in ultra high resolution mode with a 5 kV accelerating voltage.

4.2.4 Atomic Force Microscopy

The surface topography of the films was quantified using atomic force microscopy (Digital Instruments Nanoscope IIIa, Veeco Metrology, Tucson, AZ) in wet mode in PBS with a scan rate of 0.500 Hz. Several areas 25 um2 in size were imaged and analyzed with Digital Instruments software version 4.22r2. Topographical roughness parameters Ra (average deviation from arithmetic centerline), Rmax (maximum peak height), and Rq (root mean square deviation of the surface) were determined.

4.2.5 Cell Proliferation and Morphology

Plasma etched and untreated PET substrates were placed in a 24-well cell culture plate then sterilized by immersion in 70% alcohol for 30 min followed by washing with phosphate-buffered saline (PBS) followed by medium. This process was repeated for all

PET disk used for cell culture. Cells were seeded onto sterilized untreated and treated disks at a density of 1.5x104 cells/well and cultured for 7 days. Media was exchanged every other day. On days 1, 2, 3, 5 and 7 cell number was examined with an MTS assay

112 ® (CellTiter 96 AQueous Non-Radioactive Cell Proliferation Assay, Promega, Madison

WI). For each time point duplicate disks were used and experiments were repeated three times. Cell morphology on the surfaces was studied using SEM. Cells on PET disks were fixed with 3% glutaraldehyde in a 0.1M phosphate buffer containing 0.1M sucrose (pH

7.4) for 2 h. They were then washed with the sucrose-phosphate buffer for 30 min. The samples were osmicated in 1% OsO4 for 1 h and washed again. After dehydration in a graded ethanol series, the samples were then subjected to a graded ethanol- hexamethyldisilazane (HMDS, Ted Pella, Redding, CA) series (3:1 ETOH:HMDS to

100% HMDS). The disks were immersed in 100% HMDS and dried overnight by evaporation. Disks were then sputter coated with gold/palladium and examined using

SEM at 5 kV(Sirion, FEI). General morphology was noted; cell size and the extent of cell spreading at each time point was analyzed by categorizing each cell as rounded, spread, or elongated.

4.2.6 Immunocytochemistry

The expression of F-actin and cytoplasmic proteins dephospho- and phospho-FAK were examined by phalloidin staining and immunocytochemistry, respectively. Briefly, untreated and treated PET disks were seeded with cells (1.5x104cells/well) and cultured for 0-72 hours with the zero point considered to be the time at which approximately 30% of the cells had attached to the surface. A “zero hour” time point was taken to analyze cells at the onset of attachment and spreading. This time was determined to be 45 min after initial seeding and all subsequent experiments were performed with the zero point occurring at 45 min. The cell-seeded disks were then rinsed with PBS and fixed with 4%

113 paraformaldehyde in PBS for 1 hr. After fixation, the samples were rinsed gently with

PBS and permeabilized with 0.5% Triton X-100 PBS for 15 min. Samples were pretreated for 1 hr with 5% horse serum (HS) PBS followed by incubation overnight at

4ºC with the primary antibodies of FAK and p-FAK Tyr 397 or FAK and p-FAK Tyr-925 to double-label using anti-FAK and anti-phospho FAK (all 1:40, Santa Cruz

Biotechnology, Santa Cruz, CA)). The disks were thoroughly rinsed with PBS then stained with the appropriate secondary antibody for 1 hour at room temperature. Disks were subsequently incubated with phalloidin (1:40, AlexaFluor® phalloidin, Molecular

Probes, Eugene, OR) for 1 hr at room temperature. After rinsing with PBS, the triple labeled disks were examined via confocal microscopy (Zeiss 510 META). Negative controls were prepared following the same procedures but omitting the primary antibodies.

4.2.7 Western Blotting

Cells from unmodified and modified PET disks at the completion of the experiments were rinsed with PBS and lysed with RIPA buffer (1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, 0.0004% sodium azide in 1X TBS, Santa Cruz

Biotechnology, Santa Cruz, CA) supplemented with 0.1 mM PMSF, aprotinin (10

µg/ml), and 100 mM sodium orthovandate for 60 minutes on ice. Lysates were centrifuged at 10,000 rpm for 10 min and the protein content was analyzed using the

BioRad protein assay (Bio-Rab, Hercules, CA) read in a plate reader (Dynex

Technologies). Protein lysates were then boiled for 2 minutes in SDS-PAGE loading buffer prior to loading 30 µg of protein/lane onto 10% SDS-polyacrylamide gels for

114 electrophoresis. Proteins were transferred to Hybond C Extra™ nitrocellulose membranes (Amersham Biosciences, Little Chalfont, Bucks U.K.) probed with anti-FAK, anti-phospho FAK Tyr 397 and anti-phospho FAK Tyr 925. Briefly, membranes were blocked in 5% Blotto (5 wt% nonfat dry milk 1X TBS-T; 1.54M NaCl, 1 M Tris buffer,

1% Tween-20) for 1 hour prior to overnight 4°C incubation with 1:200 dilution of antibodies against FAK (Santa Cruz Biotechnologies, Santa Barbara, CA), p-FAK (Tyr

925) (Santa Cruz), and 1:1000 dilution of p-FAK (Tyr 397) (Upstate Cell Signaling,

Lake Placid, NY). Following repeated washes with TBS-T, membranes were incubated for 60 min with either bovine anti-goat or bovine anti-rabbit secondary antibodies (Santa

Cruz Biotechnologies, Santa Barbara, CA) corresponding to the primary antibody.

Membranes were washed again and immunoreactivity was visualized using SuperSignal

West Femento Maximum Sensitivity Substrate solution (Pierce, Rockford, IL) in a

VersaDoc imaging system (model 3000, BioRad). Individual band intensity was quantified using Quantity One software version 4.31.

4.2.8 Adhesion Studies

The relative adhesion strength of cells grown on both untreated and treated PET rectangular coverslips was tested using a parallel plate device constructed following

Kawamoto et al.[24] (Figure 4.1). To ensure that this arrangement did not create considerable turbulence, a tracking dye was added to the inlet port prior to initial media flow. The flow of dye was tracked visually and as desired, the flow pattern appeared to be

dominated by laminar flow. For adhesion experiments, cells were seeded at a density of

115

Figure 4.1. Parallel plate device used for adhesion studies. Cells are seeded onto a PET coverslip for 24 hours then placed face down in parallel plate device. Media is flowed though plate producing shear stress on the cells.

116 2.4 x 103 cells/mm2 on the etched and non-etched PET coverslips for 24 hours to ensure that cell confluence would be less than 60%. PET coverslips were then placed into the parallel plate device with the cell side oriented toward the gasket (Figure 4.2) and exposed to a laminar flow of DMEM supplemented with 25 mM HEPES. The apparatus was maintained at 37°C in a water bath. Cells were exposed to shear stress (0-500 dynes/cm2) for 5 min. Following exposure to shear stress the remaining cells were removed from the coverslips with trypsin and counted in a coulter counter. Each experiment was repeated six times and expressed as average + SEM.

4.2.9 Statistical analysis

For cell growth and adhesion experiments, a one-way analysis of variance (ANOVA), followed by the Turkey-Kramer multiple comparison analysis, was performed. The data were presented as either mean + standard deviation or mean + SEM, and p<0.05 was considered significant.

4.2.10 X-ray Photoelectron Spectroscopy (XPS)

XPS analysis was performed on untreated and treated disks after rinsing with PBS followed by distilled water. Disks were analyzed for C, N and O with a mono Al source at 13 kV, 10 mA in hybrid lens mode (AXIS Ultra). Surfaces were analyzed, etched for 1 min with an Ar ion beam, removing approximately 10-15 Å, the analyzed again. Mass concentration percentage is reported

4.3 Results

4.3.1 Oxygen plasma etching generates nanofibrillar surfaces

A shown previously [23], reactive ion etching of PET with oxygen for 30 min

117 produced a nanofibrillar surface topography with fibrils roughly 300 nm in length and 20 nm in diameter (Figure 1A). Upon exposure to water for 24 hrs at 37°C, the surface fibrils became unstable as is evident by the appearance of rafts of fractured nanofibrils and the remaining fibril bases (Figure 1B, C).

To produce thicker, more stable nanostructures, partially cooled structures were used [23,25]. This technique generated shorter (~200 nm) clustered fibrils which provided additional strength (Figure 1D), while retaining its distinctive nanoscalar appearance.

Examination after exposure to media at 37°C for 24 hours revealed that the bonded nanofibers were stable and neither raft formation nor fractured fibrils were observed. This surface was then characterized and quantified under PBS utilizing wet mode AFM. The average roughness, Ra, of the thermally bonded etched surface was 51 + 6 nm and the maximum peak height was 402 + 156 nm (Figure 2) compared to an Ra of 3.5 + 0.8 nm for the untreated surfaces.

XPS analysis revealed a slight change in chemical composition between the two surfaces (Table 4.1), with the etched surfaces becoming enriched with nitrogen following an oxygen plasma etch. This fortification with nitrogen comes at an expense to oxygen content in the sample.

4.3.2 Proliferation

To asses growth on untreated and treated PET, non-radioactive MTS assays were performed at 1-7 days after plating cells. We detected a statistically significant difference in 3T3-L1 growth on unmodified and modified surfaces for day 2 (p < 0.001) and day 5

(p < 0.05) only, with cell number on the unmodified surface greater than the modified

118

Figure 4.2. SEM images of the PET coverslip surface A) after etching for 30 minutes in oxygen plasma B) after etching followed by soaking in media at 37°C for 24 hours (low magnification) C) after etching followed by soaking in media at 37°C for 24 hours (high magnification) D) after etching to produce thermal bonding followed by soaking in media at 37°C for 24 hours (low magnification). The latter structure was used for all subsequent experiments. Small ‘rafts’ of broken fibrils are evident after the exposure producing (C).

119

Figure 4.3. Wet mode AFM conducted in PBS. A) 2D and B) 3D images of the thermally bonded PET structure.

120

Figure 4.4. MTS assay of 3T3-L1 cells grown on both as-is and plasma etched PET. Data presented on mean + standard deviation

121 surface (Figure 3). The differences in cell number were not substantial, however, and were not significant for all time points suggesting that the differences in proliferation found on day 2 and 5 were not due to treatment rather to another source of error.

4.3.3 Nanofeatures alter cell morphology and the actin cytoskeleton

Cell morphology was evaluated via SEM at time points ranging from 0-72 hrs.

Cells were grouped categorically as either round (spherical shape), spread onto substrate

(cell has spread but retains equi-axed shape) and elongated (cells has spread onto surface and is elongated into fibroblastic configuration). After 4 hrs in culture 78 + 21% of the cells on smooth surfaces were rounded in appearance whereas only 6.9 + 6.7% of cells on the nanoscalar surfaces remained rounded. By 24 hrs in vitro, all cells had spread onto the growth surface and elongated. When grown on nanoscalar surfaces, fibroblasts appear to be slightly larger (Figure 4A & B) and extend a greater number of nanoscalar lamellapodia to the surface than cells on untreated substrates (Figure 4D, E). These lamellapodia also appear to subdivide and ‘grab’ onto the nanofibrils (Figure 4E). A lamellapodium was followed out to its entire length to track it interaction with the naofibrils (Figure 4E). As shown in the inset, the initial lamellapodium extends a number of much smaller filapodia from its original “stem”. These minute filapodia appear to interact directly with the nanofibrils (Figure 4E arrowheads). Attempts to visualize these interfacial structures using TEM proven technically challenging. However, our initial data indicate the very fine tips of the lamellapodial extensions turn downward and appear to contact the tips of the nanofibrils (Figure 5).

122

Mass Conc Mass Conc UT % E %

O 39.14 T 39.17 N 0.97 C 0.63 C 59.89 H 60.2

Mass Conc Mass Conc T % E % O 33.17 T 28.05 N 4.68 C 3.69 C 62.15 H 68.26

Table 4.1. Chemical composition of untreated and treated surfaces via XPS on sample surface and 15 angstroms below the surface.

