Polymer Biomaterial Constructs for Regenerative

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POLYMER BIOMATERIAL CONSTRUCTS FOR REGENERATIVE

MEDICINE AND FUNCTIONAL BIOLOGICAL SYSTEMS

LINGHUI MENG

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Dissertation Advisor: Dr. Gary E. Wnek

Department of Macromolecular Science and Engineering

CASE WESTERN RESERVE UNIVERSITY

May, 2012

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Linghui Meng s candidate for the Ph.D. degree *.

(signed) Dr. Gary E. Wnek s

(chair of the committee)

Dr. David Schiraldi s

Dr. Alexander M. Jamieson s

Dr. Horst Von Recum s

(date) 12/05/2011 s

*We also certify that written approval has been obtained for any proprietary material contained therein.

Table of Contents

Chapter I Crosslinking and Modification of Collagen-based Materials ...... 1 1.1 Introduction ...... 1 1.2 Characteristics of Collagen as a Biomaterial ...... 2 1.2.1 Collagen Native Structures ...... 2 1.2.2 Collagen Natural Crosslinks ...... 6 1.3 Isolation and Purification of Collagen ...... 7 1.4 Exogeneous Crosslinking of Collagen ...... 10 1.4.1 Chemical Crosslinking ...... 11 1.4.2 Physical Crosslinking ...... 17 1.4.3 Enzymatic Crosslinking ...... 18 1.5 Collagen in Tissue Engineering ...... 19 1.5.1 Electrospinning Technique ...... 19 1.5.2 Electrospinning of Collagen ...... 21 1.5.3 Electrospinning of Collagen and Natural Polymer Blends ...... 24 1.5.4 Electrospinning of Collagen and Synthetic Polymer Blends ...... 26 1.6 Dissertation Overview ...... 29 Chapter II Collagen-PCL Sheath-Core Bicomponent Fibers ...... 30 2.1 Introduction ...... 30 2.2 Coaxial Electrospinning ...... 31 2.3. Fabrication and Characterization of Collagen-PCL Sheath-Core Fibers ...... 33 2.3.1 Materials ...... 33 2.3.2 Coaxial Electrospinning of Collagen and PCL ...... 34 2.3.3 Post-crosslinking Treatment ...... 35 2.3.4 Characterization of Fiber Morphology ...... 35 2.3.5 Characterization of Mechanical Properties ...... 36 2.4 Results and Discussion ...... 37 2.4.1 Core-Shell Fiber Formation ...... 37 2.4.2 Post-Crosslinking of Collagen-PCL Hybrid Fibers ...... 44 2.4.3 Mechanical Properties of Collagen-PCL Scaffolds ...... 45 2.5 Conclusions ...... 48 Chapter III In-situ Crosslinked Collagen Nanofibers ...... 50 3.1 Introduction ...... 50

3.2 Fabrication and Characterization of In-situ Crosslinked Collagen Nanofibers ...... 53 3.2.1 Materials ...... 53 3.2.2 Preparation of In-situ Crosslinking Collagen Solution ...... 54 3.2.3 Rheological Experiment ...... 54 3.2.4 Electrospinning of In-situ Crosslinked Collagen Nanofibers ...... 55 3.2.5 Characterization of Fiber Morphology ...... 55 3.2.6 Fourier Transform Infrared (FTIR) Spectroscopy ...... 56 3.2.7 Characterization of Mechanical Properties ...... 56 3.3 Results and Discussion ...... 57 3.3.1 In-situ Crosslinking Approach ...... 57 3.3.2 Formation of In-situ Crosslinked Collagen Nanofibers ...... 59 3.3.3 Mechanical Properties of In-situ Crosslinked Collagen Scaffolds ...... 66 3.3.4 FTIR Spectra of In-situ Crosslinked Collagen Scaffolds ...... 68 3.3.5 Plausible Mechanisms of In-situ Crosslinking Reactions ...... 70 3.4 Applications of In-situ Crosslinked Collagen Fiber Scaffolds ...... 72 3.4.1 Cell Culture Studies ...... 72 3.4.2 Open Wound Healing Test ...... 74 3.4.3 Drug Release Studies ...... 77 3.5 Conclusions ...... 81 Chapter IV In-situ Crosslinked Collagen Gels and Sponges ...... 84 4.1 In-situ Crosslinked Collagen Gels ...... 84 4.1.1 Formation and Characterization of In-situ Crosslinked Collagen Gels ...... 85 4.1.2 Results and Discussion ...... 87 4.1.3 Conclusions ...... 94 4.2 In-situ Crosslinked Collagen Sponges ...... 95 4.2.1 Formation and Characterization of In-situ Crosslinked Collagen Sponges ...... 96 4.2.2 Results and Discussion ...... 97 4.2.3 Conclusions ...... 101 4.3 Biomineralization of In-situ Crosslinked Collagen Gels ...... 102 4.3.1 Electrophoresis Experiments ...... 103 4.3.2 Results and Discussion ...... 104 4.3.3 Conclusions ...... 107 Chapter V Phase Transition of Poly (Acrylic Acid) Nanofibers ...... 109 5.1 Introduction ...... 109

5.2 Formation and Characterization of PAA Nanofiber Tubes ...... 111 5.2.1 Materials ...... 111 5.2.2 Preparation of PAA Ethanol Solution ...... 111 5.2.3 Fabrication of PAA Nanofibers ...... 111 5.2.4 Thermo-crosslinking of PAA Nanofibers ...... 112 5.2.5 Neutralization of PAA Nanofiber Tubes ...... 112 5.2.6 Characterization of Fiber Morphology ...... 112 5.2.7 Titrations of PAA Tubes ...... 113 5.2.8 Elemental Analysis of PAA Tubes ...... 114 5.3 Results and Discussion ...... 114 5.3.1 PAA Nanofiber Formation ...... 114 5.3.2 Crosslinking of PAA Nanofibers ...... 116 5.3.3 Neutralization of PAA Nanofibers ...... 118 5.3.4 Calcium Ion-induced Phase Transition ...... 120 5.3.5 Magnesium Ion-induced Phase Transition ...... 128 5.3.6 Potential Biomimicking Applications ...... 132 5.4 Conclusions ...... 134 Chapter VI Conclusions and Future Work ...... 136 6.1 Conclusions ...... 136 6.2 Future Work ...... 141 References ...... 144

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List of Tables

Chapter I Crosslinking and Modification of Collagen-based Materials

Table 1.1 The amino acid composition of type I collagen from calf-skin (the value in parentheses are the residues contributed by the non-helical telopeptide regions) [14]…………..……………..3

Chapter II Collagen-PCL Sheath-Core Bicomponent Fibers

Table 2.1 Tensile properties of electrospun collagen-PCL bicomponent scaffolds…...…….…...47

Chapter III In-situ Crosslinked Collagen Nanofibers

Table 3.1. Mechanical properties of in-situ crosslinked collagen scaffolds…..…………………68

Table 3.2 Comparative study of FTIR spectrum wavelength in the amide I and amide II region for Kensey Nash collagen, post-crosslinked electrospun collagen and in-situ crosslinked collagen.

……………………………………………………………………………………………………69

Chapter IV In-situ Crosslinked Collagen Gels and Sponges

Table 4.1 Solubility of type-I collagen in the in-situ crosslinking solution.……………..….…90

List of Figures

Chapter I Crosslinking and Modification of Collagen-based Materials

Figure 1.1 Chemical structure of type I collagen. (a) Primary amino acid sequence, (b) secondary left-handed helix and tertiary right-handed triple-helix structure and (c) staggered quaternary structure. [9]...... ………………………………….………………………………………………4

Figure 1.2 Covalent aldol cross-links form between two lysine or hydroxylysine residues at the non-helical telopeptide regions. [23]………………………………………………………………6

Figure 1.3 Chemical structures of natural collagen crosslinks. (a) Intramolecular aldol condensation type crosslink, (b) intermolecular aldimine (Schiffs’ base) type crosslink, (c) condensation of aldol condensation type crosslink with hydroxyproline and (d) hydroxypyridinium type crosslink. [24]……...……………………………………………………8

Figure 1.4. Schematic representation of the crosslinking reaction of glutaraldehyde (GA) with collagen. [37]…..…………………………………………………………………………………12

Figure 1.5 Schematic representation of the cross-linking reaction of an epoxy compound with collagen. [45]…..…………………………………………………………………………………13

Figure 1.6 Schematic representation of the cross-linking reaction of EDC and NHS with collagen. [47]..………………………………………………………………………………………………15

Figure 1.7. Schematic representation of a traditional electrospinning setup with a horizontal orientation. [64]..…………………………………………………………………………………20

Figure 1.8 Scanning electron micrographs (SEM) of electrospun collagen type I (A) [69], type II (B) [70], type III (C) [71], and type IV (D) [63] from HFIP solution. SEM scale bars = 1 µm (A, B, C) and 10 µm (D)..……………………………………………………………………………22

Figure 1.9 SEM image of electrospun collagen fibers from PBS (20x)/ethanol (1:1). [72]….……………………………………………………………………………………………23

Figure 1.10 SEM micrographs of electrospun scaffolds: (a) collagen fibers, (b) collagen + 10% nanoHA (uncrosslinked), (c) collagen + 10% nanoHA (crosslinked), and (d) collagen + 20%HA (crosslinked). Scaffolds become dense due to chemical crosslinking.[75]…………….…………25

Figure 1.11 Fabricated bilayered tubular construct of SPU/collagen. (1) Appearance of the tube. Scale bar: 1 mm. (2) Scanning electron micrograph of the tube. (3) Magnified image of region 3 in photo (2). (4a–c) Magnified images of the outer layer of region 4 in photo (2). (5a–c) Magnified images of the inner layer of region 5 in photo (2). [79]..………………………..……27

Chapter II Collagen-PCL Sheath-Core Bicomponent Fibers

Figure 2.1 Schematic of coaxial electrospinning setup …………………………………………32

Figure 2.2 Schematic illustration of compound Taylor cone formation (A: Surface charges on the sheath solution, B: viscous drag exerted on the core by the deformed sheath droplet, C: Sheath- core compound Taylor cone formed due to continuous viscous drag)…..………………………33

Figure 2.3 Appearances of electrospun pure collagen mats from water-alcohol-salt benign solvent system before (a) and after (b) soaking in water…………………………………………………37

Figure 2.4 Appearances of collagen-PCL sheath-core bicomponent scaffolds (a, c) deposited on glass slides and soaking in chloroform (b) and water (d) for 30 min, respectively...……………40

Figure 2.5 SEM images of the coaxial electrospun collagen-PCL bicomponent fibers before (a) and after (b) water-treatment. (c) Distribution of resulting collagen-PCL nanofiber diameter….41

Figure 2.6 SEM images of the freeze-fractured collagen-PCL sheath-core nanofibers…………42

Figure 2.7 SEM images of electrospun mono-component collagen (a) and PCL (b) fibers…….43

Figure 2.8 SEM image of the post-crosslinked collagen-PCL sheath-core fiber scaffolds..……45

Figure 2.9 Tensile stress-strain profiles of electrospun scaffolds……………..…………………46

Chapter III In-situ Crosslinked Collagen Nanofibers

Figure 3.1 Scanning electron microscopy micrographs: samples cross-linked with (A) 20-mM 1- ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC) (B) 200-mMEDC (C) 20- mM EDC/N-hydroxysuccinimide (NHS) (D) 200-mMEDC/NHS (E) 3% glutaraldehyde (room temperature), (F) 50% glutaraldehyde (heated), (G) 100% ethanol-soaked, and (H) no treatment (dry); scale bar is 10 mm. [108]…..………………………………………………………………52

Figure 3.2 Plot of the apparent gelation time as a function of the molar ratio of NHS and EDC……………………………………………………………………………………………….58

Figure 3.3 Typical SEM images of electrospun in-situ crosslinked collagen fiber at low (a) and high (b) magnification. (c) Distribution of fiber diameter. (d) Diameter of in-situ crosslinked collagen fibers increased during electrospinning processing………..……………………………60

Figure 3.4 SEM images of electrospun in-situ crosslinking collagen nanofibers collected on the glass slides at various time points with low (a, c, e, g) and high (b, d, f, h) magnifications..……61

Figure 3.5 Effect of relative humidity on the morphology of collagen fibers. SEM images of electrospun in-situ crosslinking collagen fibers at humidity of 33% for 3 days (a), 43% for 3 days (b) and 53% for 1day (c). SEM image of electrospun collagen fibers without EDC and NHS stored at humidity of 53% for 7days (d)……………………………………………………….…63

Figure 3.6 Snapshots of electrospun collagen scaffolds post-crosslinked (a, b) and in-situ crosslinked (c, d) with EDC/NHS. The scaffolds were placed on the glass slides before (a, c) and after (b, d) water treatment…….…………………………………………………………………65

Figure 3.7 SEM images of electrospun in-situ crosslinking collagen scaffolds after water- treatment at low (a) and high (b) magnifications……………...... ……………65

Figure.3.8. Representative plots of stress-strain curves to demonstrate the difference in behavior of in-situ crosslinked collagen dry and hydrated samples……….………………………………67

Figure 3.9 FTIR spectra of electrospun collagen scaffolds crosslinked by an in-situ method (a) and by a post method (b)…………………………………………………………………………69

Figure 3.10 Reactions of a carboxyl groups on a collagen molecule with EDC to result in esterification (possible but probably infrequent), transesterification, or a stable amide bond with another collagen molecule. This figure was adapted from [108]…………………..……………71

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Figure 3.11 CHO cells growing for 5 days on electrospun in-situ crosslinked collagen nanofibers. The image was taken at 20x magnification………………………………………………………74

Figure 3.12 Appearance of wound healing at day 0, 5 and 10 after patching with electrospun collagen scaffolds (in-situ crosslinked and non-crosslinked). Circular wounds of 6mm diameter were prepared on each of the mice’s back. ………………………………………………………76

Figure 3.13 Chemical structures of silicon phthalocyanine Pc4 (a) and doxycycline (b)………78

Figure 3.14 Optical images of electrospun collagen mats containing Pc4 (a, b) and BSA-Texas Red (c, d). Images were taken at bright field (a, c) and laser fluoresce light (b, d)…………..…79

Figure 3.15 SEM image of electrospun collagen fibers containing doxycycline (a). In vitro release of doxycycline from collagen scaffolds (b)………………………………………………80

Chapter IV In-situ Crosslinked Collagen Gels and Sponges

Figure 4.1 Appearances of collagen hydrogel plate (a), tubes (b) and hemisphere (c) prepared by the in-situ crosslinking method……………………………...……………………………………91

Figure 4.2 Snapshot of a CWRU logo printed by using in-situ crosslinking collagen solution…………………………………………………………………………………………....92

Figure 4.3 CHO cells 24 hours after seeding. The image (a) shows cells seeded onto the collagen gel. The image (b) shows CHO cells seeded on a control tissue culture polystyrene surface. The magnification for both images is 10x. In this study the collagen gels were dried at room temperature overnight and then seeded with CHO cells……………………….…………………93

Figure 4.4 CHO cells 48 hours after seeding. The image (a) is the cells on the collagen hydrogel surface, and the image (b) contains CHO cells plated on a control tissue culture polystyrene surface. Images are at 20x magnification…………………………………………………………94

Figure 4.5 Snapshots of cylindrical and tubular in-situ crosslinked collagen sponges prepared by freeze drying of collagen gels………………………………….…………………………………98

Figure 4.6 SEM images of in-situ crosslinked collagen sponges prepared in refrigerator (a), dry- ice ethanol bath (b) and liquid nitrogen (c, d)………………….…………………………………99

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Figure 4.7 FTIR spectrums of in-situ crosslinked collagen sponges…………..……….………100

Figure 4.8 Appearance of wound healings at day 0, 5 and 10 after incision……...……………101

Figure 4.9 Collagen hydrogels before (a) and after (b) electrophoretic mineral deposition for one hour...……………………………………………………………………………………………104

Figure 4.10 In-situ crosslinked collagen sponges after freeze drying. SEM imaging (A and B) show no particulates on surface indicating prior mineral residue. EDX analysis (C) indicates C and O as the primary detected elements…………………………………………………………105

Figure 4.11 SEM images of collagen hydrogel after 1 hour of electrophoretic mineral deposition (A and B). EDX (C and D) confirms that there is much phosphate and calcium in the region, and the calcium + phosphate overlay shows very good correspondence with the particulate, indicating calcium phosphate buildup……………………106

Chapter V Phase Transition of Poly (Acrylic Acid) Nanofibers

Figure 5.1 SEM images of PAA nanofibers electrospun from different concentrations at a flow rate of 0.8 mL/h: (a) 2 wt%, (b) 4 wt%, (c) 6 wt%, and (d) 4wt% at a flow rate of 1.0 mL/h….116

Figure 5.2 Thermal esterification of polyacrylic acid with ethylene glycol……………………117

Figure 5.3 SEM images and snap shots (inserted) of crosslinked PAA nanofiber tubes before (a) and after (b) immersing in water…………………...……………………………………………117

Figure 5.4 Proposed structure of PAA-Na polymer network…………...………………………118

Figure 5.5 pH responsive of crosslinked PAA fiber tubes………………...……………………120

Figure 5.6 Typical ESEM image of neutralized PAA fibers in water………….………………120

Figure 5.7. (a) Length changes of neutralized PAA nanofibrous tubes with increasing the concentration of Ca2+ in solution. (b) Snap shots of PAA tubes before and after the titration of

CaCl2 solution. ……………………………………………….…………………………………121

Figure 5.8 Representative schematic of calcium induced cross-bridge between carboxylate groups in PAA tubes……………………………….....…………………………………………122

Figure 5.9 Proposed structure of the compacted PAA-Ca polymer……………………………124

Figure 5.10 (a) Length changes of compacted PAA-Ca tubes with increasing concentration of EDTA in solution. (b) Snap shots of PAA tubes before and after the titration of EDTA………125

Figure 5.11 SEM images of PAA fiber network after calcium crosslink (a) and EDTA chelation (b)…………………………..……………………………………………………………………127

Figure 5.12 Swelling behaviors of the compacted PAA-Ca tubes in citrate and EDTA solutions, respectively…………...…………………………………………………………………………128

Figure 5.13 Length changes of neutralized PAA tubes with the increasing concentrations of calcium ad magnesium ions in solution, respectively…………..………………………………129

Figure 5.14 Phase transitions of the neutralized PAA tubes in different conditions. Representative shrinking-swelling cycle responses in calcium-EDTA titration (a), magnesium-EDTA titration (b), calcium-citrate titration (c), and magnesium-citrate titration (d). All the curves were generated based on the same PAA samples. ………………………………………………………………131

Figure 5.15 Proposed models of structural dynamics and the action potential. Initially (A) the network is collapsed as strands are bridged by calcium. The network expands as sodium replaces calcium (B). But increasing sodium eventually neutralizes surface charge, weakens water structure and allows the polymer-retractive force to collapse the network (C), at which stage calcium may easily bridge the strands once again. [172]………………….……………………133

List of Abbreviations

Arg – Arginine

Asp – Aspartic Acid

BSA – Bovine Serum Albumin

CD – Circular Dichroism

CHO – Chinese Hamster Ovarian cells

CS – Chondroitin Sulfate

DHT – Dehydrothermal Treatment

DMA – Dynamic Mechanic Analysis

DMEM – Dulbecco's Modified Eagle Medium

DPPA– Diphenylphosphoryl Azide

ECM – Extracellular Matrix

EDC – 1-ethyl-3-(3-dimethylaminopropyl) Carbodiimide

EDTA – Ethylenediaminetetraacetic Acid

EDX – Energy Dispersive X-Ray

EG – Ethylene Glycol

FTIR – Fourier Transform Infrared Spectroscopy

GA – Glutaraldehyde

GAG – Glycosaminoglycan

Gly – Glycine

HA – Hyaluronic acid

HFIP – 1,1,1,3,3,3 Hexafluoro-2-propanol

Hyp – Hydroxyproline

ICP-OES – Inductively Coupled Plasma Optical Emission Spectroscopy

NHS – N-hydroxysuccinimide

NMR – Nuclear Magnetic Resonance

PAA – Poly (acrylic acid)

PBS – Phosphate Buffered Saline

Pc4 – Silicon Phthalocyanine

PCL – Polycaprolactone

PDO – Polydioxanone

PEO – Poly (ethylene oxide)

PGA – Poly (glycolic acid)

PLA – Poly (lactic acid)

PLCL – Poly (lactide-co-caprolactone)

PLGA – Poly (lactic-co-glycolic acid)

Pro – Proline

RGD – Arginine-Glycine-Aspartic Acid

SAXS – Small-Angle X-ray Scattering

SEI- Secondary Electron Imaging mode

SEM – Scanning Electron Microscopy

SPU – Segmented Polyurethane

TEM – Transmission Electron Microscopy

TFE – 2,2,2-Trifluoroethanol

TGs – Transglutaminases

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Polymer Biomaterial Constructs for Regenerative Medicine and Functional

Biological Systems

Abstract

LINGHUI MENG

The use of collagen as a biomaterial is currently undergoing a renaissance in the tissue engineering field. The excellent biocompatibility and safety due to its biological characteristics, such as biodegradability and weak antigenicity, make collagen a primary

material resource in medical applications. Described herein is work towards the

development of novel collagen-based matrices, with additional multi-functionality

imparted through a novel in-situ crosslinking approach.

The process of electrospinning has become a widely used technique for the creation of fibrous scaffolds for tissue engineering applications due to its ability to rapidly create structures composed of nano-scale polymer fibers closely resembling the architecture of the extracellular matrix (ECM). Collagen-PCL sheath-core bicomponent fibrous scaffolds were fabricated using a novel variation on traditional electrospinning, known as co-axial electrospinning. The results showed that the addition of a synthetic polymer core into collagen nanofibers remarkably increased the mechanical strength of collagen matrices spun from the benign solvent system.

A novel single-step, in-situ collagen crosslinking approach was developed in order to solve the problems dominating traditional collagen crosslinking methods, such as

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dimensional shrinking and loss of porous morphology, and to simplify the crosslinking

procedure for electrospun collagen scaffolds. The excess amount of NHS present in the

crosslinking mixture was found to delay the EDC/collagen coupling reaction in a

controlled fashion. Fundamental investigations into the development and characterization

of in-situ crosslinked collagen matrices such as fibrous scaffolds, gels and sponges, as

well as their biomedical applications including cell culture substrates, wound dressings,

drug delivery matrices and bone regeneration substitutes, were performed. The preliminary mice studies indicated that the in-situ crosslinked collagen matrices could be good candidates for wound healing and skin regeneration.

Polyelectrolyte fibrous tubes of highly-crosslinked poly (acrylic acid) were fabricated by means of electrospinning as polymer models for functional biological systems, with special attention to the axon cortical layer and its cation-exchange properties. The processing parameters of fiber formation and the reversible phase transitions of PAA tubes according to monovalent-divalent ion exchange in solution were systematically investigated. The results showed that the neutralized PAA tubes were responsive to calcium ions, exhibiting significant shrinkage that could be reversed with a chelator such as citrate. Study of such phase transitions may help to better understand the electrophysiological processes known as nerve excitation and conduction in the nervous system, and the resulting PAA tubes might be used as polymer models of artificial axons for potential tissue engineering and nerve repair applications.

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Chapter I Crosslinking and Modification of Collagen-based Materials

1.1 Introduction

Restoration of the function of a failed organ due to injury or disease often requires transplantation. This approach has been largely limited by the scarcity of well-suited donors [1, 2], which has necessitated the development of alternative surgical techniques that would obviate the need for donor organs. ‘Tissue engineering’ has emerged as a promising approach to address this problem with the goal of developing biological substitutes that can be implanted into the body to restore, maintain, or improve tissue function [3]. In this technique, the cells from the patient’s body are isolated, expanded, and cultured onto a three-dimensional porous structure called ‘scaffold.’ It is assumed that the cells will adhere to the scaffold, proliferate and produce the natural tissue substitute [4].

The use of collagen as a biomaterial is currently undergoing a renaissance in the tissue engineering field [5, 6]. The excellent biocompatibility and safety due to its biological characteristics, such as biodegradability and weak antigenicity, make collagen a primary material resource in medical applications [7, 8]. However, collagen, like many natural polymers, once extracted from its original source and then reprocessed, suffers from weak mechanical properties, thermal instability and ease of proteolytic breakdown [9].

To overcome these problems, collagen crosslinking is needed [10]. Basic knowledge about the native structure and crosslinking is necessary to understand the properties of isolated collagen materials and the effects achieved by potential modifications.

Accordingly, this chapter reviews the native characteristics of collagen and the methods to crosslink collagen after extraction from native tissue. Also reviewed are the studies

published to date about applications of collagen in tissue engineering, especially collagen

fibrous scaffolds fabricated by the electrospinning technique.

1.2 Characteristics of Collagen as a Biomaterial

1.2.1 Collagen Native Structures

Structural order in collagen, as in other proteins, occurs at a several discrete levels of the structural hierarchy [11]. The collagen in the tissues of a vertebrate occurs in at least 20 different forms, each of these being predominant in a specific tissue. Structurally, these collagens share the characteristic triple helix, and variations among them are restricted to the length of the non-helical fraction, as well as the length of the helix itself and the number and nature of carbohydrates attached on the triple helix [12]. Here the discussion will be limited to type I collagen which is by far the most abundant and represents the majority of collagen materials for tissue engineering applications.

Type I collagen is predominant in higher order animals especially in the skin, tendon, and bone where extreme forces are transmitted [13]. It is a macromolecular complex of three chains, two of which are identical, termed α1 (I) chains, and one α2 (I) chain with different amino acid composition. Each α chain consists of more than 1000 amino acids.

The composition of the α1 (I) and α2 (I) chains of calf-skin collagen are given in Table

1.1 [14]. There are only minor differences between type I collagens from different vertebrate species [15].

Table 1.1 The amino acid composition of type I collagen from calf-skin (the value in

parentheses are the residues contributed by the non-helical telopeptide regions) [14]

The primary structure of Type I collagen denotes the complete sequence of amino acids along each of three polypeptide chains [16]. Approximately one-third of the residues are glycine (Gly) and it repeats at every third position in the sequence. About 35% of the non-glycine positions in the repeating unit Gly-X-Y are occupied by proline (Pro), found almost exclusively in the X-position, and 4-hydroxyproline (Hyp), predominantly in the

Y-position, as shown in Figure 1.1a.

The secondary structure is the local configuration of a polypeptide chain that results from satisfaction of stereo-chemical angles and hydrogen-bonding potential of peptide residues

[17]. In collagen, the repeating proline and hydroxyproline residues play key

conformational roles. Their very rigid ring structures enable polypeptide chains to fold

into left-handed helices with 3.3 residues per turn and a pitch of 0.87nm as shown in

Figure 1.1b [18]. On the other hand, the absence of a side chain in glycine permits close

approach of polypeptide chains in the collagen triple helix.

Figure 1.1 Chemical structure of type I collagen. (a) Primary amino acid sequence, (b)

secondary left-handed helix and tertiary right-handed triple-helix structure and (c)

staggered quaternary structure. [9]

The tertiary structure refers to the fundamental unit originally known as tropocollagen:

three polypeptide chains intertwined to form a right-handed triple-helix which can exist

as a physicochemically stable entity in solution, namely, the triple-helical collagen molecule [17]. The hydroxyl groups of hydroxyproline residues are involved in hydrogen bonding which are important for stabilizing the triple-helix structure [19]. Two hydrogen bonds per triplet are found, which are: one between the NH-group of a glycine residue and the CO-group of the residue in the second position of the triplet in the adjacent chain,

and one via the water molecule participating in the formation of additional hydrogen

bonds with the help of the hydroxyl group of hydroxyproline in the third position [20].

Furthermore, model studies show that glycine, proline and hydroxyproline are the triple-

helix forming amino acids and only those molecules which contain the triplets Gly-Pro-

Hyp are able to form a helical structure [21]. Therefore, the collagen triple helical

domains have an amino acid sequence (primary structure) that is rich in glycine, proline

and hydroxyproline residues.

The rod-shape triple helix has an average molecular weight of approximately 300 kDa, a

length of about 300 nm with a diameter of 1.5 nm, as shown in Figure 1.1c. This extreme

ratio of the dimensions gives rise to high viscosity in solutions and high mobility in

electric fields [22]. In addition, the helical structure extends over approximately 1000 of the residues in each of the three chains, leaving the remaining residues at the both ends in a non-helical configuration [21]. These non-helical regions are denoted as telopeptides.

