NATURE INSPIRED COMPOSITE NANOFIBERS

TEO WEE EONG

B. ENG.(HONS.) NUS

A THESIS SUBMITTED FOR THE DEGREE OF MASTER

OF ENGINEERING

DEPARTMENT OF MECHANICAL ENGINEERING

NATIONAL UNIVERSITY OF SINGAPORE

2009

Acknowledgement Completing this Masters Program is not easy while holding a full time job. Work

commitment often brings my mind away from thinking about my Masters project. I am

fortunate to have an understanding boss and supervisor, Prof Seeram Ramakrishna, who I

would like to express my thanks for his encouragement and guidance. Being an engineer,

it is certainly my wish to see my research work being translated into a useful product. For

this, I have to thank my co-supervisor, Prof Casey Chan for seeing the potential of my

project for development into biomedical products. Hopefully, in a few years time, the

work that begins from this Masters Project is used as a successful clinical product.

I would like to thank my beloved wife, Karen, for her understanding when I have to

spend time studying or reading up research materials during our courtship days. To my

parents who have shared the ups and downs of my research work, thank you for lending

your ears although it must have been really tough for you to understand my work.

Finally, I have to thank my friends and colleagues in the lab. This project will have been

much tougher and will probably take longer to complete without you sharing knowledge

and experience.

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Table of Contents Acknowledgement ............................................................................................................. 2

Table of Contents ............................................................................................................... i

Summary ........................................................................................................................... iii

List of Tables ..................................................................................................................... v

List of Figures ................................................................................................................... vi

List of Publications ........................................................................................................ viii

Chapter 1: Introduction ................................................................................................... 1

1.1 Composite Nanofibers .............................................................................................. 1 1.2 Nanofiber fabrication techniques .............................................................................. 2 1.3 Biomedical applications ............................................................................................ 3 1.4 Motivation ................................................................................................................. 4 1.5 Objective ................................................................................................................... 4

Chapter 2: Literature Review .......................................................................................... 6

2.1 Bone Structure and Organization .............................................................................. 6 2.2 Current Technology .................................................................................................. 9 2.3 Electrospinning ....................................................................................................... 11

Chapter 3: Dynamic fluid method to reorganize nanofibers ...................................... 13

3.1 Introduction ............................................................................................................. 13 3.2 Experimental Procedures ........................................................................................ 13 3.3 Results and Discussions .......................................................................................... 16

Remarks .................................................................................................................... 19 3.4 Biomedical applications .......................................................................................... 23 3.5 Conclusion .............................................................................................................. 25

Chapter 4: Fabrication of 3D Assemblies using fluid properties ............................... 26

4.1 Introduction ............................................................................................................. 26 4.2 Experimental Procedures ........................................................................................ 26 4.3 Results and Discussions .......................................................................................... 28

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4.3.1 Physical structure observation ......................................................................... 28 4.3.2 Effect of hydrophobicity .................................................................................. 31 4.3.3 Effect of cooling rate ....................................................................................... 35

4.4 Application in Tissue Engineering .......................................................................... 37 4.5 Conclusion .............................................................................................................. 38

Chapter 5: Mineralization of three-dimensional matrix ............................................. 40

5.1 Introduction ............................................................................................................. 40 5.2 Experimental Procedures ........................................................................................ 40 5.3 Results and Discussions .......................................................................................... 43 5.4 Conclusion .............................................................................................................. 56

Chapter 6: Research Summary and Future Recommendations ................................. 57

Reference ......................................................................................................................... 60

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Summary

Natural extracellular matrices are usually constructed from collagen nanofibers and this

made nanofibers an attractive candidate for use in biomedical applications in particular

regenerative scaffold. Since electrospun collagen nanofibers are weak and degrade

rapidly in vivo, synthetic polymers are often incorporated to form composite nanofibers.

Extensive studies carried out on electrospun nanofibrous two-dimensional mesh have

shown that the architecture provides a favorable environment for cells to thrive on.