123

A B

C D

Figure 4.5. SEM micrographs of fibroblasts grown on control (A&B) and treated (C&D) disks. E.) At higher magnifications the lamellapodia is ‘grabbing’ onto the nanofibrils via generation of successively smaller and smaller extensions.

124

A B

C D

Figure 4.6. F-actin staining of cells grown on untreated disks at A) 0 hrs and B) 24hrs. C) treated 0 hrs D) treated 24 hrs. Scale bars = 50 µm

125

A B C

FAK 10µm p-FAK 397 10µm F-actin 10µm D E F FAK p-FAK 397 F-actin

5µm 5µm 5µm G H I

FAK 20µm p-FAK 925 20µm F-actin 20µm

J K L

FAK 20µm p-FAK 925 20µm F-actin 20µm

Figure 4.7. Confocal images of cells seeded onto untreated and treated substrates. 3T3L-1 cells grown on unetched substrates at the zero time point stained for A) FAK B) P-FAK 397 and C) F-actin. At 0hrs, cells on nanofibrillar surfaces exhibit less punctuate FAK (D) and p-FAK 397 (E) at the periphery with a disorganized actin cytoskeleton (F). After 72 hours in culture FAK (G), P-FAK 397 (H) distribution on smooth surfaces is similar to that on treated surfaces (J and K respectively). On the contrary, F-actin has formed stress fibers on the untreated sufaces (I) while none can be seen on the nanoscale substrated (L). 126 Using phalloidin-NBD-FITC as a marker for F-actin, we detected dramatic differences in the organization of the actin cytoskeleton on cells grown on etched surfaces compared to cells seeded onto untreated PET disks. Cells on treated and untreated surfaces formed a cortical ring of actin as the cells began to attach to the substrate as evidenced from actin staining at 0 hrs (Figure 6 A, C). As the cells began to spread on the unmodified surface, the actin cytoskeleton organized into a network of well-defined filaments, stress fibers (Figure 6B). In contrast, cells grown on treated substrates exhibited far less F-actin organization and did not contain visible stress fibers.

By 24 hours in vivo the 3T3-L1 cells seeded on modified surfaces had no evidence of filamentous actin and or stress fibers (Figure 6D).

4.3.4 Substrate nanofibrils alter FAK expression and activation

The expression and localization FAK and phospho-FAK was determined via immunocytochemistry on cells seeded for 0-72 hrs on both untreated and nanofibrillar surfaces. The localization of dephospho- and phospho-FAK on tyrosines 397 and 925 appear to be more concentrated on the periphery of the cell at low time points on the untreated substrate compared to the treated substrates (Figure 7 A,B & D&E). However by 24 and 72 hrs the expression of FAK and p-FAK is quite similar (Figure 7 G,H &

J,K).

127

A. B.

Figure 4.8. Densitometry of western blots performed for A) FAK B) p-FAK 397 and C) p-FAK 925. Blots were repeated for three different experiements with similar trends found in all experiments. Note FAK epression is similar in cells on treated and untreated surfaces and does not vary dramatically with time. On the contrary phosphor-FAK on both tyrosine residues (397 and 925) is slightly greater in cells on untreated surfaces and increases with increasing culture duration.

128 While the localization and intensity of the FAK and p-FAK appear to be similar in

ICC, western blot analyses were performed to determine the relative amounts of FAK, p-

FAK 397 and p-FAK 925. Overall FAK intensity in immunoblots was similar regardless of substrate treatment and time whereas both phospho-FAK on both tyrosine increases with time(Figure 8 A, B, and C). Densitometry of the western blots revealed that there is a slightly greater proportion of the phosphor-FAK on the 397 tyrosine residue within cells grown on smooth versus rough surfaces. Due to technical issues each graph represents densitometry from one set of blots. Each experiment was repeated three times with similar trends seen in each experiment.

4.3.5 Substrate etching alters relative strength of adhesion

Having established that FAK autophosphorylation and actin cytoskeletal organization into stress fibers were diminished when cells were grown on etched substrates, we next examined the relative strength of cell-polymer adhesion. To accomplish this, we used a parallel plate device to establish a shear stress across a non- confluent layer of 3T3-L1 cells on untreated and etched substrates. Quantification of the cells remaining on substrates after exposure to shear stress revealed that there is a trend of decreasing cells remaining as shear stress increased on both surfaces. Average percentage of cells remaining on unmodified surfaces is significantly greater than on modified surface for all time points (Figure 4.9).

129

Figure 4.9. Relative strength of adhesion of cells grown on treated and untreated surfaces for 24 hours prior to exposure to a given shear stress for 5 minutes. Data reported as mean + SEM

130 4.4 Discussion

Comparison of cells grown on these surfaces might lead to the conclusion that the actin cytoskeleton of the cells on “thermally bonded” nanofibrillar surfaces has been disrupted. It is important to consider if this disruption was caused by initial surface interference, never allowing stress fibers to form, or if stress fibers were able to form but were later disassembled. The disruption of the actin cytoskeleton and disassembly of stress fibers can be produced with the addition of various drugs to the culture media.

Mermelstein et al used a retinol treatment to induce active reorganization of the F-actin in hepatic stellate cells [26]. The major stress fibers fragmented and generated diffuse or granular actin in the perinuclear area. The behavior we see is dissimilar in that there were no observations of either a polygonal meshwork, large clumps in the perinuclear area, solid spheres or larger spheres that do not retain actin in their center. The cells on treated surfaces at no time point contain stress fiber fragments or large lumps of actin. Along with the relatively short-term observations of these cells this data suggests that stress fibers never form in these cells. Thus an earlier cellular process is likely responsible for the disruption of the actin network. or large lumps of actin. Along with the relatively short-term observations of these cells this data suggests that stress fibers never form in these cells. Thus an earlier cellular process is likely responsible for the disruption of the actin network.

In formulating mechanistic insights and general principles regarding the influence of nanoscalar topographies, it is important to compare the scale of the nanofibrials to that of the cell itself. The width of these nanofibrils is approximately 1/500 of the cell

131 diameter or about 15 integrins wide, thus only a finite number of integrins can be bound to the surface of the fibril. In addition, the density of the nanofibrils likely affects the spacing of the integrin ligands. From the TEM data (Figure 5), the cells appear to only attach at the tips of the fibrils, the integrin clusters will be spaced according to the density of the nanofibrils. From this it is easy to realize that specific cell morphologies- e.g. cellular alignment with specific nanofibrils- are impossible when the differences in size of the cell and the nanoscale are so large that each cell sits above many individual nanofibrils. These data indicates, however, a fundamental influence of contact by these nanoscalar features on cellular behavior.

Initial cell adhesion and subsequently cytoskeletal organization is heavily dependent on integrin-surface ligand binding [17,18, 27,28]. Previous work by Massia and

Hubbell [29] observed that at low surface ligand densities, cells attach but do not spread.

At higher densities, when the spacing between peptides reaches ~440 nm, cells attach and spread but do not form focal adhesions. However, at still higher densities of RGD peptides, with a spacing of ~140 nm or less, spreading is accompanied by development of focal adhesions and stress fibers [29]. Many more modern observations of the same effect in the field of RGD clustering exist [30,31]; however, this first observation of the need for a high density of integrin ligand interactions proved that a critical number of interacting integrins must be clustered for visible FAs and stress fibers to form. By adapting this observation and an existing model relating integrin assembly to stress fiber development

[19], we can begin to explain the observed effects of these nanoscalar surfaces on cell behavior, specifically the inhibition of stress fiber formation. Figure 4.10 (adapted from

132 [19]) shows a model for cellular attachment to both nanofibrillar and untreated surfaces.

The spacing of the nanofibers is approximately 200 nm, a distance greater than the 140 nm ligand density seen to facilitate focal adhesion formation. This low ligand density could, to some extent, inhibit the formation of focal adhesions and the ensuing generation of stress fibers (Figure 4.10b) [19]. Due to the fact that focal adhesion kinase is seen via

ICC in cells grown on both surfaces, the prospect of nanofibril spacing completely hindering focal adhesion formation is doubtful. It is probable that focal adhesions form on both surfaces with a spatial inhibition of large scale focal adhesion interaction on nanofibrillar surfaces. Therefore the absence of a highly organized F-actin cytoskeleton may be linked to the spatial separation of integrin cluster (Figure 4.10a) which could prevent groups of integrins from clustering and subsequently organizing bundles of F- actin together to form stress fibers.

In addition to influencing the formation of stress fibers, this nanoscalar surface promotes very slight alterations in the amount of FAK present and the phosphorylation of

FAK at tyrosine residues 397 and 925. Complete disruption of the actin cytoskeleton has been observed to inhibit the phosphorylation of FAK[32, 33] as well as various downstream signaling events [33]. When cells are plated, inhibitors of contractility can either block the formation of FAs or result in very small FAs that have not consolidated into their typical organized state [19]. For example, the disruption of the actin cytoskeleton by treatment with cytochalasin D completely has been observed to inhibit the phosphorylation of FAK as well as various downstream signaling events. From the discussion, total disruption of the actin cytoskeleton and focal adhesion formation should prevent FAK activation, and

133 would promote loss of adhesion. In contrast, the physical adhesion data shows that cells cultured on these nanostructured maintain approximately 50% of the untreated anchorage, suggesting that anchorage is weaker but is certainly not absent. In fact, the percentage of adherent cells beings to merge at the highest levels of shear stress. In addition, total disruption of the actin cytoskeleton blocks integrin aggregation and leads to a measurable inhibition of FAK activity. Given the very modest effects we observed on FAK, p-FAK and proliferation, the nanoscale clearly does not disrupt the formation of

F-actin rather it inhibits the close association of F-actin into bundles or stress fibers.

Alterations in actin organization and focal adhesion formation have been linked to observed changes on cell adhesion and migration. Integrin-linked focal adhesion complexes provide the main sites of cell adhesion to extracellular matrix and associate with the actin cytoskeleton to control cell shape and movement. Dynamic regulation of focal adhesions and reorganization of the associated actin cytoskeleton are crucial determinants of cell motility [34,17-19]. The mechanics of cell movement depends on cell- substrate adhesion, remodeling of the actin cytoskeleton and cell-substrate detachment [17-

19, 28]. These integrin-linked focal adhesions or focal complexes are the primary sites of adhesion between cells and the surrounding ECM. The dynamic regulation of their assembly and disassembly is therefore an important determinant of how quickly cells can move. Thus it is feasible to reason that the absence of stress fibers and lower expression of FAK and FAK autophosphorylation results in weaker cell adhesion and more migratory cells. On nanofibrillar substrates we observed a decrease in activation of FAK and assembly of the actin cytoskeletal along with a reduction in relative adhesion

134 compared to cells plated on untreated surface. Accordingly, cells on treated surfaces also appear to be more migratory as evidenced from the presence of “migratory ruffles” as seen by Kapur and Rudolph (Figure 11) [34]. Thus the alterations in cell behavior induced by surface roughness have been shown to affect many functions from initial adhesion to the organization of the actin cytoskeleton.

To our knowledge, this is the first observation of nanostructural instability in this context. Brownian motion of the individual nanofibrils likely leads to oscillation and then localized agglomeration followed by fracture, producing the nanofibril ‘rafts’ visible on the SEM images even before they are influenced by cell-surface interactions. This is an unexpected consequence of attempting to make fully synthetic surface adopts a more biological appearance having realistic length scales.

In reviewing these results, we observed striking similarities to those of recent reports in which cellular behavior was explained on the basis of mechanical interaction with the underlying substrate surface [35-37]. One might treat the nanoscaled surface as a

‘soft’ layer intervening between the cell and the relatively ‘hard’ non-nanoscaled PET below; verification would require exacting measurements of the mechanical properties of this discontinuous, nanoscalar layer under physiologically relevant conditions.