On the fourth level of order termed quaternary structure, the triple-helical molecules

stagger longitudinally and bilaterally into fibrils with distinct periodicity [22]. The

collagen molecules aggregate through fibrillogenesis into microfibrils consisting of four

to eight collagen molecules and further into fibrils. Those fibrils reach from 10 to 500 nm

in diameter depending on tissue type and stage of development. The triple-helices are

staggered by 67 nm with an additional gap of 40 nm between succeeding molecules [17].

These collagen fibrils organize into fibers, which can form even larger fiber bundles.

1.2.2 Collagen Natural Crosslinks

The systematic packaging of the triple-helices provides strength and resilience to the collagen fibers. Additional mechanical and chemical stability derives from intra- and intermolecular crosslinks. Initially, the formation of crosslinks is mediated by lysyl oxidase during fibril formation [23]. The non-helical telopeptide regions at both ends of the polypeptide chains contain the unusual amino acid hydroxylysine which is of particular importance in the formation of collagen fibrils. The extracellular enzyme lysyl oxidase catalyzes the conversion of selective lysyl and hydroxylysyl residues to the corresponding aldehydes allysine and hyroxyallysine, as shown in Figure 1.2. When the fibrils approach, the aldehydes can spontaneously react to form aldol crosslinks [24].

H N N H

H C (CH2)2 CH2 CH2 NH3 H3N CH2 CH2 (CH2)2 C H

O C Lysine Residues C O

Lysyl oxidase O2

H N N H O O

H C (CH2)2 CH2 C C CH2 (CH2)2 C H H H O C C O Aldehyde Derivatives

Spontaneous

H N N H

H C (CH2)2 CH2 C C (CH2)2 C H H O C C C O O H Aldol Cross-link

Figure 1.2 Covalent aldol cross-links form between two lysine or hydroxylysine residues

at the non-helical telopeptide regions. [23]

Intramolecular crosslinks occur between two α-chains in the non-helical section of the same collagen molecule by aldol condensation of two aldehydes [23]. Intermolecular crosslinks form between the telopeptide region of one collagen molecule and the helical region of a quarterly staggered, adjacent molecule [24]. These bridges between two different tropocollagen molecules result from aldimine or Schiff base formation between aldehyde resides and ε-amino groups presented by lysine and hydroxylysine residues which are shown in Figure 1.3. The interchain bifunctional crosslinks are still reactive and continue to form polyfunctional crosslinks through multiple condensations with histidine, lysine, or hydroxylysine residues [25-27].

Therefore, through specific self-aggregation and crosslinking, collagen can form fibers of unusual strength and stability, which is the primary reason for the usefulness of collagen in native tissues. The degree of crosslinking increases with age and stress and consequently changes the properties of collagen material [28].

1.3 Isolation and Purification of Collagen

The major impediment to isolation of type I collagen from tissue is the presence of covalent crosslinks between molecules. Collagen is insoluble in most common organic solvents, and water-soluble collagen represents only a small fraction of total collagen in body, the amount depending on the age of the animal and the type of tissue extracted [28].

The nature of the crosslinks prevalent in different tissues determines the particular solvent to be used and the corresponding yields.

The most commonly used solvents are neutral salt solution (0.15 - 2 M NaCl) or dilute acetic acid [29]. Neutral salt solutions will extract freshly synthesized and negligibly cross-linked collagen molecules present in the tissue. Modifications in temperature, shaking rate, and volume ratio of extractant to tissue will inevitably change the composition of the collagen derived [29]. The extracted material is purified by dialysis, precipitation, and centrifugation. However, most tissues have little or no salt-extractable collagen.

Dilute acidic solvents, e.g. 0.5 M acetic acid, citrate buffer, or hydrochloric acid pH 2-3

are more efficient than neutral salt solutions. The intermolecular crosslinks of the

aldimine type can be dissociated by the dilute acids and the repulsive like charges on the

triple-helices lead to swelling of fibrillar structures [30]. However, dilute acids will not

dissociate strong crosslinks such as keto-imine bonds. Therefore collagen from tissues containing higher percentages of keto-imine bonds, i.e. bone, cartilage, or materials from

older animals, has a lower solubility in dilute acid solvents [31].

Additional collagen material can be solubilized by treating connective tissue with an

aqueous solution comprised of alkali hydroxide and alkali sulfate, e.g. approximately 10%

of sodium hydroxide and 10% of sodium sulfate [32]. Fat associated with the insoluble

collagen will be saponified, and non-helical telopeptide regions are truncated and the

collagen fibers finally disintegrate. The size and molecular weight of the resulting

collagen material depends on the time of treatment and alkali concentration. The presence

of alkali sulfate controls the swelling of the collagen structures and protects the native

triple-helical characteristics [33].

Much higher yields compared with acidic extraction can be achieved by using enzymes such as pepsin, ficin or chymotrypsin below approximately 20 °C [14, 17]. The efficacy of enzymatic treatment arises from selective cleavage in the terminal non-helical regions

breaking peptide bonds near crosslinks and releasing molecules which can dissolve.

Some crosslinks presumably remain, attaching small peptide remnants to the solubilized molecules. Thus, the telopeptide ends of the polymer chains are dissected but under appropriate conditions the helices remain essentially intact [12].

Soluble collagen is purified mainly by precipitation after pH, salt concentration or

temperature adjustment [34]. The high viscosity of even dilute solutions interferes with

purification methods such as chromatography, electrophoresis and differential

sedimentation. Collagen solutions contain varying proportions of monomer and higher

molecular weight covalently linked aggregates, depending on the source and method of

preparation.

1.4 Exogeneous Crosslinking of Collagen

Natural crosslinking renders high tensile strength and proteolytic resistance to collagen.

Due to dissociation of natural crosslinks in the aforementioned isolation and purification

processes, reconstituted forms of collagen such as fibers, films or sponges can lack

sufficient strength and may disintegrate upon handling or collapse under the pressure

from surrounding tissue in vivo [10]. In addition, the rate of biodegradation has to be

customized based on the specific applications. For example, as a hemostat, collagen has

accomplished its mission once a blood clot has formed, whereas for tissue augmentation

an implant has to maintain its scaffolding properties while it is gradually replaced by host

collagen [35]. Therefore, it is often necessary to impart mechanical firmness and

collagenase resistance to the reconstituted collagen structures by introduction of

exogeneous crosslinking.

1.4.1 Chemical Crosslinking

1.4.1.1 Glutaraldehyde

The predominant chemical agent that has been investigated for the treatment of collagen- based materials is the bifunctional molecule glutaraldehyde (GA) [36, 37], which gives materials with the highest degree of crosslinking when compared with other known methods such as formaldehyde, epoxy compounds, cyanamide and the acyl-azide method

[38]. The reactions involving crosslinking of proteins with glutaraldehyde have been extensively studied, but the reaction mechanism is very complex and still not completely

understood [35]. Aqueous solutions of glutaraldehyde contain a mixture of free aldehyde,

mono- and dihydrated glutaraldehyde, and monomeric and polymeric hemiacetals. Due to

the ease of hydration and cyclization, the concentration of free, monomeric aldehydes in

the commercial solutions is usually low [37].

Because of the complexity of the reaction solutions, many reactions can occur during crosslinking, as shown in Figure 1.4 [37]. In general, glutaraldehyde (II) preferably reacts with ε-amino groups (I) of lysine or hydroxylysine residues of the collagen at neutral pH, yielding a Schiff base (III). Cheung et al. suggested that Schiff bases are stable under the crosslinking conditions and crosslinking involves the formation of GA polymers (IV) due to aldol condensation reactions [39]. In addition, the formation of an

unsaturated Schiff base intermediate (V) followed by Michael addition of a collagen

amine group with the unsaturated group of (V) to give (VI) results in the formation of a

crosslink. Furthermore, formation of a crosslink is possible by the reaction of amine

groups with free aldehyde groups of (V) to give (VII), or a free aldehyde group remaining

after aldol polymerization of (V) to give (VII) [37].

O N Coll IV Coll N ( ) n

O O

O O Coll N Coll N - Coll NH2 + Coll-N (VI)

HN oll O O C (I) (II) (III) V ( ) O

Coll N O N Coll (VII)

Figure 1.4. Schematic representation of the crosslinking reaction of glutaraldehyde (GA)

with collagen. [37]

Treatment with glutaraldehyde has been shown to reduce the immunogenicity of

collageneous materials while increasing their resistance to enzymatic degradation [36].

However, glutaraldehyde-treated materials calcify to a large extent [40]. For instance,

calcification is the major cause in the failure of bio-prosthetic heart valves [41].

Moreover, depolymerization of polymeric glutaraldehyde crosslinks has been reported, which releases monomeric and highly cytotoxic glutaraldehyde into the recipient [42].

Researchers embarked on attempts to replace glutaraldehyde as a crosslinking agent. The

obvious strategy would be the use of other bifunctional reagents. Epoxy and diisocyanate

compounds dominate this approach. Another strategy is to activate the carboxylic acid

groups of collagen, followed by reaction with adjacent amine groups. This method is the

basis for the carbodiimide and the acyl azide reactions.

1.4.1.2 Epoxy Compounds

Epoxy compounds have been extensively used in the past decade for the stabilization of

collagen-based materials [43-45]. Generally mixtures of bi- and tri-functional glycidyl

ethers based on glycerol are applied. In addition, a broad range of multifunctional crosslinkers containing epoxy can be used [44]. Due to its highly strained three-membered ring, epoxide groups are susceptible to nucleophilic attack. Predominantly, a reaction with amino groups of lysine or hydroxylysine residues will occur as shown in Figure 1.5.

NH 2 HN

H C O CH CH CH H C O CH CH CH 2 2 2 pH > 8.0 2 2 2 O HC OR HC OR OH

H2C O CH2 CH CH2 H2C O CH2 CH CH2 O OH NH NH2

Figure 1.5 Schematic representation of the cross-linking reaction of an epoxy compound

with collagen. [45]

Additionally, epoxide groups can react with the secondary amine groups of histidine, and

the carboxylic groups of aspartic and glutamic acid, which increase the versatility of the

crosslinking [45]. In general, biological materials are crosslinked in basic solutions (pH >

8.0) containing relatively high concentrations of epoxy compounds ranging from 1 to 5

wt%. A lower shrinkage temperature was obtained compared to GA crosslinked materials

but the in-vitro stability of the cross-linked tissue was similar [44]. Epoxy treatment

makes collagen materials more flexible and decreases the incidence of calcification in

vivo. The cytotoxicity of the polyepoxy components has been shown to be acceptable in

in vitro studies [46]. Besides epoxides, other bifunctional crosslinkers have been applied in the crosslinking of collagen, such as hexamethylene diisocyanate (HDC), dimethyl

suberimidate (DMS) and bis-N-hydroxysuccinimide ester derivatives, which will not be detailed in this overview.

1.4.1.3 Carbodiimides

Crosslinking with carbodiimides, especially 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC), offers a main advantage over aldehydes and epoxy compounds in that these carbodiimides only facilitate the formation of amide bonds between carboxylic and amino groups on the collagen molecules without becoming part of the actual linkage

[47-48]. Thus, bifunctional crosslinking agents are obviated and amide bonds formed with this crosslinking reaction provide a natural linkage [49].

N OH (IV) R' R' O N O N O O O COOH + C C O C C O N + R' NH C NH R" N N V VI ( ) O ( ) (I) R" (III) R" (II) NH2 (VII) O R' = CH3 O CH 3 (VIII) C N + N OH (IV) R" = HN CH Cl 3 H O

Figure 1.6 Schematic representation of the cross-linking reaction of EDC and NHS with

collagen. [47]

EDC crosslinking involves the activation of the carboxylic acid groups from aspartic acid or glutamic acid residues (I) by EDC (II) to form an amine-reactive O-acylisourea intermediate (III) [47]. This intermediate may react with an amine group of lysine or hydroxylysine residues (VII), yielding a conjugate of the two molecules joined by a stable amide bond (VIII). However, the intermediate (III) is also susceptible to hydrolysis, making it unstable and short-lived in aqueous solution. In order to suppress the hydrolysis reaction of O-acylisourea groups, N-hydroxysuccinimide (NHS) (IV) is added to convert

the O-acylisourea group into an NHS-activated carboxylic acid group (V), which is

considerably more stable than the O-acylisourea intermediate but remains reactive toward

amino groups of lysine or hydroxylysine (VII), yielding a so-called zero-length cross-link

(VIII). EDC is not incorporated in the matrix but is converted to the soluble 1-ethyl-3-(3- dimethyl-aminopropyl)-urea (VI) [48]. Within the triple helical structure of collagen, there is a large number of opportunities for lysine and hydroxylysine interactions with

15 aspartic acid and glutamic acid residues. Thus, crosslinks created using EDC can be formed within an α chain, between α chains, between collagen molecules, or between collagen fibrils [49].

Crosslinking of dermal sheep collagen using EDC/NHS results in materials having a higher shrinkage temperature and enzymatic resistance than GA cross-linked collagen

[50]. Furthermore, rat sub-dermal implantation studies showed that the EDC/NHS crosslinked collagen samples had a low tendency to calcify and a good biocompatibility

[51]. It appears that the amide crosslinks formed may be beneficial in terms of the anti- calcification properties by limiting calcium binding sites. Additionally, EDC and NHS can be used in combination with diamine or diacid compounds to introduce 'extended' cross-links. The carboxylic acid groups of either collagen or diacid molecules will be activated with EDC/NHS followed by reaction with the amine groups of diamine molecules or collagen, respectively [52].

1.4.1.4 Acyl Azide Method

The acyl azide method is another crosslinking procedure in which the acid groups become activated followed by reaction with an adjacent amine group. The acyl azide method is a multistep reaction in which the carboxylic acid groups are first esterified with methanol under acidic conditions for 7 days [53]. Then the methyl esters are converted to a hydrazide by reaction with hydrazine. Finally, the hydrazide is reacted with sodium nitrite to give the acyl azide, which can subsequently react with an amine group of an adjacent polypeptide chain. A variation of this method has been developed by Petite et al.

[54]. They used diphenylphosphoryl azide (DPPA) to convert the carboxylic acid group into an acyl azide group in one step. DPPA and the acyl-azide crosslinked materials were

16 found to be less toxic than GA counterparts and a marked reduction in calcification was obtained compared to GA crosslinked controls after 90 days of subcutaneous implantation in rats [54].

1.4.2 Physical Crosslinking

The primary advantage of physical treatments is that they do not introduce chemicals that cause potential harm. Typical processes such as heating, drying and irradiation have been applied to collagen [55]. Short wave length UV irradiation (254 nm) can introduce crosslinks in the collagen. Formation of crosslinks during UV-irradiation is thought to be initiated by free radicals formed on aromatic amino acid residues which indicate a rather limited maximum degree of crosslinking due to the small number of tyrosine and phenylalanine residues present in collagen [56] as seen in Table 1.1. Polypeptide chain scission may become a substantial side reaction resulting in denaturation of the collagen molecules.

Dehydrothermal treatment (DHT) increases the shrinkage temperature of collagen by removing water from collagen [57]. However, collagen molecules become partially denatured by this physical treatment. In order to keep degradation of the triple helices to a minimum it is crucial for DHT treatment to reduce the water content via vacuum as thoroughly as possible prior to heating. Even small amounts of residual moisture can cause breakdown of the helical structures and proteolysis. Severe dehydration itself already induces amide bond formation and esterification between carboxyl and free amino and hydroxyl groups, respectively. But the effect is insignificant and typical DHT conditions are 110 °C for several hours up to a few days. Generally, the degree of crosslinking is considerably lower than obtained by chemical methods. Sometimes, DHT

treatment is followed by a chemical treatment (cyanamide) to increase the stability of the

treated material [58].

1.4.3 Enzymatic Crosslinking

Transglutaminases (TGs) are a family of enzymes that can catalyze the formation of a

covalent bond between a free amine group (e.g., protein- or peptide-bound lysine) and the

gamma-carboxamid group of protein- or peptide-bound glutamide [59]. Bonds formed by transglutaminase exhibit high resistance to proteolytic degradation. TGs are also able to covalently attach primary amine-containing compounds to peptide-bound glutamine, facilitating modification of the physical, chemical and biological properties of proteins

[60]. For these reasons, TGs have been utilized by the commercial sector in many different processes and have attracted much attention from the research community. Chau et al. demonstrated that on treating native bovine type I collagen with both tissue and microbial transglutaminase leads to an enhancement in cell attachment, spreading and proliferation of human osteoblasts and human foreskin dermal fibroblasts when compared to culturing on native collagen [61]. The transglutaminase-treated collagen substrates also showed a greater resistance to cell-mediated endogenous protease degradation than the native collagen.

1.5 Collagen in Tissue Engineering

The attractiveness of collagen as a biomaterial rests largely on the view that it is a natural

material of low immunogenicity and low inflammatory and cytotoxic responses, and is

therefore seen by the body as a normal constituent rather than foreign matter [17].

Collagen has been processed into a number of forms such as fibrous scaffolds, gels, sponges, and hybrid composites containing other naturally derived polymers, biodegradable synthetic polymers, or inorganic biomaterials.

Among them, collagen fibrous scaffolds are of particular interest in tissue engineering because these scaffolds can mimic the biochemical and ultra-structural properties of the native extracellular matrix (ECM) of tissue, which is known to influence cell behavior

[62]. In the past decade the process of electrospinning has become a widely used technique for the creation of fibrous scaffolds for tissue engineering applications. With its ability to rapidly create structures composed of nano-scale collagen fibers closely resembling the architecture of the ECM, the electrospinning technique has experienced a revival in recent years and continues to find new niches in tissue engineering [63].

1.5.1 Electrospinning Technique

Electrospinning is a process that utilizes a strong electrostatic field to produce ultrafine fibers from a polymer solution [64]. In its simplest form the process of electrospinning requires little specialized equipment. Everything that is needed includes a high voltage power supply, a grounded target, and a small-diameter conductive capillary (typically a needle with a blunt end) to emit a polymer solution in a controlled fashion, as shown in

Figure 1.7.

Figure 1.7. Schematic representation of a traditional electrospinning setup with a

horizontal orientation. [64]

Polymers are dissolved in a solvent, everything from water to organic solvents depending upon the polymer, at a concentration high enough for polymer chain entanglement to occur. The polymer solution is then fed through a capillary at a constant flow rate, with the polymer solution within the capillary charged to a high electrical potential opposite a grounded target to create a static electric field. When the electric field produces a force strong enough to overcome the surface tension of the polymer solution collected at the capillary tip, a Taylor cone forms [63]. This Taylor cone is then stretched until a jet of polymer is drawn from its tip and attracted to the grounded collecting target. During travel through the air, the jet undergoes a bending instability and follows a back-and-forth whipping trajectory, during which the solvent gradually evaporates and nano to micron- scale fibers are deposited on the collector [64]. The charge from the fibers dissipates into the surrounding environment, and a non-woven fiber mat is formed.

The ability for electrospinning to create tissue-specific scaffolds arises from the

adaptability of the process and the system control provided by a number of tunable

processing parameters, which include solution properties (viscosity, conductivity, and

surface tension), processing conditions (voltage, capillary diameter, distance from

capillary orifice to grounded target), and environmental conditions (temperature,

humidity and static electricity) [62-65]. Each of the aforementioned parameters, separately or in conjunction with one another, can affect fiber deposition, fiber diameter, and scaffold porosity.

Various biodegradable polymers, both synthetic and natural, have been used to construct nanofiber scaffolds for tissue engineering by means of the electrospinning technique [64].

For example, electrospinning of polymers of synthetic origin including absorbable

polyesters such as polyglycolic acid (PGA), polylactic acid (PLA), polylactic-co-glycolic acid (PLGA), polycaprolactone (PCL), and polylactide-co-caprolactone (PLCL), and those of natural origin including proteins such as collagen, gelatin, elastin, and silk, and polysaccharides such as hyaluronic acid (HA), dextran, and chitosan, have been reported to form fibrous scaffolds for tissue engineering applications [63].

1.5.2 Electrospinning of Collagen

The electrospinning of collagen was first reported with the use of poly (ethylene oxide)

(PEO) as a carrier from acetic acid solution by Huang et al. [66]. Acetic acid or formic acid can dramatically increase the solubility of collagen and facilitate the electrospinning process, but high acidity can lead to degradation of collagen structure [67]. The slow evaporation rate of the acid and its strong affinity with collagen leads to the deposition of wet fibers on the target, which can partially weld together and lessen the porosity of the

electrospun mat [68]. Furthermore, the polymer carrier PEO is required, and formation of

pure collagen fibrous scaffolds is difficult by this method.

Figure 1.8 Scanning electron micrographs (SEM) of electrospun collagen type I (A) [69],

type II (B) [70], type III (C) [71], and type IV (D) [63] from HFIP solution. SEM scale

bars = 1 µm (A, B, C) and 10 µm (D).

In the past decade, 1,1,1,3,3,3 hexafluoro-2-propanol (HFIP) appeared to be the only solvent that can be used to electrospin collagen without the aid of carrier polymers [62].

Collagen type I, II, III and IV can be solubilized in HFIP and electrospun into a porous scaffold from concentrations as low as 0.05g/mL [69-71]. The high evaporation rate and moderate affinity of HFIP for collagen makes it a very good solvent for electrospinning.

Bowlin et al. demonstrated that the electrospinning of collagen from HFIP has the potential to produce collagen fibers that mimic the structural and biological properties of

the natural collagen ECM [71]. Electrospun fibers of type I, II and III collagen dissolved

in HFIP exhibited a linear relationship between polymer concentration and average fiber

diameter, with the lower fiber size range falling within the 50-500nm seen in native tissue

[69-71]. Electrospun type I collagen fibers have been fabricated with diameters ranging from 100nm-5µm [69], with diameters of 20-200 nm for type II collagen [70], diameters of 115-612 nm for collagen type III [71], and fibers of 100-2 µm for collagen type IV

[63], as shown in Figure 1.8. However, HFIP is rather toxic, expensive and highly volatile, which makes it very dangerous and may cause some serious health problems during the electrospinning process.

Figure 1.9 SEM image of electrospun collagen fibers from PBS (20x)/ethanol (1:1). [72]

Recently, our laboratory reported a novel water-alcohol-salt “benign” solvent system in which type I collagen is quickly solubilized and from which collagen nanofibers can be easily electrospun [72]. Since hydrophobic interactions and hydrogen bonds between amino acid residues that are oriented toward the outside of the collagen triple helix maintain the fibril structure [20-22], we reasoned that the buffer-ethanol binary solvent played a dual role in collagen solubilization, with the ethanol apparently facilitating the disruption of hydrophobic interactions, and the buffer assisting in breaking hydrogen

bonding in collagen. The water-alcohol-salt benign solvent presents two additional advantages for the electrospinning process: (i) it helps to overcome the high surface tension of water that usually contributes in the poor electrospinnability of water-based

polymer solutions, and (ii) it increases the conductivity of the solution thus facilitating

the formation of a stable Taylor cone. Electrospun type I collagen fibers with diameter of

210± 60nm have been successfully prepared through this approach and the resulting

fibers are shown in Figure 1.9 [72]. The higher the salt concentration in the benign

mixture, the lower the fiber diameter average is, as expected due to the increase in the

conductivity properties of the mixture with salt content.

1.5.3 Electrospinning of Collagen and Natural Polymer Blends

The incorporation of glycosaminoglycans (GAG) into collagen fibrous scaffolds during

electrospinning could potentially be an important aspect in truly mimicking the native

ECM [73]. GAGs serve a variety of functions including linking collagen structures and

binding growth factors. The specific GAGs of physiological and tissue engineering

scaffold significance includes hyaluronic acid, dermatan sulfate, chondroitin sulfate

(most abundant GAG in tissues), heparin, heparan sulfate, and keratan sulfate [73]. For

example, blends of collagen and GAGs such as chondroitin sulfate (CS) have been found

to improve tissue growth and regeneration over the use of collagen alone. Zhong et al.

developed blended scaffolds of collagen and CS, crosslinked with glutaraldehyde. These

scaffolds allowed for increased proliferation of rabbit conjunctiva fibroblasts [74].

Nanofibrous biocomposite scaffolds of type I collagen and nanohydroxyapatite (nanoHA)

of varying compositions (wt%) were prepared by Thomas et al. for bone tissue

engineering applications, as shown in Figure 1.10 [75]. The surface roughness, fiber

diameter and the tensile strength of the scaffold was increased with the presence of

nanoHA. Chemical cross-linking with glutaraldehyde further enhanced the mechanical

properties of the scaffold.

Figure 1.10 SEM micrographs of electrospun scaffolds: (a) collagen fibers, (b) collagen

+ 10% nanoHA (uncrosslinked), (c) collagen + 10% nanoHA (crosslinked), and (d) collagen + 20%HA (crosslinked). Scaffolds become dense due to chemical crosslinking.

[75]

Recently, Yeo et al. used collagen and silk fibroin to create two types of electrospun

scaffolds, a blend scaffold and a hybrid scaffold [76]. The blended scaffold was created

by mixing the two polymer solutions prior to electrospinning, while the hybrid scaffold

was electrospun simultaneously from two syringe needles. They concluded that the

hybrid matrix was better for wound dressing applications as cell attachment and the

spreading of normal human epidermal keratinocytes was better on the hybrid matrix as

compared to the blend matrix.

1.5.4 Electrospinning of Collagen and Synthetic Polymer Blends

Blending collagen with other natural and/or synthetic polymers has enabled tissue

engineers to fine-tune the desired properties of the electrospun scaffolds [63]. Collagen acts as an adhesion protein in the native ECM, enhancing cell attachment and proliferation through specific interactions between domains in collagen molecules and integrin receptors in the cell membrane such as RGD (Arg-Gly-Asp) [77]. Thus, the presence of collagen in a synthetic polymer scaffold is intended to impart biocompatibility and bioactivity to the scaffolds, with the synthetic polymer providing mechanical integrity to the structure.

Compared with other approaches for introducing proteins into nanofibrous structures, such as coating or grafting, electrospinning of a blended collagen mixture is not only simpler, but also solves the problem of slow mass transfer process dominating many traditional approaches and uses lesser amounts of chemical reagents [63]. Moreover, the existence of collagen on the surface and inside the structure provides sustained cell recognition signals with polymer degradation, which is crucial for cell function and development [78].

Scaffolds composed of collagen and synthetic polymers have been widely used for cardiovascular tissue engineering applications. In a study done by He et al. [78], it was demonstrated that type I collagen blended with poly (L-lactic acid)-co-poly (ε- caprolactone) [(PLLA-CL), 70:30] biohybrid scaffolds could enhance the viability,

26 spreading and attachment of human coronary artery endothelial cells (HCAECs) and preserve their phenotype.

In another study, a bilayer tubular construct composed of a thick segmented polyurethane

(SPU) microfiber mesh as an outer layer and a thin type I collagen nanofiber mesh as an inner layer was fabricated by Kidoki et al. as a prototype scaffold of artificial grafts, and was visualized by scanning electron microscopy [79] as shown in Figure 1.11. The authors speculated that such hierarchically-designed artificial grafts may provide compliance matching that of native arteries, along with good cell and tissue ingrowth and transient antithrombogenicity in the early phase of implantation.

Polycaprolactone (PCL) is an inexpensive, bioresorbable polymer with excellent

mechanical properties, lack of toxicity and slow degradation time [80, 81]. Lee et al. [80]

developed PCL/collagen type I composite scaffolds with the ability to resist high degrees

of pressurized flow over long durations, while still providing a favorable environment for

the growth of vascular cells. The scaffolds were also found to be conducive to bovine EC

and SMC attachment and proliferation. These PCL/collagen electrospun scaffolds were

also investigated for their in-vivo stability in a rabbit aortoiliac bypass model [80]. It was

observed that endothelialized grafts resisted adherence of platelets when exposed to

blood. Also, it was demonstrated by ultrasonography that these scaffolds were able to

retain their structural integrity over one month of implantation [81]. Moreover, these

scaffolds continued to maintain biomechanical strength at retrieval that was comparable

to native artery.