However, the ability to construct other electrospun architectures has been challenging. A

modified electrospinning setup using water as a working substrate has been demonstrated

here to be capable of fabricating composite nanofibrous yarn and three-dimensional (3D)

nanofibrous architectures. Nanofiber and yarn diameter has been shown to be affected by

the solution feed-rate and concentration of the solution. Higher solution feed-rate and

concentration gave rise to fiber and yarn of larger diameter. The microstructure of the 3D

nanofibrous scaffold is affected by the hydrophobicity of the polymer and the drying and

freezing condition of the scaffold. Using a modified alternate dipping method, calcium

phosphate minerals can be deposited throughout the 3D nanofibrous scaffold. While

conventional static alternate dipping mineralization method yields minerals mainly on the

surface of the 3D scaffold and uneven distribution of the minerals on the nanofibers, a

flow mineralization method is able to achieve mineralization both at the surface and the

core of the 3D scaffold with improved mineral distribution on the nanofibers. Despite

static mineralized scaffold having greater mineral contents, its compressive strength and

modulus is much lower than flow mineralized scaffold. Therefore, the mechanical

strength of the mineralized nanofibrous scaffold is significantly affected by how the

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minerals are deposited on the nanofibers. The ability to construct nanofibrous yarn, 3D

nanofibrous scaffold and 3D mineralized nanofibrous scaffold has opened up new

opportunities for the construction of biomimetic regenerative scaffold. The ability to

incorporate biological materials into these architectures that mimics the physical

characteristic of natural extracellular matrix gave it the potential to replace autologous

implants. The next step of development will be to test these constructs in in vivo studies.

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List of Tables Table 3.1 Polymer solution and average fiber diameter. Table 3.2 Fiber diameter with respect to feed rate for PVDF-co-HFP.

vi

List of Figures

Figure 1.1 Hierarchical organization of bone from the lowest level on the left to the bone macrostructure. Figure 2.1 Schematic of collagen fibril and crystal growth. The left column shows the arrangement of the collagen triple helix molecules in a fibril in 2-dimension and the right column in 3-dimension. [A] Staggered arrangement of the collagen triple helix molecules. The 3-dimensional distribution shows the channels resulting from the orderly arrangement of the fibrils. [B] Nucleation of the minerals in the gap between the collagen molecules. [C] Growth of the minerals between the gaps and along the channel results in a parallel array of coplanar crystals. Figure 3.1 Setup used to create a flowing water system for the manipulation of deposited nanofibers. Figure 3.2 Electrospun PVDF-co-HFP fibers deposited on a aluminum foil. Figure 3.3 SEM images of the collected yarn. a) Without going through the drawing process in the air. b) After going through the drawing process in the air. Figure 3.4 Graph of average PVDF-co-HFP yarn diameter against feed-rate. The vertical line for each point depicts the scatter in the yarn diameter for each feed rate. Figure 3.5 Nanofibrous yarn used as intra-luminal guidance channel. [A] Overview of nerve guidance channel consisting of a nanofibrous conduit and nanofibrous yarn in the lumen. [B] SEM image of conduit extracted from rat sciatic nerve after 3 months. [Picture courtesy of H. S. Koh] Figure 4.1 [a] Polycaprolactone (PCL) 3D mesh dried in room condition with no visible pores. [b] PCL 3D mesh pre-frozen at -86 oC and freeze-dried in a cylinder with visible pores [c] PCL/col 3D mesh pre-frozen at -86 oC and freeze-dried in a hemispherical container showing visible pores [d] PCL/collagen 3D mesh pre-frozen at -86 oC and freeze-dried in a cylinder with visible pores. Figure 4.2 [a] PCL mesh dried under room condition showing yarn stacked closely on top of one another. [b] Freeze-dried PCL mesh pre-frozen at -86 oC showing distinct and isolated yarns made out of aligned nanofibres. [c] PCL/collagen mesh pre-frozen at -86 oC before freeze-drying showing ridge-like structures. [d] Disordered nanofibres that formed the ridges. All samples were packed in 15 mm diameter cylinder. Figure 4.3 Schematic of the ice crystals pushing the entangled yarn to form disordered fibres and ridges on the PCL/col scaffold. [a] Stage 1. Nanofibrous yarns were suspended in the water. [b] Stage 2. As the water cools, ice nucleates on the wall of the cylinder and