Unfortunately nanoscalar characterization of these soft/hard surfaces has not been carried out in these cellular-based investigations [35-37]. Motile 3T3 fibroblasts [35] and vascular smooth muscle cells [36] show an apparent ability to detect and respond to gradients in bis-acrylamide cross-linker concentration/hardness in polyacrylamide gels. However,

Suzuki et al [38] previously showed that under aqueous conditions, AFM indicates that

135 these acrylamide gel surfaces consist of “undulations on the order of a few micrometers, large holes and projections” similar to what we observe for plasma treated PET (Figure

2). They also show an increase in bis-acrylamide concentration produces a considerably smoother surface and an increase in height of the average topographical feature with increases in temperature (from 25 to 38 ºC) [38]. Chondrocytes cultured on alignate gels of varying stiffness have been shown to alter cell behavior with cells attaching more rapidly on stiffer surfaces [39]. A recent review of AFM performed on polysaccaride gels finds that all polysaccharide gels have complex nanoscalar structures [40]. As a substrate for studying cell-susbtrate interactions independent of topography, both of these gel systems are far from featureless thus it is likely that the surface morphology also played a role in altering cell function/

These intriguing observations suggest that rather than utilizing mechanical properties to explain of our results, the mechanism in Figure 4.10 provides a compelling explanation of how the presence of nanoscalar topography can interact with intracellular and extracellular processes.

It is important to note that our focus has been limited to the nanoscale. Larger scale (i.e. micron-sized) features would not be as dimensionally matched; the cells could

136

Figure 4.10. Model for integrin-ligand binding on smooth versus nanofibrillar surfaces. A) on nanofibrillar surfaces surface ligands are present both on tips of fibrils and in valleys, however cells are believed to bind only at tips of fibrils where ligands are spatially restricted. F-actin forms but numerous F- actin fibers cannot cluster due to nanofibril density. B) On the contrary, surface ligands on smooth surfaces are able to promote formation of F-actin along with F-actin bundling into stress fibers

137

FAK p-FAK 397

F-actin Merged

Figure 4.11. Fluorescence image of a cell grown on a nanofibrillar surface for 24 hours. Note the ruffled leading edge of the cells (white arrow). These structures have been seen previously [Kapur and Rudolph] associated with fibroblasts that were migratory.

138 easily gather sufficient integrins via internal contraction to form large focal adhesions on either side of a micron-sized feature. Whereas, smaller features have been shown to have little effect on the actin cytoskeleton [41].

4.5 Conclusions

Nanoscalar surfaces can clearly alter actin cytoskeleton organization and influence cell-substrate signaling and adhesion. These results advance the possibility that the nanoscale can be used to alter cell behavior in a predictable manner on a given substrate without a considerable change in chemical composition. Such ability provides the opportunity to modify tissue engineering scaffolds in a manner which can tailor subsequent levels of cell adhesion, cell shape and even phenotype. These advances in understanding the cell-material interaction have future implications on adhering, maintaining, and differentiation of cells in vivo.

Acknowledgements

HMP would like to thank the National Defense Science and Engineering Graduate

Fellowship for financial support. The authors would also like to the Campus Electron

Optics Facility, Campus Microscopy and Imaging Facility and the Ohio MicroMD

Laboratory at the Ohio State University for use of their facilities.

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143

CHAPTER 5

NANOTOPOGRAPHY ALTERS PREADIPOCYTE DIFFERENTIATION

5.1 Introduction

The interaction between a cell and its substrate regulates a complex milieu of cellular events from initial adhesion to cell proliferation, differentiation and apoptosis.

The concept that surfaces affect cell function has been recognized for many years [1]. Of recent debate is the role nanoscalar features play in altering cell behavior. Fibroblasts grown on 13-nm high polymer demixed islands have been reported to up-regulate proliferation, cell signaling and extracellular matrix formation compared to smooth counterparts [2]. In contrast, Vance et al. observed a significant decrease in fibroblast density on nanoscalar PLGA, PU and PCL versus their unaltered controls. Nanoscale grooved surfaces have also been shown to possess the ability to orient cells, however this contact guidance was largely dependent on cell type and cell-cell interactions [3]. In addition, previous studies show disruption of the cytoskeleton in fibroblasts grown on nanoscale surfaces [4]. Although nanoscalar surfaces have been extensively studied and shown to alter cell-substrate adhesion and cytoskeletal organization, few reports investigate the downstream consequences of such alterations on cell function[2,5].

144 Primary contact of a cell to a surface initiates cell adhesion which is essential in all aspects of development. The process of adhesion is tightly linked to a complex web of signaling pathways leading to cell division, cell, migration, differentiation and apoptosis.

Thus alterations in initial adhesion caused by surface topography may lead to differences in cell-substrate signaling and ultimately differentiation. For example, Schnieder et al have seen significant increases in osteoblast differentiation on rough implants [5]. In addition formation and organization of the cytoskeleton has been shown to play an essential role in signal transduction and differentiation [6-9]. McBeath et al. demonstrated that the disruption of the cytoskeleton and reduction in cell tension via cytochalasin D lead to increased adipogenesis in human mesenchymal stem cells [10]. Previous work in our lab suggests nanofibrillar surfaces trigger a disruption of the actin cytoskeleton, decreased strength of adhesion and reduced levels of cell-substrate signaling versus non- treated substrates [4]. These alterations in early cell function may cause significant downstream effects on cell behavior.

In this study, the ability of nanoscalar surfaces to alter the differentiation of 3T3-

L1 pre-adipocytes into adipocytes and production of cAMP in response to β-adrenergic stimulation was investigated. High resolution scanning electron microscopy and atomic force microscopy were used to characterize the untreated and nanoscalar surfaces prior to culture as well. Cells will be cultured on the surfaces and monitored for cytoskeletal re- arrangement and lipid body formation via phalloidin and bodipy staining respectively along with presence of PPAR-γ via immunocytochemistry (ICC) and Western blotting.

145 5.2 Materials and Methods

5.2.1 Cell Culture and Differentiation

Mouse 3T3-L1 preadipocytes obtained from the American Type Culture

37°C .

For differentiation experiments, plasma etched and untreated PET substrates were placed in a 24-well cell culture plate then sterilized by immersion in 70% EtOH/ 30%

H2O for 30 min followed by washing with medium. Each well containing a PET substrate

4 was seeded with 1.5 X 10 cells/well and incubated in a humidified 5% CO2−95% air incubator at 37 °C for 24hrs. After 24 hours, differentiation was induced with 10 µg/ml insulin (Life Technologies, Grand Islands, NY), 1 µM dexamethasone (Sigma, St. Louis,

MO) and 500 µM isobutyl-methylxanthine (IBMX) (Sigma, St. Louis, MO) in DMEM containing 10% FBS (IDX mixture). After two days of differentiation, cells were incubated in DMEM containing 10% FBS and replaced every other day. At days 2-30 after differentiation, a portion of the cell-seeded disks were fixed for ICC while the remaining cells were lysed for western blot analysis. Each experiment was done in no less than triplicate and repeated at least once.

146 5.2.2 Surface Modification via Reactive Ion Etching

Polyethylene terephthalate (PET), Thermanox™ plastic coverslips (Nalgene Nunc

International, Rochester, NY) were used in this study. The thickness and diameter of the

PET films were 13 mm and 200 µm respectively. PET substrates were modified utilizing a thermally bonded reactive ion etching technique, following a method described previously [11] using a Technics Micro-RIE Series 800-II (Technics, Concord, CA) at 50

W, 160 mTorr with oxygen supplied to the system at a rate of 25 ccm. Thermally bonded

PET coverslips were etched continuously for 30 min with oxygen plasma then placed upright in a multi-well cell culture plate to prevent subsequent contact of the nanoscalar features with any other surface. Surfaces were analyzed via scanning electron microscopy and shown to have the same morphology as previously reported (Figure 5.1)

5.2.3 Immunocytochemistry

Re-arrangement of the actin cytoskeletal and lipid body formation were imaged via phalloidin and bodipy staining respectively while the presence and localization of

PPAR-γ was determined using immunocytochemistry (ICC). Briefly, untreated and treated PET disks were seeded with cells (1.5x104cells/well) and cultured for 1 day after initial seeding and 2-30 days after differentiation. The cell-seeded disks were then rinsed with PBS and fixed with 4% paraformaldehyde in PBS for 1 hr. After fixation, the samples were rinsed gently with PBS and permeabilized with 0.5% Triton X-100 PBS for

147

Figure 5.1. SEM of thermally bonded nanofibrillar surface

148 15 min. Samples were pretreated for 1 hr with 5% horse serum (HS) PBS followed by incubation overnight at 4ºC with the primary antibody for PPAR- γ (Santa Cruz

Biotechnology, Santa Cruz CA) and phalloidin (AlexaFluor phalloidin, Molecular

Probes, Eugene, OR). The disks were subsequently probed for lipid with a BODIPY neutral lipid stain (Molecular Probes, Eugene, OR) Disks were thoroughly rinsed with

PBS then stained with the appropriate secondary antibody for 1 hour at room temperature. After rinsing with PBS, the triple labeled disks were examined via confocal microscopy (Zeiss 510 META). Negative controls were prepared following the same procedures but omitting the primary antibody.

5.2.4 Image Analysis

Lipid droplet size was evaluated via image analysis (Image J Software) on treated and untreated surfaces as a function of time and surface treatment. For each sample set no less than 3,000 individual lipid droplets were analyzed. Data is reported area of lipid versus total area. Area rather than diameter was selected as the output parameter due to the deviation in lipid droplet shape from roughly spherical to elongated globules as time progressed.

5.2.5 Western Blotting

Biotechnology, Santa Cruz, CA) supplemented with 10 ρg/ml PMSF, aprotinin

(10µg/ml), and 100 mM sodium orthovandate for 60 minutes on ice. Lysates were

149 centrifuged at 10,000 rpm for 10 min and the supernant was analyzed using the BioRad protein assay (Bio-Rab, Hercules, CA) read at 570 nm in a plate reader (Dynex

Technologies). Protein lysates were then boiled for 2 minutes in SDS-PAGE loading buffer prior to loading 30 µg of protein from each sample into a 12% SDS- polyacrylamide gel for electrophoresis. Proteins were transferred to Hybond C Extra nitrocellulose membranes (Amersham Biosciences, Little Chalfont, Bucks U.K.) were immunoassayed using an immunodetection protocol previously described. Briefly, membranes were blocked in 5% Blotto (5 wt% nonfat dry milk 1X T/TBS; 1.54M NaCl,

1 M Tris buffer, 1 vol/wt% Tween-20) for 1 hour prior to overnight 4°C incubation with

1:200 dilution of an antibody against PPAR-γ (Santa Cruz Biotechnologies, Santa

Barbara, CA). Following repeated washes with 1X T/TBS, membranes were incubated for 60 minutes with bovine anti-mouse IgG1 secondary antibody (Santa Cruz

Biotechnologies, Santa Barbara, CA). Membranes were washed again and immunoreactivity was visualized using SuperSignal West Femento Maximum Sensitivity

Substrate solution (Pierce, Rockford, IL) in a VersaDoc imaging system (model 3000,

BioRad). Individual band intensity was quantified using Quantity One software version

4.31.

5.2.6 Cyclic AMP (cAMP) Production

The extracellular presence of cAMP was evaluated in cells cultured on treated and untreated substrates which were incubated for 8 days after differentiation and in a control set of cells at the same time points which had not been differentiated. At the end of the culture period, cells were exposed 10µM IBMX for 30 min prior to drug addition to

150 block cAMP breakdown. Isoproteranol (Iso), norepinephrine (Nor), prostaglandin E2

(PGE2) and ethanol (Et) as a control were then added to the media at a 1 µM concentration for 30 minutes. Subsequently media was removed and analyzed with a cAMP EIA kit (Cayman Chemicals, Ann Arbor, MI). A protein analysis was performed on each cell seeded disk to report levels of cAMP as pmol/mg of protein. Each experiment was repeated three times.

5.2.7. Statistical analysis

For PPAR-γ densitometry and cAMP production, a one-way analysis of variance

(ANOVA), followed by the Tukey-Kramer multiple comparison analysis, was performed.

The data were presented as the mean and standard deviation, and p<0.05 was considered significant.