Polydioxanone (PDO) is another synthetic polymer widely used in biomedical fields. It is

a highly crystalline polymer, commonly used as a commercially available suture that has

been shown to exhibit excellent mechanical properties, elasticity, shape memory, a low

inflammatory response, and a slower rate of degradation than other resorbable suture

materials [82]. Barnes et al. created electrospun blended scaffolds of PDO and collagen

types I and III in various ratios [63]. The addition of collagen to PDO resulted in average

fiber diameter measurements comparable to those seen in the native ECM fibers (210-340

nm), surprisingly with no significant difference in fiber diameters with the addition of

increasing amounts of collagen. Preliminary in-vitro cell culture with human dermal fibroblasts demonstrated favorable cellular interactions on all constructs containing

collagen, with prominent cell migration into the thickness of the scaffolds compared to

simple surface spreading with no penetration on pure PDO scaffolds.

1.6 Dissertation Overview

Detailed herein are investigations into the development of collagen-based scaffolding

produced by novel variations of crosslink processing for use in tissue engineering,

regenerative medicine and other biomedical applications. Discussions will begin in

Chapter II with fabrication of collagen-PCL sheath-core fibrous materials through a novel

co-axial electrospinning processing to improve the mechanical properties of collagen

fibers spun from the benign solvent system. Chapter III introduces a novel, in-situ

crosslinking approach to solve the problems of dimensional shrinking and loss of porous

morphology dominating the traditional collagen crosslinking methods. Fundamental

investigations into the development and characterization of in-situ crosslinked collagen fibrous scaffolds as well as their biomedical applications including cell culture substrates, wound dressings and drug delivery matrices, will be detailed. Chapter IV describes the development of collagen gels and sponges prepared through the in-situ crosslinking method and discusses their potential applications in wound healing and bone regeneration.

Chapter V illustrates the development of polymeric fibrous tubes for potential axon- mimicking and nerve repair applications based on a novel crosslinked poly (acrylic acid), where the electrospun polyelectrolyte tubes exhibit reversible contraction behavior according to monovalent-divalent ion exchange in solution. Finally, future work and efforts to continue the development of these materials is detailed in the concluding chapter.

Chapter II Collagen-PCL Sheath-Core Bicomponent Fibers

2.1 Introduction

In recent years, it has been shown that the nanofiber structures prepared using the electrospinning technique can serve as near ideal substrates for engineering of tissues [83,

84]. Various biodegradable polymers of natural and synthetic origin have been used to

construct the nanofiber scaffolds [85-87]. The use of natural polymers is important in that

they are inherently capable of binding cells since they carry specific cell recognition sites,

for example the RGD (arginine/ glycine/ aspartic acid) amino acid sequence in collagen

[88, 89]. Synthetic biodegradable polymers, on the other hand, can provide necessary

mechanical properties and their degradation rate can be controlled over a broad range

[90]. When used alone, however, neither can readily provide an ideal structure for long-

term development of tissues [88]. This is because the regenerated natural polymers,

although greatly biocompatible, are weak and degrade rapidly and uncontrollably, while

the synthetic polymers, although mechanically more stable, are not as biocompatible.

Accordingly, the focus of this chapter is to combine natural and synthetic polymers and

to produce materials that have novel hybrid properties at the nano level.

Although the blending of synthetic and natural polymers could improve cell growth

behavior on such a blended scaffold [80-82], a more appropriate setting could be thought

of as a bicomponent nanofiber with natural polymer sheath and synthetic polymer core

[91, 92]. It is hypothesized that the natural polymer will aid in cell adhesion and

proliferation while the synthetic polymer will impart strength and elasticity. The sheath

polymer, after initiating cell adhesion and growth, will disintegrate first. This will expose

the core, which will continue to support the developing tissue over a longer time until it

becomes capable of sustaining the further growth on its own. At this point, the core will

degrade and a completely biological tissue substitute will be formed. The most suitable

polymers visualized for this purpose are collagen as the sheath and polycaprolactone

(PCL) as the core. To produce such bicomponent structures, a ‘co-axial electrospinning’

process was developed.

2.2 Coaxial Electrospinning

The general set up of coaxial electrospinning is quite similar to that used for conventional

electrospinning [93, 94]. A modification is made in the spinneret by inserting a smaller

(inner) capillary that fits concentrically inside the larger (outer) capillary resulting in a co-axial configuration (Figure 2.1). The outer needle is attached to the reservoir containing the sheath solution and the inner is connected to the one holding the core solution. The arrangement could be horizontal as shown in Figure 1.7 or vertical as illustrated in Figure 2.1. The feed rates of the solutions are controlled using either metering pumps [95] or air pressure [93]. In some studies, the sheath solution was exposed to atmospheric pressure and allowed to flow due to gravity [96, 97]. The arrangement required in these cases was vertical. Co-axial spinning could also be

conducted using polymer melts, for which a heating system is used that surrounds the

reservoir [98].

Figure 2.1 Schematic of coaxial electrospinning setup

The process of co-axial electrospinning is conceptually similar to that of the single jet electrospinning. When the polymer solutions are charged using a high voltage, the charge accumulation occurs predominantly on the surface of the sheath liquid coming out of the outer co-axial capillary [94]. The pendant droplet of the sheath solution elongates and stretches due to charge-charge repulsion to form a conical shape and once the charge accumulation reaches a certain threshold value due to the increased applied potential, a fine jet extends from the cone. The stresses generated in the sheath solution cause shearing of the core solution via “viscous dragging” and “contact friction” [99]. This causes the core liquid to deform into the conical shape and a compound co-axial jet develops at the tip of the cones. This is illustrated in Figure 2.2. On the way to the collector, as happens in the single fluid electrospinning, the jet undergoes a bending

instability and follows a back-and-forth whipping trajectory, during which the two

solvents evaporate and core-sheath nanofibers are formed [93, 95].

Figure 2.2 Schematic illustration of compound Taylor cone formation (A: Surface charges on the sheath solution, B: viscous drag exerted on the core by the deformed sheath droplet, C: Sheath-core compound Taylor cone formed due to continuous viscous drag)

2.3. Fabrication and Characterization of Collagen-PCL Sheath-Core Fibers

In this study, collagen-PCL sheath-core bicomponent fiber scaffolds were prepared through the coaxial electrospinning technique and the detailed experimental procedure is described as follows.

2.3.1 Materials

The collagen (semed S, acid-soluble), principally collagen type I with ca. 5% type III from bovine dermis, was a generous gift from Kensey Nash Corporation. PCL (Mw

80,000), chloroform, ethanol, 1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride (EDC), and N-hydroxysuccinimide (NHS) were purchased from Sigma-

Aldrich Co. Ltd. and used as-received without further purification. Potassium chloride, sodium chloride, monobasic potassium phosphate and dibasic sodium chloride heptahydrate were purchased from Fisher Scientific and used without further purification.

Deionized (DI) water was obtained using a Barnstead Nanopure low pressure, reverse

osmosis purification system. Phosphate buffered saline (PBS) 20x buffer was prepared

according to the literature [100] by dissolving 4 g potassium chloride, 160 g sodium

chloride, 4 g potassium phosphate monobasic, and 43.2 g sodium dibasic heptahydrate in

1 L deionized (DI) water.

2.3.2 Coaxial Electrospinning of Collagen and PCL

A collagen solution was prepared by dissolving Type I collagen into water-alcohol-salt

benign solvent system mentioned in Chapter I. Briefly, 0.4g of Type I collagen was

added into a mixture of 1mL ethanol and 1mL 20x PBS buffer. Complete dissolution was

observed after 10 minutes of mixing with a magnetic stir bar at room temperature. The

collagen concentration was calculated to be 16% by weight.

A PCL solution was prepared by dissolving 1.65g of PCL into 10mL chloroform to give a

concentration of 10% by weight. The PCL-chloroform mixture was stirred gently for 24 hours at room temperature in order to obtain a homogeneous solution. Both the PCL and collagen solutions were then loaded into 5ml BD plastic syringes, respectively.

A co-axial electrospinning approach was used to produce the hybrid collagen-PCL

sheath-core structures. A compound spinneret to hold solutions for the sheath (collagen)

and the core (PCL) was constructed as shown in Figure 2.1. The larger capillary (OD 2.0

mm, ID 1.5 mm) was connected to the syringe containing collagen solution, and the

smaller capillary (OD 1.0mm, ID 0.5 mm) was connected to the PCL solution syringe to

create the “co-axial” configuration at the tip. The syringes were independently driven by

syringe pumps (Kd Scientific Pump Systems) to control the flow rates of the two

materials. A high-voltage supply (Series EL, Glassman High Voltage, Inc.) was

connected between the co-axial capillary and the mandrel. The distance between the end of the capillary and the point of collection was kept constant at 15 cm. The applied

voltage and the flow rates of the solutions were adjusted as needed to achieve the

maximum stability of the compound Taylor cone. The voltage was maintained at 20 kV

and the flow rates for the collagen sheath and PCL core solutions were maintained at 0.5

mL/h and 0.3 mL/h, respectively.

For control purposes, pure collagen nanofibers from the benign solvent system and pure

PCL fibers from chloroform solution were also prepared through single jet

electrospinning using the same processing parameters as that of co-axial electrospinning.

2.3.3 Post-crosslinking Treatment

Crosslinking of electrospun collagen-PCL core-sheath scaffolds was done by soaking the collected mats in 200 mM of 1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride (EDC) and 200 mM of N-hydroxysuccinimide (NHS) ethanol solution for four hours. After crosslinking, the samples were rinsed with water to remove any non- crosslinked collagen and EDC / NHS residues.

2.3.4 Characterization of Fiber Morphology

Morphological characterization was performed using an optical microscope (Nikon

Eclipse TS100) and by scanning electron microscopy (SEM, JEOL JSM-6510 LV). For the optical microscope observation, the electrospun fibers were collected on glass slides and the corresponding photos were taken at 200 x magnification by using QCapture Pro software. For SEM imaging, all samples were sputter-coated with 10 nm gold layer using

an Anatech Hummer 6.2 sputtering system and observed at 5 kV in the SEM. Fiber diameters and distributions were assessed using NIH Image J software. The image pixels were calibrated using the pixel length of a magnification bar in micrometers generated on the SEM image. To measure the fiber diameter, a line was drawn on the fiber perpendicular to its longitudinal axis. The length of the line was automatically converted into nanometers by the software, producing a fiber diameter value at that location. From each image, 50 readings were taken to calculate the average value of the fiber diameter for each electrospun scaffold and to determine the standard deviation.

Sheath-core morphology within the fibers was confirmed by a freeze-fracturing technique.

The fiber mats were immersed in liquid nitrogen for approximately 10 seconds. While still in the liquid nitrogen bath, the scaffold fibers were fractured at various locations using sharp tweezers to induce differential fibrillation between the collagen sheath and the PCL core. The fractured fibers were then sputter-coated and viewed using SEM.

2.3.5 Characterization of Mechanical Properties

Mechanical property analysis of the electrospun core-sheath fiber mats was performed using dynamic mechanical analysis (DMA). A sample of each electrospun scaffold with approximately a 5:1 length: width ratio was cut and mounted in a tension film clamp.

Using the controlled force mode, the samples were loaded in tension at 2 MPa/min at room temperature until failure to obtain the stress-strain profile. Universal Analysis software (TA Instruments) was used to determine the elastic modulus, and stress and strain at break, of all samples. Samples of pure collagen and pure PCL electrospun scaffolds were also evaluated under identical loading conditions for comparison with the collagen-PCL bicomponent scaffolds.

2.4 Results and Discussion

2.4.1 Core-Shell Fiber Formation

Previous studies have shown that the collagen nanofiber scaffolds reconstituted from the water-alcohol-salt benign solvent system inevitably have insufficient resistance in water and poor mechanical firmness to resist handling [72]. This is because a large amount of salt and alcohol components present in the benign solvent system may have some hydrophobic and hydrophilic interactions with collagen bundles which resulted in weakly bonded fragments of collagen molecules. Figure 2.3 shows that the pure collagen nanofiber scaffolds processed from the benign solvents by the single jet electrospinning technique disintegrated and dissolved almost immediately after soaking in water. Their mechanical weakness limits the applications of such collagen scaffolds.

Figure 2.3 Appearances of electrospun pure collagen mats from water-alcohol-salt

benign solvent system before (a) and after (b) soaking in water.

Effective use of polymer nanofibers for tissue engineering relies not only on the biochemical characteristics of the fibers, but also on the mechanical integrity of the scaffolds to withstand manual manipulation [91]. The incorporation of PCL into collagen

37 nanofibers aims to impart the strength and elasticity of a synthetic polymer into the fibrous scaffolds without offsetting the excellent biocompatibility of collagen. Normally, functionalizing collagen nanofibers with PCL can be achieved through two methods. One approach involves mixing of PCL with collagen solutions to prepare bioactive composite nanofibers by means of electrospinning [81, 82, 101]. Due to the different nature of the synthetic polymer PCL and the biological polymer collagen, their co-solvents are limited.

To date, 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP) [81, 82] and 2,2,2-trifluoroethanol

(TFE) [101] appear to be the only solvents that can be used to dissolve collagen and PCL together without the aid of surfactants. However, given the corrosive nature and toxicity of HFIP and TFE, extra care is needed while handling it, which limits its applications.

Moreover, recent research has suggested that collagen fibers electrospun from HFIP and

TFE lack native ultra-structures [102].

The second method used to functionalize collagen nanofibers with PCL is to fabricate a collagen-PCL sheath-core bicomponent structure. This approach can be broadly viewed as combining materials such that the two materials maintain their separate identities, with the core material completely surrounded by the sheath material. Core-sheath fiber formation could also be viewed as a one-step process for obtaining a surface-modified or a coated product. It is hypothesized that a bicomponent structure would be differentially biodegradable in that the natural polymer sheath would degrade relatively faster, after initiating cell activity, and the synthetic polymer core would degrade at a much slower rate and continue to support the cell growth over a longer period of time. Therefore, the collagen-PCL sheath-core bicomponent structure was the goal of current work to enhance

the properties of the collagen nanofibers which are electrospun from the benign solvent

system.

In order to produce such a desirable collagen-PCL sheath-core bicomponent structure, a coaxial electrospinning setup was developed in this study. With optimized electrospinning parameters, bead-free and relatively uniform ultrafine bicomponent fibers of collagen and PCL were successfully fabricated as shown in Figure 2.4. An initial assay of the resultant sheath-core structure was carried out by a stability test of co-axial electrospun scaffolds in different solvents. As we know, the synthetic polymer PCL can readily dissolve in chloroform. The solubility test in chloroform (Figure 2.4 a, b) shows little change of fiber morphology, if any, in the electrospun scaffolds which suggests that the PCL component is not exposed at the fiber surface and received protection from the outer collagen layer. As mentioned above, the electrospun collagen is water-soluble.

Figure 2.4 c and d show a major morphology change after soaking the coaxial electrospun scaffolds in the water for approx. 30 minutes. This is because the removal of collagen sheath by water resulted in larger spaces left between the electrospun fibers and a loose scaffold was formed. Therefore, these solubility tests in chloroform and water may indicate a “coating” of collagen onto individual PCL nanofibers.

Figure 2.4 Appearances of collagen-PCL sheath-core bicomponent scaffolds (a, c) deposited on glass slides and soaking in chloroform (b) and water (d) for 30 min, respectively.

Fiber morphology was also characterized by scanning electron microscopy. Typical SEM images of collagen-PCL bicomponent fibrous scaffolds are shown in Figure 2.5, and it can be seen that the coaxial electrospun fibers possess a common feature of a random array and a very porous structure. The average fiber diameter is in the range of 0.95 ±

0.23 µm. After dissolving the collagen sheath in water, the average fiber diameter decreases to 0.65 ± 0.15 µm as shown in Figure 2.5c. Therefore, the thickness of the collagen layer coated on the PCL core is roughly calculated to be approximately 300 nm.

Figure 2.5 SEM images of the coaxial electrospun collagen-PCL bicomponent fibers before (a) and after (b) water-treatment. (c) Distribution of resulting collagen-PCL nanofiber diameter.

The technique of freeze-fracturing was employed to confirm the presence of the core- sheath structure. In this technique, the fibers were frozen using liquid nitrogen and were fractured at various places to induce differential fibrillation in the sheath and the core.

The fibers were subsequently viewed under SEM. Figure 2.6 shows the resulting SEM images which clearly give the evidence of the presence of core-sheath arrangement of the two components. This technique exploited the differences in the elastic properties of the

41 sheath and the core materials in order to demonstrate their arrangement. Collagen electrospun from the benign solvent system became rigid and highly fragile after freezing and could be fractured. The more elastic PCL could not be fractured as easily and remained intact shown in Figure 2.6.

Figure 2.6 SEM images of the freeze-fractured collagen-PCL sheath-core nanofibers.

For comparison purposes, the electrospun mono-component collagen and PCL fibrous scaffolds were also prepared through single jet electrospinning technique using the same processing parameters as that of co-axial electrospinning, and the typical SEM images are shown in Figure 2.7. The average diameter of electrospun collagen nanofibers was in the range of 0.24 ± 0.06 µm, while mono jet electrospinning of PCL solution surprisingly produced micron-size fibers with average diameter of 3.12 ± 0.23 µm. The fiber scaffolds obtained using two electrospinning approaches (traditional and coaxial) lead to a big difference in fiber diameters, with the bicomponent fibers having a diameter that is in between those of the two mono-components. It seems that the combination of a PCL core and a collagen shell results in increasing the size of the collagen nanofibers and shrinking the PCL core.

Figure 2.7 SEM images of electrospun mono-component collagen (a) and PCL (b) fibers

A plausible reason for this phenomenon is the formation of a compound Taylor cone.

Although the complex electrohydrodynamics involved in the coaxial electrospinning is

yet to be determined, the difference between the sheath and core solvents is believed to

have a major impact on the spinning process [92]. In coaxial electrospinning, the

compound jet formation from the sheath and the core solutions is the result of the transfer

of shearing forces from the sheath solution to the core solution through “viscous dragging”

and “contact friction,” as mentioned in the former section [99]. High interfacial tension

between the solutions hampers this mechanism and the compound Taylor cone does not

form. It has been shown by Diaz et al. that by increasing the similarity of the two

solutions, a compound Taylor cone can be formed [103]. In this study, a stable compound

Taylor cone was observed when using a collagen water-alcohol-salt solution as the sheath and a PCL-chloroform solution as the core under the conditions described in the experimental section. This may indicate some compatibility between the benign solvents and chloroform. The alcohol component present in the collagen solution may not only facilitate the dissolution of collagen, but also help to minimize the interfacial tension between sheath and core by increasing the affinity to chloroform. Furthermore, the large

amount of phosphate salts in the benign solution ensures a high conductivity of the

solution and strong electrical forces occurring on the sheath. Therefore, in the resultant

compound Taylor cone, the shearing forces generated between the collagen and PCL

solutions may be strong enough to shrink down the size of the PCL core to the nano scale.

2.4.2 Post-Crosslinking of Collagen-PCL Hybrid Fibers

Although the incorporation of the PCL core increased the size of hybrid collagen fibers, the electrospun collagen shell was still too weak for tissue engineering applications. In order to enhance the stability of collagen sheath in water and further improve the

mechanical durability of collagen-PCL sheath-core bicomponent scaffolds, post- crosslinking of hybrid collagen-PCL fibers was performed by immersing the scaffolds into EDC and NHS in ethanol for 4h, followed by rinsing with water to remove the crosslinker residues and non-crosslinked collagen. As discussed in the Chapter I, EDC and NHS can induce the formation of an amide bond by activation of the side chain carboxylic acid groups of aspartic and glutamic amino acids in collagen, followed by aminolysis of the intermediate o-isoacylurea by the primary amino groups of lysine or hydroxylysine residues on the adjacent collagen fibrils, forming intra- and inter-helical crosslinks. A typical SEM image of the crosslinked collagen-PCL sheath-core scaffolds is shown in Figure 2.8, which exhibits a highly porous structure with some adhesion occurring between the fibers. This high porosity may have a big potential in biomedical applications, because it may enhance nutrient delivery and the removal of metabolic wastes, while simultaneously providing space for cell penetration and migration into the structure [83, 84].

Figure 2.8 SEM image of the post-crosslinked collagen-PCL sheath-core fiber scaffolds

2.4.3 Mechanical Properties of Collagen-PCL Scaffolds

To assess the mechanical properties of the bicomponent scaffolds, the tensile response of the electrospun constructs and the impact of post-crosslinking on the mechanical response were evaluated and compared to that of the mono-component scaffolds fabricated by the single jet electrospinning technique. Due to the extremely poor mechanical strength, the electrospun pure collagen scaffolds did not allow DMA testing, and thus no valuable data was obtained. The addition of the PCL core into collagen nanofibers definitely increased mechanical properties and the representative stress-strain curves of collagen-PCL sheath-core bicomponent fiber scaffolds are shown in Figure 2.9.

Pure PCL PCL/Collagen (Crosslinked) 3.0 PCL/Collagen (Uncrosslinked)

2.5

2.0

1.5

Stress (MPa) 1.0

0.5

0.0 0 20 40 60 80 100 Strain (%)

Figure 2.9 Tensile stress-strain profiles of electrospun scaffolds.

It is not surprising that the electrospun pure PCL scaffolds render the best mechanical behavior in comparison. The stress-strain profile for the PCL scaffolds has a shape characteristic that is similar to that of many other synthetic polymers, with a linear elastic region followed by a period of plastic deformation until failure. The tensile response of the uncrosslinked collagen-PCL bicomponent scaffolds, however, only shows a weak elastic behavior before failure. Since the uncrosslinked collagen layer is fragile and has little contribution to the tensile properties, the tensile response of uncrosslinked collagen-

PCL bicomponent fibers mainly reflects the mechanical properties of the PCL core. The unexpected low tensile properties of the uncrosslinked collagen-PCL hybrid fibers may indicate a non-uniform dispersion of PCL core in the collagen nanofibers. In other words,

some discontinuity of the PCL core may occur inside the collagen fibers leading to the weak tensile behavior. Post-crosslinking of collagen markedly improves the mechanical strength of collagen-PCL scaffolds, and a similar linear elastic region and plastic deformation were presented on the corresponding stress-strain profile.

Table 2.1 Tensile properties of electrospun collagen-PCL bicomponent scaffolds.

Elastic Ultimate Tensile Ultimate Samples Modulus (MPa) Strength (MPa) Elongation (%)

PCL 10.5 2.98 99.9

PCL/Collagen 4.1 0.77 89.4 (Crosslinked)

PCL/Collagen 0.5 0.30 95.7 (Uncrosslinked)

The elastic modulus, as well as the ultimate tensile strength (the stress at failure) and the

ultimate elongation (the strain at failure) of coaxial electrospun collagen (crosslinked and

uncrosslinked)-PCL sheath-core fiber scaffolds and pure electrospun PCL, are shown in

Table 2.1. It can be seen that post-crosslinking treatment improves the mechanical

properties of collagen-PCL bicomponent scaffolds. The elastic modulus and tensile

strength increase dramatically after post-crosslinking of the collagen layer, and the

ultimate elongation decrease slightly. These results are expected, as the collagen layer

became more robust after post-crosslinking and made contributions to the tensile

properties of the scaffolds.

2.5 Conclusions

In this study, a novel variation on traditional electrospinning was made by arranging two

capillaries coaxially to spin the core-sheath bicomponent nanofibers of a natural and a

biodegradable synthetic polymer. A collagen-PCL sheath-core structure was generated using the collagen benign solvent solution and PCL in chloroform, and such a hybrid structure was assayed in solubility tests with different solvents (i.e. water and chloroform) and a core-shell structure was confirmed in the freeze-fracture experiments. The collagen-PCL sheath-core bicomponent fibers had a diameter of 0.95 ± 0.23 µm and the collagen sheath was of approximately 0.30 µm in thickness.

The alcohol component present in the collagen benign solution was assumed to increase the similarity between the collagen solution and the PCL solution, and thus reduced the interfacial tension between the core and the sheath, which facilitated the formation of a compound Taylor cone. Meanwhile, the phosphate salts in the benign solvent ensured a high conductivity of the collagen solution and strong shearing forces occurring on the sheath, which assisted the coaxial electrospinning process.

The addition of the synthetic polymer PCL core into collagen nanofibers increased the mechanical properties of collagen scaffolds electrospun from the benign solvent system.

Further improvement was done by post-crosslinking of the collagen-PCL sheath-core

bicomponent fibers in an EDC/NHS ethanol solution. A highly porous structure was

retained after crosslinking.

It has been reported that a proper match of the mechanical properties and the loosely

laced fibro-porous structure could favor the cells modulating and the artificial ECM

48 toward favorable interactions for synthesizing a “real” tissue [9]. At this point, collagen-

PCL sheath-core bicomponent fibrous scaffolds prepared in this study could be potentially effective in creating bioactive scaffolds because they may mimic the natural

ECM to a large extent in terms of the physical structure and biochemical characteristics.

Chapter III In-situ Crosslinked Collagen Nanofibers

3.1 Introduction

Collagen fibers are of particular interest and importance in tissue engineering and

regenerative medicine [62, 104]. Owing to high porosity and large surface area, non-

woven scaffolds of electrospun collagen nanofibers can serve as an ideal engineering

scaffold to mimic the biochemical and ultra-structural properties of the native

extracellular matrix (ECM) of tissues [75, 105]. However, when pure collagen is

regenerated by means of electrospinning, no matter what solvent (e.g. acidic water, HFIP,

or benign mixture) is used, the resulting fibers inevitably present insufficient resistance in

water and poor mechanical firmness to resist handling [66, 69, 72]. This is because the

lack of native inter- and intramolecular crosslinks in the reconstituted collagen leads to weakly bonded fragments of collagen molecules [9].

As discussed in Chapter I, much of the effort and the energy in the past has been focused

on crosslinking of collagen, such as exposure to ultraviolet light [56], enzymatic

treatment using transglutaminase [61], and the use of chemical crosslinking agents such

as glutaraldehyde (GA) [36, 37], epoxy compounds [44], 1-ethyl-3-(3-dimethylaminopropyl)-1-carbodiimide hydrochloride (EDC) and N-hydroxysuccinimide (NHS)

[72]. Among them, chemical crosslinking normally produces the materials with the highest degree of crosslinking when compared with photochemical and enzymatic methods [106]. Generally, a complementary, exogenous crosslinking agent is required if pure electrospun collagen is to be used as tissue engineering scaffolds.

The choice of crosslinking agent requires knowledge of the reactive groups present,

typically an amino acid side chain, and appropriate ambient conditions (e.g., pH,

temperature, solvent) that do not negatively affect the protein [10]. Van Wachem et al.

compared the four chemical cross-linking agents GA, epoxy, EDC and acyl azide with respect to biocompatibility and tissue regenerating capacity [107]. The most promising

results were obtained with EDC-crosslinked material which resulted in a mild cellular response after implantation with increased neutrophil and macrophage infiltration. In combination with the slow degradation rate it allowed replacement by newly formed collagen and tissue regeneration. Thus, EDC has been widely used to crosslink reconstituted collagen fibrils (i.e. collagen molecules removed from tissues via acid or enzyme extraction that spontaneously form fibrils), insoluble collagen, and collagen fibers from intact tissues.

However, typical protocols for crosslinking electrospun collagen with EDC frequently suffer a negative result when the structural aspect of the scaffold is considered [72, 108].

It has been reported that the collagen scaffold shrinks during crosslinking and the open

structure which is desirable for cell repopulation on the scaffold is almost obstructed by

film-like structure on the surface of the electrospun mats [108], as shown in Figure 3.1,

hence defeating the purpose of electrospinning collagen in the first place. Although

electrospun blends of collagen with other natural or synthetic polymers (Chapter I) or

fabrication of core-sheath collagen fibers through coaxial electrospinning (Chapter II)

have been proven successful in maintaining fibrous structure and initial cell studies, it is

desirable to be able to work with collagen-only electrospun scaffolds for such

applications as cartilage tissue engineering or skin replacement.