vii

on the nanofibres. [c] Stage 3. Growing ice front from the cylinder wall pushes against the nanofibres resulting in [d] ridges made out of random nanofibres. Figure 4.4 Cross section of the PCL/col scaffold cooled at -86 oC showing inhomogeneous pores and a boundary layer between the displaced nanofibers on the surface and compacted fibers at the inner core. Figure 4.5 Freeze-dried PCL/col [a] prefrozen at -86 oC showing “spikes”, [b] mesh pre-frozen in liquid nitrogen has a relatively smoother surface. [c] Higher magnification showing microstructure of scaffold pre-frozen at -86 oC where the “spikes” were ridges formed by disordered nanofibres and [d] mesh pre-frozen in liquid nitrogen with distinct yarns. [e] Cross-section of the mesh pre-frozen at -86 oC showing the ridges are only found on the surface while [f] the mesh pre-frozen in liquid nitrogen did not exhibit any apparent differences in the microstructure from the surface to the interior. Figure 4.6 Schematic of the ice nucleation and growth on PCL/col mesh cooled at -196 oC. [a] Stage 1, nanofibrous mesh is suspended in water. [b] Numerous and rapid ice nucleation on the surface of the nanofibers while ice nucleation and growth commence from the cylinder wall. [c] Complete freezing of mesh before the ice growing from the cylinder wall reaches it. Figure 5.1 Setup for mineralization of the 3D scaffold.

Figure 5.2 Spectra of PLLA and PLLA/col blended 3D nanofiber scaffold.

Figure 5.3 SEM images of mineralized [A] PLLA sample and [B] PLLA/col sample.

Figure 5.4 Distribution of minerals on across the cross-section of static mineralized scaffold. Figure 5.5 Distribution of the minerals in the cross-section of the scaffolds using flow mineralization. Figure 5.6 XRD showing the presence of amorphous calcium phosphate crystals in the mineralized scaffold. Figure 5.7 Ash remains of mineralized scaffold after sintering at 500 oC. [A] Static mineralized scaffold showing a hollow core. [B] Flow mineralized scaffold showing a solid core. Figure 5.8 Mechanical properties of scaffold mineralized under different condition [A] Compressive strength. [B] Compressive Modulus Figure 5.9 Comparison of mineral nanoparticles distribution in [A] mineralized nanofibrous scaffold and [B]cancellous bone.

viii

List of Publications

W.E Teo, S. Liao, C.K. Chan, S. Ramakrishna (2008) Remodeling of Three-dimensional

Hierarchically Organized Nanofibrous Assemblies. Current Nanoscience vol. 4 pg. 361-

369.

Wee-Eong Teo, Renuga Gopal, Ramakrishnan Ramaseshan Kazutoshi Fujihara, Seeram

Ramakrishna (2007) A dynamic liquid support system for continuous electrospun yarn

fabrication. Polymer vol. 48 pg. 3400-3405.

Teo WE, He W, Ramakrishna S (2006) Electrospun scaffold tailored for tissue-specific

extracellular matrix. Biotechnology Journal vol. 1 pg. 918-929. (Top download in

Biotechnology Journal in September 2006)

Wee-Eong Teo, Seeram Ramakrishna (2009) Electrospun nanofibers as a platform for

multifunctional, hierarchically organized nanocomposite Compos. Sci. Technol. Vol. 69

pg. 1804-1817.

Wee Eong Teo, Kazutoshi Fujihara, Casey Kwan-Ho Chan, Seeram Ramakrishna. Fiber

Structures and Process for their preparation. WO 2008/036051. 27 March 2008.

Wee Eong Teo, Kazutoshi Fujihara, Seeram Ramakrishna. Method & Apparatus for

producing Fiber Yarn. WO 2007/013858 A1. 01 February 2007.