5.3 Results

5.3.1 Cytoskeletal Organization

As reported previously, the organization of F-actin cytoskeleton was substantially altered by nanoscale roughness after one day in culture[4]. On smooth surfaces, well formed stress fibers are generated while no stress fibers can be seen in cells cultured on nanofibrillar surfaces after 24 hrs in culture (Figure 5.2). Two days after initial differentiation typical fibroblastic shape was retained on both surfaces with stress fibers still seen only within cells on untreated surfaces (Figure 5.3a). As time progresses a proportion of cells begin to take on a more rounded shape with a thin bright line of F-

151 a.

a. b.

Figure 5.2 Phalloidin F-actin stain of cells grown on a) untreated and b) treated surfaces for 24 hrs. Note lack of stress fiber generation of treated surfaces

152

a. b.

c. d.

e. f.

g. h.

Figure 5.3 F-actin staining for cells on a) untreated and b) treated surfaces for 2 days c) untreated and d) treated substrates for 4 days, e) untreated and f) treated surfaces for 6 days and g) untreated and h) treated surfaces for 8 days

153

a. b.

c. d.

e. f.

Figure 5.4 F-actin staining of cells grown on a) untreated and b) treated surfaces for 15 days, c) & d) for 21 days and e) & f) for 30 days.

154 b.

a. b.

c. d.

e. f.

g. h.

Figure 5.5 PPAR-γ (red), neutral lipid (green) and F-actin staining (pink) of cells grown on a) untreated and b) treated surfaces for 2 days, c) & d) for

6 days, e) & f) for 15 days and g) & h) 21 days.

155 actin surrounding each round cell (5.3 b & c). With time the percentage of rounded cells increase until a majority of cells are circular (Figures 5.3 & 5.4). Eventually the cells assume a typical adipocyte morphology; large rounded cells with cage-like F-actin structures surrounding the lipid droplets (Figure 5.4 a-c). This phenomenon was seen in cells on both surfaces with no substantial differences in the process.

5.3.2 Lipid droplet formation and coalescence

The appearance, size and coalescence of lipid droplets was analyzed via BODIPY staining as an indication of the differentiation state of the cell. At early time points, 2-6 days after induction of differentiation, lipid droplets are extremely small and relatively sparse. The localization and relative amount of lipid droplets within cells on both treated and untreated surfaces is strikingly similar at these time points (Figure 5.5). By day 8, larger lipid droplets appear which appear to surround the nucleus. On untreated surfaces, the “large” lipid droplets seen in days 6 and 8 seem to coalescence into massive lipid globules at days 15 and higher (Figure 5.5 a, c, & e). However on the treated surfaces this coalescence appears to be impeded. On these surfaces two populations of lipid droplet sizes emerge; a small population of larger droplets with a more numerous amount of undersized droplets (Figure 5.5 b, d, & f). This observation was quantified by image analysis and held to be true (Figure 5.6a&b). Treated surfaces have a lower percentage of large droplets compared to smooth surfaces (Figure 5.6 a&b) yet the total amount of neutral lipid is comparable on both surfaces.

156

(µm)

Figure 5.6. Distribution of lipid droplet size on untreated versus treated surfaces.

157

Figure 5.7 Production of cAMP in differentiated cells cultured on untreated and treated substrates for 8 days.

158 5.3.3 PPAR-γ Expression

PPAR-γ, a functional nuclear receptor commonly associated with adipogenesis

[13], was probed for via both ICC and immunoblotting. In ICC, PPAR- γ was not detected at 2 days (Figure 5.5 a & b) and only a small number of cells on days 4-6 after differentiation expressed PPAR- γ (Figure 5.5 c-f). Cells on both surfaces at day 8-30 showed an increasing amount of PPAR- γ within the nucleus with no detectable difference in intensity seen between surfaces. Western blot analysis of PPAR- γ also indicated similar levels of the receptor on both surfaces with an increasing amount of

PPAR- γ as culture time increased.

5.3.4 cAMP Production

To assess cell metabolism on etched and untreated surfaces, non-differentiated and differentiated 3T3-L1 cells were exposed to either a dose of norepinephrine (Nor), isoproteranol (Iso), prostaglandin E2 (PGE2) or ethanol (Et) as a control. From Figure 5.7, one can see a significant increase in cAMP production on response to exposure to Nor on untreated surfaces while on treated surfaces this drug had only a small increase in cAMP production over the control. Conversely, the addition of Iso to each well produced the opposite effect. cAMP was increased dramatically on treated surfaces while it only generated a 4-fold increase in cAMP compared to control on smooth surfaces. These results are surprising and due to undetectable readings on several samples the experiment will be performed again to obtain a n > 3 for each sample.

159

5.3.6 .Protein Coatings on Lipid Bodies

The surface of lipids bodies are coated by several proteins, many of which are thought the play a role in lipolysis and other enzymatic processes [14-17]. Western blots were performed to asses if there were quantitative difference in the expression of two proteins which commonly coat the surface of lipid bodies; adipophilin which is associates with small lipid droplets and perilipin which coats the surface of large lipid droplets [16].

From these analyses we have determined that adipophilin expression increases with time reaching a relative maximum at 15 days (Figure 5.8) and no difference in adipophilin expression between smooth and nanoscalar surfaces. Perilipin western blots have similar trends with little to no perilipin expression prior to the 8 day time point followed by a temporal increase in expression with culture duration (data not shown).

5.4 Discussion

In these experiments, surface treatment has not altered the total volume of lipid droplets accumulated in differentiating preadipocytes nor did it influence the quantity of

PPAR- γ within the nucleus. However, alterations in lipid droplet size distribution were detected. In vivo white adipose tissue is largely comprised of large lipid droplets where as brown adipose tissue contains vast amounts of small lipid droplets. These two types of adipose tissue play different regulatory roles in the body thus knowledge of the mechanisms by which lipid droplet size distributions arise is of great importance.

160

30 D T

30D UT

21D T

21D UT

15D T

15D UT

8D T

8D UT

6D T

6D UT

4D T

4D UT

2D T

2D UT

Figure 5.8 Western blot of ADRP on untreated and treated surfaces.

161

During differentiation there is a marked change in cellular morphology from a flat fibroblastic shape to a nearly spherical shape [18, 19]. This change is a programmed event not just a passive response to lipid accumulation as evidences by the shape change that occurs even when lipid accumulation is blocked in cells [20]. Adipose conversion is also associated with a decrease in the level of actin from 12.7% of total soluble protein to

3.2% and the synthesis of tubulin is decreased by over 95% [18,21]. These changes in cytoskeletal protein synthesis occur very early in differentiation. This decrease in actin production is also correlated with a loss of thick actin filaments [21]. Early decreases in synthesis of cytoskeletal protein suggest cytoskeleton disassembly influences subsequent biosynthesic events specific to adipocytes differentiation [21]. This is of particular importance in our studies because this type of nanotopography has been previously shown to alter the actin cytoskeleton. Specifically nanotopography has inhibited the formation is stress fibers. During adipocyte differentiation actin synthesis decrease and the actin cytoskeleton disassembles [18,21]. This would lead one to believe that cells with a previously disrupted cytoskeleton, as in cells grown in nanoscalar surfaces, would differentiate more readily. However this trend was not seen with preadipocytes grown in nanofibrillar surfaces possessing indistinguishable amounts of PPAR-γ when compared to cells on control surfaces.

Lieber and Evans suggest a more physical role of the cytoskeleton in adipocytes differentiation [12]. They have suggested that intermediate filaments form cage-like structures around lipid droplets stabilizing them and protecting them from lipolysis. A

162 disruption in vimentin organization was shown to significantly decrease the lipid-droplet forming capacity of 3T3-L1 cells [12]. This disruption in cytoskeleton stability inhibits lipid droplet accumulation. Thus it is unlikely that changes in initial cytoskeletal arrangement alone lead to difference in lipid droplet size. It is more probably that an array of biological events leads to this change.

Lipid droplets are coated with several different proteins that are thought to play a regulatory role in lipid droplet formation and lipolysis. Adipose differentiation related protein (ADRP, adipophilin in humans) is a 50 kDa fatty acid binding protein which coats the surface of newly formed lipid droplets and is part of the perilipin family of lipid storage proteins [15]. ADRP is transcriptionally activated when preadipocytes differentiate into mature adipocytes. The accumulation of lipid storage droplets is correlated with amount and localization of ADRP in mice [15]. In human macrophages, adipophilin played a role in lipid accumulation [14]. Overexpression in adipophilin induces significant increases in triglycerides whereas inhibition of endogenous adipophilin expression dramtically reduced lipid accumulation. ADRP overexpression stimulated lipid accumulation and lipid droplet formation without inducing other adipocytes specific genes. Adipophilin contributes to further lipid accumulation by inhibiting cellular cholesterol efflux. [14]

Other members of the perilipin family encase intracellular neutral lipid droplets in adipocytes. Specifically perilipin has been shown to coat lipid droplets and is thought to play a regulatory role in triacylglycerol hydrolysis [16]. To determine the role of perilipin in adipose differentiation, studies with perilipin knockout mice have been performed.

163 Knockout mice possessed elevated basal lypolysis because of loss of protective coating.

Perilipin -/- mice had greater lean body mass and elevated metabolic rate. Their adipose tissue was large comprised of many small lipid droplets coated by ADRP rather than perilipin. Thus perilipin coated droplets are characteristically large which ADRP-coated neutral lipids are smaller in size. [16]

These previous studies would indicate that there should be a higher proportion of perilipin in cells grown on untreated surfaces as their lipid droplets tend to be larger while cells grown on treated surfaces should have both ADRP and perilipin because there is both a population of small and large lipid droplets within those cells. Evaluation of these two proteins with western blot analysis does not reveal this trend. A possible explaination for this phenomenon would be the extremely large number of lipid droplets samples in western blot analyses tend to homogenize the date thus the relative amounts of these proteins appear to be equivalent. Currently we are investigating the coatings on individual droplets via ICC to determine is in fact the small droplets on the treated surfaces are coated with ADRP and the large are encased by perilipin.

5.5 Conclusions

The alteration in lipid droplet size distribution on these surfaces suggest that small changes in initial cell function on nanoscalar surfaces have a downstream effect on late stage cellular processes such as differentiation. It is clear that the mechanisms of lipid droplet coalescence are not well known. From these studies it has been shown that cytoskeletal disruption does not inhibit adipocyte conversion when a biochemical stimulus is present. It would be of interest to determine if adipogenesis caused by

164 endogenous signals such as cell confluence is altered by this disruption in actin organization. In addition the biochemical similarity of these lipid droplets must be known to elucidate the mechanism which allows droplets to coalesce on smooth surfaces while it is inhibited on treated surfaces.

165 References

1. Weiss, P. (1934). In vitro experiments on the Factors Determining the course of the outgrowing Nerve Fiber. Journal of Experimental Zoology 68, 393-448.

2. Dalby, M. J., Yarwood, S. J., Riehle, M. O., Johnstone, H. J. H., Affrossman, S., and Curtis, A. S. G. (2002). Increasing Fibroblast Response to Materials Using Nanotopography: Morphological and Genetic Measurements of Cell Response to 13- nm-High Polymer Demixed Islands. Experimental Cell Research 276, 1-9.

3. Clark, P., Connoly, P., Curtis, A. S. G., Dow, J. A. T., and Wilikinson, C. D. W. (1991). Cell guidance by ultrafine topography in vitro. Journal of Cell Science 99, 73- 77.

4. Powell, H.M., D.A. Kniss, J.J. Lannutti (2004) Topographical Control of Cytoskeletal Organization (Data not published)

5. Schneider, G. B., Zaharias, R., Seabold, D., Keller, J., and Stanford, C. (2004). Differentiation of preosteoblasts is affected by implant surface microtopographies. Journal of Biomedical Materials Research 69A, 462-468.