Figure 3.1 Scanning electron microscopy micrographs: samples cross-linked with (A)

20-mM 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC) (B) 200- mMEDC (C) 20-mM EDC/N-hydroxysuccinimide (NHS) (D) 200-mMEDC/NHS (E) 3% glutaraldehyde (room temperature), (F) 50% glutaraldehyde (heated), (G) 100% ethanol- soaked, and (H) no treatment (dry); scale bar is 10 mm. [108]

Furthermore, conventional methods of creating crosslinked collagen fibers, based on

physical or chemical treatments, are always two-step methods: collagen fibrous scaffold

preparation followed by post-production crosslinking. Such post-crosslinking procedures frequently increase the complexity of the collagen production and can be technically limited by a high cost-benefit ratio for large scale implementation [106]. Therefore, it is

highly desirable to simplify the procedures and solve the aforementioned negative

influences brought by EDC post-crosslinking for all applications where collagen fiber scaffolds would be used. Accordingly, the focus of this chapter is to develop an advanced method to combine fiber formation and post-crosslink treatment in a single step to fabricate water-stable collagen scaffolds and maintain the desirable fibrous morphology.

3.2 Fabrication and Characterization of In-situ Crosslinked Collagen Nanofibers

In-situ crosslinked collagen is prepared by incorporating specific crosslinking agents into

fibers during electrospinning, thereby enabling collagen to crosslink spontaneously after

fiber formation. The experimental steps are detailed as follows.

3.2.1 Materials

Lyophilized collagen (semed S, acid-soluble), principally collagen type I with ca. 5% type III, from bovine dermis was kindly gifted by Kensey Nash Corporation. Ethanol,

EDC, NHS, magnesium nitrate hexahydrate and potassium carbonate were purchased from Sigma-Aldrich Co. Ltd. and used without further purification. Potassium chloride, sodium chloride, monobasic potassium phosphate, dibasic sodium chloride heptahydrate and magnesium chloride hexahydrate were purchased from Fisher Scientific and used

without further purification. Phosphate buffered saline (PBS) 20x buffer was prepared

according to the literature [100].

3.2.2 Preparation of In-situ Crosslinking Collagen Solution

In-situ crosslinking collagen solution was prepared by mixing the crosslinkers and

collagen in a very particular order. First EDC and NHS were added to ethanol in pre-

calculated quantity and ratio. The ethanol mixture was then magnetically stirred until the

EDC and NHS were fully dissolved. Then 20X PBS buffer was added to the ethanol

mixture in a buffer: ethanol ratio of 50:50 by volume. The collagen was then added to the

benign mixture at a concentration of 16% by weight. It was important to add the collagen

quickly after mixing the buffer and ethanol, as otherwise phase separation occurred and

the salts in buffer precipitated in ethanol. Once all of the collagen had been added, it was

magnetically stirred for roughly 5 minutes in order to make a uniform solution which was

then placed in a syringe and used either for electrospinning or rheological experiments.

3.2.3 Rheological Experiment

Dynamic oscillatory shear experiments were carried out on the in-situ crosslinking

collagen solutions with different molar ratio of NHS to EDC at room temperature. The

concentration of EDC was fixed at 200mM in ethanol. A 40 mm diameter, 2 steel cone

and plate geometry was used with a solvent trap which helped to prevent the ⁰evaporation

of the alcohol from the sample. The oscillatory mode had a frequency of 6.283 rad/s and

a strain of 0.2% and longtime sweep experiments were conducted. The storage (G’) and

loss (G”) modulus as well as the loss tangent, tan σ (= G”/G’), were recorded as a

function of time on TA Instruments AR-2000ex Rheometer. The apparent gelation time

was rheologically determined by the time at which G’ was equal to G”.

3.2.4 Electrospinning of In-situ Crosslinked Collagen Nanofibers

The freshly-made in-situ crosslinking collagen solution was loaded into a 5mL BD syringe with a 21 gauge blunt needle, which was then placed in a syringe pump. The process parameters (e.g. flow rate, potential field, and needle-collector distance) and environmental conditions (e.g. humidity and temperature) were varied to optimize the stability of the electrostatic jet. Electrospinning was usually carried out at 20 kV with a pump rate of 0.5mL/h at a relatively low humidity (20%) controlled using Drierite desiccant (anhydrous calcium sulfate). A drum rotating at 5m/s was placed 12cm from the needle and the electrospun fibers were collected at room temperature. The resulting collagen fiber scaffolds were then stored in desiccators under controlled humidity.

Saturated solutions of magnesium chloride, potassium carbonate and magnesium nitrate were used to provide the relative humidity of 33%, 43% and 53% in desiccators [110],

respectively.

3.2.5 Characterization of Fiber Morphology

In order to monitor the change of fiber morphology during the electrospinning process,

small amount of the electrospun collagen fibers were collected on glass slide at various

time points and observed using an optical microscope (Nikon Eclipse TS100).

Morphological characterization was also performed on a JEOL JSM-6510LV scanning electron microscope (SEM) in secondary electron imaging (SEI) mode. Electrospun collagen mats were sputter-coated with a 5 nm gold layer and observed in SEM at 5 kV.

Data treatment was done with free Image J software, as described in Chapter II. The images were acquired at three magnifications: 1K, 5K and 10K. The lower magnification allowed the examination of the morphology of a larger sample of the fiber mat. The

higher magnification images were used for examination of local features and estimation

of the fiber diameters.

Water-treated, in-situ crosslinked electrospun collagen samples were prepared by soaking

the pre-made crosslinked collagen scaffolds in PBS for approx. 30 min followed by

rinsing with water, and then dried in vacuo for sputter-coating and SEM imaging.

3.2.6 Fourier Transform Infrared (FTIR) Spectroscopy

An ABB Bomem MB-104 Spectrophotometer was used to perform the FTIR analysis of

the samples. A small piece was cut from the electrospun collagen sample and was fixed

onto the FTIR holder. A spectrum showing the absorbance at 600-2000 cm-1 was acquired

using the Bomem Grams software. The FTIR spectrum of Kensey Nash collagen and

post-crosslinked collagen were also recorded as controls.

3.2.7 Characterization of Mechanical Properties

Mechanical testing of the electrospun collagen scaffolds was performed on a dynamic mechanical analyzer (DMA-Q800, TA Instruments Inc.) in the ramp force mode.

Collagen samples were carefully cut into rectangular strips with approximately a 5:1 length: width ratio. The width and thickness of the fibrous specimen were measured and loaded into the tensile testing fixture, and a ramp force of (0.5 x width x thickness) N/min was applied at room temperature to obtain the stress-strain profile. Universal Analysis software (TA Instruments) was used to determine elastic modulus, and stress and strain at break of all samples.

Mechanical properties of the crosslinked mats were also measured in the hydrated state prepared by soaking the mats in PBS for approximately 30 minutes at room temperature.

Uniaxial tensile testing of these mats was performed as described above.

3.3 Results and Discussion

3.3.1 In-situ Crosslinking Approach

Chemical crosslinking of collagen with EDC and NHS is one of the best methods to produce non-toxic, water-stable collagen products [106, 107]. A literature survey about collagen crosslinking shows that very different concentrations of EDC and NHS have been used, and the molar ratio of EDC and NHS varies from one study to another [47-51,

106-109]. Generally, a smaller amount of NHS, compared to EDC, is used because it acts as a catalyst and can theoretically be fully recovered after the crosslinking reaction.

However, for the first time, an excess amount of NHS present in crosslinking reaction has been found by our laboratory to be able to delay the total coupling reaction in a controlled fashion. A series of rheological analyses has been performed on the collagen benign solutions containing different molar ratios of NHS and EDC at room temperature. The apparent gelation time, which is the time for the collagen solution to become a gel, is determined rheologically in dynamic oscillatory shear experiments and the relationship between the apparent gelation time and the molar ratio of NHS and EDC is plotted in

Figure 3.2.

When the concentration of EDC is fixed at 200mM, the apparent gelation time is highly dependent on the amount of NHS in solution. It can be seen in Figure 3.2 that the addition of NHS to the EDC solution has little influence on the apparent gelation time

57 initially. The collagen benign solution turned to a gel in minutes when the concentration of NHS was relatively low. However, at the point where the ratio rose up to about 1.5, there was a prompt increase in the apparent gelation time, and then it increased almost linearly with the molar ratio of NHS to EDC in the range higher than about 2. Therefore, if the amount of NHS added to collagen solution exceeds twice as much as that of EDC, the apparent gelation time will increase from minutes to hours, which is very valuable for engineering design. During this desirable period, the collagen benign solution can be processed into a variety of shapes and morphologies. Significantly, electrospinning of collagen fibers from the benign solvent system became possible, with crosslinking occurring in-situ afterward.

200

150

100

50 Apparent Gelation Time (min) 0

0.5 1.0 1.5 2.0 2.5 3.0 3.5 Ratio of NHS/EDC

Figure 3.2 Plot of the apparent gelation time as a function of the molar ratio of NHS and EDC.

3.3.2 Formation of In-situ Crosslinked Collagen Nanofibers

In-situ crosslinking fibers were successfully fabricated from a collagen benign solution containing 200mM EDC and 400mM NHS through a traditional single jet electrospinning setup described in Chapter I. Typical SEM images of the electrospun collagen fibers are given in Figure 3.3, which show cylindrical and continuous nanofibers possessing a common feature of random arrays and a very porous structure. The average fiber diameter calculated for in-situ crosslinking collagen fiber scaffolds is in the range of 0.42

± 0.11 µm, which is almost as twice as the size of the collagen fibers generated from the benign solution without crosslinkers, as shown in Figure 2.7. This is expected, because the addition of crosslinker EDC into collagen solution inevitably increased the viscosity even though gelation process was postponed by the presence of excess amount of NHS, and higher viscosity of collagen solution resulted in larger fibers formed. Similar results were also found in rheological experiments. The storage modulus (G’) of in-situ crosslinking collagen solution kept increasing until the dynamic oscillatory shear testing finished, which indicated that the viscosity of solution gradually increased after mixing the collagen and the crosslinkers together.

Figure 3.3 Typical SEM images of electrospun in-situ crosslinked collagen fiber at low

(a) and high (b) magnification. (c) Distribution of fiber diameter. (d) Diameter of in-situ

crosslinked collagen fibers increased during electrospinning processing.

According to the rheological experiments, the in-situ crosslinking collagen solution containing 200mM EDC and 400mM NHS became a gel in about one and a half hours after the preparation of crosslinking solution. During this period, the viscosity of solution increased with time. Accordingly, the electrospun collagen fibers should become thicker and thicker. Figure 3.4 shows typical SEM images of the electrospun in-situ crosslinked collagen fibers collected on glass slides at different time points. The high magnification images suggest that some crosslinks apparently occurred between the collagen fibers,

60 which are not observed in the electrospinning of collagen benign solvent solution. The statistical analysis of fiber diameter was given in Figure 3.3d, which displays a slight increase from 0.355 to 0.456 µm during electrospinning processing.

Figure 3.4 SEM images of electrospun in-situ crosslinking collagen nanofibers collected

on the glass slides at various time points with low (a, c, e, g) and high (b, d, f, h)

magnifications.

The resulting electrospun, in-situ crosslinking collagen fibers were stored in the designated desiccators which had different humidity levels. The equilibrium relative humidity was controlled by the use of saturated salt solutions. These salt solutions were made up as a slushy mixture with distilled water and chemically pure salts and enclosed in the sealed glass desiccators. For humidity control purposes, saturated solutions of magnesium chloride, potassium carbonate and magnesium nitrate were used to provide the equilibrium relative humidity of 33%, 43% and 53% at room temperature [110],

respectively.

SEM images of electrospun, in-situ crosslinked collagen fibers stored at different

humidity levels for a period of time are shown in Figure 3.5. It appears that the relative

humidity is an important parameter to control the fiber morphology of in-situ crosslinked

collagen after electrospinning. Collagen fiber scaffolds stored at 33% humidity for 3days

(Figure 3.5a) still possessed individual fiber features, while some “melting” of fibers was

observed with samples stored at 53% humidity for only one day (Figure 3.5c). High

humidity might partially soften the collagen before they became crosslinked by

incorporated EDC and NHS. However, low humidity presented inside the chamber may

limit the motilities of the crosslinkers and the polymer chains in the collagen fibers which

resulted in low crosslink efficiency. The collagen fibers were found to be insufficiently

crosslinked at 33% humidity after 3 days and partially disintegrated when placed in the

water, while the sample stored at 43% for 3 days (Figure 3.5b) displayed a better result.

Therefore, a proper humidity is needed to control in-situ crosslinking collagen. In this study, 43% relative humidity provided by saturated potassium carbonate solution was used for the rest of the experiments. Furthermore, it is of interest to note that the influence of humidity on fiber morphology discussed above was not observed with the collagen fibers electrospun from benign solution without EDC and NHS, as shown in

Figure 3.5d.

EDC and NHS stored at humidity of 53% for 7days (d).

Post-crosslinking of collagen fiber scaffolds has also been performed by soaking the

collagen scaffolds electrospun from benign solution without crosslinkers into a 200mM

of EDC and NHS ethanol solution for 4 hours. After the post-crosslink process, the

resulting collagen scaffolds shrank to 40% of the original dimensions when immersed in

the water, as shown in Figure 3.6 a and b. In contrast, the in-situ crosslinked collagen

scaffolds normally swelled up to two-fold of the original size when they were hydrated, as shown in Figure 3.6 c and d. The prominent difference in water stability of collagen scaffolds crosslinked post-electrospinning vs. the in-situ method may arise from the distribution of crosslinkers in the collagen fibers. It has been reported that the coupling reaction in the traditional two-step crosslinking approach is mainly concentrated on the outer surface of collagen fibers due to the slow penetration of EDC and NHS into dense fibers, which is increasingly retarded as crosslinking continues at/near the surfaces [111].

Conversely, it is believed that EDC and NHS are more uniformly distributed in the collagen fibers by using the in-situ method developed in this work and that crosslinking reactions take place throughout the fibers, resulting in better crosslinked collagen scaffolds as shown in Figure 3.6.

The surface morphology of electrospun, in-situ crosslinked collagen scaffolds after immersing in water was characterized by scanning electron microscopy, and typical low and high magnification images are displayed in Figure 3.7. It can be seen that the water- treated, in-situ crosslinked collagen scaffolds still possess a very porous structure, which is one of the most important features of the scaffold to begin with. Compared to the mats shown in Figure 3.1, the porous collagen scaffolds prepared in this study may have better

64 cell penetration and interaction between cell and substrates, which makes them potentially more promising in biomedical applications.

Figure 3.6 Snapshots of electrospun collagen scaffolds post-crosslinked (a, b) and in-situ crosslinked (c, d) with EDC/NHS. The scaffolds were placed on the glass slides before

(a, c) and after (b, d) water treatment.

Figure 3.7 SEM images of electrospun in-situ crosslinking collagen scaffolds after water- treatment at low (a) and high (b) magnifications.

3.3.3 Mechanical Properties of In-situ Crosslinked Collagen Scaffolds

To evaluate the mechanical properties of the electrospun, in-situ crosslinked collagen

scaffolds, the uniaxial tensile response of the resultant mats were measured in both dry

and hydrated states. The hydrated samples were used to show tensile behavior of

electrospun in-situ crosslinked collagen fibers under approximate physiological

conditions.

Uniaxial tensile testing performed on the in-situ crosslinked collagen scaffolds indicated

a significant difference in tensile behaviors of the dry and hydrated samples. A

representative plot of the stress-strain relationship of the dry scaffolds exhibited a shape characteristic similar to that of many synthetic polymers, with a linear elastic region followed by a period of plastic deformation until failure, as shown in Figure 3.8. In contrast, the hydrated collagen scaffolds demonstrated a uniaxial J-shaped stress-strain

curve which is similar to that of many native tissues.

In-situ crosslinked collagen (dry) 16

6 Stress (MPa)

0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 Strain (%)

In-situ crosslinked collagen (hydrated) 0.25

0.20

0.15

0.10 Stress (MPa) 0.05

0.00

0 20 40 60 80 100 120 140 Strain (%)

Figure.3.8. Representative plots of stress-strain curves to demonstrate the difference in

behavior of in-situ crosslinked collagen dry and hydrated samples.

The typical tensile properties in terms of peak stress and strain at break are summarized in Table 3.1, in which the hydrated crosslinked collagen scaffolds show a significantly greater strain and a dramatically smaller stress than those of the dry samples. This is not surprising, because the electrospun in-situ crosslinked collagen scaffolds did not dissolve in water but swelled to approximately twice of the original dimensions due to the crosslink network, as discussed above. This expansion would increase the space between the collagen fibers and result in a loose porous structure formed, which makes for a low peak stress shown on the stress-strain curve of the hydrated samples. Furthermore, a large amount of water present in the hydrated crosslinked collagen scaffolds might increase the mobility of polymer chains. Consequently, a high strain was expected from the hydrated samples.

Table 3.1. Mechanical properties of in-situ crosslinked collagen scaffolds

Samples Peak Stress (MPa) Strain at break (%)

In-situ crosslinked collagen (dry) 14.53 ± 1.61 3.28 ± 0.12

In-situ crosslinked collagen (hydrated) 0.22 ± 0.02 134.5 ± 10.0

3.3.4 FTIR Spectra of In-situ Crosslinked Collagen Scaffolds

In order to explore the collagen structure present in electrospun in-situ crosslinked collagen scaffolds, FTIR spectra of the resulting collagen were generated using a Bomem

MB-104 spectrophotometer, and are compared to those of post-crosslinked collagen and

Kensey Nash collagen, as shown in Figure 3.9. According to the literature, native

collagen exhibits a peak at ca. 1655 cm-1 in the amide I region, which arises predominantly from protein amide C=O stretching vibrations, and a peak at 1545 cm-1 in

the amide II region, which is made up of amide N-H bending vibrations and C-N

stretching vibrations [112]. These two characteristic peaks are indicative of the collagen

triple-helix and the amide I peak is associated with the triple helix structure. When

collagen exhibits a triple helix structure, the amide group is used in hydrogen bonding in

each triplet (Gly-Pro-Hyp) and is characterized by a peak around 1655 cm-1 instead of the

typical 1620 cm-1 for gelatin, which is the denatured collagen [19]. The representative

amide I and II bands of Kensey Nash collagen, post-crosslinked collagen and in-situ

crosslinked collagen are summarized in Table 3.2, and show very similar data. As

expected, the FTIR data preliminarily indicate that the EDC/NHS crosslinking does not

impact the nature of the collagen in the mat and the triple helix structure present in the

electrospun in-situ crosslinked collagen fiber scaffolds. More detailed experiments, such

as transmission electron microscopy (TEM) and circular dichroism (CD), are in progress.

Table 3.2 Comparative study of FTIR spectrum wavelength in the amide I and amide II region for Kensey Nash collagen, post-crosslinked electrospun collagen and in-situ crosslinked collagen.

Sample Amide I band (cm-1) Amide II band (cm-1)

Kensey Nash Collagen 1655 1545

Post-crosslinked Collagen 1656 1547

In-situ crosslinked Collagen 1653 1541

1.2

1.0

0.8

0.6 Absorbance 0.4

(a) 0.2 (b) 0.0 2000 1800 1600 1400 1200 1000 800 600 Wavenumber (cm-1)

Figure 3.9 FTIR spectra of electrospun collagen scaffolds crosslinked by an in-situ method (a) and by a post method (b).

3.3.5 Plausible Mechanisms of In-situ Crosslinking Reactions

An initial attempt to explain the in-situ crosslinking collagen reaction is as follows.

According to literature, the EDC crosslinking reaction begins with the formation of an amide bond by activation of the side chain carboxylic acid groups of aspartic and glutamic acids, followed by aminolysis of the intermediate o-isoacylurea by the primary

amino groups of lysine or hydroxylysine residues on the adjacent collagen fibrils,

forming intra- and inter-helical crosslinks [108], as shown in Figure 3.10. However, the

amine-reactive O-acylisourea intermediate is susceptible to hydrolysis, making it unstable

and short-lived in aqueous solution. The addition of a small amount of NHS stabilizes the

amine-reactive intermediate by converting it to an amine-reactive NHS ester, thus

increasing the efficiency of EDC-mediated coupling reactions [109].

It is possible that the amine-reactive NHS ester is much more stable and thus less active

than the intermediate o-isoacylurea. Consequently, an excess of NHS present in reaction

mixture may convert most of active EDC intermediates to the less active NHS esters,

which inevitably slows down the coupling to a second collagen molecule. Moreover, the

newly-formed NHS esters may undergo nucleophilic attack from either a primary amino

group of second collagen molecule or the N-OH group of the NHS. The crosslinking rate

would then depend on competition between NHS and amino groups of collagen

molecules to react with an NHS ester already linked to a collagen molecule. Since excess

NHS molecules are small and more accessible to the amine-reactive NHS esters, an

‘NHS-ester exchange’ might dominate the reaction with the NHS esters. Therefore, the

excess amount of NHS present in the crosslink mixture retards the rate of amide bond

formation and eventually postposes the whole crosslinking reaction.

3.4 Applications of In-situ Crosslinked Collagen Fiber Scaffolds

Wound dressings from electrospun collagen nanofibrous scaffolds potentially offer more

advantages over conventional wound care materials [113]. With its huge surface area and

microporous structure, the nanofibrous scaffold could quickly initiate a signaling

pathway and attract fibroblasts to the dermis layer, which can excrete important

extracellular matrix components such as host collagen and growth factors to repair

damaged tissue [114, 115]. To assay the cytocompatibility and cell behavior of

electrospun in-situ crosslinked collagen nanofibers, cell attachment and spreading of

Chinese Hamster Ovarian (CHO) cells on the electrospun collagen mats were studied.

Additionally, the effect of electrospun in-situ crosslinked collagen scaffolds on open

wound healing in mice was briefly examined. Furthermore, the release of antibiotic drugs

from the collagen mats was also preliminarily investigated.

3.4.1 Cell Culture Studies

The cell culture studies were performed as a part of the joint project with the Biomedical

Engineering Department (BME) by Ms. Sonia Merritt (graduate student) under the

guidance of Dr. Horst von Recum (Associate Professor, BME).

3.4.1.1 Materials and Methods

The electrospun in-situ crosslinked collagen scaffolds were prepared as described above and sterilized under UV light for 30 minutes on each side prior to cell seeding. Chinese

Hamster Ovarian (CHO) cells were selected in this study due to ease of availability.

Initially, CHO cells were cultured in high-glucose Dulbecco's modified eagle medium

(DMEM) supplemented with 1% pennicillin/strempomycin and 10% fetal bovine serum

in a 10cm diameter tissue culture dish. The media was changed every three days until 80%

confluence was reached. For the cell culture study, the electrospun collagen mats were

submerged into the normal growth media, and cells were seeded onto the mats at 40,000

cells per cm2 (n=3). Cell morphology on the mats was observed by an inverted optical

microscope and images were taken 5 days after cell seeding.

3.4.1.2 Results and Discussion

A representative image of CHO cells seeded on electrospun in-situ crosslinked collagen

scaffolds is shown in Figure 3.11. Although some of the cells are out of the plane of

focus due to issues with getting the electrospun fibrous mat to lay flat in the center of the

cell culture dish, the image clearly shows a favorable cell attachment with spindle-shape

CHO cells spreading over the surface of in-situ crosslinked collagen scaffolds. Such

result indicates that trace amounts of crosslinker EDC/NHS and the crosslink by-product

1-ethyl-3-(3-dimethyl-aminopropyl)-urea present in the collagen fibers have little, if any, influence on the cellular behavior and the electrospun collagen mats prepared by in-situ crosslinking approach have expectantly good in-vitro biocompatibility and support proliferation and normal functions of CHO cells.

Figure 3.11 CHO cells growing for 5 days on electrospun in-situ crosslinked collagen

nanofibers. The image was taken at 20x magnification.

3.4.2 Open Wound Healing Test

A preliminary in-vivo mouse study for wound healing applications of electrospun in-situ

crosslinking collagen scaffolds was carried out as a joint project with the Dermatology

Department by Dr. Minh Lam (Assistant Professor, Dermatology) and Dr. Elma Baron

(Associate Professor, Dermatology).

3.4.2.1 Materials and Methods

The electrospun collagen mats (in-situ crosslinked and non-crosslinked) were prepared as described above and sterilized in 100% ethanol. BALB/c mice were chosen. Typically, hair on the backs of mice was removed with Nair two days before creating incisions.

After anesthetization, three full thickness wounds (i.e. tissue destruction extending through dermis) with a diameter of 6mm were cut from the back of the BALB/c mice using a surgical scissor. The electrospun collagen mats were cut and applied to extend

over the entire wound area. The “patched” mice were then monitored for 14 days,

specifically comparing the rate of wound closure (patched vs. control) and photos were

taken at pre-set time points using a SONY DSC-W310 camera.

3.4.2.2 Results and Discussion

In open wound-healing tests, three full-thickness circular wounds were made on the back

of each mouse. Figure 3.12 shows representative animals at day 0, and at days 5 and 10

after patching with electrospun collagen mats. The influence of collagen mats on wound

healing were empirically studied by comparing the wound treated with in-situ crosslinked

collagen with the one treated with non-crosslinked collagen and the control with respect

to the closing rate.

As expected from previous mechanical testing, the non-crosslinked electrospun collagen

scaffold exhibited a typical property of weak strength and quickly dissolved in the wound

area. In contrast, the in-situ crosslinked collagen mat became semi-transparent and

covered the wound for a prolonged period. One can see in Figure 3.12 that both in-situ

crosslinked and non-crosslinked collagen disappeared from the wound area at Day 5.

This maybe because the collagen scaffolds either disintegrated or remodeled by the local

cells, and these collagen components present in the wounds might also help cells excrete

important extracellular matrix such as dermal collagen and growth factors during the

wound healing process. It is also of interest to note that the wound treated with in-situ crosslinked collagen displayed a smaller scar when compared to the other two sample wounds. The suppression of scar synthesis, with the simultaneous synthesis of physiological tissues, might be achieved by implanting such porous collagen scaffolds

75 within the site of injury in order to host cells and guide their behavior toward regeneration.

At Day 10, it is clearly observed that the wound patched with in-situ crosslinked collagen sample displayed a pinkish and scarless feature, which indicated a complete closure, while the control wound was still open with a scar on it. These promising results preliminarily suggest that the crosslinkers EDC/NHS and urea byproduct have no harmful effect on the wound healing process and the electrospun in-situ crosslinked collagen scaffolds prepared in this study could be a good candidate to promote quick and scarless wound healing. Detailed histological examination of epithelialization and granulation are in progress.

3.4.3 Drug Release Studies

Drug delivery systems based on collagen have been tested in a variety of applications [9,

116]. The positive effect of collagen on cell interactions is the main reason for the

particular interest in local treatment of affected tissue for wound healing and skin

regeneration [117]. In this section, a preliminary study on drug release from in-situ crosslinked collagen scaffolds is summarized. Various types of drugs, such as the photodynamic therapy drug silicon phthalocyanine Pc4 (Figure 3.13), the antibiotic doxycycline hyclate, and bovine serum albumin (BSA) were added to the benign collagen solution and electrospun followed by in-situ crosslinking.

3.4.3.1 Materials and Methods

The photodynamic therapy drug Pc4 was a kind gift from Dr. Malcolm E.Kenney

(Professor, Chemistry) and dissolved in prepared EDC/NHS ethanol solution at a

concentration of 0.6 mg/mL. Bovine serum albumin conjugated with Texas Red (BSA-

TR) was purchased from Invitrogen and diluted to 1 mg/mL in 20x PBS buffer.

Doxycycline hyclate was purchased from Sigma-Aldrich Co. Ltd. and dissolved in 20x

PBS buffer at a concentration of 2 mg/mL.

Electrospun in-situ crosslinked collagen scaffolds containing different drugs were prepared using a procedure similar to that described in section 3.2. A few modifications were made by using EDC/NHS-Pc4 ethanol solution, BSA-PBS buffer and doxycycline-

PBS buffer to replace the corresponding parts in collagen mixture preparation, respectively. The resultant collagen mats containing Pc4 were examined by a laser scanning confocal microscope and the images were generated with Volocity software.

The fiber morphology of scaffolds having BSA-TR and doxycycline hyclate was

visualized by a fluorescence microscope (Nikon Eclipse TS100) and a scanning electron

microscope (JOEL JSM-6510LV), respectively.