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Chapter 1: Introduction

1.1 Composite Nanofibers

Nature has been the inspiration for many materials design and architectures and the

strongest materials are often made out of composites. Closer examination of these

composites reveals an intricate hierarchical organization from the nano-scale level

which gives it superior properties to meet the functional requirement. Hierarchical

organization has been shown to play an important role in maximizing the

performance of the structure using abundant elements. The organic matrix which all

natural materials are made of is mainly carbon. Collagen, cellulose and calcium

phosphate have low strength and stiffness individually. However, through

organization of these compounds hierarchically as a composite, the resultant materials

such as wood, bone and tendons are much higher strength. A closer examination of

these structures showed that nanofibers play an important role in giving them the

superior mechanical properties (Fratzl and Weinkamer 2007). In the human body,

many organs are constructed from nanofibers organized in a hierarchical structure.

Figure 1.1 shows the hierarchical organization of bone. Studies have shown that cells

are influenced by nano-textures (Wan et al. 2005). Thus construction of tissue

regenerative scaffold may require special consideration of the hierarchical

organization of the organ from the nano-level.

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Figure 1.1 Hierarchical organization of bone from the lowest level on the left to the bone macrostructure.

1.2 Nanofiber fabrication techniques

For the last few decades, man has been inspired by the structure of nature’s design

and this has been used in macro-level designs. It is only in recent years where

advances in nanotechnology have allowed us to mimic nature’s nano-structure design

(Meyers et al. 2008). Many different techniques for fabrication of nanofibers, both at

industry level and laboratory scale level, have been developed. At the laboratory

scale, direct drawing from liquid polymer using AFM tip allows controlled and

precise placement of the nanofiber strand. However, this method is laborious and

there is a limitation on the length of the fiber that can be drawn (Harfenist et al.

2004). Molecular self-assembly is another method often used to fabricate biological

nanofibrous material. The nanofibers produced has diameter in the tens of nanometer

and they are often assembled from peptides (Zhang 2002). However, the length of the

nanofibers is short and the resultant scaffold is often a disordered mesh of nanofibers.

At the industrial level, development of melt-blowing has been shown to be capable of

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fabricating long continuous nanofibers at high speed (Suzuki and Tanizawa 2009).

The last decade has seen the adoption of a process known as electrospinning by

researchers to construct various nanofibrous assemblies and composite nanofibrous

structures. This is due to the ease of fabricating nanofibers from viscous solution of

polymers, blends and mixtures that allows researchers to construct hierarchically

organized structures (Teo and Ramakrishna In press). This process has been used at

the commercial level to produce nanofibers by various companies such as Donaldson

Inc. and eSpin Inc.

1.3 Biomedical applications

Since natural extracellular matrix is consist mainly of nanofibers, researchers are

using electrospinning to construct biomimicking tissue scaffolds (Teo et al. 2006).

Bone is perhaps one of the most widely studies composite structure found in human

body. Although the structure of the bone has been studied for decades, its precise

hierarchical structure has proven to be extremely difficult to replicate. Today, bone

repairs are still commonly carried out using allograft and autograft. However, the risk

of disease transmission and donor site morbidity respectively, has led researchers to

look for synthetic substitutes. Synthetic grafts currently represent only about 10% of

the bone graft market and they are made out of ceramics, composites (Bucholz 2002)

or polymers. To replace autograft in bone repair treatments, researchers are trying to

mimic natural bone in terms of its (Burg et al. 2000) physical structures, mechanical

properties, pore-dimensions and chemical cues. While the shape of specific parts of

the bone has been constructed for load bearing applications, the ability to replicate the

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micro and perhaps nano-structure of the bone are important criteria for the bone graft

to be fully integrated into the body.

1.4 Motivation

In construction of bone regenerative scaffold, some researchers have developed

scaffolds that satisfy some of the micro-structural characteristics. Others are able to

create some of its nano-structural organization. However, the organization in these

synthetic grafts is far from the hierarchical organization of natural bone. Although

many researches on biomedical application of electrospun nanofibers have been

carried out, these are mainly focused on flat nanofibrous mesh. New methodology

and modification of the conventional electrospinning process needs to be developed

to mimic the higher order of nanofibrous organization found in many organs such as

bone. It is therefore necessary to develop methods of organizing the nanofibers such

that other structures can be fabricated. Of particular importance will be the ability to

construct a three-dimensional block scaffold that is able to fill in a bone defect. Since

the targeted application is in bone regeneration, minerals found in native bone should

be added to the scaffold so as to mimic the physical structure and the biochemistry of

bone.