6. Juliano, R. L. and Haskill, S. (1993). Signal-Transduction from the Extracellular Matrix. The Journal of Cell Biology 120, 577-585

7. Lee, J. and Gotlieb, A. (2003). Understanding the role of the cytoskeleton in the complex regulation of endothelial repair. Histology and Histopathology 18, 879-887.

8. Morely, S. and Bierer, B. (2001). The actin cytoskeleton, membrane lipid microdomains, and T-call signal transduction. Advances in Immunology 77, 1-43.

9. Juliano, R. L. (2002). Signal transduction by cell adhesion receptors and the cytoskeleton: Functions of integrins, cadherins, selectins, and immunoglubulin- superfamily members. Annual Review of Pharmacology and Toxicology 42, 283-323.

10. McBeath, R., Pirone, D. M., Nelson, C. M., Bhadriraju, K., and Chen, C. S. (2004). Cell Shape, Cytoskeletal Tension, And RhoA Regulate Stem Cell Lineage Commitment. Developmental Cell 6, 483-495.

11. Powell, H. M. and Lannutti, J. J. (2003). Nanofibrillar Surface Via Reactive Ion Etching. Langmuir 19, 9078

12. Lieber, J.G. and R.M. Evans (1996). Disruption of vimentin intermediate filament system during adipose conversion of 3T3-L1 cells inhibits lipid droplet accumulation

166 13. Puigserver, P. and Speigelman, B. M. (2003). Peroxisome proliferator-activated receptor-γ coactivator 1α (PGC-1α): Transcriptional coactovator and metabolic regulator. Endocrine Reviews 24, 78-90.

14. Larigauderie, G., Furman, C., Jaye, M., Lasselin, C., Copin, C., Fruchart, J.-C., Castro, G., and Rouis, M. (2004). Adipophilin enhances lipid accumulation and prevents lipid efflux from THP-1 macrophages: potential role in atherogensis. Arteriosclerosis and Thrombosis in Vascular Biology 24, 504-510.

15. Mishra, R., Emancipator, S. N., Miller, C., Kern, T., and Simonson, M. S. (2004). Adipose differentiation-related protein and regulators of lipid homeostasis identified by gene expression profiling in the murine db/db diabetic kidney. American Journal of Physiology: Renal Physiology 286, F913-F921.

16. Tansey, J. T., Sztalryd, C., Bruia-Gray, J., Roush, D. L., Zee, J. V., Gavrilova, O., Reitman, M. L., Deng, C.-X., Li, C., Kimmel, A. R., and Londos, C. (2001). Perilipin ablation results in a lean mouse with aberrant adipocyte lipolysis, enhancs leptin production and resistance to diet-induced obesity. PNAS 98, 6494-6499.

17. Murphy, D. J. and Vance, J. (1999). Mechanisms of lipid-body formation. Trends in Biochemical Scienced 24, 109-115

18. Speigelman, B. M. and Farmer, S. R. (1982). Decrease in tubulin and actin gene expression prior to morphological differentiation of 3T3 adipocytes. Cell 29, 53-60.

19. Speigelman, B. M. and Ginty, C. A. (1983). Fibronectin Modulation of cell shape and lipogenic gene expression in 3T3-Adipocytes. Cell 35, 657-666.

20. Kuri-Harcuch, W, Wise, L.S. and Green, H. (1978). Interruption of the adipose comversion of 3T3 cells by biotin deficiency: differentiation without triglyceride accumulation. Cell 14, 53-59.

21. Speigelman, B. M. and Green, H. (1981). Cyclic AMP-mediated control of lipogenic emzyme synthesis during adipose differentiation of 3T3 cells. Cell 24, 503-510.

167

CHAPTER 6

ELECTROSPINNING REVIEW

6.1 Introduction

In any tissue engineering effort the end goal is functional tissue. While in vitro cultures on two-dimensional scaffolds can provide a great deal of insight into cell-material interactions, ultimately the principles gleaned from such experiments must be applied to three dimensions. Surfaces with nanotopographies provide a substrate which can be considered pseudo-three dimensional, however cell behavior on true three dimensional scaffolds must be investigated. It was our objective to engineering three dimensional meshes with nanometric size fibers to continue our investigations on the role of the nanoscale in tissue engineering..

6.1.1 Mimicking the ECM

Much effort has been directed toward creating scaffolds with chemistries and architectures that closely resemble natural structures. Such surface-based biomimesis is intended to provide cells with structural cues absent current from the generation of biomaterials. For example Gadegaard et al prepared nickel replicas of bovine type I collagen via physical vapor deposition of a small-grained metal layer followed by galvanic plating. Thermoplastic polymers were injection molded into the nickel replica to create surfaces with biomimetic topography[1]. The fibrous nature of the natural ECM has

168 lead researchers to utilize fiber based scaffolds in hopes that by mimicking the natural

ECM the synthetic scaffold will be more likely to induce normal tissue organization and function.

Fiber based scaffolds have been of current interest due to their close similarity to natural ECM. Fiber matrices offer a wide range of fiber diameters, orientations, porosity and knitted and non-woven architectures [2][3][4]. Non-woven matrices are manufactured by entangling fibers or filaments into an isotropic 3-D matrix leaving a vast amount of empty space (typically between 80-99%) [5], ideal for forming three dimensional structures. Early studies have found that pore size, pore orientation, fiber structure and fiber diameter influence cell behavior and were important factors in the development of the tissue. Polyethylene terephthalate (PET) nonwoven fiber matrices with varying amounts of porosity and pore sizes have been used as scaffolding for in vitro placental models[6][7]. It was shown that scaffolds with high porosity (93%) had poor surface accessibility for cells to attach and spread due to the distance between adjacent fibers.

ED27 metabolic activity and proliferation rates were higher initially for scaffolds with

84% porosity and 30 µm average pore diameter when compared to scaffolds with 93% porosity. Polylactide-co-glycolide fleeces soaked in alginate were implanted subcutaneously into nude mice for 8 weeks and formed a few millimeters of new cartilage[8]. Although these scaffolds have had limited success their fiber diameters are orders of magnitude larger than fibrous components of natural ECM thus the ability to form nanoscale polymer fibers is essential.

169

Figure 6.1. Schematic diagram of the melt spinning process [9] for forming fibers

170 6.2 Electrospinning

Conventional fiber forming processes for synthetic polymers involve spinning and heat treatment. A common method, melt spinning, is used for polymers that can be extruded in the melt state. The melt is ejected from small capillary holes in a spinneret to form filaments which are cooled to solidify (Figure 6.1). Thus the size of the capillary holes directly limits the fiber diameter. Forcing polymer melts or solutions through the small capillaries requires an enormous amount of pressure which makes the process of forming micron to submicron sized fibers either too expensive or impossible. There are a number of techniques that can produce fibers without the use of a spinneret. Continuous fibers can be produced by centrifugal spinning, tack spinning [10] or electrostatic spinning also known as electrospinning [11][12][13] The latter has become extremely useful to the tissue engineering community due to it ability to form submicron, ECM-like fibers. Fiber diameters produced by electrospinning are typically hundreds of nanometers which is one to two orders of magnitude smaller than any other conventional technique.

6.2.1 Theory of Electrospinning

Electrospinning transforms a millimeter diameter fluid stream into solid fibers four orders of magnitude smaller in diameter. Although the theory of electrospinning has been known for over 100 years and the process has been patented for almost 70 years [11][14], is has just recently been exploited to produce fabrics for biocatalysts, filtration applications and optical sensor devices due to its ability to make

171

Figure 6.2. Photographs of a) a droplet hanging at capillary tip, which b) elongates as the liquid is exposed to a charge. As the intensity of the charge increases the droplet forms c) a Taylor cone. When the charge is further increased d) an electrified jet is expelled from the apex of the cone [18,]

172

Needle

High-Voltage Power Supply Syringe Pump Tubing

Grounded Target

Figure 6.3 Typical experimental set up for electrospinning involves a polymer solution which is either forced through a syringe pump (shown) or through a capillary tube. The solution at the tip of the needle or capillary tube is electrified by applying a voltage to the needle or tube. An electrified jet is then expelled from the tip of the needle or tube towards a ground where the electrospun fibers are collected.

173 fibers with nanoscale diameters[15][16][17].In electrostatic spinning or electrospinning, a polymer solution of polymer melt is forced through a syringe or a capillary tube. The polymer solution is held by its surface tension at the end of the capillary tube (Figure

6.2a) or syringe needle which will be subjected to an electric field. A charge is induced on the liquid surface by the electric field. Charge repulsion causes a force directly opposite to surface tension. As the intensity of electric field increased the hemi-spherical surface of the solution elongates to a conical shape known as the Taylor cone (Figure

6.2c). When the electric field reaches a critical the repulsive electric force overcomes the surface tension causing a charged jet of solution to be ejected from the tip of the Taylor cone (Figure 6.2d).

The ejected polymer jet is accelerated towards the target which is oppositely charged or grounded (Figure 6.3). As the jet travels towards the target the diameter decreases due a number of processes including the evaporation of the solvent/cooling and the effect of stretching. The end result of this process is a dense nonwoven mesh composed of nanometric sized fibers. However the dominant thinning mechanism is under debate. One of the proposed mechanisms is termed “splaying” because it involves the splitting of the primary jet into thinner secondary jets [19] (Figure 6.4). As the diameter of the jet decreases the surface charge density increases. The high repulsive force within the jet forces it to split and splay. Those further decrease in jet diameter of the secondary jets cause those to split and splay causing a cascade of reduced diameters.

Thus with this mechanism the final fiber diameter is determined primarily by the number

174

Figure 6.4. Photograph showing the splaying phenomenon just beginning to occur at the end of the jet (arrow)[18]

175 of subsidiary jets. [13,20]. Because the jet is a charged entity there is a strong repulsion of the jet with itself; the jet tries to avoid contact with any of its other surfaces. In intense electric fields the jet becomes unstable after traveling just a short distant from the nozzle.

The unstable region is similar to an inverted cone (Figure 6.5). In reality the cone which appears to be comprised of many jets is a single rapidly whipping jet [22,24,25] oscillating at such a high frequency that it appears to be multiple filaments. In this mechanism the final fiber diameter is determined by the type and magnitude of the instable jet.

It is likely that both of the mechanisms come into play during the electrospinning process. In order to tailor the nonwoven meshes to custom design each mesh the competition between each of the modes of instability need to be more fully understood.

However there are other variables that are known to alter the final mesh. Parameters that affect the electrospinning include the solution properties such as viscosity, conductivity and surface tension, controlled variables such as feed velocity of pressure in the capillary, electric potential at the tip, distance from the tip and the target, temperature, humidity and air velocity in the chamber. By altering these variables a wide variety of fiber shapes, diameters and mesh porosities can be made.

6.2.2 Variability in Fiber Diameter

The architecture of the electrospun mesh is highly dependent on fiber diameter; porosity, pore size and the pore structure are intimately linked to fiber diameter. Fiber diameter can be altered by the rate of flow through the syringe pump or pressure on the capillary tube. Higher flow rates produce larger diameter fibers. Fiber

176

Figure 6.5 Frame by frame images showing the evolution of bending/whipping instability on a jet of 15% PCL in acetone. Note the inverted cone shape of the region of instability. [21]

177

a. b.

Figure 6.6. Scanning electron micrographs of the morphology of fibers versus solution viscosity. a) 527 centipoise PEO-water solution and b) 1835 centipoise PEO-water solution. Note the slightly larger fiber diameter from the higher [24] viscosity solution. The horizontal edge of each image is 20 µm long.

178

Figure 6.7. SEM of an electrospun, low viscosity solution of PEO and water. The low viscosity hinders the electrospinning process leading to the formation of fine beads rather than continuous fibers. [24] The horizontal edge of the micrograph is 20 µm long.