To assay doxycycline hyclate released from the fibers, samples of the electrospun mats

were cut and placed in 10 mL of PBS at room temperature. At each time point, 100 µL of

PBS was removed and replaced with fresh PBS. The amount of doxycycline released was

quantified from the absorbance measured at 347 nm on a UV-VIS spectrometer (Perkin-

Elmer Lambda 800), compared to a standard doxycycline curve.

3.4.3.2 Results and Discussion

The hydrophobic compound Pc4 and hydrophilic protein BSA-TR were employed as

model drugs in this study to demonstrate the drug loading capability of electrospun in-

situ crosslinked collagen scaffolds. Typical laser optical images of electrospun mats

bearing these two drugs were shown in Figure 3.14. It can be seen that electrospun fibers

containing both drugs exhibit a uniform fluorescence light along the fiber direction,

which indicates a successful incorporation of both hydrophobic and hydrophilic drugs

into collagen fibers respectively, and suggests that an even distribution can be achieved.

Figure 3.13 Chemical structures of silicon phthalocyanine Pc4 (a) and doxycycline (b).

Figure 3.14 Optical images of electrospun collagen mats containing Pc4 (a, b) and BSA-

Texas Red (c, d). Images were taken at bright field (a, c) and laser fluoresce light (b, d).

The influence of addition drugs on the electrospun fiber morphology and the release behavior from collagen scaffolds were examined by using an antibiotic doxycycline hyclate as a model drug molecule. The SEM image of electrospun collagen and doxycycline hyclate fibrous scaffold is shown in Figure 3.15a, which exhibits a typical feature of crosslinked fibers with a very porous structure. The average fiber diameter was analyzed to be around 0.89 µm, which is larger than the normal in-situ crosslinked collagen fibers fabricated above.

Figure 3.15 SEM image of electrospun collagen fibers containing doxycycline (a). In

vitro release of doxycycline from collagen scaffolds (b).

The in vitro release of doxycycline from the collagen scaffolds in PBS at room temperature is depicted in Figure 3.15b. As can be seen in the release profile, there is a significant burst release, in which approximately 70% of the total drug incorporated into the scaffold is eluted. A plausible reason for this is the swollen state of electrospun collagen mat when placed in aqueous solution. It has been reported that drug release from collagen is governed by diffusion from the swollen matrices [117]. Therefore, the porous

structure and huge surface area (Figure 3.7) present in the hydrated collagen fibrous scaffold may facilitate the diffusion of doxycycline molecules through the fiber and enable a quick liberation into PBS buffer.

3.5 Conclusions

In order to solve the problems presented by traditional collagen crosslinking methods,

such as dimensional shrinking and loss of porous morphology, and to simplify the

crosslinking procedure for electrospun collagen scaffolds, a novel single-step, in-situ

crosslinking approach was developed in this study.

The excess amount of NHS present in the crosslinking mixture was found to delay the

EDC coupling reaction for several hours. Water-stable collagen nanofibers with a

diameter of 0.42 ± 0.11 µm were successfully generated by electrospinning an in-situ

crosslinking collagen solution during this period. The diameter of electrospun collagen

fibers slightly increased during electrospinning, which suggested the onset of in-situ

crosslinking.

Humidity was found to be an important factor to control the in-situ crosslinking rate and

the subsequent fiber morphology, and the humidity of 43% was found to be preferable to

afford a sufficiently-crosslinked collagen network within 3 days.

The in-situ crosslinked collagen scaffolds exhibited swelling behavior when placed in

water, in contrast to the dimensional shrinking frequently observed in post-crosslinked

collagen materials. The porous surface morphology of the resulting mats is preserved in

water-treatment which is one of the most important features of the in-situ crosslinked

scaffolds.

The in-situ crosslinked collagen demonstrated a similar uniaxial tensile behavior of

native tissue in mechanical testing and displayed a characteristic triple-helical structure in

FTIR spectra, which suggested the in-situ crosslinking approach did not significantly compromise the nature of collagen. This in-situ crosslinking technique may also have applicability with other biomaterials including polypeptides, proteins and polysaccharides, to produce versatile structures for tissue engineering.

Two plausible reasons for the delayed crosslinking reaction in the presence of excess

NHS are discussed in this chapter: one proposes a large amount of less active amine-

reactive NHS ester formed before the stable amide bond formation; the other concerns a

competition between the amino groups of second collagen molecule and the mobile NHS

molecules to attack the aforementioned NHS ester.

A preliminary study of the use of electrospun in-situ crosslinked collagen scaffolds in a

wound healing application showed very promising results. Trace amount of EDC / NHS

and subsequent urea byproducts incorporated in collagen scaffolds apparently did not

display a negative influence on CHO cell culture and open-wound closing tests. The collage scaffolds produced by the in-situ crosslinking approach may be good candidates to promote quick and scarless wound healing.

Preliminary drug release studies were carried out by loading both hydrophobic and

hydrophilic drugs into in-situ crosslinked collagen scaffolds through electrospinning. The

release profile of antibiotic doxycycline from the collagen mats exhibited a burst release

82 in PBS buffer due to the swelling behavior of crosslinked collagen. The combination of collagen and specific drugs may represent a convenient system for the local delivery of drug for wound healing and skin regeneration.

Chapter IV In-situ Crosslinked Collagen Gels and Sponges

4.1 In-situ Crosslinked Collagen Gels

Collagen gels have many favorable attributes for tissue engineering because they can act

as a “cage” to retain cells and carry bioactive components such as growth factors [118].

As early as 1962, Grillo and Gross described the preparation of collagen gels from

neutralized acidic collagen solutions, and proposed their use as medical implants [119].

Since then collagen gels have been identified as an excellent three-dimensional substrate for the adhesion, proliferation and differentiation of numerous cell types [8, 9]. The method of collagen gel preparation easily allows the direct injection of cells prior to gelling, thereby enhancing seeding efficiency and the homogeneous distribution of cells within the scaffold [120]. Furthermore, type I collagen gel exhibits a low degree of immunogenicity and the capacity for endogenous enzymatic degradation in the absence of adverse effects towards surrounding tissue. Collagen hydrogels are currently in clinical use for dura mater replacement [121] and cellular hydrogels have proven effective for the treatment of skin ulcers, burns and oral mucosal defects [122, 123].

Although collagen exhibits excellent biological properties, the application of highly hydrated collagen gels in tissue engineering is limited due to low mechanical strength

[123] and structural instability arising from mechanical tension imposed by the constituent cells [124]. Several approaches have been taken to overcome the mechanical and geometrical drawbacks of collagen based hydrogels, including cell-induced remodeling, a process that can be time consuming and unpredictable [125]. A variety of crosslinking methods using either chemical [37, 47], photochemical [126, 127], or

enzymatic agents [61, 128] have also been investigated. However, each of these methods

imparts some degree of cytotoxicity [129].

As mentioned in the previous chapter, the crosslinking of collagen by using EDC and

NHS is one of the best methods to produce non-toxic, water-stable collagen-based products [106, 107]. Preliminary cell culture study on the in-situ crosslinked collagen fibrous scaffolds suggested that the trace amount of EDC and NHS had no harmful influence on the cell attachment. Moreover, the in-situ crosslinking approach using EDC and an excess of NHS delays the reaction and provides a valuable capability to control the collagen gelation time. Various shapes and morphologies of collagen-based gels can be produced during this period. Accordingly, the current section focuses on the production of different collagen gels through the in-situ crosslinking approach, and preliminary cyto-compatibility and cell behavior of the resulting collagen gels were also investigated.

4.1.1 Formation and Characterization of In-situ Crosslinked Collagen Gels

4.1.1.1 Materials

Lyophilized collagen (semed S, acid-soluble), principally collagen type I with ca. 5% type III, from bovine dermis was kindly gifted by Kensey Nash Corporation. Ethanol,

EDC and NHS were purchased from Sigma-Aldrich Co. Ltd. and used without further purification. Potassium chloride, sodium chloride, monobasic potassium phosphate and dibasic sodium chloride heptahydrate were purchased from Fisher Scientific and used without further purification. Phosphate buffered saline (PBS) 20x buffer was prepared as

described previously. Any PBS buffer with lesser salt content than PBS 20x was prepared

by diluting PBS 20x accordingly to obtain the desired salt concentration.

4.1.1.2 Collagen Solubility Test in In-situ Crosslinking Solution

Collagen solubility testing was performed by mixing collagen with a combination of PBS

buffer with different salt concentrations (1x, 10x, 20x) and different amounts of ethanol.

The PBS 1x and 10x buffers were obtained from the dilution PBS 20x buffer by 20 fold

and 2 fold, respectively. 38mg of EDC and 46mg of NHS were dissolved in different

amounts of ethanol according to Table 4.1. The PBS buffer and alcohol were mixed

before the collagen was added. The total volume for each mixture composition was kept

at 1 mL. The Kensey Nash collagen presents itself in the form of flakes that are insoluble

in water. Therefore, the solubility of collagen in the in-situ crosslinking solution was

tested simply by naked eye with respect to the homogeneity and gelation of the prepared

mixtures.

4.1.1.3 Formation of In-situ Crosslinked Collagen Gels

Collagen gels were prepared by simply allowing in-situ crosslinking collagen solution to

complete crosslinking overnight without any disturbance at room temperature. Then the formed gels were placed in DI water for an additional three days to replace ethanol and buffer solutions with DI water for hydrogel formation. The in-situ crosslinking collagen

solution was prepared as described in the previous chapter. Briefly, 16 wt% of Type-I

collagen was dissolved in PBS buffer 20x and ethanol with the volume ratio of 1:1. Prior

to adding the buffer, 200 mM of EDC and 400 mM of NHS were added to the ethanol. A

homogenous collagen solution was formed after magnetically stirring the mixture for

approximately 5 minutes. Different shapes of collagen gels were prepared by using

different molds, such as a small vial, tube, and hemisphere ball. Photos of the resulting

gels were taken using a Canon PowerShot SD1400 camera.

4.1.1.4 Characterization of Cell Biocompatibility

Preliminary cell culture studies were carried out as a part of the joint project with the

Biomedical Engineering Department (BME) by Ms. Sonia Merritt (graduate student)

under the guidance of Dr. Horst von Recum (Associate Professor, BME).

Typically, the in-situ crosslinking collagen solution was prepared as described above and transferred into a 24-well cell culture plate with approximately 1mm thickness for each

sample. Then the collagen gels were crosslinked for two days and rinsed with sterile

water. After an initial rinsing, the gels remained swelled in the sterile water for an

additional 7 days to remove residual solvent and crosslinkers as much as possible.

Chinese Hamster Ovarian (CHO) cells were cultured in high-glucose DMEM media

supplemented with 1% pennicillin/strempomycin and 10% fetal bovine serum in a 10cm

diameter tissue culture dish. The media was changed every three days until 80%

confluence was reached. Then cells were transferred and seeded onto the collagen gels at

40,000 cells per cm2 (n=3). Cell morphology and behavior on the gels was observed by

an inverted optical microscope and the images were taken at pre-set time points.

4.1.2 Results and Discussion

4.1.2.1 Solubilization of Collagen in In-situ Crosslinking Solution

As discussed in Chapter I, collagen is insoluble in most common solvents due to complex

interactions between collagen molecules [8, 9]. In order to dissolve collagen, single

solvents or solvent mixtures need to interact with the collagen triple helix side groups to

dissociate its bundle structures. Recently, our laboratory reported a benign solvent system,

in which there is only buffer and alcohol, to easily prepare a collagen solution [72]. In the

benign solvent system, it is supposed that the alcohol component facilitates the disruption

of hydrophobic interactions, and the buffer assists in breaking hydrogen bonding and

ionic interactions in the collagen bundles. However, the collagen constructs generated

from the benign solvents are inevitably water-soluble and mechanically weak for tissue engineering applications and are in need of crosslinking. Accordingly, a novel in-situ

crosslinking collagen solution was developed in Chapter II by adding specific ratio of

crosslinkers EDC and NHS into the benign solvent system.

The influence of the crosslinkers on collagen solubility and the concentrations of buffer

and ethanol on collagen gelation were examined by mixing collagen with a mixture of

PBS buffer with different salt concentration (1x, 10x, 20x) and different amounts of

ethanol with the same crosslinkers. The preliminary results are shown in Table 4.1,

which indicate that the addition of crosslinkers decreased the solubility of collagen in

benign solvents. A collagen paste was formed instead of a uniform solution when 1x PBS

buffer was used in the mixture. This is expected, because the crosslinking reaction

between EDC and collagen started almost instantly upon mixing. The dissociation effect

of collagen bundles induced by the mixture of ethanol and 1x PBS buffer might not be

kinetically comparable with network formation brought about by the crosslinkers.

Therefore, collagen might be partially crosslinked before complete dissolution.

However, the solubility of collagen in the in-situ crosslinking solution increased with the salt concentration. When 10x PBS buffer was employed in the mixture, a white-colored collagen suspension was formed and subsequently a uniform gel was generated a few

hours later. Upon further increasing the salt concentration to 20x PBS buffer, collagen

was easily dissolved in the in-situ crosslinking solution and accordingly a semi-

transparent, colorless collagen gel was produced within a few hours. The difference in the

color of collagen gels prepared from 10x and 20x PBS buffers might suggest that the

different structures of collagen were present in each solution. Higher salt concentration

may result in more effective dissolution of Kensey Nash collagen, which led to smaller

fragments of collagen bundles and subsequently lower light scattering.

Furthermore, the ethanol component also played an important role in dissolving collagen

in the in-situ crosslinking solution. A similar optical change was also found with an

increasing concentration of ethanol in the mixture. For example, when 20x PBS buffer

was used, the appearance of the collagen gels shifted from white to semi-transparent with

increasing concentration of ethanol, which might indicate a higher dissociation of collagen in a high concentration of ethanol.

For gel formation, there is a limitation found based on the collagen concentration. A 10

wt% collagen solution was not able to form a gel after the solution preparation, while 20

wt% collagen solutions produced a dense gel. Therefore, a proper amount of collagen in

the in-situ crosslinking solution is needed to form a regular gel. A collagen concentration

of 16 wt% was selected in this study to produce the hydrogel for the remainder of

experiments.

4.1.2.2 Various Forms of In-situ Crosslinked Collagen Gels

The in-situ crosslinking method easily allows the formation of various shapes and morphologies of collagen gels. By simply injecting the in-situ crosslinking collagen solution into different molds, collagen plates, tubes and hemispherical hydrogels were successfully produced as shown in Figure 4.1. Each of the different physical forms of collagen gel has advantages for specific applications. For example, the collagen plate may be useful for cartilage implantation and the hydrogel tubes may be valuable as nerve guides. The hemispherical gel was an initial trail to eventually fabricate collagen hydrogel contact lenses for corneal wound repair. Furthermore, it is of importance to note that the in-situ crosslinked collagen gels would swell in water after removing the ethanol and salts.

Figure 4.1 Appearances of collagen hydrogel plate (a), tubes (b) and hemisphere (c)

prepared by the in-situ crosslinking method.

Another interesting application of in-situ crosslinking collagen gel is collagen printing.

Due to the fluid nature of the in-situ crosslinking solution, it can be processed as an “ink” and printed on particular substrates. A CWRU logo was printed on a glass slide using in-

situ crosslinking solution and became a water-stable collagen stamp within a few hours,

as shown in Figure 4.2.

Figure 4.2 Snapshot of a CWRU logo printed by using in-situ crosslinking collagen

solution.

4.1.2.3 Cell Culture Studies

An initial cell culture study was performed on the in-situ crosslinked collagen gels to gain

insight on any factors in the cross-linking and washing steps that would lead to cell detachment. The result showed an unexpected poor cell attachment on the collagen gels prepared from a mixture of 20xPBS buffer and ethanol with a volume ratio of 50:50. One plausible reason for this is the large amount of residual ethanol present in the gel which would be unfavorable for cell growth. Although the collagen gels were placed in the sterile water for an additional 7 days before cell culture, the exchange of ethanol by the

bulk water might require multiple soak-rinse cycles.

In order to remove the ethanol residual solvent from the collagen gel, two different

approaches were adopted. First, the newly-formed collagen gels were dried at room

temperature overnight to evaporate the ethanol component and then swelled in cell media during cell seeding. The resulting images of cells seeded on such dried collagen gels are shown in Figure 4.3, which indicates improved cell attachment. However, the cells did grow on the collagen surface near the center of the gel but not the edges.

The second approach to reduce the alcohol content in the collagen gel was to use less ethanol in the preparation of the collagen solution. According to collagen solubility and gelation tests, an in-situ crosslinked collagen gel could be made from a mixture of 20x

PBS buffer and ethanol with a volume ratio of 90:10. Such low ethanol content gels were then immersed in sterile water for another 7 days followed by the cell seeding experiment.

CHO cells showed a significant affinity to this collagen gel, and multiple cell layers growing in the gel were surprisingly observed in 48 hours after seeding, as shown in

Figure 4.4. Therefore, these preliminary results indicate that ethanol could induce an unfavorable cell attachment, and lowering the content of ethanol resulted in a higher cell biocompatibility of the in-situ crosslinked collagen gel.

Figure 4.4 CHO cells 48 hours after seeding. The image (a) is the cells on the collagen

hydrogel surface, and the image (b) contains CHO cells plated on a control tissue culture

polystyrene surface. Images are at 20x magnification.

4.1.3 Conclusions

Collagen gels can be formed from the in-situ crosslinking collagen solution introduced in

the previous chapter. Various physical forms of collagen gels, such as plates, tube,

hemispheres and collagen stamps, demonstrated the capability of the in-situ crosslinking method to produce customized collagen gel-based materials.

The addition of crosslinkers to the solution decreased the collagen solubility presumably due to crosslink formation, and collagen cannot dissolve in in-situ crosslinking solution made from low salt content buffer (i.e. PBS 1x). Increasing salt concentration can improve the solubility of collagen in solution and probably results in a more complete dissociation of collagen bundles. Moreover, the collagen concentration needs to be high enough (i.e., 16 wt%) to form a crosslinked gel.

Ethanol also plays an important role in collagen solubilization and gelation. However, it was found to cause an unfavorable cell detachment in the CHO cell seeding study.

Reducing ethanol content in the resulting collagen gel showed a significant improvement

of cell biocompatibility. An in-situ crosslinked collagen gel generated from the mixture of 20x PBS buffer and ethanol with volume ratio of 90:10 displayed a significantly favorable cell attachment with multiple cell layers formed inside the gel. This material could be very useful for medical implants and tissue regeneration.

4.2 In-situ Crosslinked Collagen Sponges

Collagen sponges are originally developed as wound dressings and hemostyptics [8, 9].

Major benefits of collagen sponges for these applications include their ability to easily

absorb large quantities of tissue exudate, offer smooth adherence to the wet wound area

with preservation of this moist micro climate, afford shielding against mechanical harm

and prevent secondary bacterial infections [130, 131]. Besides these physical effects, the

open and interconnected porous structure of collagen sponge can also facilitate cell

seeding, cell penetration, and distribution throughout the scaffold, and accelerate cell

proliferation and regeneration of new tissue [132, 133]. Therefore, collagen sponges have

been widely used in tissue engineering applications, such as skin replacement, bone

substitutes, and bioengineered tissues [134, 135].

Collagen sponges are generally prepared by freeze-drying aqueous acid- or alkali-swollen

collagen solutions or gels of approximately 0.1-5% dry matter content [9]. The porosity

of the lyophilized collagen sponges can be altered by varying the collagen concentration

and the freezing rate [131, 132]. In addition, collagen can be combined with other

materials like elastin [136], fibronectin [137] or glycoaminoglycans [138] and the

resulting scaffold can be crosslinked by for example, glutaraldehyde or by UV treatment

in order to achieve highly resilient materials [131, 139]. Accordingly, the production of crosslinked collagen sponge from in-situ crosslinked collagen gel is the goal of this section, and its wound dressing application is preliminarily investigated.

4.2.1 Formation and Characterization of In-situ Crosslinked Collagen Sponges

4.2.1.1 Freeze Drying of In-situ Crosslinked Collagen Gels

In-situ crosslinked collagen sponge was produced by freeze drying the crosslinked collagen gel in Virtis Advantage EL-85 lyophilizer with a condenser temperature of -

80 °C. Briefly, the collagen gels prepared in the last section were swelled in DI water for

3 days to reduce the ethanol and buffer salt residues and then frozen in refrigerator (-

6 °C), dry ice-ethanol bath (-72 °C) and liquid nitrogen (-196 °C), respectively. The frozen samples were then stored in a freezer (-20 °C) before lyophilization. The freeze- drying processing was carried out with an initial shelf temperature of -70 °C and then gradually heated up to 25°C with a rate of 0.5 °C/min at an ultimate 5 µbar vacuum.

4.2.1.2 Characterization of Porous Morphology

The pore structure of the resulting collagen sponges was characterized by means of scanning electron microscopy (SEM). Several cross-sections of each sample were sputter-coated with gold (Anatech Hummer 6.2), and visualized on SEM (JEOL JSM-

6510 LV) using secondary electron emission at 15 kV. The average pore size throughout the cross-section was assessed using NIH Image J software.

4.2.1.3 Fourier Transform Infrared (FTIR) Spectroscopy

The FTIR of the crosslinked collagen sponge was performed on ABB Bomem MB-104 spectrophotometer. A small piece was cut from each of sponge samples and fixed onto the FTIR holder. A spectrum with the absorbance from 600 to 2000 cm-1 was recorded using the Bomem Grams software.

4.2.1.4 Wound Healing Experiments

Wound healing efficacy of in-situ crosslinked collagen sponges was performed as a part of the joint project with the Dermatology Department by Dr. Minh Lam (Assistant

Professor, Dermatology) and Dr. Elma Baron (Associate Professor, Dermatology).

The surgical procedure was similar to the one described in the previous chapter. Briefly,

BALB/c mice received a hair-removal treatment using Nair two days before applying incisions. After anesthetization, full-thickness wounds (i.e. tissue destruction extending through dermis) with a 6mm diameter were cut from the backs of the mice using a surgical scissor. Then collagen sponges were cut and applied to fill in the wound area with approximately 2mm thickness. The treated mice were then monitored for 14 days, specifically comparing the rate of wound closure (patched vs. control) and photos were taken at pre-set time points using a SONY DSC-W310 camera.

4.2.2 Results and Discussion

In this study, two types of collagen-based matrices, namely cylindrical and tubular collagen crosslinked sponges, were produced by freeze-drying the respective collagen gels to demonstrate the ease of customizing the sponge to a desired shape. The representative images are given in Figure 4.5. Prior to the freeze-drying process, ethanol

97 and salt residues present in the gels need to be replaced by water because the freezing point of the buffer and ethanol mixture is much lower than the operating temperature and became unpredictable after mixing with collagen.

Figure 4.5 Snapshots of cylindrical and tubular in-situ crosslinked collagen sponges

prepared by freeze drying of collagen gels.

Since the porosity of collagen sponges obtained through the freeze-drying process is the negative of the ice crystal structure formed during the freezing phase, the control of the pore size is easily performed by varying selected parameters affecting ice formation during freezing, such as the freezing temperature. Generally, with regard to the effect of different freezing temperatures on the ice crystal size, lower temperatures of freezing are expected to yield faster rates of ice crystal nucleation and lower rates of diffusion, and thus significantly smaller pores [140]. In this study, the in-situ crosslinked collagen gels were frozen under three different conditions, namely in a refrigerator, a dry ice-ethanol bath, and liquid nitrogen. It was found that complete freezing of collagen gels in refrigerator occurred in hours, while it took only a few minutes for freezing in dry-ice ethanol and liquid nitrogen.

Figure 4.6 SEM images of in-situ crosslinked collagen sponges prepared in refrigerator

(a), dry-ice ethanol bath (b) and liquid nitrogen (c, d).

The SEM images of cross-sections of each sample are shown in Figure 4.6. As expected,

the sample frozen in refrigerator displayed the largest pores with diameters of

approximately 150 ± 50 µm (Figure 4.6a). The pore size decreased to about 80± 30 µm

when the collagen gel was frozen in a dry ice-ethanol bath (Figure 4.6b), and the

smallest pores of approximately 35 ± 10 µm were observed in the samples frozen with

liquid nitrogen (Figure 4.6c). Moreover, it is of interest to note that axially-oriented pores were also discovered in the liquid nitrogen-frozen samples, as shown in Figure

4.6d, which might arise from an axial gradient of temperature presented in the collagen gels [132].

In order to investigate the influence of freeze-drying process on the collagen structure,

FTIR spectra of the collagen sponge were generated on Bomem MB-104 spectrophotometer and one is shown in Figure 4.7. As mentioned in a previous chapter, if collagen contains a triple helix structure, the amide group is used in hydrogen bonding in each triplet (Gly-Pro-Hyp) and is characterized by a peak around 1655 cm-1 instead of

the typical 1620 cm-1 band for gelatin [19]. The Kensey Nash collagen used in this study

has a peak at 1655 cm-1, and the amide I peak obtained from in-situ crosslinked collagen

sponge is located at 1647 cm-1. The shifting toward a wavelength more characteristic of

gelatin preliminarily suggests a mixture of native-like collagen (triple helical structure) and denatured collagen (random coil-like gelatin) presented in the resultant freeze-dried sponges [141].

Amide I 1647

0.5 Amide II 1540

0.4 Absorption

0.3

2000 1800 1600 1400 1200 1000 800 600 Wavenumber (cm-1)

Figure 4.7 FTIR spectrum of in-situ crosslinked collagen sponges.

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Wound dressing application of in-situ crosslinked collagen sponge was tested on BALB/c

mice. Full thickness wounds with a diameter of 6mm were cut from the back of mice and

patched with collagen sponges. The representative images of animals at day 0, 5 and 10 after incision are shown in Figure 4.8, which indicate collagen sponge was able to accelerate the healing rate. A significant difference in the wound size between the control and the treated one was observed at day 5 with a much smaller size of treated wound compared with the control. At day 10, the patched wound was pinkish and scarless, which indicated rather complete wound healing, while the control wound still exhibited a large scar. This promising result suggests that the in-situ crosslinked collagen sponge prepared in this study could be a potentially useful material for wound healing and skin regeneration applications.

Figure 4.8 Appearance of wound healings at day 0, 5 and 10 after incision.

4.2.3 Conclusions

Collagen sponge was prepared from the in-situ crosslinked collagen gels by means of freeze-drying. The porosity was controlled through different freezing processes. Lower freezing temperatures provided smaller pore sizes. The smallest pores with a diameter of

35 ± 10 µm were observed in collagen sponges frozen with liquid nitrogen. According to

101 the FTIR spectrum, freeze-drying process could partially denature collagen. Preliminary in vivo wound healing tests indicate that the in-situ crosslinked collagen sponges facilitate wound closure and may be a good candidate for wound healing and skin regeneration.

4.3 Biomineralization of In-situ Crosslinked Collagen Gels

Calcium phosphate biomineralization is one of the most important processes in the mammalian kingdom, contributing to the structural support of tissues such as bone, teeth and enamel [142, 143]. Collagen represents a naturally-occurring group of proteins found mostly in mammalian connective tissue and bone [8, 9]. Thus the integration of calcium phosphates and collagen gives rise to one of the most important biological composites.

Electrophoresis has been successfully demonstrated in many systems to induce fast development of calcium carbonate or calcium phosphate within the interior of a hydrogel

[144]. The advantage presented by electrophoresis is considerably speeding up mineralization times by using an electrical current to drive the positive and negative ions into the hydrogel interior to form minerals first within a material [145]. This approach also solves the problem of surface mineralization dominating many direct mineralization routes while also giving large flexibility to the physical dimensions of the mineralized hydrogel [146].

Accordingly, the current section focuses on using electrophoresis to produce biomineralized collagen hydrogels which are seeded with calcium phosphate and highly porous composite collagen sponges that could be used for direct bone substitutes. This is

102 a joint project with Jack Johnson (graduate student) under the guidance of Dr. David

Schiraldi (Professor) in the Department of Macromolecular Science and Engineering.

4.3.1 Electrophoresis Experiments

4.3.1.1 Collagen Gel Sample Preparation

Collagen hydrogels were prepared as described previously. Briefly, 16wt% of Type-I collagen was dissolved in 20x PBS and ethanol with a volume ratio of 50:50. Prior to adding the buffer, the crosslinkers EDC and NHS were added to the ethanol to equal 200 mM of EDC and 400 mM of NHS in the solution. Samples were prepared directly in 1 mL B-D plastic syringes at a target volume of 0.2-0.3 mL per syringe, and then stored at room temperature for 3 days in to allow full crosslinking to occur. The syringes were then cut at both ends and placed in DI water for an additional 3 days to replace ethanol and buffer solutions with DI water for hydrogel formation.