1.5 Objective

The objective of this project is to develop a method of fabricating composite

nanofibrous structures for biomedical applications. Particular attention is paid to the

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construction of bone regenerative scaffold. To achieve this, a novel dynamic fluid

flow method has been developed to organize the electrospun nanofibers (Chapter 3).

This method is further modified to demonstrate its ability to form a 3D scaffold with

distinct microstructures made from nanofibers (Chapter 4). Finally, for application as

bone regenerative scaffold, the nanofibrous scaffold was incorporated with calcium

phosphate nanoparticles using a customized and newly developed setup (Chapter 5).

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Chapter 2: Literature Review

2.1 Bone Structure and Organization

It is estimated that 77% of the mineral is found outside the fibril and 23% of the

mineral is found inside the fibril (Sasaki et al. 2002). This is in agreement with the

estimation by Bonar et al who suggested that at most 35% of the mineral in mature,

fully mineralized bovine tibia to be inside the bone collagen fibrils (Bonar et al.

1985). Collagen Type I is the principal component of the organic matrix of the bone

accounting for approximately 30% of the dry mass of bone matrix while non-

collagenous proteins account for approximately 5% of the dry bone matrix. Reducing

the amount of collagen in the bone will significantly increase the modulus of dry

bone. However, the toughness and the strength of the bone will be reduced (Currey

2003;Fantner et al. 2004). A study by Martin and Ishida has shown that the

orientation of the fiber in cortical bone is the best predictor of tensile strength and is

at least as important as density, porosity and mineralization in determining tensile

strength (Martin and Ishida 1989).

Hodge and Petruska described in details the arrangement of the collagen molecules

that give the fibrils a 67 nm periodicity. They deduced that “holes” of about 40 nm

must be present between the collagen molecules for the formation of native type

periodicity in the fibrils and suggested that this might be sites of mineralization

(Hodge and Petruska 1963). From the distribution of minerals in calcified tendon

using high-voltage electron microscope, Landis et al suggests nucleation of small

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minerals may occur within the holes of the fibrils. Crystals may also form in the gap

between the layers of collagen molecules. Growth of the mineral crystals would

proceed beyond the holes and into adjacent holes and gaps between the molecules

layers (Landis et al. 1993). Traub et al have also observed that crystals in mineralized

turkey tendon extend over several fibrils and are arranged in several parallel layers in

a coplanar distribution (Traub et al. 1989). In a later study, observation of embryonic

chick bone using high voltage electron microscopic tomography revealed that the

mineral nanoparticles are staggered with a periodicity similar to that of collagen

molecules at 67 nm (Landis et al. 1996). Observations using electron microscope on

the dispersion and orientation of apatite nanoparticles on the surface of the collagen

fibrils by Rubin et al (Rubin et al. 2003)) and Su et al (Su et al. 2003) showed that the

apatite crystals are aligned with their length (c-axis) parallel to the longitudinal axis

of the collagen fibrils. An illustration of the collagen molecules and the organization

of the minerals within the fibrils are shown in Figure 2.1.

Figure 2.1 Schematic of collagen fibril and crystal growth. The left column shows the arrangement of the collagen triple helix molecules in a fibril in 2-dimension and the right column in 3-dimension. [A] Staggered arrangement of the collagen triple

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helix molecules. The 3-dimensional distribution shows the channels resulting from the orderly arrangement of the fibrils. [B] Nucleation of the minerals in the gap between the collagen molecules. [C] Growth of the minerals between the gaps and along the channel results in a parallel array of coplanar crystals.