179 diameter can also be altered by viscosity of the polymer solution of melt. Thinner or less viscous solutions tend to produce thinner fibers (Figure 6.6). There are limitations to altering the solution viscosity in general solutions under 8 wt.% (polymer to solvent ratio) are too thin to electrospin and result in what is termed an electrospray where instead of continuous fibers, fine beads of polymer are deposited (Figure 6.8). Extremely thick solutions (>20 wt.%) are too viscous to electrospin. Fiber diameter can also be altered by varying the distance between the tip of the syringe/capillary tube and the target. Larger distances produce thinner fibers; the whipping instability has more time to thin the fiber on its path to the target and as the fiber diameter decreases there is an increased probability of splaying. Longer trajectory paths also lead to breakage of the fiber which decreases the tensile strength of the mesh when compared with a mesh of a continuous fiber. Each of these parameters can be altered together or separately to make very specific fiber diameters leading to tailorable porosities. This is extremely beneficial for tissue engineering applications where different cell types require varies levels of porosity to migrate into the construct.

6.2.3 Materials for Electrospinning

In addition to high variability in fiber diameter, the electrospinning process can be used for a wide array of polymeric materials from synthetics to natural and hybrid polymers. One of the earliest electrospun polymers was poly(ethylene oxide) (PEO).

180

Figure 6.8. SEM micrographs of electrospun PEO fibers. [25]

181 PEO-water solutions (10 wt%) were spun at a voltages from 5 to 15 kV to create fibers which were 300 nm in diameter (Figure 13) [25].Poly(lactide-co-glycolide) (PLGA), a polymer commonly used for biomedical applications, was electrospun to form a mesh comprised of fibers 500 to 800 nm in diameter. The pore diameter distribution, total pore volume, total pore area and porosity of the structure were calculated for the resulting meshes [26]. The porosity of the PLGA meshes was 91.63% with a pre size distribution ranging broadly from 2 to 465 microns. Meshes of PLGA also exhibited excellent mechanical properties; the tensile modulus of the structure was 323.15 MPa while the ultimate tensile stress of the construct was 22.67 MPa [26]. These tensile properties are comparable to the properties of skin and cartilage. (Table 6.1) [26]. An extensive amount of research has gone into investigating the structure of electrospun polycaprolactone

(PCL) [21,27,28]. PCL is advantageous to many biomedical applications due to its slow degradation rate. Another feature that makes PCL so beneficial to electrospin is its tendency to loop during the whipping instability [21]. This causes the mesh to be much loftier and contain larger pore diameters than other electrospun meshes. Twenty nanometer to 5 micron diameter PCL meshes were produced from 10wt% PCL in chloroform solutions [28]. Electrospinning of PCL in a variety of solutions from methylene chloride, to dimethylformamide and toluene have been investigated [29].

Toluene-PCL solutions have been very difficult to electrospin due to their high viscosities and low conductivity. Dimethylformamide-PCL solutions produce small fiber diameters

182

Electrospun Cartilage Skin

PLGA

Tensile Modulus 323 130 15-

(MPa) 150

Ultimate Tensile 23 19 5-30

Stress(MPa)

Ultimate Tensile 96 20-120 35-

Strain (%) 115

Table 6.1 Comparison of Tensile Properties of Electrospun PLGA meshes with Natural Tissues [26]

183 when methylene chloride is mixed into the solution but solutions without a methylene chloride are difficult to spin verifying the importance of solution properties. A wide range of polymers including acrylic[22], nylon[16], poly(ethylene glycol) [30], poly(methyl methacrylate) [31] and poly(ethylene terephthalate) [30] have been successfully electrospun and produce constructs each with a specific set of mechanical and chemical properties for a wide range of applications.

Natural polymers have also been electrospun to overcome the biocompatibility issues inherent to some synthetic polymers. Considerable effort has been invested in contrasting scaffolds out of natural materials to closely mimic the in vivo environment.

Because so much of the natural ECM is composed of collagen, collagen seems an ideal material for scaffolding. Lyophilized collagen was dissolved in hexafluoro-2-propanol at various concentrations and electrospun into matrices with average fiber diameter of 100 nm + 40 nm [32]. The electrospun type I collagen even exhibited the 67 nm banding typical of native collagen (Figure 6.9) suggesting that the body will react to the synthetically manufactured collagen as it would to native collagen [32]. After the initial success of electrospun collagen other biopolymers were investigated. Bovine fibrinogen- hexafluoro-2-propanol solutions were electrospun at various concentrations from

0.083g/ml to .167g/ml to produce meshes with average fiber diameters from 80 + 20 nm to 700 + 110 nm [33].

To match the mechanical properties of native tissues hybrid solutions containing both synthetic and biopolymers have been spun. Collagen-PEO fabrics have been spun

(Figure 6.10) and have tensile strengths much greater than either polymer alone [34]. A

184

Figure 6.9 a) SEM micrograph of electrospun type I collagen from the human placenta b) Transmission electron micrograph of electrospun type I calfskin collagen exhibiting the 67 nm banding typical of native collagen. Scale bar on b is 100 nm [30]

185

Figure 6.10. TEM micrograph of fibers spun from 2 wt% solutions of collagen-PEO at a 1:2 collagen/PEO ratio. Scale bar is 2 µm [34]

186 pure PEO fabric has a tensile strength of 90kPa and a modulus of 7MPa while 1:2 ratio blends of collagen-PEO had a tensile strength of 270 kPa and a modulus of 8 MPa [34].

These strategies to alter electrospun architecture, chemistry and mechanical properties allow the electrospinning process to be applied to many applications from textiles to membranes. However one of the most innovative applications for this technique is scaffolding for tissue engineering.

6.2.4 Tissue Engineering Applications

Although it has been postulated that these meshes are ideal for tissue engineering applications, few studies have taken place which examine the resultant scaffolds ability to provide an environment onto which a cell can attach and proliferate. Despite the meshes obvious similarity to the ECM, it was not until 2001 that this technology was exploited for tissue engineering applications. Kenawy et al. have shown that poly(ethylene-co-vinyl alcohol) mats can support the culturing of smooth muscle cells and fibroblasts. Human dermal fibroblasts and aortic smooth muscle cells were seeded into the mats at a density of 10^6 cells per 12 mm diameter mat. The cells were kept in culture for 7 days with replacement of media supplemented with 15% fetal bovine serum every other day.

Although the seeding process had low efficiency the cells that did seed attached to the scaffold in a natural fashion (i.e. web-like, flattened structures) (Figure 6.11 [35]).

Unfortunately other than attachment no other biological functions were investigated.

PLGA electrospun meshes with average fiber diameter ranging from 500 to 800 nm were

187

Figure 6.11 Smooth muscle cells interacting with poly(ethylene- [35] co-vinyl alcohol) mat after 7 days in culture.

188

Figure 6.12 SEM of smooth muscle cells seeded into electrospun collagen cylindrical constructs that were placed into [32] a bioreactor for 7 days of culture.

189 seeded with either human derived bone-marrow-derived mesenchymal stem cells or

BALB/c C7 mouse fibroblasts and cultured for up to 10 days. Mesenchymal stem cells showed increased cell proliferation till day 10 and exhibited a total fivefold increase.

Both cell types exhibited the ability to migrate into the core of the scaffold [26] . Collagen matrices were also evaluated for their bio compatibility. Aortic smooth muscle cells were seeded onto the collagen mats and cultured for seven days. At day 7 the scaffolds were densely populations with smooth muscle cells. Cross-sectional analysis also revealed the electrospun collagen promoted extensive infiltration into the fibrillar network (Figure

6.12) [32] One of the most promising studies has shown the ability of PCL electrospun scaffolds to support the growth of bone-marrow derived neonatal rat mesenchymal stem cells and their differentiation [28]. The mesenchymal stem cells were seeded onto the PCL mats and given osteogenic differentiation medium. The stem cells differentiated into osteoblast as expected but those cells were able to penetrate the scaffold and lay down abundant ECM after only one week. In addition mineralization and type I collagen were clearly observed at 4 weeks [28]. While these studies are few in number, it is clear that electrospinning has enormous implications for the tissue engineering community.

190 References

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6. Ma, T., Li, Y., Yang, S. T., and Kniss, D. A. Tissue engineering human placenta trophoblast cells in 3-D fibrous matrix: spatial effects on cell proliferation and function. Biotechnology progress 1999.15(4), 715-724.

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16. Gibson, P. W., Schreuder-Gibson, H. L., and Rivin, D. Electrospun fiber mats: transport properties. AIChE Journal 1999.45(1), 190-195.

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18. Doshi, J.A. The electrospinning process and application of electrospun fibers. Thesis (Ph.D.), University of Akron, Department of Polymer Science 1994

19. Reneker, D.H., Yarin, A.L., Fong, H., and Koombhongse, S. Bending instability of electrically charged liquid jets of polymer solutions in electrospinning. Journal of Applied Physics 2000. 87(9, Pt. 1), 4531-4547.

20. Hohman, M.M., Shin, M., Rutledge, G., and Brenner, M.P. Electrospinning and electrically forced jets. II. Applications. Physics of Fluids 2001.13(8), 2221-2236.

21. Reneker, D. H., Kataphinan, W., Theron, A., Zussman, E., and Yarin, A. L. Nanofiber garlands of polycaprolactone by electrospinning. Polymer 2002.43(25), 6785-6794.

22. Baumgarten, P.K. Electrostatic spinning of acrylic microfibers. Journal of Colloid and Interface Science 1971.36(1), 71-79.

24. Fong, H., Chun, I., and Reneker, D. H. Beaded nanofibers formed during electrospinning. Polymer 1999.40(16), 4585-4592.

25. Deitzel, J. M., Kleinmeyer, J. D., Hirvonen, J. K., and Beck Tan, N. C. Controlled deposition of electrospun poly(ethylene oxide) fibers. Polymer 2001.42(19), 8163- 8170.

192 26. Li, W, Laurencin, C.T., Caterson, E.J., Tuan, R.S., and Ko, F.K. Electrospun nanofibrous structure: a novel scaffold for tissue engineering. Journal of biomedical materials research 2002.60(4), 613-621. 6-15-

27. Lee, SH, Yoon, J.W., and SUh, M.H. Continuous nanofibers manufactures by electrospinning technique. Macromolecular Research 2002.10(5):282-285.

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32. Matthews, J.A., Wnek, G.E., Simpson, D.G., and Bowlin, G.L. Electrospinning of Collagen Nanofibers. Biomacromolecules 2002.3(2), 232-238.

34. Huang, L., Nagapudi, K., Apkarian, R. P., and Chaikof, E. L. Engineered collagen- PEO nanofibers and fabrics. Journal of biomaterials science.Polymer edition 2001.12(9), 979-993.

35. Kenawy, E.R., Layman, J.M., Watkins, J.R., Bowlin, G.L., Matthews, J.A., Simpson, D.G., and Wnek, G.E. Electrospinning of poly(ethylene-co-vinyl alcohol) fibers. Biomaterials 2003.24(6), 907-913.

193

CHAPTER 7

FABRICATION AND CHARACTERIZATION OF 3D POLYCAPROLACTONE ELECTROSPUN SCAFFOLDS

7.1 Introduction

Scaffold properties including scaffold dimension, configuration and pore structure have been shown to greatly affect cell behavior [1]. In general 3-D scaffolds can sustain a higher cell density and a longer proliferation period compared to 2-D substrates.

Increased maintenance of differentiated phenotype and function has also been linked to 3-

D culture [2]. Pore size has also been found to influence differentiation with hepatocytes differentiating more readily when grown on foamed scaffolds with larger pore diameters

[3]. Thus scaffold properties are an integral part of generating a synthetic tissue with properties analogous to the natural tissue. Electrospinning provides a technique which can produce a wide variety of 3-D scaffold architectures for tissue engineering [4-6]. In the following experiments nonwoven PCL meshes with varying morphologies were fabricated via electrospinning and characterized via SEM. “Ideal” scaffolds were then evaluated for degradation and subsequent change in pH of the media.

194 7.2 Materials and Methods

7.2.1 PCL solutions

There are four experimental variables that are known to strongly affect the properties of electrospun materials: solution viscosity, applied electrical field, flow rate of polymer and distance of the syringe needle from the collection plate [7-10]. Thus PCL scaffolds were generated by varying these parameters following. To achieve this, polycaprolactone (40,000 MW, Sigma Aldrich) beads were put into solution in acetone at a range of weight concentrations. The solutions were then electrospun following the parameter combinations outlined in Table 7.1.