4.3.1.2 Electrophoretic Mineralization

For electrophoretic mineralization, a FisherBiotech® Horizontal Electrophoresis System with a 7 X 10 cm gel size was used in conjunction with a 120 V FisherBiotech® Power

Supply. A mold of the well was made using the TAP Urethane Mold Making System and cut with a band saw to accommodate the 1 mL syringes without leakage from one side to the other. 0.1 M CaCl2 solution was made in DI water and placed in the anode reservoir, along with a 0.1 M Na2HPO4 solution that was placed in the cathode reservoir. The voltage was set to 100 V and the amperage was set at 50 mA, with the electrical current running for various times, up to 4 hr. All resulting samples were then placed in DI water to wash any free ions out, cut to expose the inside of the samples, and then frozen in

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liquid nitrogen and transferred to a VirTis AdVantage® EL-85 freeze-dryer and subjected to an ultimate vacuum of <10µBar to sublime the ice for 5 days.

4.3.1.3 Characterization

Digital photographs were obtained using a Nikon D80 DSLR camera with a Nikon

Micro-Nikkor 55mm lens. Scanning Electron Microscopy (SEM) and Energy Dispersive

X-Ray (EDX) elemental mapping was performed on Pd-coated samples using a FEI

Quanta 200 3D ESEM/FIB system.

4.3.2 Results and Discussion

The electrophoretic mineralization process is relatively simple, using an electrical

potential to force ions of opposite charge into the interior of the gel [36-38]. While initial

electrophoretic mineralization attempts used very low times for mineralization, longer

times were desired in this approach in an attempt to more homogeneously mineralize the

gels. The collagen hydrogels started as very transparent materials and a change over time

in transparency was observed. After one hour of electrophoretic mineral deposition, the

collagen hydrogel exhibited marked whitening in the interior, as shown in Figure 4.9.

More general opaqueness was also observed throughout the sample suggesting that a

great amount of mineral material had built up inside the gel.

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Figure 4.9 Collagen hydrogels before (a) and after (b) electrophoretic mineral deposition

for one hour.

To determine the extent of mineralization in the hydrogels, the samples were dissected and freeze-dried for SEM analysis. Energy dispersive x-ray (EDX) spectroscopy was also employed to examine the chemical nature of any particulate inside the dried gels and to construct an elemental map of the material to determine if particulates are calcium phosphate.

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SEM analysis of the freeze dried collagen hydrogel, namely collagen sponge, is given in

Figure 4.10, which shows a typical open network of collagen as discussed in the previous

section. A high-magnification image indicates no particulate detected in the dried collagen hydrogel. Elemental mapping shows regions of carbon and oxygen, but little calcium or phosphorous were detected that would be indicative of insufficient mineral dissolution in the collagen used.

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Figure 4.11 SEM images of collagen hydrogel after 1 hour of electrophoretic mineral

deposition (A and B). EDX (C and D) confirms that there is much phosphate and

calcium in the region, and the calcium + phosphate overlay shows very good

correspondence with the particulate, indicating calcium phosphate buildup.

SEM and EDX analysis of mineralized hydrogels showed completely different results.

Figure 4.11 presents an analysis of a sample mineralized for one hour. Figure 4.11A

shows the SEM analysis of the interior of the dried hydrogel, with many small particles

being revealed. Figure 4.11B shows the exposed edges of collagen with integrated

calcium phosphate in the collagen, indicating that the mineralization is not limited to the

surface of the hydrogel, but occurs within the collagen gel during electrophoresis. This

important discovery shows that the biomineralization is occurring much like natural bone

growth. Figures 4.11 C and D confirm that the particles grown in collagen are indeed

calcium phosphate, with an overlay of Ca and P elemental maps on the SEM image

showing perfect correspondence with the particles on the surface. Therefore, these

preliminary results suggest that the biomineralized collagen gels and composite collagen

sponges were successfully prepared through the electrophoretic processing of in-situ

crosslinked collagen gels.

4.3.3 Conclusions

This initial study indicates that in-situ crosslinked collagen gels and sponges could be promising candidates for bone substitute applications. Combining the in-situ crosslinking

method with an electrophoretic approach to mineralization can provide many advantages

to existing systems and could be a viable approach to bone replacement with further

107 refinement. Collagen was in-situ crosslinked to form a hydrogel, which was subjected to electrophoretic mineralization that substantially seeded the hydrogel with calcium phosphate. Freezing drying these materials provided a very porous network of mineralized collagen sponges, similar to what can be found in natural bone, and this discovery may be a step towards bone replacement materials.

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Chapter V Phase Transition of Poly (Acrylic Acid) Nanofibers

5.1 Introduction

It has been suggested that a common underlying mechanism of cellular functions is a polymer-gel phase transition, which means a major structural change is prompted by a subtle environmental change [147, 148]. These phase transitions are thought to be involved in many cell behaviors, such as cellular solute exchange and transport, actin filament shortening, helix-coil transitions, and several other force-dependent mechanisms

[149-151]. Among them, an analogous phase transition in living nerve fibers has been extensively investigated in the last two decades [152-156]. The existence of cation- exchange processes involving Ca2+ and abrupt structural changes associated with these cation-exchange processes in the cortical layer of nerve fibers have been detailed in prior literature [153, 154]. Tasaki and the others have shown that the electrophysiological processes known as nerve excitation and conduction are basically manifestations of abrupt phase-transitions of the cytoskeleton in the cortical gel layer of the axon and can be reproduced by using synthetic polyanionic hydrogels in place of living nervous tissues

[155, 156].

Negatively-charged synthetic polyelectrolyte hydrogels are well known to exhibit a reversible transformation from their swollen state to the compact state when the monovalent counter ions (e.g. Na+, K+, etc.), in the gels are exchanged with divalent cations (e.g. Ca2+, Mg2+, etc.) [157-159]. This structural transformation can be initiated and completed by a small change in the salt composition of the surrounding solution

[157]. Since the superficial gel layer of a nerve fiber undoubtedly carries fixed negative charges under physiological conditions [152], polyelectrolyte hydrogels can be used as

109

synthetic model systems for the study of the process of divalent-monovalent cation

exchange in the cortical layer of nerve fibers [153, 154].

In this study, polyelectrolyte poly (acrylic acid) (PAA) nanofibers were generated by means of electrospinning, and the influence of the preparation parameters during the electrospinning process such as PAA concentration and feeding rate on the formation of

PAA nanofibers was systematically investigated. The PAA nanofibers contained ethylene glycol (EG) and were rendered water-stable by cross-linking via thermally-induced esterification. Since an axon in nerve fibers possesses an elongated conduit morphology, non-woven PAA fibrous tubes were fabricated and their reversible swelling behavior was studied in CaCl2 and MgCl2 solutions, respectively. This work can serve as a model study

to explore the development of an abiotic, polymeric mimic of an axon.

Electrospinning can generate fibers with diameters of several tens to hundreds of

nanometers from many polymers and is a technique with renewed interest and

development in the last decade [160-162]. The electrospun scaffolds containing ultra-fine fibers have many exciting properties absent in traditional fibers or films, due principally to the much higher surface area and wider range of inter-fiber pore structures [163-165].

Although the stimuli-responsive behavior of PAA hydrogels has been reported

extensively [166-168], to the best of our knowledge our group is the first to create water-

stable PAA nanofibrous tubes that exhibit reversible phase transitions induced by

divalent-monovalent cation exchange in aqueous solution.

Previous studies on the swelling of polymer gels commonly report the mass uptake of

liquids due to the ease of measurement [161, 168]. For applications like artificial axons or

110

muscles, dimensional transformation during swelling is of primary concern. Accordingly,

actual length changes of PAA tubes during phase transitions were examined, in addition

to mass uptake, to provide further insight into the swelling process.

5.2 Formation and Characterization of PAA Nanofiber Tubes

5.2.1 Materials

Poly (acrylic acid) (average Mv = 450,000), ethylene glycol (anhydrous), calcium chloride (anhydrous) and sodium citrate were purchased from Sigma-Aldrich and used

without further purification. Ethylenediaminetetraacetic acid (EDTA) disodium salt

dehydrate, sodium hydroxide, sodium chloride, magnesium chloride hexahydrate, sulfuric

acid, hydrochloric acid and water (HPLC grade) were purchased from Fischer Scientific

and used as received without further purification.

5.2.2 Preparation of PAA Ethanol Solution

Poly (acrylic acid) solutions were prepared by dissolving the appropriate amount of

polymer in ethanol with various concentrations varied from 2 to 6 wt%. Ethylene glycol

was added to each sample as a crosslink agent at the concentration of 16 wt% relative to

the polymer. Complete dissolution was observed after 24 hours of mixing with a

magnetic stir bar at ambient temperature. 1M sulfuric acid was added to the PAA-EG

solution right before electrospinning processing at a concentration of 50µL/mL.

5.2.3 Fabrication of PAA Nanofibers

Poly(acrylic acid) nanofibers were produced through single jet electrospinning technique

as introduced in Chapter I. Typically, the experimental set-up consisted of a high voltage

power supply (0-30kV Spellman CZE1000R; Spellman High Voltage Electronics

111

Corporation), syringe pump (KD Scientific), and a variable speed, rotating, stainless steel grounded counter-electrode mandrel (4.0 mm OD x 15.0 cm L). Briefly, PAA solution was loaded into a syringe affixed with a blunt-tip 18-guage needle placed 20 cm away from the leading face of the rotating collection target. The solutions were electrified by applying a positive potential (15kV) to the syringe needle by means of an alligator clamp.

The volumetric flow rate of the solution was varied from 0.6 to 1.0 ml/hr. Under such spinning conditions (voltage, distance, and volumetric flow rate), PAA nanofibers impregnated with residual ethylene glycol were collected on the mandrel.

5.2.4 Thermo-crosslinking of PAA Nanofibers

Freshly made PAA nanofibers sitting on the mandrel were crosslinked upon heat treatment in a vacuum oven at 130 and 25 in Hg of vacuum for 30 minutes, and then cooled down to room temperature.℃ Crosslinked PAA fibrous tubes were obtained by carefully removing polymer fibers from the mandrel without breaking them after thermo- crosslink treatment.

5.2.5 Neutralization of PAA Nanofiber Tubes

PAA fiber tubes were neutralized by placing in a base solution containing 1M NaOH and

1M NaCl for approximately one hour and then rinsing with water to remove residual salts.

The mass of PAA dry tubes and the hydrated tubes after neutralization were recorded on a balance (Mettler Toledo AL104).

5.2.6 Characterization of Fiber Morphology

The morphologies of electrospun PAA nanofibers were visualized by means of a scanning electron microscope (SEM). PAA nanofiber tubes were sputter-coated with a

112

10nm gold layer and observed using a JOEL JSM-6510LV SEM with an operating

voltage of 15 kV. Data treatment was done with free Image J software.

Morphological characterization was also performed on the hydrated neutralized PAA tubes. For this purpose, small pieces of PAA samples were carefully cut in water and mounted on the standard SEM holder with the help of conductive carbon tapes. Then the samples were directly examined in an environmental SEM (FEI Quanta 200 3D

ESEM/FIB system) without sputter-coating. The operating voltage was 20 kV with a chamber pressure of 7.67 torr.

5.2.7 Titrations of PAA Tubes

Calcium chloride and magnesium chloride were selected to prepare an aqueous solution of Ca2+ and Mg2+, respectively. The phase transitions of PAA fibrous tubes were

investigated by changing the cation ion concentration in solution through a titration

approach.

Typically, in batch experiments, each neutralized PAA tube was placed in an Erlenmeyer

flask with 50 mL DI water at room temperature. 1M CaCl2 or 1M MgCl2 solutions were

carefully added through a burette at the rate of 0.5 mL each time. The length of PAA tube

in solution was carefully measured after the addition of salt solution. The resulting

compacted PAA tubes were withdrawn from the titration solution and rinsed with DI

water.

In order to examine the reversibility of PAA phase transitions, cation ion chelators (e.g.

EDTA and citrate) were employed to remove Ca2+ or Mg2+ from the compacted PAA

tubes. Briefly, EDTA and citrate solutions with a concentration of 0.2M were prepared

113

by dissolving EDTA disodium dehydrate or sodium citrate in water, respectively.

Compacted PAA tubes were immersed in 50mL water and then titrated with EDTA or

citrate solution. The changes of tube length were carefully measured each time until they

were constant in solution. Then the resultant PAA tubes were withdrawn and stored in DI

water for the next round of titration.

5.2.8 Elemental Analysis of PAA Tubes

The content of sodium element in the neutralized PAA tubes and the calcium / sodium

elements in the resulting PAA compacted tubes were quantitatively analyzed by

inductively coupled plasma optical emission spectroscopy (ICP-OES) at Galbraith

Laboratories, Inc. The analytical method was GLI procedure ME-70 accredited by the

American Association for Laboratory Accreditation (A2LA), certificate number 2777.01.

Before each test, the PAA samples were dried according to GLI procedure 3.2.3.6.

5.3 Results and Discussion

5.3.1 PAA Nanofiber Formation

Electrospinning of PAA/EG/ethanol solutions at a series of concentrations produced a stable polymer jet under proper spinning conditions, but yielded different morphologies of nanofibers. Figure 5.1 (a, b, c) show the SEM images of electrospun PAA nanofibers at the concentration of 2 wt%, 4wt%, and 6wt% with a flow rate of 0.8ml/h when an operating voltage of 15 kV was applied. It can be seen that at a concentration of 2wt%, some spindle-like structures formed along the fibers. With an increase of the solution concentration, the spindle-like structure disappeared. When the concentration was increased up to 4wt%, a fine fiber structure with an average diameter of 890 ± 90 nm was

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clearly observed, as shown in Figure 5.1b. Further increasing concentration led to the

formation of ribbon-like structures as well as thicker fibers, and distinct adhesion

between the fibers appears to be dominant, resulting in a rough mat. It appears that the

polymer concentration is one of the most critical parameters to control the quality of the

nanofibers. The concentration is directly related to the viscosity, surface tension, and

conductivity of the polymer solution [160]. Lower concentrations result in lower

viscosity and higher surface tension, which favors the formation of bead structures,

whereas increasing the concentration could result in increased solution viscosity, which is

essential for the formation of fiber structures [163-165].

The flow rate of the polymer solution is another factor influencing the morphology of

PAA nanofibers. When the PAA concentration was kept constant at 4wt%, some ribbon- like fibers were formed with increasing flow rate from 0.8 to 1.0 mL/h, as shown in

Figure 5.1d. This is because with very high flow rates, fibers may not dry completely before reaching the collector, which leads to the formation of ribbon-like or flattened structures. In addition, it is worthwhile to note that PAA nanofibers can be formed with a flow rate under 0.8 ml/h, but in these cases the distribution of fiber diameters became broader. This may be because the Taylor cone at the tip of the capillary cannot be maintained when the flow of solution through the capillary is insufficient to replace the solution ejected as the fiber jet.

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Figure 5.1 SEM images of PAA nanofibers electrospun from different concentrations at a flow rate of 0.8 mL/h: (a) 2 wt%, (b) 4 wt%, (c) 6 wt%, and (d) 4wt% at a flow rate of

1.0 mL/h.

5.3.2 Crosslinking of PAA Nanofibers

Since both PAA and EG are water-soluble, the electrospun PAA nanofibers were dissolved in water almost immediately. In order to retain their unique fiber structure, electrospun PAA nanofibers were treated at 130 within a vacuum for 30 minutes to form intermolecular crosslinks between the carboxyl℃ groups of the PAA and the hydroxyl groups of the EG, as shown in Figure 5.2. A little sulfuric acid was added as a catalyst in the thermal esterification.

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Figure 5.2 Thermal esterification of polyacrylic acid with ethylene glycol.

After thermal crosslinking, a water-stable PAA fiber tube was formed and demonstrated in Figure 5.3, which shows the common feature of a random array and a very porous structure presented in the aqueous solution. It is of interest to note that no changes of tube morphology (i.e. length and width) were detected after immersing in water for one week, indicating sufficient crosslinking occurred inside the PAA fibers.

Figure 5.3 SEM images and snap shots (inserted) of crosslinked PAA nanofiber tubes

before (a) and after (b) immersing in water.

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5.3.3 Neutralization of PAA Nanofibers

In order to investigate the efficiency of such PAA and EG thermal esterification, the crosslinked PAA fibers were neutralized and converted to sodium salts in a mixture of

1M NaOH and 1M NaCl solution. The samples were rinsed with DI water extensively to

remove the residual salts from the PAA-Na fibers and then were analyzed by ICP-OES at

Galbraith Laboratories, Inc. The results indicated the content of sodium in the dry PAA-

Na sample was 19.8% of the total weight. Based on an assumption that all the

uncrosslinked carboxylic acid groups in PAA were transferred into sodium salts, and the

neutralized PAA-Na polymer chains were composed only of two kinds of segments, as

shown in Figure 5.4 (in which segment I stood for the uncrosslinked carboxyl groups and

segment II represented the crosslinked ones), the content of uncrosslinked PAA was

roughly calculated to be 80% of total carboxyl groups present. In other words, the

crosslink content of PAA was equal to or less than 20% after thermal esterification and

approximately 80% of carboxylate groups in PAA fiber tubes were available for the

following phase transitions induced by divalent-monovalent exchange in solution.

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Figure 5.4 Proposed structure of PAA-Na polymer network.

For phase transition tests, it is important to know the swelling behavior of the PAA

nanofiber tubes in aqueous solution. The swelling ratio of the crosslinked tubes was

calculated according to following equation:

Swelling ratio = (Ws – Wo)/Wo

where Wo is the dry weight of the PAA samples, and Ws is the weight of the fiber tubes

immersed in solution for one hour at room temperature. Based on a statistical analysis,

the swelling ratio of the crosslinked PAA tubes in water was discovered to be 4.39 ± 0.14.

Lightly-crosslinked PAA is a principal commercial super-absorbent material and a

prototypical pH-responsive polyelectrolyte [160, 161], the swelling behavior of PAA tube

can be dramatically varied with the different values of pH. Figure 5.5 shows a significant

difference in the tube thickness of crosslinked PAA samples soaked in a base (NaOH)

and an acid (HCl) solution, respectively. PAA tubes were neutralized in base solution and

the swelling ratio increased up to 28.13 ± 1.23, while the swelling ratio of tubes in acid

solution was similar to the ones in water. This result is expected because the acid dissociation constant (pKa) of PAA is about 4.28 [169]. Carboxyl groups (COOH) of

PAA were largely dissociated to form carboxylate groups (COO-) in NaOH solution and the hydration of COO- is larger than that of COOH. Therefore, a swollen and porous fiber network was expected in the hydrated PAA-Na tubes, as shown in Figure 5.6. It is also interesting to note that the changes of PAA tube thickness were switchable in base and acid solution, and the change of tube length was negligible.

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Figure 5.5 pH responsive of crosslinked PAA fiber tubes

Figure 5.6 Typical ESEM image of neutralized PAA fibers in water.

5.3.4 Calcium Ion-induced Phase Transition

In the calcium titration, the neutralized PAA fiber tubes became compacted when the concentration of divalent cations Ca2+ increased in solution. The length of neutralized tubes was carefully measured each time when CaCl2 solution was titrated in. Figure 5.7a

120 shows the changes of tube length according to the Ca2+ concentration in solution. It can be seen that the Ca-salt added to the solution had little influence on the tube length initially. However, at the point where the Ca2+ concentration rose up to about 20mM, there was a prompt decrease in the tube length, and this shrinkage was completed in the range of Ca2+ concentration just beyond about 55mM.

Figure 5.7. (a) Length changes of neutralized PAA nanofibrous tubes with increasing the concentration of Ca2+ in solution. (b) Snap shots of PAA tubes before and after the titration of CaCl2 solution.

It is well known that the COO- groups in macromolecules overwhelmingly prefer Ca2+ to

Na+ [157, 158]. The shrinkage of tubes can be ascribed to the formation of a complex between Ca2+ and carboxyl groups of PAA tubes which cross-bridge the COO- group of one chain with another COO- group in a neighboring chain, as shown in Figure 5.8. A

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typical SEM image of compacted PAA-Ca fibrous tubes is shown in Figure 5.11a, which

clearly exhibits a strong fiber network due to the complex formation.

Ca O Ca C O

CH2CH CH2CH CH2CH C O C O a O C O CH2 Ca CH Ca 2 Ca O O O C O C O C O

CH2CH CH2CH CH2CH

Figure 5.8 Representative schematic of calcium induced cross-bridge between

carboxylate groups in PAA tubes.

Furthermore, it is of interest to mention that the shrinkage of neutralized PAA fibrous

tubes was not observed until the Ca2+ concentration increased up to some critical point, e.g. 20mM. Figure 5.7a indicates that the tube length stayed constant with addition of

CaCl2 solution initially. This may be because the carboxyl groups of PAA tubes were

widely separated from one another at very low concentration of Ca2+ in solution. The

Ca2+ cross-bridge formed in this situation was readily broken by the incessant thermal

motion of the polymer chains. By increasing the Ca2+ concentration in solution, the

cations surrounding the polymer fibers were predominantly Ca2+, and the formation of

one Ca2+ cross-bridge would enhance the probability of neighboring COO- groups of polymer chains to form a second cross-bridge. That is, the free energy required to form one cross-bridge might be smaller when the neighboring sites were already cross-linked

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than when they were not. The process of Ca2+ cross-bridge formation would then proceed

in a cooperative fashion until the entire polymer fiber was involved in the process.

Finally, at the equilibrium stage, a large number of water molecules around the

carboxylate group and calcium ion within the fibers were displaced and compacted PAA

fibrous tubes were formed.

The shrinkage of neutralized PAA fibrous tubes was also calculated according to the

following equation:

Shrinkage = Wca / Wn

where Wn is the weight of the neutralized fibrous tube, and Wca is the weight of the tube

treated with CaCl2 solution at room temperature. The shrinkage of the neutralized PAA

fibrous tubes was determined to be 0.16 ± 0.02, which indicates that a large amount of

water was expelled due to the formation of cross-bridges between Ca2+ and COO- groups

of PAA. After calcium titration, the neutralized PAA tube decreased to 65% of the

original length, as shown in Figure 5.7.

In order to examine the effciency of such calcium ion coupling reaction, the compacted

PAA-Ca tubes were withdrawn from the titration solutions and rinsed thoroughly with water to remove the free cations and then were quantitatively analyzed by ICP-OES at

Galbraith Laboratories, Inc. The results showed that amounts of calcium and sodium present in the PAA-Ca dry samples were 7.86% and 0.179% of the total weight, respectively. Compared to the content of sodium in the neutralized PAA, which was 19.8% by weight, the results of calcium analysis suggested sodium ions were almost completely removed from the PAA fibers after exchanging with calcium ions. Based on the results of

123 calcium and sodium elemental analysis, another assumption was made, which was the compacted PAA-Ca polymers consist of three parts, as shown in Figure 5.9. Part I represents the crosslinked PAA, Part II is the free carboxylate groups without bonding calcium ions, and Part III represents the PAA crosslinked with calcium. According to the previous results, which indicated that the crosslinked PAA (Part I) is approximately 20% of total carboxylate groups and calcium element occupies 7.86% of the total weight in the compacted PAA-Ca polymers, the contents of Part II and Part III were calculated to be about 48% and 32% of total carboxyl groups present in PAA polymer, respectively.

Expressly, after the thermal esterification of PAA and EG, approximately 80% of carboxyl groups were left and available for the calcium ion-induced phase transition.

Among them, about 32% of total carboxyl groups were bonded with calcium ions, and approximately 48% of them were still free after complete calcium titration. These preliminary results indicated an equilibrium existing in the calcium ion coupling reactions, and apparently such reversible equilibrium favored the dissociation side.

Figure 5.9 Proposed structure of the compacted PAA-Ca polymer.

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It was hypothesized that the shrinking behavior of PAA tubes is reversible and the tube

length could be fully restored after removing calcium ions from the PAA fibers. In order

to test this assumption, a widely-used metal ion chelator, EDTA, was employed to remove Ca2+ from the PAA fibers and break the cross-bridges between two carboxylate groups. Accordingly, the swelling behavior of compacted PAA-Ca tubes was expected with increasing concentration of EDTA in solution, and a representative curve is displayed in Figure 5.10.

Figure 5.10 (a) Length changes of compacted PAA-Ca tubes with increasing concentration of EDTA in solution. (b) Snap shots of PAA tubes before and after the titration of EDTA.

When the concentration of EDTA in solution was increased from zero, an interesting plateau was clearly observed at the early stage of swelling behavior, which may indicate a lower limit of EDTA concentration existing in the calcium chelation. It is well known

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that the complex of Ca2+ and COO- groups is relatively strong [158]. A small amount of

EDTA present in solution might not be sufficient to compete with the polyanionic groups

of PAA in combining with Ca2+. When the concentration of EDTA increased up to some critical value (12mM), the first complex of EDTA and Ca2+ was formed. This may help to break up the Ca2+ cross-bridge from the neighboring sites, and more EDTA-Ca2+

complexes were formed and the swelling behavior of PAA fibrous tubes began.

Therefore, the chelation between EDTA and Ca2+ in PAA fibers might also occur in a

cooperative fashion until the entire polymer is involved as mentioned above. Eventually,

the PAA fibrous tubes swelled back to about 92% of the original length, as shown in

Figure 5.10a. The mass of PAA tubes after EDTA chelation was restored to only about

40% of the weight when initially swollen before Ca2+ titration, even though the length of

the tubes was almost back to original values. It is also worthwhile to note that the length

and weight of PAA nanofibrous tubes could be fully restored when the tubes were placed

in water overnight after the EDTA treatment, which indicated that the reconstruction of

the swollen fiber network took time and water-uptake occurred in a slow fashion. A typical SEM image of PAA tubes restored after EDTA titration is given in Figure 5.11b,

which shows a significant fibrous structure with larger pores compared to the compacted

porous surface of PAA-Ca fibers (Figure 5.11a).

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Figure 5.11 SEM images of PAA fiber network after calcium crosslink (a) and EDTA

chelation (b).

An alternative chelator, citrate, was also used to remove calcium ions from the PAA fibers and restore the compacted PAA-Ca tubes. Unlike titration with EDTA, the PAA-

Ca tubes exhibited a quick response to the addition of citrate in solution, as shown in

Figure 5.12. The tubes started to expand almost instantly when the citrate was titrated in and the tube length was fully restored at a citrate concentration of about 10mM. The different swelling behaviors of PAA-Ca tubes in EDTA and citrate solutions may arise from different calcium chelate structures. As is known, EDTA chelation is typically hexadentate (“six-toothed”) form, and the four ligands are required in conjugating metal ions [170]. Since EDTA disodium salt was used in the study to prepare the standard solution and the pKa value (1.99, 2.67, 6.16 and 10.26) of the fourth proton in EDTA is fairly large, it is possible that the dissociation of EDTA in water was not sufficient and the four ligands were not available at the same time for chelation. In contrast, citrate solution was prepared from citrate tri-sodium salt. The three ligands could be simultaneously utilized to conjugate calcium ions in solution, which resulted in a quick expansion of PAA tubes.

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Citrate 100

95 EDTA 90

80 Length (%) 75

0.000 0.005 0.010 0.015 0.020 0.025 0.030 0.035 0.040 Concentration (M)

Figure 5.12 Swelling behaviors of the compacted PAA-Ca tubes in citrate and EDTA

solutions, respectively

5.3.5 Magnesium Ion-induced Phase Transition

In order to investigate the response of neutralized PAA tubes to the other divalent cation

ions such as Mg2+, magnesium chloride was used to prepare the titration solution. Since

magnesium is one of the most abundant elements in the human body and its ions are

essential to all living cells, the study of a magnesium ion-induced phase transition is of special interest.