In cancellous bone, the fibrils are aligned in a general direction to form bundles of

collagen fibers (Fantner et al. 2006). Using a scanning small angle X-ray scattering

analysis, Rinnerthaler et al deduced that the minerals and collagen fibers are oriented

to the direction of the trabeculae (Rinnerthaler et al. 1999). However, AFM study by

Hassenkam showed that the fibrils arrangement in the trabecula differ in different

parts. On one end of the trabecula, the collagen fibrils did not seem to have a

preferred orientation, in the middle of the trabecula are arranged in a crisscross

pattern and at the other end of the trabecula, the fibrils formed huge bundles of

collagen aligned in the same direction (Hassenkam et al. 2004). It is likely that some

of the collagen fibrils in the crisscross pattern are cross-linked (Landis et al. 1996)

giving the structure greater strength.

On top of the collagen fibrils and mineral particles, there is a coating of non-fibrillar

organic material. The distribution of the non-fibrillar organic material and the extent

of mineralization of the fibrils are non-uniform. When a fracture occurs, the organic

matrix would form filaments that span the micro-crack thereby resisting the growth

and propagation of the micro-crack (Fantner et al. 2006).

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2.2 Current Technology

In the development of synthetic bone graft, most efforts are directed towards creating

macro and microporous scaffolds. Pore size and porosity of bone graft are critical for

the successful regeneration of bone. It is generally accepted that a pore size of about

100 µm is recommended for new bone formation (Karageorgiou and Kaplan 2005). In

polymer and polymer-mineral mixture, common techniques of creating macroporous

scaffold include using a combination of gas foaming (Nam et al. 2000;Mathieu et al.

2006) and particulate leaching (Chen et al. 2000;Tu et al. 2003) with freeze-drying or

phase separation (Zhang and Ma 1999). To fabricate macroporous ceramic scaffold

with structure identical to cancellous bone, Tancred et al used a multi-stage process

involving the formation of a negative wax mould following by infiltration of the wax

mould using a ceramic slip, removal of the wax and ends with firing to produce a

positive replica of the cancellous bone (Tancred et al. 1998).

The macroporous scaffold mimics the highest level of the bone hierarchical structure.

At the lowest level, bone consists of collagen molecules and hydroxyapatite (HA)

nanocrystals. Zhang et al aims to create a bone scaffold through the assembly of its

most basic structure, nanofibrils and nanoparticles. By controlling the pH of a

solution containing collagen molecules, phosphate ions and calcium ions, the collagen

molecules can self-assemble to form a fibril surrounded by a layer of HA

nanocrystals with its c-axis aligned with the longitudinal axes if the collagen fibrils

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(Zhang et al. 2003). However, a three-dimensional assembly with sufficient volume

for bone repair has not been reported using this technique till date.

In recent years, the benefits of using nanocomposites in bone graft have been

highlighted by several researchers (Stylios et al. 2007;Chan et al. 2006;Murugan and

Ramakrishna 2005). Webster et al has shown that osteoblast adhere better on

nanophase ceramics than microphase ceramics (Webster et al. 1999). Kikuchi et al

demonstrated in vivo that the integration of HA/collagen nanocomposites with bone

remodeling is similar to the osteointegration of autografted bone (Kikuchi et al.

2001). The superior performance of nanocomposite as compared to monolithic and

conventional composite may be attributed to its resemblance to natural bone which

itself is a nanocomposite consisting mainly of HA nanocrystallites and collagen

nanofibers. Similarly, new development of engineered scaffolds for tissue

replacement is moving towards replicating the natural extracellular matrix (ECM) of

the organ or tissue it is replacing. To replicate the hierarchical structure of bone, Wei

et al used sugar sphere and phase separation to create nanofibrous scaffolds with

micro-pores (Wei and Ma 2006). Chen et al used reverse solid freeform and phase

separation to fabricate nanofibrous scaffold with controlled and complex geometries

with micro and macro-pores. Comparing the degree of differentiation of preosteoblast

between nanofibrous and solid-walled scaffolds, preosteoblast cultured on

nanofibrous scaffolds exhibited greater differentiation with evidence of greater

mineral production, higher expression of osteocalcin and bone sialprotein mRNAs

(Chen et al. 2006). Therefore, it may be advantageous that scaffolds be made out of

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nanofibers with organization and properties similar to natural extracellular matrix.

Although phase separation is possible to fabricate nanofibers, the fibers are generally

short and the organization of the fibers is limited. At the beginning of this century,

many researchers are using a century old fiber processing method known as

electrospinning to fabricate nanofibrous scaffold.