7.2.2. SEM

The resultant scaffolds were mounted onto an Al stub with carbon adhesive tape

(Ted Pella, Reading, CA) and sputter coated with Au-Pd. Scaffolds were then characterized via scanning electron microscopy (ESEM, FEI) at 10kV. Average fiber diameter and range were calculated via image analysis.

7.2.3. Degradation

To analyze the degradation rate of the scaffolds, a scaffold with the most ideal properties was chosen and places into deionized water at 37ºC. pH readings were taken from days 0-60. Samples were removed from the water at days 1,7,15 and 60, dried and characterized via SEM for any morphological signs or degradation.

195

Solution Flow Rate wt% (ul/min) Voltage Height 8 700 28 20 8 500 24 20 8 400 26 20 8 300 26 20 8 250 24 20 8 200 24 20 8 150 23 20 8 300 24 35 8 200 20 25 9 400 26 20 9 300 228 20 9 250 27 20 9 200 26 20 9 150 25 20 10 700 23 20 10 500 31 20 10 400 29 20 10 300 22 20 10 250 24 20 10 200 26 20 10 150 21 20 10 700 20 25 10 250 20 30 10 300 20 35 12 700 32 20 12 500 32 20 12 400 29 20/28.5 12 300 28 20 12 250 29 20 12 200 25 20 12 150 23 20 12 400 25 25 12 400 15 15

Table 7.1 PCL scaffold fabrication parameters

196

a. b.

c. d.

Figure 7.1 SEM micrographs of PCL electrospun with a) 12 b) 10 c) 9 and d) 8 wt% PCL in acetone. Note the decrease in fiber diameter as solution concentration decreases. Scale bar is 10µm in all images.

197 7.3 Results and Discussion

7.3.1 Scaffold Architecture

Characterization of the electrospun PCL with SEM reveals a trend of decreasing fiber diameter with decreasing weight percentage of PCL in the solution (Figure 7.1). A reduction in PCL concentration from 10 wt% to 8 wt% at the same flow rate, diminishes the average fiber diameter from 500 nm to 250 nm while also decreasing the fiber diameter range, 2µm-500 nm and 1 µm-150 nm respectively (Figure 7.2). Unfortunately this reduction in fiber diameter is often accompanied by the emergence of beaded fibers

(Figure 7.3) when a pulse of polymer solution is ejected from the needle. The presence of beads is typically not desired in the final outcome because they will likely be targets for stress concentration, weakening the scaffold. They also generate a site of disorder in the cell organization which can affect the function of the tissue

To eliminate beaded fibers, a balance between flow rate and jet ejection must be reached. It was seen that at high voltages a reduction in flow rate increased the quantity of beads (Figure 7.4a&b). At low voltages the polymer ejection rate is balanced by reducing the flow rate (Figure 7.4 b&c). At higher voltages the polymer is ejected more rapidly thus at too low a flow rate, thin fibers are swiftly ejected followed by a thicker fiber when another drop of solution reaches the tip. In agreement, at low applied voltages the polymer is being ejected at a slower rate thus lower flow rates would prevent beads from forming.

198

Figure 7.2. Reduction in fiber diameter with lower solution concentrations a) 12 wt % 500 µl/min 22 kV b)8 wt% 500µl/min 24 kV

199

Figure 7.3 Presence of beaded fibers in meshes spun with low wt% PCL solutions. An 8 wt% solution is shown here.

200

a. b.

c. d.

Figure 7.4 SEM microgrpahs of PCL meshes produces with a) 12 wt%, 700 µl/min, 32 kV b) 12 wt % 500 µl/min, 32 kV c) 12 wt%, 400 µl/min 28 kV d) 12 wt % 400 µl/min, 20 kV. Scale bar is 10 µm in each image.

201

Figure 7.5 Scaffolds spun with 12 wt% PCL-acetone and a flow rate of 400 µl/min at varying heights a) from 20 cm above the collection plate and b) from 15 cm above the plate.

202 Distance between the tip of the needle and collection plate has also been shown to alter the scaffold morphology. Needles placed only 15 cm away from the collection plate generate scaffolds containing fibers that appear to have been bonded together (Figure

7.5). It is likely that the reduced travel distance of the polymer jet is too small to allow complete evaporation of the acetone thus when the fiber reaches the plate is essentially

“sticky” and can bond to the fiber below it.

7.3.2 Scaffold Degradation

7.3.2.1 pH

A scaffold, thought to be of the correct architecture for cell growth and proliferation, was chosen (Figure 7.5 a) for the subsequent degradation studies. Measures of pH over a time period of 60 days indicated that the PCL scaffold relatively stable over the first three weeks of exposure to water with a change in pH of only 0.395 from day 1 to day 21 (Figure 7.6). However upon exposure to water for 60 days, the pH begins to shift towards the acidic region suggesting that at this point the scaffold has degraded to a point which can be detected.

7.3.2.2 Architecture of Degraded Scaffold

Scaffolds were characterized via SEM at days 1-60. No significant alteration in fiber morphology or diameter was seen from day 1-7. At day 15 fibers have roughened on the surface and are slightly smaller in diameter (Figure 7.7a). Suggesting that some surface degradation has taken place. By 60 days of incubation a proportion of fibers are non-continuous and have even greater surface roughness (Figure 7.7b). These

203

Figure 7.6 pH of water used to incubate PCL scaffolds at 37ºC for days 0-60. Each time point was done in triplicate.

204

Figure 7.7 Electrospun scaffold after a) 15 and b) 60 days of exposure to water at 37ºC. Note the increased surface roughness of the fibers (a, red arrow) and broken fibers (b, red arrows).

205 indicate the scaffold had now begun to undergo considerable degradation. When this study is carried out for longer periods of time, the fibers will continue to degrade until no fibers remain. However that time point has yet to be determined.

7.4 Conclusions

Electrospinning is capable of producing nonwoven meshes of varying architectures and fiber diameters. The ease of fiber generation and ability to tailor the process for a specific need makes this technology of particular interest for tissue engineering. We have seen that nanofibrous scaffolds can be manufactured and are stable for relatively long periods of time thus providing a somewhat long term synthetic ECM upon which cell can attach.

206 References

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207

CHAPTER 8

ELECTROSPUN TUBULAR SCAFFOLDS FOR VASCULAR TISSUE ENGINEERING

8.1 Introduction

Arteriosclerosis, the net effect of vascular processes lending to narrowing of the arteries, is the most common cause of premature mortality in the western world [1]. For coronary arteries this narrowing induces a weakening of the myocardium and, ultimately, a heart attack (myocardial infarction) [2]. If narrowing progresses to the point where a heart attack is either imminent or has already occured the diseased portion of the artery must be replaced to restore normal blood flow. The most common treatment is coronary artery bypass grafting (CABG). In CABG, the diseased artery is often replaced with vasculature from other regions of the body such as the internal mammary artery or the saphenous vein [1,2,3]. Unfortunately, these native vessels may not be available due either to disease or prior utilization. In addition, native vessels display a pronouced tendency toward reocclusion and thus tissue engineering alternatives are a high priority for the US healthcare system.

208 Early attempts at circumventing the use of autologous tissue utilized tubular woven polymeric meshs comprised of polyethylene terephthalate (Dacron) or ePTFE

(expanded polytetrafluoroethylene) [1,2,3,4]. While these grafts function reasonably well in specific cardiovascular applications where the vessel is large (> 6-10 mm in diameter), small diameter grafts are swiftly blocked by thrombotic events [2]. To alleviate this problem, tissue engineering concepts are being employed to create cell-scaffold constructs which mimic the structure of natural blood vessels. Of great importance is the vasoactivity provided by organized layers of smooth muscle cells and the non- thrombogenic surface supplied by a layer of endothelial cells lining the lumen of the vessel.

Many techniques have been used to form vessel replacements [5,6,7,8]. Dacron and ePTFE grafts seeded with endothelial cells constituted an attempt to mimic the natural endothelial lining [9,10,11]. Unfortunately, the endothelial cells are often dislodged from the graft upon exposure to hemodynamic flow. Collagen gels shaped into tubular structures are often seeded with endothelial cells and utilized as a replacement vessels [12]. Although this structure benefits from reduced thrombogenicity its poor mechanical properties do not lend themselves to in vivo applications. Cellular self-assembly techniques roll confluent sheets of vascular smooth muscle cells onto a soluble mandrel followed by the addition of a fibroblastic layer and subsquent endothelialization of the lumen [13]. While these grafts exhibit reduced thrombotic events the strength of the vessel is still not high enough to support blood flow. While many techniques have been proposed, each has distinct advantages and weaknesses, and none possesses all of the desired

209 physiomechanical properties. Thus scaffolds that possess appropriate levels of vasoactivity, resistance to thrombotic events and mechanical properties need to be engineered.

In addition, major morphological differences exists between the natural

“scaffolding” of the vasculature and all previous attempts to form synthetic grafts. The fiber diameter in these synthetic grafts is up to twenty times as large as natural collagen fibers. The inherent pore diameters are also much larger than those found in vivo. In addition, these vessels have morphologies and mechanical properties that differ greatly from those of the native extracellular matrix. To provide improvements in vasoactivity or nonthrombogenicity, it is our goal to engineer a biodegradable, biocompatible scaffold that mimics the architecture of a vessel’s own extracellular matrix. This biomimicry would ideally cause the cells to organize themselves in a manner that more faithfully reproduces in vivo robustness and function. To achieve this, we turned to a process known as electrospinning or electrostatic spinning. Electrospinning transforms a fluid stream approximately a millimeter in diameter into polymer nanofibers that are orders of magnitude smaller [14]. A polymer solution is initially forced through a syringe or a capillary tube. The polymer solution is held by its surface tension at the end of the capillary tube or syringe needle which will be subjected to an electric field. A charge is induced on the liquid surface by this applied electric field. Charge repulsion causes a force directly opposite to surface tension. As the intensity of electric field increased the hemi-spherical surface of the solution elongates to a conical shape known as the Taylor

210 cone. When the electric field reaches a critical the repulsive electric force overcomes the surface tension causing a charged jet of solution to be ejected from the tip of the Taylor cone. The ejected polymer jet is accelerated towards the target which is oppositely charged or grounded. As the jet travels towards the target the diameter decreases due a number of processes including the evaporation of the solvent/cooling and the effect of kinetic stretching. The end result of this process is a nonwoven mesh composed of nanometer sized fibers [14,15,16].

8.2 Materials and Methods

8.2.1 Tubular Scaffolds

In this study a 12 wt.% polycaprolactone (PCL, 40,000 MW, Sigma Aldrich) in acetone solution was electrospun onto a grounded stainless steel rod rotating at 30 rpm (Figure

8.1) to create closed-end tubes approximately 3 mm in diameter and 30 mm long having a wall thickness of 40 µm (Figure 8.2). The tubes were then sterilized in 70% ethanol/30% water solution for 10 min followed by rinsing in phosphate-buffered saline (PBS) prior to cell seeding.

8.2.2 Cell culture experiments

Primary cultures of human aortic vascular smooth muscle cells (VSMCs, Clonetics) were maintained in growth media (Clonetics SMGM2 supplemented with 5% fetal bovine serum). Two million VSMCs were loaded into the tubular scaffold under low

211

Figure 8.1. Experimental set-up for electrospinning PCL tubes. PCL acetone solution was loaded into the syringe and set to flow at 400 ul/min. The acetone-PCL solution was carried through the extension set tubing to the needle where it was electrified by an applied voltage. The charged polymer jet then travelled to a grounded stainless steel rotating drum to form a closed-end tubular scaffold.

212

Figure 8.2 Macroscopic image of closed end PCL electrospun tube

213 serum (0.5%) conditions. The open end of the tube was tied off with suture and the cell- scaffold construct placed into a 6-well culture plate containing 5% serum. This generated a chemo-attractant gradient across the scaffold to stimulate VSMC migration through the

8.2.3 Cellular morphology scaffold walls. The 6-well plates containing the cell seeded tubes were then incubated in a humidified 5% CO2−95% air incubator (37 °C) for two days. Each experiment was repeated at least once.