The representative shrinking behavior of the neutralized PAA tubes in the presence of the various concentrations of Mg2+ in solution is plotted in Figure 5.13, which shows a

completely different result comparing with the calcium ion-induced curve. The onset of shrinkage occurred as long as Mg2+ was added, and eventually the tube shrank to 77% of

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the original length. Such different responses of PAA tubes to the calcium and magnesium

divalent ions may be ascribed to the different hydration states of the PAA-cation

complexes. Ikegami et al. has reported that calcium ion is bound to carboxylate groups

mainly in the form of COO-Ca-OOC which is relatively strong and makes the polymer

more hydrophobic, while magnesium ion takes the form of (COO-Mg2+ + COO-) and the

resulting complex ion is still hydrophilic [171]. Since the latter form is more hydrophilic,

the formation of Mg2+ and PAA complex may be relatively easier but much weaker than the one formed between Ca2+ and PAA. Therefore, the shrinking response of PAA tubes

to magnesium titration was relatively quick with a smaller change of tube length (i.e.

23%), as shown in Figure 5.13.

100

90 Calcium 85

Length (%) 75 Magnesium

60 0.00 0.01 0.02 0.03 0.04 0.05 0.06 0.07 0.08 Concentration (M)

Figure 5.13 Length changes of neutralized PAA tubes with the increasing concentrations

of calcium ad magnesium ions in solution, respectively.

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Similar to the compacted PAA-Ca tubes, the magnesium-compacted PAA tubes could be restored by removing Mg2+ from the complex and breaking crosslinks between carboxylate groups of PAA. The magnesium chelation was carried out in both EDTA and

citrate solutions, respectively, and the resulting plots are summarized in Figure 5.14. The

swelling characteristics of compacted PAA-Mg tubes in EDTA and citrate solution are

shown in Figure 5.14c and d, which display a similar result to the one discussed above.

Apparently, citrate was more efficient than EDTA disodium salt used in the study with

respect to re-swelling the PAA-Mg tubes. Furthermore, it can be seen that the initial length (i.e. 83%) of PAA tubes in EDTA or citrate titration was different from the final length (i.e. 77%) of the tubes in MgCl2 titration. The reason for this inconsistency was that the PAA-Mg complex is relatively unstable in solution and partially dissociated after rinsing with water before titration with EDTA or citrate.

130

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5.3.6 Potential Biomimicking Applications

As an example of functional biological systems, the neutralized PAA nanofiber tubes can

be used as polymer models to mimic the axon cortical layer and its cation-exchange properties. It has been suggested that the action potential occurring in the peripheral cytoskeleton of the axon cortical layer could be initiated by replacing calcium with a monovalent ion, Na+ [152-154]. In a proposed model [172], sodium ions flow into the peripheral cytoskeleton and begin displacing calcium ions, which loosens the whole network and enables the layer expand, as shown in Figure 5.15. Meanwhile, the entering sodium ions drive the cell potential to a more positive value, establishing the action potential’s rising phase. At some point, the increase of adsorbed sodium would impair the hydrated structure of the cortical layer to the point that the network’s retractive force dominates against the swelling force, and the network begins to collapse. With anionic sites brought into closer apposition, calcium ions can displace the bound sodium, restoring both volume and cell potential back to their initial values. Tasaki and coworkers have reported that the propagating nerve impulse is a running wave of reversible structural change which represents a continuous displacement of the boundary between the site of swelling and the site of shrinkage of the cortical gel layer [155, 156].

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Figure 5.15 Proposed models of structural dynamics and the action potential. Initially (A)

the network is collapsed as strands are bridged by calcium. The network expands as

sodium replaces calcium (B). But increasing sodium eventually neutralizes surface

charge, weakens water structure and allows the polymer-retractive force to collapse the network (C), at which stage calcium may easily bridge the strands once again. [172]

In the previous sections, it has been shown that neutralized PAA nanofibrous tubes fabricated in this study can undergo similar reversible and abrupt structural changes with divalent-monovalent cation exchange. These structural changes (shrinking and swelling) are so sharp that they can be initiated and completed by a small change of Ca2+ or EDTA

concentration in solution. In addition, the fibrous structure of PAA tubes may also

resemble the unique fibrous and porous characteristics of the peripheral cytoskeleton in

the cortical layer of nerve fibers. Therefore, neutralized PAA nanofibrous tubes could be

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promising candidates to construct an abiotic, polymer mimic of an axon. Furthermore, the

reversible contracting behavior according to Ca2+ / Na+ concentrations could be utilized in

artificial muscle applications. Further studies focusing on the mechanism of the phase

transition of PAA nanofibrous tubes are in progress.

5.4 Conclusions

In summary, PAA nanofibers electrospun under various processing parameters were

systematically investigated. The results showed that uniform fine fibers with an average

diameter of 890±90nm were obtained at a polymer concentration of 4wt% with a flow

rate of 0.8mL/h. The samples were rendered water-insoluble by heat-induced

esterification and the fiber structure was preserved in water.

For the axon-mimic applications, PAA nanofibrous tubes were fabricated and neutralized

in a base solution. Neutralized PAA tubes consisted of a swollen and porous polymer

network in water. Sodium analysis of neutralized PAA tubes indicated a crosslink content

of approximately 20% after the thermal esterification, and about 80% of carboxylate

groups in PAA were available for phase transition experiments.

Divalent-monovalent ion exchange-induced shrinking behavior of neutralized PAA tubes was investigated using Ca2+ and Mg2+ as model species. The contraction of PAA tubes was caused by complex formation of divalent ions and carboxylate groups. Calcium analysis of PAA-Ca tubes indicated approximately 32% of total carboxylate groups were bonded with Ca2+ and about 48% of carboxylate groups remained in PAA fibers after

complete calcium titration.

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Two different shrinking behaviors of crosslinked PAA tubes were discovered. The cross- bridge between Ca2+ and two COO- groups occurred in a cooperative fashion, which

means the first complex assists the formation of a second one, while ionic crosslinks with

Mg2+ and COO- were formed directly without evidence of cooperative behavior. The

complex of PAA-Ca was found to be relatively stronger than the PAA-Mg complex.

The compacted PAA tubes were restored in EDTA or citrate solution by removing Ca2+

or Mg2+ from the complexes. Citrate was found to be more efficient than EDTA disodium

salt with respect to competing for divalent metal ions in the compacted tubes.

This reversible, abrupt structural change of neutralized PAA nanofibrous tubes triggered

by divalent-monovalent ion exchange may resemble the phase transition of the peripheral

cytoskeleton of the axon cortical layer in nerve fibers. Studies of PAA tubes in this work

may help to better understand the electrophysiological processes known as nerve

excitation and conduction in the nervous system, and the PAA tubes might be used as

polymer models of artificial axons for potential tissue engineering and nerve repair

applications.

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Chapter VI Conclusions and Future Work

6.1 Conclusions

Collagen, as basic structural element for native extracellular matrix (ECM), has been

used as a biomaterial in a variety of tissue engineering applications. Electrospinning of

collagen has been demonstrated to be a facile method of producing nanofibrous collagen

constructs that resemble the native state. Not surprisingly, the electrospun collagen fibers

do not appear to fully reconstitute the structural or mechanical properties of the parent

material, mainly due to the lack of native inter- and intramolecular crosslinks between collagen molecules.

In order to improve the mechanical properties of collagen scaffolds electrospun from the benign solvent solution, collagen-PCL sheath-core bicomponent fibers with diameters of

0.95 ± 0.23 µm were successfully fabricated through a novel co-axial electrospinning approach. The buffer-ethanol binary solvent was thought to facilitate the formation of a compound Taylor cone, with the ethanol apparently increasing the similarity of the core and sheath solutions, and the buffer increasing the conductivity of collagen solution thus increasing the shearing force generated at the interface. As expected, the addition of a

PCL core into collagen nanofibers markedly increased the mechanical strength of collagen scaffolds and further improvement was achieved upon post-crosslinking of the collagen-PCL sheath-core bicomponent fibers in an EDC/NHS ethanol solution.

In order to solve the problems dominating traditional collagen crosslinking methods, such as dimensional shrinking and loss of porous morphology, and to simplify the crosslinking procedure for electrospun collagen scaffolds, a novel single-step, in-situ crosslinking

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approach was developed. The excess amount of NHS present in the crosslinking mixture

was found to delay the EDC coupling reaction for several hours. Two plausible reasons

for the delayed crosslinking reaction in the presence of excess NHS are discussed: one

proposes a large amount of less active amine-reactive NHS ester formed before the stable

amide bond formation; the other concerns a competition between the amino groups of the

second collagen molecule and the mobile NHS molecules to attack the aforementioned

NHS ester.

Water-stable collagen nanofibers with a diameter of 0.42 ± 0.11 µm were successfully

generated by electrospinning an in-situ crosslinking collagen solution. The diameter of

electrospun collagen fibers slightly increased during electrospinning, which indicated the

onset of in-situ crosslinking. Humidity was found to be an important factor to control the

in-situ crosslinking rate and the subsequent fiber morphology, and the humidity of 43%

was discovered to be preferable to afford a sufficiently-crosslinked collagen network

within 3 days.

The in-situ crosslinked collagen scaffolds exhibited swelling behavior when placed in

water, in contrast to the dimensional shrinking frequently observed in post-crosslinked collagen materials. The porous surface morphology of the resulting mats was preserved in water-treatment which is one of the most important features of the in-situ crosslinked scaffolds. The in-situ crosslinked collagen demonstrated a similar uniaxial tensile behavior of native tissue in mechanical testing and displayed a characteristic triple-helical structure in FTIR spectra, which suggested the in-situ crosslinking approach did not significantly compromise the nature of the original collagen.

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A preliminary study of the use of electrospun in-situ crosslinked collagen scaffolds in a

wound dressing application showed they had potential to promote quick and scarless

wound healing. Trace amounts of EDC / NHS residues and subsequent urea byproducts

present in collagen scaffolds apparently did not display a negative influence on CHO cell

culture and open-wound closing tests. Preliminary drug release studies were carried out

by loading both hydrophobic and hydrophilic drugs into in-situ crosslinked collagen

scaffolds through electrospinning. The release profile of the antibiotic doxycycline from

the collagen mats exhibited a burst release in PBS buffer due to the swelling behavior of

crosslinked collagen.

Various physical forms of collagen gels, such as plates, tube, hemispheres and collagen

stamps, were also generated from the in-situ crosslinking collagen solution to demonstrate the capability of the in-situ crosslinking method to produce customized collagen gel-based materials. The addition of crosslinkers was found to decrease the collagen solubility in solution presumably due to crosslink formation. Increasing salt concentration can improve the solubility of collagen and probably results in a more complete dissociation of collagen bundles. Moreover, the collagen concentration needs to be high enough (i.e., 16 wt%) to form a crosslinked gel. Ethanol also plays an important role in collagen solubilization and gelation. However, it was found to cause an unfavorable cell detachment in the CHO cell seeding study. Reducing ethanol content in the resulting collagen gel showed a significant improvement of cell biocompatibility. An in-situ crosslinked collagen gel generated from the mixture of 20x PBS buffer and ethanol with volume ratio of 90:10 displayed a significantly favorable cell attachment with multiple cell layers formed inside the gel.

138

Porous collagen sponges were prepared by freeze-drying of the in-situ crosslinked collagen gels. The porosity was controlled through different freezing processes. Lower freezing temperatures provided smaller pore sizes. The smallest pores with a diameter of

35 ± 10 µm were observed in collagen sponges initially frozen with liquid nitrogen.

Preliminary in-vivo wound healing tests indicate that the in-situ crosslinked collagen sponges facilitate wound closure and may be a good candidate for wound healing and skin regeneration.

An initial study of electrophoretic mineralization indicates that in-situ crosslinked collagen gels and sponges could be promising candidates for bone substitute applications.

Combining the in-situ crosslinking method with an electrophoretic approach to mineralization can provide many advantages to existing systems and could be a viable approach to bone replacement with further refinement. Collagen was in-situ crosslinked to form a hydrogel, which was subjected to electrophoretic mineralization that substantially seeded the hydrogel with calcium phosphate. Freezing-drying these materials provided a very porous network of mineralized collagen sponges, similar to what can be found in natural bone, and this discovery may be a step towards bone replacement materials.

Poly (acrylic acid) (PAA) is a well-known polymer because of its two major properties: extreme water absorption because of the ionic nature of neutralized PAA and interaction with metal ions. PAA nanofibers containing ethylene glycol (EG) electrospun under various processing parameters were systematically investigated. The results showed that uniform fine fibers with an average diameter of 0.89 ± 0.09 µm were obtained at a polymer concentration of 4wt% with a flow rate of 0.8mL/h. The samples were rendered

139

water-insoluble by heat-induced esterification and the fiber structure was preserved in

water.

For the axon-mimic applications, PAA nanofibrous tubes were fabricated and neutralized

in a base solution. Neutralized PAA tubes consisted of a swollen and porous polymer

network in water. Sodium analysis of neutralized PAA tubes indicated a crosslink content

of approximately 20% after thermal esterification, and about 80% of carboxylate groups

in PAA were available for phase transition experiments.

Divalent-monovalent ion exchange-induced shrinking behavior of neutralized PAA tubes

was investigated using Ca2+ and Mg2+ as model species. The contraction of PAA tubes was caused by complex formation of divalent ions and carboxylate groups. Calcium analysis of PAA-Ca2+ tubes indicated approximately 32% of total carboxylate groups

were bonded with Ca2+ and about 48% of carboxylate groups remained in PAA fibers

after complete calcium titration.

Two different shrinking behaviors of crosslinked PAA tubes were discovered. The cross-

bridge between Ca2+ and two COO- groups occurred in a cooperative fashion, which

means the first complex assists the formation of a second one, while ionic crosslinks with

Mg2+ and COO- were formed directly without evidence of cooperative behavior. The

complex of PAA-Ca2+ was found to be relatively stronger than the PAA-Mg2+ complex.

The compacted PAA tubes were restored in EDTA or citrate solution by removing Ca2+

or Mg2+ from the complexes. Citrate was found to be more efficient than EDTA disodium salt with respect to competing for divalent metal ions in the compacted tubes.

140

This reversible, abrupt structural change of neutralized PAA nanofibrous tubes triggered

by divalent-monovalent ion exchange may resemble the phase transition of the peripheral

cytoskeleton of the axon cortical layer in nerve fibers. Studies of PAA tubes in this

research may help to better understand the electrophysiological processes known as nerve

excitation and conduction in the nervous system, and the PAA tubes might be used as

polymer models of artificial axons for potential tissue engineering and nerve repair

applications.

6.2 Future Work

The use of the in-situ crosslinking approach to produce water-stable, non-toxic collagen- based materials is novel and unique. Although the preliminary FTIR spectra of in-situ crosslinked collagen fibers suggested that the characteristic triple helical structure of collagen were preserved in the resultant scaffolds, the actual collagen structures present in the in-situ crosslinking solution or the benign solvent system are still unclear. The exploration of the influence of the ethanol-buffer binary solution on the collagen hierarchical structures may help to better understand the underlying solubilization mechanism of the benign solvent system and the EDC / NHS delayed crosslinking reactions, and help to predict the effects achieved by potential modifications. For this purpose, small-angle X-ray scattering (SAXS), circular dichroism (CD), and nuclear magnetic resonance (NMR) may be the right tools to acquire such structural information.

Preliminary in-vivo wound healing testing indicated that both in-situ crosslinked collagen electrospun fiber scaffolds and freeze-dried sponges were able to accelerate wound closure and could be good candidates for wound healing and skin regeneration. Detailed histological examination of epithelialization and granulation are needed in order to

141

understand the functions of collagen scaffolds in the wound healing process and

accordingly improve their performance.

Being a part of the native extracellular matrix (ECM), the collagen used in this study

showed excellent biocompatibility and biodegradability in cell culture and wound healing

experiments. The ECM also contains other compounds that play critical roles to maintain

cellular functions. Such compounds include elastin, laminin, vitronectin, fibronectin, and

glycosaminoglycans (GAGs). The inclusion of such materials in the collagen scaffolds

through the in-situ crosslinking approach may further improve cell growth behavior on

the scaffolds.

Preliminary drug release studies have been carried out on in-situ crosslinked collagen

fiber scaffolds and the release profile of hydrophilic doxycycline exhibited a burst release

in PBS buffer due to the swelling behavior of crosslinked collagen. It is of interest to

investigate the release of different drugs (e.g. hydrophobic drugs such as Silicon

phthalocyanine Pc4) from different types of collagen constructs (e.g. gels and sponges).

The combination of collagen and specific drugs may represent a convenient system for

the local delivery of drugs for wound healing and skin regeneration.

Divalent-monovalent ion exchange-induced phase transitions of neutralized PAA tubes were demonstrated in the current research using Ca2+ and Mg2+ as model species, which showed surprisingly different results, with Ca2+-crosslinking occurring in a cooperative fashion and Mg2+-crosslinking without evidence of cooperative behavior. Further studies

of PAA phase transitions induced by other cations, such as Zn2+ or Fe3+, are also of

142

special interest and may render intriguing insights regarding natural physiological

activities.

The combination of collagen and PAA is extremely interesting because the presence of

PAA in composite scaffolds may impart the divalent-monovalent ion exchange-induced reversible contraction behaviors to the scaffolds with the collagen providing biocompatibility and bioactivity to the structure. The carboxylic acid groups of PAA may be involved into the EDC/NHS delayed crosslinking with collagen, and thus the thermal crosslinking of PAA may be not needed. The resultant the composite fiber scaffolds may be useful in skin replacement and nerve repair applications.

143

References

1. Chen, G.; Ushida, T.; Tateishi, T. “Development of biodegradable porous scaffolds

for tissue engineering.” Mater. Sci. and Engg. 2001, C17, 63-69.

2. Chapekar M. “Tissue Engineering: Challenges and Opportunities.” J. Biomed.

Mater.Res. (Appl. Biomater.) 2000, 53, 617-620.

3. Langer, R. “Tissue Engineering.” Molecular Therapy 2000, 1, 12-15.

4. Chen, G.; Ushida, T.; Tateishi, T. “Scaffold Design for Tissue Engineering.”

Macromol.Biosci. 2002, 2, 67-77.

5. Bareil, R.P.; Gauvin, R.; Berhod, F. “Collagen-Based Biomaterials for Tissue

Engineering Applications.” Materials 2010, 3, 1863-1887.

6. Cen, L.; Liu, W.; Cui, L.; Zhang, W.; Cao, Y. “Collagen Tissue Engineering:

Development of Novel Biomaterials and Applications.” Biomaterials 2008, 63, 492-

496.

7. Glowacki, J.; Mizuno, S. “Collagen Scaffolds for Tissue Engineering.” Biopolymers

2007, 89, 338-344.

8. Lee, C.H.; Singla, A.; Lee, Y. “Biomedical applications of collagen.” Int. J. Pharm.

2001, 221, 1-22.

9. Friess, W. “Collagen-biomaterial for drug delivery.” Euro. J. Pharm. Biopharm. 1998,

45, 113-136.

10. Koob, T.J. “Collagen fixation” In: Wnek, G.E.; Bowlin, G.L., eds. Encyclopedia of

Biomaterials and Biomedical Engineering. New York: Marcel Dekker, 2004, 335-347.

11. Kielty, C. M., and Grant, M. E. “The collagen family: structure, assembly, and

organisation in the extracellular matrix. Connective tissue and its heritable disorders.”

144

Molecular Genetics in Medical Aspects, 2nd ed., Wiley Liss, New York, 2002, 159-

221.

12. Miller, E.J. “Collagen types: structure, distribution and functions.” in: M.E. Nimni

(Ed.), Collagen Vol. I-Biochemistry, CRC Press, Boca Raton, FL, 1988, 139-157.

13. Kucharz, E.J. “The Collagens: Biochemistry and Pathophysiology.” Springer-Verlag,

Berlin, 1992, 7-29.

14. Piez, K.A. “Collagen” in: J.I. Kroschwitz (Ed.), Encyclopedia of Polymer Science and

Engineering, Wiley, New York, 1985, 699-727.

15. Timpl, R. “Immunology of the collagens.” in: K.A. Piez, A.H. Reddi (Eds.),

Extracellular Matrix Biochemistry, Elsevier, New York, 1984, 159-190.

16. Kucharz, E.J. “The Collagens: Biochemistry and Pathophysiology.” Springer-Verlag,

Berlin, 1992, 34-39.

17. Piez, K.A. “Molecular and aggregate structures of the collagens.” in: K.A. Piez, A.H.

Reddi (Eds.), Extracellular Matrix Biochemistry, Elsevier, New York, 1984, 1-40.

18. Brodsky, B.; Tanaka, S.; Eikenberry, E.F. “X-ray diffraction as a tool for studying

collagen structure.” in: M.E. Nimni (Ed.), Collagen Vol. I - Biochemistry, CRC Press,

Boca Raton, FL, 1988, 95-112.

19. Doyle, B.B.; Bendit, E.G.; Blout, E.R. “Infrared spectroscopy of collagen and

collagen-like polypeptides.” Biopolymers 1975, 14, 937-957.

20. Persikov, A.V.; Ramshaw, J.A.M.; Kirkpatrick, A.; Brodsky, B. “Amino Acid

Propensities for the Collagen Triple-Helix.” Biochemistry 2000, 39, 14690-14697.

145

21. Shah, N.K.; Brodsky, B.; Kirkpatrick, A.; Ramshaw, J.A.M. “Structural

Consequences of D-Amino Acids in Collagen Triple-Helical Peptides.” Biopolymers,

1999, 49, 297-302.

22. Nimni, M.E.; Harkness, R.D. “Molecular structures and functions of collagen.” in:

M.E. Nimni (Ed.), Collagen Vol. I- Biochemistry, CRC Press, Boca Raton, FL, 1988,

1-79.

23. Yamauchi, M.; Mechanic, G. “Crosslinking of collagen.” in: M.E. Nimni (Ed.),

Collagen Vol. I - Biochemistry, CRC Press, Boca Raton, FL, 1988, 157-172.

24. Reiser, K.; McCormick, R.J.; Rucker, R.B. “Enzymatic and nonenzymatic cross-

linking of collagen and elastin.” FASEB J. 1992, 6, 2439-2449.

25. Graham, L.; Gallop, P.M. “Covalent protein crosslinks: general detection,

quantitation, and characterization via modification with diphenylborinic acid.” Anal.

Biochem. 1994, 217, 298–305.

26. Hanson, D.A.; Eyre, D.R. “Molecular site specificity of pyridinoline and pyrrole

crosslinks in type I collagen of human bone.” J. Biol. Chem. 1996, 271, 26508-26516.

27. Pratt, D.A.; Daniloff, Y.; Duncan, A.; Robins, S.P. “Automated analysis of the

pyridinium crosslinks of collagen in tissue and urine using solid-phase extraction and

reversed-phase high-performance liquid chromatography.” Anal. Biochem.1992, 207,

168-175.

28. Palokangas, H.; Kovanen, V.; Duncan, A.; Robins, S.P. “Age-related changes in the

concentration of hydroxypyridinium crosslinks in functionally different skeletal

muscles.” Matrix 1992, 12, 291-296.

146

29. Fielding, A.M. “Preparation of neutral salt soluble collagen.” in: D.A. Hall (Ed.), The

Methodology of Connective Tissue Research, Joynson-Bruvvers, Oxford, 1976, 9-12.

30. Trelstad, R.L. “Immunology of collagens.” in: H. Furthmayer (Ed.),

Immunochemistry of the Extracellular Matrix, Vol. I Methods, CRC Press, Boca

Raton, FL, 1982, 32–39.

31. Bazin, S.; Delaumay, A. “Preparation of acid and citrate soluble collagen.” in: D.A.

Hall (Ed.), The Methodology of Connective Tissue Research, Joynson–Bruvvers,

Oxford, 1976, 13-18.

32. Cioca, G. “Process for preparing macromolecular biologically active collagen.” US

Patent 4279812 (1981).

33. Roreger, M. “Collagen preparation for the controlled release of active substances.”

PCT WO 95/28964 (1995).

34. Li, S.-T. “Tissue-derived biomaterials (collagen).” in: J.D. Bronzino (Ed.), The

Biomedical Engineering Handbook, CRC Press, Boca Raton, FL, 1995, 627-647.

35. Gao, T.J.; Lindholm, T.S. “Searching for a novel carrier for bioactive delivery of

bone morphogenetic protein.” in: T.S. Lindholm (Ed.), Bone Morphogenetic Proteins:

Biology, Biochemistry and Reconstructive Surgery, R.G. Landes, Austin, TX, 1996,

121-130.

36. Jayakrishnan, A.; Jameela, S.R. "Glutaraldehyde as a fixative in bioprosthetic and

drug delivery matrices." Biomaterials, 1996, 17, 471-484.

37. Olde Damink, L.H.H.; Dijkstra, P.J.; Luyn, M.J.A.; Wachem, P.B.; Nieuwenhuis, P.;

Feijen, J. "Glutaraldehyde as crosslinking agent for collagen based biomaterials." J.

Mat. Sci.: Mat in Med. 1995, 6, 460-472.

147

38. Rault, I.; Frei, V.; Herbage, D.; Abdul-Marak, N.; Huc, A. "Evaluation of different

chemical methods for cross-linking collagen gels, films and sponges." J. Mat. Sci:

Mat in Med., 1996, 7, 215-222.

39. Cheung, D.T.; Perelman, N.; Ko, E.C.; Nimni, M.E. "Mechanism of cross-linking of

proteins by glutaraldehyde III. Reaction with collagen in tissues." Conn. Tissue

Res.1985, 13, 109-115.

40. Girardot, M.N.; Torrianni, M.; Dillehay, D.; Girardot, J.M. "Role of glutaraldehyde in

calcification of procine heart valves: comparing cusp and wall." J. Biomed. Mat.

Res.1995, 29, 793-801

41. Wachem P.B.; Luyn, M.J.A.; Olde Damink, L.H.H.; Dijkstra, P.J.; Feijen, J.;

Nieuwenhuis, P. "Biocompatibility and tissue regenerating capacity of crosslinked

dermal sheep collagen." J. Biomed. Mat. Res.1994, 28, 353-363.

42. Grimm, M.; Eybl, E.; Grabenwöger, M.; Spreitzer, H.; Jäger, W.; Grimm, G.; Böck,

P.; Müller, M.; Wolner, E. "Glutaraldehyde affects biocompatibility of bioprosthetic

heart valves." Surgery 1992, 111, 74-78.

43. Lee, J.M.; Pereira, C.A.; Kan, L.W.K. "Effect of molecular structure of poly (glycidyl

ether) reagents on crosslinking and mechanical properties of bovine pericardial

xenograft materials." J. Biomed. Mat. Res.1994, 28, 981-992.

44. Sung, H.-W.; Hsu, H.-L.; Shih, C.-C.; Lin, D.-S. "Cross-linking characteristics of

biological tissue fixed with monofunctional or multifunctional epoxy compounds."

Biomaterials 1996, 17, 1405-1410.

148

45. Tu, R.; Quijano, R.C.; Lu, C.L.; Shen, S.; Wang, E.; Hata, C.; Lin, D. "A preliminary

study of the fixation mechanism of collagen reaction with a polyepoxy fixative." Int.

J. Art. Org. 1993, 16, 537-544.

46. Nishi, C.; Nakajima, N.; Ikada, Y. "In vitro evaluation of cytotoxicity of diepoxy

compounds used for biomaterials." J. Biomed. Mat. Res.1995, 29, 829-835.

47. Olde Damink, L.H.H.; Dijkstra, P.J.; Luyn, M.J.A.; Wachem, P.B.; Nieuwenhuis, P.;

Feijen, J. "Crosslinking of dermal sheep collagen using a water-soluble

carbodiimide." Biomaterials 1996, 17, 765-773.

48. Graberek Z.; Gergely, J. "Zero-length cross-link procedure with the use of active

esters." Analyt. Biochem.1990, 185, 131-135.

49. Nimni, M.E.; Cheung, D.; Strates, B.; Kodana, M.; Sheikh, K. "Chemically modified

collagen: A natural biomaterial for tissue replacement." J. Biomed. Mat. Res.1987, 21

741-771.

50. Wachem, P.B.; Luyn, M.J.A.; Olde Damink, L.H.H.; Dijkstra, P.J.; Feijen, J.;

Nieuwenhuis, P. "Tissue regenerating capacity of carbodiimide-crosslinked dermal

sheep collagen during repair of the abdominal wall." J. Artif. Organs 1994, 17, 230-

239.