2.3 Electrospinning

The ability to fabricate a variety of structures (Teo and Ramakrishna 2006) with little

restriction in the type of polymer and mixture that can be used (Zhong et al.

2007;Zhong et al. 2005;He et al. 2005;He et al. 2006) has seen the growing popularity

of using electrospinning in the fabrication of biomimetic scaffold. Tailoring the

structure and composition of the scaffold using this process to match the respective

tissue/organ ECM it is replacing (Teo et al. 2006) has contributed to the

understanding of the interaction between cells and nanofibers (He et al. 2006;Zhong

et al. 2006).

Bone grafts made out of electrospun fibers have been developed from a variety of

materials and compositions. Study by Wutticharoenmongkol et al showed that pre-

osteoblast expressed higher amount of osteocalcin gene and protein and secreted

more minerals when cultured on polycaprolactone/nanohydroxyapatite composite

than on pure polycaprolactone (PCL) nanofibers although the amount of

nanohydroxyapatite was just 1% w/v of the blended solution (Wutticharoenmongkol

et al. 2007). Sefcik et al demonstrated more than one-fold up-regulation of osteogenic

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genes after 21 days when human adipose stem cells were cultured on collagen

electrospun fibers compared with 2D collagen coatings (Sefcik et al. 2008).

Current electrospinning process is still mainly restricted to the fabrication of two-

dimensional mesh. Although it is possible to fabricate tubular scaffold or to stack

numerous two-dimensional mesh to form a three-dimensional structure, it is still a

challenge to fabricate a truly three-dimensional architecture. Thus there is a need to

develop new methodologies and understanding of the electrospinning process

especially in the fabrication of three-dimensional nanofibrous architectures.

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Chapter 3: Dynamic fluid method to reorganize nanofibers

3.1 Introduction

To fabricate a nanofiber composite for bone repair, it is necessary to develop a

method to construct 3D structure. The minute size of nanofibers means that they are

not mechanically strong enough to withstand conventional mechanical manipulation

to form higher order assemblies. Thus, a different manipulation technique is required

to construct a 3D structure. Unlike a rigid solid substrate, a micro-component on a

liquid substrate can be easily maneuvered and this has been used in the self-assembly

of micro-structures (Syms et al. 2003). Fluid properties such as fluid interfaces and

hydrodynamic interaction have also been used for the assembly of particles

(Grzybowski and Whitesides 2002). Liquid properties such as surface tension,

viscosity, interfaces and hydrodynamic interactions, may be used to control the

positioning of electrospun fibers. In this chapter, the hypothesis is a dynamic liquid

substrate can be used as a working platform and tool for the manipulation of

nanofibers. The effect of feed-rate and solution concentration on the electrospun

fibers was examined.

3.2 Experimental Procedures

Poly(vinylidene flouride-co-hexafluoropropylene) (PVDF-co-HFP) (Aldrich, Mw

455,000), polycaprolactone (PCL) (Aldrich, Mn ca. 80,000), collagen type I (col)

(Koken) and poly-L-lactide (PLLA) (Polyscience Inc, Mw 300,000) was used. PVDF

14

pellets were dissolved in a mixture of 40% dimethylacetamide and 60% acetone,

heated to 60 oC, to give a concentration of 0.12 g/ml. To investigate the effect of

solution concentration on fiber diameter, two other concentrations of 0.08 g/ml and

0.1 g/ml were prepared. PCL and PCL/col (50:50 w/w), PLLA/col (70:30 w/w) and

PLLA/col (50:50 w/w) were dissolved in 1,1,1,3,3,3-hexafluo-2-propanol (HFP) to

give a solution of 0.08 g/ml each. PLLA/col (50:50 w/w) were also dissolved in

1,1,1,3,3,3-hexafluo-2-propanol (HFP) at two other concentrations of 0.05 g/ml and

0.03 g/ml. All chemicals were used as received without further modification.