Cell morphology and orientation with respect to the individual nanofibers was characterized with scanning electron microscopy (SEM). Cells on the PCL tubes were fixed with 3% glutaraldehyde in a phosphate buffer (pH 7.4) for 2 h. They were then washed with the phosphate buffer for 30 min. The samples were osmicated in 1% OsO4 for 1 h and washed again. After dehydration in a graded ethanol series, the samples were then subjected to a graded ethanol-hexamethyldisilazane (HMDS, Ted Pella, Redding,

CA) series (3:1 ETOH:HMDS to 100% HMDS) to avoid any swelling of the fibers caused by the critical point drying process. After the disks were immersed in 100%

HMDS, they were dried overnight by evaporation. The dry disks were mounted onto Al stubs with carbon adhesive tape, sputter coated with gold/palladium and examined using

SEM (Sirion, FEI) under 5 kV.

8.2.4 Immunocytochemistry

Cytoskeletal arrangement and cell orientation as a function of position within the tube wall were characterized with immunocytochemistry (ICC) and laser scanning confocal

214 microscopy (LSCM). The retrieved cell-seeded tubes were rinsed with PBS and fixed in

4% paraformaldehyde in PBS for 1 hr. After fixation, the samples were rinsed gently with

PBS and then permeabilized with 0.5% Triton X-100 in PBS for 15 min. Samples were then pretreated for 1 hr with 5% horse serum in PBS followed by incubation with antibodies for F-actin and nucleic acid (F-actin- AlexaFluor phalloidin, Molecular

Probes, nucleus- Sytox, Molecular Probes). The tubes were then thoroughly rinsed and examined under a fluorescence microscope (Zeiss 510 META) utilizing the z-series capability. In addition, a portion of the fixed sample was embedded in optimum cutting temperature (O.C.T.) compound and crysectioned into 2 µm slices. The cross-sections were then mounted to a glass slide and prepped for ICC following the procedure above.

The sections were imaged with fluorescence microscopy.

8.3 Results

SEM revealed that the electrospun tubes were comprised of nonwoven fibers ranging from 50 nm to 2 µm in diameter with an average pore size of approximately 7

µm (Figure 8.3). After two days in culture cells were elongated and formed a confluent sheet covering the lumen of the tube. Cells oriented themselves helically around the diameter of the tube lumen as evidenced from SEM (Figure 8.4). VSMCs were observed outside of the tube, suggesting that cells had migrated through the scaffold and into the surrounding well. LSCM observations indicated that the VSMCs populated the entire vessel wall thickness utilizing a layered morphology and orientation that changes from

215

Figure 8.3. SEM image of the fiber architecture within the electrospun PCL tube.

216

Figure 8.4. SEM image of cells lining the lumen of the PCL scaffold. Note the extended nature and longitudinal orientation of the cells.

217

Figure 8.5. Gallery of LSCM images taken from the lumen of the artificial blood vessel towards the outer surface (schematic inset). Images are presented as a sequential gallery (1.5 µm apart). Light blue fluorescence represents Alexa568-phalloidin, which labels F-actin. Red fluorescence is a nuclear stain. Human vascular smooth muscle cells (VSMs) were present in layers throughout the “vessel”. Interestingly, the orientation of the VSMs changes from “circumferential” towards a more ‘longitudinal” orientation as you progress through the vessel

218

Figure 8.6. Gallery of LSCM images taken from mid-thickness of the artificial blood vessel towards the outer surface. Images are presented as a sequential gallery (1.5 µm apart). Light blue fluorescence represents Alexa568-phalloidin, which labels F-actin. Red fluorescence is a nuclear stain. The VSMCs assume a longitudinal orientation as they move towards the outer surface of the scaffold.

219

Figure 8.7. SEM image of VSMCs (red arrows) migrating through the PCL scaffold.

220 the inner to the outer wall . The cells on the inner wall were oriented circumferentially

(Figure 8.5). However, toward the outer layer the scaffold, the cells were highly elongated and oriented along the long axis of the tube (Figure 8.6). Immunofluorescence of the cryo-sectioned samples confirmed that the VSMCs maneuvered into and through the scaffold while adopting this longitudinal orientation as they reached the outer tube.

Cross-sections of the tube also show singular VSMCs migrating through the matrix and evidence of tunneling (Figure 8.7).

8.4 Discussion

These results indicate that electrospun PCL scaffolds provide an environment upon which human VSMCs can attach and migrate. As the first step in any tissue engineering operation, cell attachment is essential for all downstream cellular events.

Evidence of cell attachment and migration indicates that proteins from the media adsorbed onto the PCL surface and have mediated strong attachment of the cells. The adsorption of these proteins and subsequent cell attachment without prior modification of the surface is an appealing property of these scaffolds compared to those which must be either chemically modified or coated with collagen in order to promote cell adhesion

[17,18,19]. In addition, the presence of cells within the thickness of the scaffold and on the outside of the tube suggests that a simple, inexpensive serum gradient effectively encouraged migration through the membrane. The combined effects of scaffold architecture, cell number and cell-cell signaling likely caused the VSMCs to orient themselves in a manner highly reminiscent of the in vivo tissue without any prior surface modification or application of mechanical stress. The time scale at which the cells

221 assumed these orientations is, on average, 30 times shorter than studies that use mechanical force to orient the cells and improve the strength of the vessel [20,21,22]. In experiments where mechanical stresses are used to orient the cells, the cells must first be cultured within the scaffold to sufficient cell density. The resultant cell-scaffold construct is then often placed under cyclic stress, for up to 20 weeks [20] to orient the cells and produce high levels of elastin and collagen. Although these mechanical methods successfully produce oriented structures, the typical cardiac patient awaiting a CABG cannot tolerate a 20 week waiting period. The rapid cellular alignment afforded by the use of the nanoscale makes this technique clinically attractive. While these encouraging results provide us with important preliminary observations further investigations are underway to determine migration pathways, scaffold mechanical integrity versus degradation and the ability of pulsatile flow to alter or enhance the orientation of these seeded VSMCs.

222

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225

CHAPTER 9

CONCLUSIONS AND RECOMMENDATIONS

The ability to predict and control a cells function on a tissue engineering scaffold is essential to proper tissue morphogenesis. It is clear that current tissue engineering scaffolds do not provide cells with the proper environmental cues for the generation of complex tissue structures, thus approaches to “synthetically” alter a cells behavior are needed. Surface topography has been identified as one the classic factors which can control cell behavior. Nanoscalar surfaces are of particular interest due to the similarity of scale to important biological molecules. In this research the techniques to produce nanoscalar two-dimensional and three-dimensional scaffold were investigated. After scaffold fabrication, cell function, including cellular organization, morphology, cytoskeletal organization, hormone production, apoptosis and differentiation, were evaluated on the surfaces and matrices. A summary of important findings is summarized below.

226 9.1 Effect of 20 nm-high protrusion

Polyethylene terephthalate tissue culture coverslips were etched with a low power

75% O2- 25% Ar plasma for a total of 30 minutes. This processing produced 20 nm high protrusions on the surface that were stable with time and had no change in surface chemistry. Surface roughness did not cause a significant alteration in cell proliferation, morphology, hormone production or cytoskeletal expression. Rates of apoptosis and differentiation were shown to slightly increase by the presence of the protrusions. It was clear that this level of nanoscalar roughness is not large enough to cause striking alterations in cell function.

9.2 Control of Surface Roughness with Reactive Ion Etching

Precise control of surface topographies has previously been achieved with micromachining and has been shown to effect cellular behavior. Unfortunately this processing is often labor intensive and costly. Reactive ion etching provides a relatively inexpensive means of altering surface topography. It was our goal to determine if reactive ion etching could be used to produce topographies of varying roughness in a reproducible manner. PET coverslips were etched with O2, N2, Ar and 1:1 combinations of each.

Qualitative and quantitative analyses reveal that the magnitude and morphology of surface roughness can be tuned by varying the gas media and etch time. In general, oxygen possess the greatest ability to etch PET and generates a distinctive nanofibrillar surface while argon has the weakest etching power and produces a fine array of rounded

227 protrusions on the surface. In addition, oxygen etching of the thermally bonded PET generates thicker and more robust fibrils. These results indicate reactive ion etching can produce a wide array of nanotopographies.

9.3 Disruption of Cytoskeletal Organization By Nanofibrillar Surfaces

The similarity in dimension of fibrils generated via oxygen plasma etching and natural collagen fibrils was the basis for selecting oxygen plasma etched surfaces for subsequent cellular growth. The stability of nanofibrillar surfaces in aqueous media was first assessed by soaking the disks for 24 hrs in DMEM. It was found that nanofibrils fractured at their base and detached themselves from the surface. Thus the more robust thermally bonded nanofibrillar structures were used. These structures were shown to cause a reduction in relative adhesion strength and cell-substrate signaling. In addition cells grown on nanofibrillar surfaces had a complete disruption of their F-actin cytoskeleton. Reports from other labs on the minimum spacing of surface ligands required for proper cell attachment and subsequently the formation of stress fibers suggest that the density of nanofibrils may be interfering with the coalescence of integrin- ligand complexes at the cell surface. Further molecular analyses must be performed to pinpoint the cause of these changes.

9.4 Alteration of Adipogenic Differentiation by Nanotopographies

To investigate the effects of these relatively early cell functions on downstream cellular events, cells were grown on nanofibrillar and control surfaces and differentiated into adipocytes. The process of differentiation was monitored by expression of PPAR-γ

228 and the formation of lipid droplets. It was seen that the presence of PPAR-γ was not affected by surface topography but the size of the lipid droplets was. In cells grown on rough surfaces lipid droplet coalescence was inhibited causing the generation of a greater proportion of small droplets compared to the few sizable droplets found within cells grown on control surfaces. This inhibition of lipid droplet coalescence is likely related to the cytoskeletal disruption seen in the previous set of experiments. The role of cytoskeletal filaments on adipocytes differentiation has been extensively studied and these filaments have been observed to play a large role in droplet formation. Others have reported a disruption in the vimentin cytoskeleton can inhibit differentiation and lipid droplet creation.

9.5 Extending these Principles to Three-Dimensions

Nanometric surfaces have been shown to alter cell function on two-dimensional surfaces. The principles collected from these experiments must be extrapolated to three dimensional scaffolds if genuine tissues are ever to be engineered. Fabrication of nanometer sized fibers by conventional methods has proved to be challenging if not impossible in most cases. Electrospinning, a technique that uses an electric field to eject a thin polymer jet from a droplet of polymer solution, is a means of producing nonwoven meshes of nanometer fiber diameters ideal for tissue engineering. Studies in our lab have shown that great variability in scaffold architecture and fiber size can be generated by altering four key electrospinning parameters: solution concentration, flow rate, applied voltage and travel distance of the jet. By altering these parameters we have constructed a

229 scaffold with an average fiber diameter of 500 nm and an average pore size of 5 µm. This scaffold morphology was generated in a tubular scaffold by spinning the polymer around a grounded rotating rod rather than a grounded plate. These tubular scaffolds are ideal for vascular tissue engineering. Therefore human aortic smooth muscle cells were cultured on these substrates for two days. Cells organized themselves in a circumferential fashion within the lumen of the tube and rotated to a more longitudinal orientation and progressed to the tube surface. This pattern of organization is highly reminiscent of natural vasculature suggesting that electrospun scaffolds have a promising future as tissue engineering scaffolds.

9.6 Recommendations

Tissue engineering has been a rapidly evolving discipline during the past thirty years with the development of more sophisticated means of scaffold fabrication, discovery of new cell sources and improvements in cell culture equipment. The knowledge gained from basic cell-material interaction studies, like the ones conducted here, can be applied to many systems and cell types. Further studies on nanometric 2-D and 3-D tissue engineering substrates should be performed to determine the effect of a wide array of topographies and architecture on various cell types.

230

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