51. Lee, J.M.; Edwards, H.L.; Pereira, C.A.; Samii, I.S. "Cross-linking of tissue-derived

biomaterials in 1-ethyl-3-(dimethylaminopropyl)-carbodiimide." J. Mat. Sci.: Mat. in

Med.1996, 7, 531-542.

52. Scholnick, F.; Liao, L.L.; Brown, E.M.; Feairheller, S.H. "Crosslinking of collagen

with dicarboxylic acids." J. Am. Leather Chem. Ass.1992, 87, 333-338.

149

53. Anselme, K.; Petite, H.; Herbage, D. "Inhibition of calcification in vivo by acyl azide

cross-linking of a collagen-GAG sponge." Matrix 1992, 12, 264-273.

54. Petite, H.; Frei, V.; Huc, A.; Herbage, D. "Use of diphenylphosphorylazide for cross-

linking collagenbased materials." J. Biomed. Mat. Res 1994, 28, 159-165

55. Khor, E. “Methods for the treatment of collagenous tissues for bioprostheses.”

Biomaterials 1997, 18, 95-105.

56. Weadock, K.S.; Miller, E.J.; Bellincampi, L.D.; Zawadsky, J.P.; Dunn, M.G.

“Physical crosslinking of collagen fibers: comparison of ultraviolet irradiation and

dehydrothermal treatment.” J. Biomed. Mater. Res. 1995, 29, 1373–1379.

57. Yannas, I.V.; Tobolsky, A.V. "Cross-linking of gelatin by dehydration." Nature 1967,

215, 509-510.

58. Law, J.K.; Parsons, J.R.; Silver, F.H.; Weiss, A.B. "An evaluation of purified

reconstituted type I collagen fibers." J. Biomed. Mat. Res. 1989, 23, 961-977.

59. Griffin, M.; Casadio, R.; Bergamini, C.M. “Transglutaminases: nature’s biological

glues.” J. Biochem. 2002, 368, 377-396.

60. Collighan, R.; Cortez, J.; Griffin, M. “The biotechnological applications of

transglutaminases.” Minerva. Biotechnol. 2002, 14,143-148.

61. Chau, D.Y.S.; Collighan, R.J.; Verderio, E.A.M.; Addy, V.L.; Griffin, M. “The

cellular response to transglutaminase-cross-linked collagen.” Biomaterials 2005, 26,

6518-6529.

62. Sell, S.A.; Wolfe, P.S.; Garg, K.; McCool J.M.; Rodriguez, I.A.; Bowlin, G.L. “The

Use of Natural Polymers in Tissue Engineering: A Focus on Electrospun

Extracellular Matrix Analogues.” Polymers 2010, 2, 522-553.

150

63. Barnes, C.P.; Sell, S.A.; Boland, E.D.; Simpson, D.G.; Bowlin, G.L. “Nanofiber

technology: Designing the next generation of tissue engineering scaffolds.” Adv.

Drug Delivery Rev. 2007, 59, 1413-1433.

64. Sill, T.J.; von Recum, H. “Electrospinning: Applications in drug delivery and tissue

engineering.” Biomaterials 2008, 29, 1989-2006.

65. Theron, S.A.; Zussman, E.; Yarin, A.L. “Experimental investigation of the governing

parameters in the electrospinning of polymer solutions.” Polymer 2004, 45, 2017-

2030.

66. Huang, L.; Nagapudi, K.; Apkarian, R.P.; Chaikof, E.L. “Engineered collagen-PEO

nanofibers and fabrics.” J. Biomater. Sci. Polym. Ed. 2001, 12, 979-993.

67. Song, J-H.; Kim, H-E.; Kim, H-W. “Production of electrospun gelatin nanofiber by

water-based co-solvent approach.” J Mater Sci: Mater Med, 2008, 19, 95-102.

68. Kia, C.S.; Baeka, D.H.; Ganga, K.D.; Leea, K.H.; Umb, I.C.; Park, Y.H.

“Characterization of gelatin nanofiber prepared from gelatin–formic acid solution.”

Polymer 2005, 46, 5094-5102.

69. Matthews, J.A.; Wnek, G.E.; Simpson, D.G.; Bowlin, G.L. “Electrospinning of

collagen nanofibers.” Biomacromolecules 2002, 3, 232-238.

70. Matthews, J.A.; Boland, E.D.; Wnek, G.E.; Simpson, D.G.; Bowlin, G.L.

“Electrospinning of Collagen Type II: A Feasibility Study.” J. Bioact. Compat. Polym.

2003, 18, 125-134.

71. Barnes, C.P.; Sell, S.A.; Knapp, D.C.; Walpoth, B.H.; Brand, D.D.; Bowlin, G.L.

“Preliminary investigation of electrospun collagen and polydioxanone for vascular

tissue engineering applications.” Int. J. Electrospun Nanofiber. Appl. 2007, 1, 73-87.

151

72. Dong, B.; Arnoult, O.; Smith, M.E.; Wnek, G.E. “Electrospinning of Collagen

Nanofiber Scaffolds from Benign Solvents.” Macromol. Rapid Commun. 2009, 30,

539-542.

73. Choi, J.S.; Lee, S.W.; Jeong, L.; Bae, S.H.; Min, B.C.; Youk, J.H.; Park, W.H.

“Effect of organosoluble salts on the nanofibrous structure of electrospun poly(3-

hydroxybutyrate-co-3-hydroxyvalerate).” Int. J. Biol. Macromol. 2004, 34, 249-256.

74. Zhong, S.; Teo, W.E.; Zhu, X.; Beuerman, R.; Ramakrishna, S.; Yung, L.Y.

“Formation of collagen-glycosaminoglycan blended nanofibrous scaffolds and their

biological properties.” Biomacromolecules 2005, 6, 2998-3004.

75. Thomas, V.; Dean, D.R.; Jose, M.V.; Mathew, B.; Chowdhury, S.; Vohra, Y.K.

“Nanostructured biocomposite scaffolds based on collagen coelectrospun with

nanohydroxyapatite.” Biomacromolecules 2007, 8, 631-637.

76. Yeo, I.S.; Oh, J.E.; Jeong, L.; Lee, T.S.; Lee, S.J.; Park, W.H.; Min, B.M. “Collagen-

based biomimetic nanofibrous scaffolds: preparation and characterization of

collagen/silk fibroin bicomponent nanofibrous structures.” Biomacromolecules 2008,

9, 1106-1116.

77. Martins-Green, M. “The dynamics of Cell-ECM interactions with implications for

tissue engineering.” In: Principles of Tissue Engineering; Lanza, R. Langer, R, Chick,

W, Eds.; R.G. Landes Company: Austin, TX, USA, 1997, 23-46.

78. He, W.; Yong, T.; Teo, W.E.; Ma, Z.; Ramakrishna, S. “Fabrication and

endothelialization of collagen-blended biodegradable polymer nanofibers: potential

vascular graft for blood vessel tissue engineering.” Tissue Eng. 2005, 11, 1574-1588.

152

79. Kidoaki, S.; Kwon, I.K.; Matsuda, T. “Mesoscopic spatial designs of nano- and

microfiber meshes for tissue-engineering matrix and scaffold based on newly devised

multilayering and mixing electrospinning techniques.” Biomaterials 2005, 26, 37-46.

80. Lee, S.J.; Liu, J.; Oh, S.H.; Soker, S.; Atala, A.; Yoo, J.J. “Development of a

composite vascular scaffolding system that withstands physiological vascular

conditions.” Biomaterials 2008, 29, 2891-2898.

81. Tillman, B.W.; Yazdani, S.K.; Lee, S.J.; Geary, R.L.; Atala, A.; Yoo, J.J. “The in

vivo stability of electrospun polycaprolactone-collagen scaffolds in vascular

reconstruction.” Biomaterials 2009, 30, 583-588.

82. Boland, E.D.; Coleman, B.D.; Barnes, C.P.; Simpson, D.G.; Wnek, G.E.; Bowlin,

G.L. “Electrospinning polydioxanone for biomedical applications.” Acta Biomater.

2005, 1, 115-123.

83. Liang, D.H.; Hisao, B.S.; Chu, B. “Functional electrospun nanofibrous scaffolds for

biomedical applications.” Adv. Drug Deliv. Rev. 2007, 59, 1392-1412.

84. Yoshimoto, H.; Shin, Y.; Terai, H.; Vacanti, J. “A biodegradable nanofiber scaffold

by electrospinning and its potential for bone tissue engineering.” Biomaterials 2003,

24, 2077-2082.

85. Nair, L.; Bhattacharya, S.; Laurencin, C. “Development of novel tissue engineering

scaffolds via electrospinning.” Expert Opin. Biol. Ther. 2004, 4, 659-668.

86. Ikada, Y. “Challenges in tissue engineering”, J. R. Soc. Interface 2006, 3, 589-601.

87. Wnek, G.; Carr, M.; Simpson, D.; Bowlin, G. “Electrospinning of Nanofiber

Fibrinogen Structures.” Nano Lett. 2003, 3, 213-216.

153

88. Chen, G.; Ushida, T.; Tateishi, T. “A hybrid network of synthetic polymer mesh and

collagen sponge.” Chem. Commun. 2000, 1505-1506.

89. Pierschbacher, M.; Ruoslahti E. “Cell attachment activity of fibronectin can be

duplicated by small synthetic fragments of the molecule.” Nature 1984, 309, 30-33.

90. Hakkarainen, M. “Aliphatic Polyesters: Abiotic and Biotic Degradation and

Degradation Products.” Adv. Polym. Sci. 2002, 157, 113-138.

91. Zhang, Y.; Venugopal, J.; Huang, Z.; Lim, C.; Ramakrishna, S. “Characterization of

the surface biocompatibility of the electrospun PCL-collagen nanofibers using

fibroblasts”, Biomacromol. 2005, 6, 2583–2589.

92. Moghe A. K.; Gupta, B. S., “Co-axial electrospinning for nanofiber structures:

preparation and applications”, Polym. Rev. 2008, 48, 353–377.

93. Sun, Z.; Zussman, E.; Yarin, A.; Wendorff J.; Greiner, A. “Compound core-shell

polymer nanofibers by co-electrospinning”, Adv. Mater. 2003, 15, 1929–1932.

94. Greiner, A.; Wendorff, J.; Yarin, A.; Zussman E. “Biohybrid nano-systems with

polymer nanofibers and nanotubes”, Appl. Microbiol. Biotechnol. 2006, 71, 387-393.

95. Yu, J.; Fridrikh, S.; Rutledge G. “Production of submicrometer diameter fibers by

two fluid electrospinning”, Adv. Mater. 2004, 16, 1562-1566.

96. Zhang, Y.; Huang, Z.; Xu, X.; Lim, C.; Ramakrishna, S. “Preparation of core-shell

structured PCL-r-Gelatin bi-component nanofibers by co-axial electrospinning”,

Chem. Mater. 2004, 16, 3406–3409.

97. Huang, Z.; Zhang, Y.; Ramakrishna, S. “Double-layered composite nanofibers and

their mechanical performance”, J. Polym. Sci., Part B: Polym. Phys. 2005, 43, 2852–

2861.

154

98. McCann, J.; Marquez, M.; Xia, Y. “Melt coaxial electrospinning: a versatile method

for the encapsulation of solid materials and fabrication of phase change nanofibers”,

Nano Lett. 2006, 6, 2868-2872.

99. Li, D.; Xia, Y. “Direct fabrication of composite and ceramic hollow nanofibers by

electrospinning”, Nano Lett. 2004, 4, 933- 938.

100. Ruzin, E. “Plant Microtechnique and Microscopy”, Oxford University Press,

Oxford 1999.

101. Lee, J.J.; Yu, H.S.; Hong, S.J.;Jeong, I.; Jang, J.H.;Kim, H.W.,"Nanofibrous

membrane of collagen–polycaprolactone for cell growth and tissue regeneration", J

Mater Sci: Mater Med 2009, 20, 1927-1935.

102. Zeugolis, D.I.; Khew, S.T.; Yew, E.S.Y.; Ekaputra, A.E.; Tong, Y.W.; Yung, L.L.;

Hutmacher, D.W.; Sheppard, C.; Raghunath, M., “Electro-spinning of pure collagen

nano-fibres - just an expensive way to make gelatin?”, Biomaterials 2008, 29, 2293-

2305.

103. Díaz, J. E.; Barrero, A.; Márquez, M.; Loscertales, I. G. “Controlled

encapsulation of hydrophobic liquids in hydrophilic polymer nanofibers by co-

electrospinning”, Adv. Funct. Mater. 2006, 16, 2110-2116.

104. Rho, K.S.; Jeong, L.; Lee, G.; Seo, B.M.; Park, Y.J.; Hong, S.D.; Roh, S.; Cho,

J.J.; Park, W.H.; Min, B.M. “Electrospinning of collagen nanofibers: Effects on the

behavior of normal human keratinocytes and early-stage wound healing,”

Biomaterials 2006, 27, 1452-1461.

105. Buttafoco, L; Kolkman, N.G.; Engbers-Buijtenhuijs, P.; Poot, A.A.; Dijkstra, P.J.;

Vermes, I.; Feijen, J. “Electrospinning of collagen and elastin for tissue engineering

155

applications,” Biomaterials 2006, 27, 724-734.

106. Torres-Giner, S.; Gimeno-Alcaniz, J.V.; Ocio, M.J.; Lagaron, J.M. “Comparative

Performance of Electrospun Collagen Nanofibers Crosslinked by Means of Different

Methods,” ACS App. Mater. Interfaces 2009, 1, 218-233.

107. Van Wachem, P.B.; Van Luyn, M.J.A.; Nieuwenhuis, P.; Koerten, H.K.; Olde

Damink, L.; Ten Hoopen, H.; Feijen, J. “In vivo degradation of processed dermal

sheep collagen evaluation with transmission electron microscopy,” Biomaterial 1991,

215-223.

108. Barnes, C.P.; Pemble, C.W.; Brand, D.D.; Simpson, D.G.; Bowlin, G.L. “Cross-

Linking Electrospun Type II Collagen Tissue Engineering Scaffolds with

Carbodiimide in Ethanol,” Tissue Engineering 2007, 13, 1593-1605.

109. Staros, J.V.; Wright, R.W.; Swingle, D.M. “Enhancement by N-

hydroxysulfosuccinimde of water-soluble carbodiimide-mediated coupling reactions,”

Anal. Biochem. 1986, 156, 220-222.

110. Rockland, L.B. “Saturated Salt Solutions for Static Control of Relative Humidity

between 5 o and 40 oC,” Anal. Chem.1960, 1375-1376.

111. Nam, K.; Kimura, T.; Kishida, A., “Controlling Coupling Reaction of EDC and

NHS for Preparation of Collagen Gels Using Ethanol/Water Co-Solvents,” Macromol.

Biosci. 2008, 8, 32-37.

112. Jakobsen, R.J.; Brown, L.L.; Hutson, T.B.; Fink, D.J.; Veis, A. “Intermolecular

interactions in collagen self-assembly as revealed by Fourier transform infrared

spectroscopy,” Science 1983, 220, 1288-1290.

156

113. Zahedi, P.; Rezaeian, I.; Ranaei-Siadat, S.; Jafari, S.; Supaphol, P. “A review on

wound dressings with an emphasis on electrospun nanofibrous polymer bandages,”

Polym. Adv. Technol. 2010, 21, 77-95.

114. Chen, J.P.; Chang, G.Y.; Chen, J.K. “Electrospun collagen/chitosan nanofibrous

membrane as wound dressing,” Colloids and Surfaces A: Physicochem. Eng. Aspects

2008, 313-314, 183-188.

115. Liu, S.J.; Kau, Y.C.; Chou, C.Y.; Chen, J.K.; Wu, R.C.; Yeh, W.L “Electrospun

PLGA/collagen nanofibrous membrane as early-stage wound dressing,” Journal of

Membrane Science 2010, 355, 53-59.

116. Radu, F. A.; Bause, M.; Knabner, P.; Lee, G.W.; Friess, W.C. “Modeling of Drug

Release From Collagen Matrices,” Journal of Pharmaceutical Sciences 2002, 91,

964-972.

117. Albu, M.G.; Popa, L.; Ghica, M.V.; Leca, M.; Trandafir, V.; Titorence, I.

“Collagen Matrices for Drug Delivery: Preparation, Characterization and Kinetics of

Release,” European Cells and Materials 2008, 16, 1.

118. Wallace, D.G.; Rosenblatt, J. “Collagen gel systems for sustained delivery and

tissue engineering.” Adv. Drug Deliv. Rev. 2003, 55, 1631-1649.

119. Grillo, H.; Gross, J. “Thermal reconstitution of collagen from solution and the

response to its heterologous implantation.” J. Surg. Res. 1962, 2, 69-82.

120. Thevenot, P.; Nair, A.; Dey, J.; Yang, J.; Tang, L.P. “Method to analyze three-

dimensional cell distribution and infiltration in degradable scaffolds.” Tissue Eng.

Part C Methods 2008, 14, 319-331.

157

121. Zerris, V.A.; James, K.S.; Roberts, J.B.; Bell, E.; Heilman, C.B. “Repair of the

dura mater with processed collagen devices.” J. Biomed. Mater. Res. B Appl.

Biomater. 2007, 83B, 580-588.

122. Falanga, V.; Sabolinski, M. “A bilayered living skin construct accelerates

complete closure of hard-to-heal venous ulcers.” Wound Repair Regen 1999, 7, 201-

207.

123. Stella, J.A.; D’Amore, A.; Wagner, W.R.; Sacks, M.S. “On the biomechanical

function of scaffolds for engineering load-bearing soft tissues.” Acta. Biomater. 2010,

6, 2365-2381.

124. Guidry, C.; Grinnell, F. “Heparin modulates the organization of hydrated collagen

gels and inhibits gel contraction by fibroblasts.” J. Cell Biol. 1987, 104, 1097-1103.

125. Feng, Z.; Matsumoto, T.; Nakamura, T. “Measurements of the mechanical

properties of contracted collagen gels populated with rat fibroblasts or

cardiomyocytes.” J. Artif. Organs 2003, 6, 192-196.

126. Chan, B.P.; Chan, O.C.M.; So, K.F. “Effects of photochemical crosslinking on the

microstructure of collagen and a feasibility study on controlled protein release.” Acta.

Biomater. 2008, 4, 1627-1636.

127. Ibusuki, S.; Halbesma, G.J.; Randolph, M.A.; Redmond, R.W.; Kochevar, I.E.;

Gill, T.J. “Photochemically cross-linked collagen gels as three-dimensional scaffolds

for tissue engineering.” Tissue Eng, 2007, 13, 1995-2001.

128. Orban, J.M.; Wilson, L.B.; Kofroth, J.A.; El-Kurdi, M.S.; Maul, T.M.; Vorp, D.A.

“Crosslinking of collagen gels by transglutaminase.” J. Biomed. Mater. Res. Part A

2004, 68A, 756-762.

158

129. Charulatha, V.; Rajaram, A. “Influence of different crosslinking treatments on the

physical properties of collagen membranes.” Biomaterials 2003, 24, 759-767.

130. Pachence, J.M.; Berg, R.A.; Silver, F.H. “Collagen: its place in the medical

device industry.” Med. Device Diagn. Ind. 1987, 9, 49-55.

131. Yannas, I.V. “Biologically active analogues of the extracellular matrix: artificial

skin and nerves.” Angew. Chem. Int. Ed. 1990, 29, 20-35.

132. Sannino, A.; Madaghiele, M. “Tuning the Porosity of Collagen-based Scaffolds

for Use as Nerve Regenerative Templates.” Journal of Cellular Plastics 2009, 45,

137-155.

133. Ko, Y.G.; Kawazoe, N.; Tateishi, T.; Chen, G.P. “Preparation of Novel Collagen

Sponges Using an Ice Particulate Template.” Journal of Bioactive and Compatible

Polymers 2010, 25, 360-373.

134. Oliveira, S.M.; Rigshia, R.A.; Legeros, R.Z.; Clark, E.; Yost, M.J.; Terracio, L.;

Teixiera, C.C. “An improved collagen scaffold for skeletal regeneration.” J. Biomed.

Mater. Res. A 2010, 94, 371-379.

135. Zhang, L.; Zhang, Y.; Jiang, B.; Fan, H.S.; Zhang, X.D. “Characterization of an

Insoluble Collagen Sponge and the Potential for Tissue Engineering Scaffold.”

Materials Science Forum 2009, 610, 1378-1381.

136. Lefebvre, F.; Gorecki, S.; Bareille, R.; Amedee, J.; Bordenave, L.; Rabaud, M.

“New artificial connective matrix-like structure made of elastin solubilized peptides

and collagens: elaboration, biochemical and structural properties.” Biomaterials 1992,

13, 28-33.

159

137. Doillon, C.J.; Silver, F.H. “Collagen-based wound dressing: effects of hyaluronic

acid and fibronectin on wound healing.” Biomaterials 1986, 7, 3-8.

138. Ellis, D.L.; Yannas, I.V. “Recent advances in tissue synthesis in vivo by use of

collagen-glycosaminoglycan copolymers.” Biomaterials 1996, 17, 291-299.

139. Chen, G.P.; Ushida, T.: Tateishi, T. “Scaffold Design for Tissue Engineering.”

Macromol. Biosci. 2002, 2, 67-77.

140. O’Brien, F.J.; Harley, B.A.; Yannas, I.V.; Gibson, L. “Influence of Freezing Rate

on Pore Structure in Freeze-Dried Collagen-GAG Scaffolds.” Biomaterials 2004, 25,

1077-1086.

141. Petibois, G.; Gouspillou, G.; Wehbe, K.; Delage, J.P.; Deleris, G. “Analysis of

type I and IV collagens by FT-IR spectroscopy and imaging for a molecular

investigation of skeletal muscle connective tissue.” Anal. Bioanal. Chem. 2006, 386,

1961-1966.

142. Mann, S.; Webb, J.; Williams, R.J.P. “Biomineralization: chemical and

biochemical perspectives.” Weinheim, Federal Republic of Germany; New York, NY,

USA: VCH 1989.

143. Baeuerlein, E. “Biomineralization: from biology to biotechnology and medical

application.” Weinheim; Chichester: Wiley-VCH 2000.

144. Akashi, M.; Watanabe, J. “Novel biomineralization for hydrogels: Electrophoresis

approach accelerates hydroxyapatite formation in hydrogels.” Biomacromolecules

2006, 7, 3008-3011.

145. Akashi, M.; Watanabe, J. “An electrophoretic approach provides tunable

mineralization inside agarose gels.” Cryst. Growth Des. 2008, 8, 478-82.

160

146. Akashi, M.; Watanabe, J. “Dual mineralization of dicalcium phosphate and

hydroxyapatite formation on both sides of porous membrane: Alternating current

induces pH-modulated crystal deposition.” J. Cryst. Growth 2009, 311, 4323-4328.

147. Tasaki, I. “Evidence for phase transition in nerve fibers, cells and synapse,”

Ferroelectrics 1999, 220, 305-316.

148. Trevors, J.T.; Pollack, G.H. “Hypothesis: the origin of life in a hydrogel

environment,” Prog. Biophys. Mol. Biol. 2005, 89, 1-8.

149. Pollack, G.H. “The role of aqueous interfaces in the cell,” Adv. Colloid Interface

Sci. 2003, 103, 173-196.

150. Pollack, G.H. “Is the Cell a Gel- and Why Does It Matter?” Japanese Journal of

Physiology 2001, 51, 649-660.

151. Pollack, G.H. “The cell as a biomaterial.” J. Mat.Sci.: Mat. In Medicine 2002, 13,

811-821.

152. Tasaki, I. “Rapid Structural Changes in Nerve Fibers and Cells Associated with

Their Excitation Processes.” Japanese Journal of Physiology 1999, 49, 125-138.

153. Tasaki, I. “Spread of Discrete Structural Changes in Synthetic Polyanionic Gel: A

Model of Propagation of a Nerve Impulse.” J. Theor. Biol. 2002, 218, 497-505.

154. Tasaki, I. “Repetitive abrupt structural changes in polyanionic gels: A comparison

with analogous processes in nerve fibers.” J. Theor. Biol. 2005, 236, 2-11.

155. Tasaki, I. “A note on the local current associated with the rising phase of a

propagating impulse in nonmyelinated nerve fibers.” Bull. Math. Biol. 2006, 68, 483-

490.

161

156. Tasaki, I. “On the Reversible Abrupt Structural Changes in Nerve Fibers

Underlying Their Excitation and Conduction Processes.” In Phase Transitions in Cell

Biology. Pollack, G.H. and Chin, W.-C., editors. Springer Science + Business Media

B.V. 2008, 1-21.

157. Horkay, F.; Tasaki, I.; Basser, P.J. “Effect of Monovalent-Divalent Cation

Exchange on Swelling of Polyacrylate Hydrogels in Physiological Salt Solutions.”

Biomacromolecules 2001, 2, 195-199.

158. Kriwet, B.; Kissel, T. “Interactions between bioadhesive poly(acrylic acid) and

calcium ions,” Int. J. Pharm. 1996, 127, 135-145.

159. Gao, F.; Reltz, F.B.; Pollack, G.H. “Potential in Anionic Polyelectrolyte

Hydrogels,” J. Appl. Polym. Sci. 2003, 89, 1319-1321.

160. Huang, C.; Soenen, S.J.; Rejman, J.; Lucas, B.; Braeckmans, K.; Demeester, J.;

Smedt, S.C. “Stimuli-responsive electrospun fibers and their applications.”

Chem.Soc.Rev. 2011, 40, 2417-2434.

161. Jin, X.; Hsieh, Y. “pH-responsive swelling behavior of poly (vinyl alcohol)/poly

(acrylic acid) bi-component fibrous hydrogel membranes.” Polymer 2005, 46, 5149-

5160.

162. Xiao, S.; Shen, M.; Ma, H.; Guo, R.; Zhu, M.; Wang, S.; Shi, X. “Fabrication of

Water-Stable Electrospun Polyacrylic Acid-Based Nanofibrous Mats for Removal of

Copper (II) Ions in Aqueous Solution.” J. Appl. Polym. Sci. 2010, 116, 2409-2417.

163. Kim, B.; Park, H.; Lee, S.H.; Sigmund, W.M. “Poly (acrylic acid) nanofibers by

electrospinning,” Mater. Lett. 2005, 59, 829-832.

162

164. Li, L.; Hsieh, Y. “Ultra-fine polyelectrolyte fibers from electrospinning of poly

(acrylic acid).” Polymer 2005, 46, 5133-5139.

165. Li, L.; Hsieh, Y. “Ultra-fine polyelectrolyte hydrogel fibers from poly (acrylic

acid) / poly (vinyl alcohol).” Nanotechnology 2005, 16, 2852-2860.

166. Safronov, A.P.; Sminova, Y. A.; Pollack, G.H.; Blyakhman, F.A. “Enthalpy of

Swelling of Potassium Polyacrylate and Polymethacrylate Hydrogels. Evaluation of

Excluded-Volume Interaction.” Macromol. Chem. Phys. 2004, 205, 1431-1438.

167. Zubarev, A.Y.; Blyakhman, F.A.; Pollack, G.H.; Gusev, P.; Safronov, A.P. “Self-

Similar Wave of Swelling/Collapse Phase Transition along Polyelectrolyte Gel.”

Macromol. Theory Simul. 2004, 13, 697-701.

168. Shklyar, T.F.; Safronov, A.P.; Klyuzhin, I.S.; Pollack, G.H.; Blyakhman, F.A. “A

Correlation between Mechanical and Electrical Properties of the Synthetic Hydrogel

Chose as an Experimental Model of Cytoskeleton.” Biophysics 2008, 53, 544-549.

169. Nge, T.T.; Yamaguchi, M.; Hori, N.; Takemura, A.; Ono, H. “Synthesis and

Characterization of chitosan/poly (acrylic acid) polyelectrolyte complex.” J. Appl.

Polym. Sci. 2002, 83, 1025-1035.

170. Johnston, M.B.; Bernard, A.J.; Flaschka, H.A. “EDTA and complex formation.” J.

Chem. Educ. 1958, 35, 601-606.

171. Ikegami, A.; Imai, N. “Precipitation of Polyelectrolytes by Salts,” J. Polym. Sci.

1962, 56, 133-152.

172. Pollack, G.H. “Cell, Gels and the Engines of Life.” Ebner & Sons 2001.

163