Figure 3.1 shows the experimental set-up to create a fluid flow system. Using a basin

with a hole of diameter 5 mm in the center, water was allowed to flow through it

thereby forming a water vortex. A pump was used to re-circulate the water from the

tank back to the basin and the water level in the basin was maintained at a constant

height. A wire was inserted into the basin to remove any residual charges on the water

surface.

A Gamma High Voltage Research HV power supply was used to generate the high

voltage for electrospinning. The spinneret used was a B-D 27G1/2 needle which was

ground to give a flat tip. A kd Scientific syringe pump was used to provide a constant

feed rate. To fabricate PVDF-co-HFP fibers, a voltage of 12 kV was applied to the

spinneret with the distance from the tip of the spinneret to the surface of the water in

the basin maintained at 12 cm. To study the effect of feed-rate on nanofiber assembly,

the feed-rates for PVDF-co-HFP were set as 1 ml/hr, 2 ml/hr, 5 ml/hr, 10 ml/hr and

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15 ml/hr for a concentration of 0.12 g/ml polymer solution. For all other PVDF-co-

HFP concentration, the feed rate was maintained at 10 ml/hr. To fabricate PCL and

PCL/col (50:50 w/w) nanofibers, a voltage of 12 kV was applied, the feed-rate was

set as 1 ml/hr and the distance from the tip of the spinneret to the surface of the water

in the basin maintained at 14 cm. To fabricate PLLA/col (70:30 w/w) and PLLA/col

(50:50 w/w), a voltage of 15 kV was applied, the feed-rate was set as 1 ml/hr and the

distance from the tip of the spinneret to the surface of the water in the basin

maintained at 14 cm.

The assembled nanofibers, which was carried by the falling water from the top basin,

was manually transferred to the rotating mandrel to initiate take-up. The speed of the

rotating mandrel was adjusted such that continuous collection of the assembled

nanofibers can be maintained. If the drawing of the yarn was not continuous, a

window (hole dimension, 4 cm by 2 cm) was passed through the falling water to

collect the assembled fibers. A scanning electron microscope (SEM), Quanta FEG

200, FEI, Netherlands, was used to observe the nanofiber assembly. Samples were

first coated with gold using a JEOL JFC-1600 Auto Fine Coater before viewing under

SEM. Diameters of the fibers were measured from the SEM images using Image J

software (National Institute of Health, USA).

16

Figure 3.1 Setup used to create a flowing water system for the manipulation of deposited nanofibers.

3.3 Results and Discussions

In this chapter, the hypothesis is that fluid can be used as a working substrate to

manipulate nanofibers. Figure 3.2 shows the typical nonwoven electrospun mesh

fabricated by collecting the fibers on a solid substrate. Given the low mechanical

strength of individual nanofiber, it is not possible to remove the nonwoven mesh from

a solid substrate and re-model it to a different form. Since the disadvantage of a solid

substrate is its rigid nature, in contrast, a fluid substrate carrying the nanofibers can be

easily manipulated. Thus in the current setup, a flowing water system was created in

the form of a vortex funneling down a sink hole at the base of basin. Since the water

17

surface converges as it flows through the hole, base on the hypothesis, the nanofibers

carried on the water surface would converge and assembled into a continuous yarn.

Figure 3.2 Electrospun PVDF-co-HFP fibers deposited on a aluminum foil.

The electrospinning process above the water continuously deposits nanofibers on the

surface of the water close to the vortex. When there was a build-up of nanofibers on

the water surface away from the vortex, the spinneret position was adjusted such that

the nanofibers were deposited on or near to the vortex such that the nanofibers were

drawn through it. Strands of fiber yarn can be observed in the water tank below the

basin. Figure 3.3a shows the SEM image of a single strand of nanofiber yarn

collected from the water tank. Thus, this shows that by creating an appropriate water

flow, deposited nanofibers can be manipulated to form other assemblies. Without any

disruption of the deposition of nanofibers on the water surface, it should be possible

to collect continuous strand of yarn. However, it was found that certain conditions

must be satisfied such that the formation and collection of nanofibrous yarn can be

18

continuous. In this study, factors affecting the continuous collection of yarn include

polymer concentration and feed-rate.

A rotating mandrel was used to collect the yarn when the drawing process was

continuous. Comparing the physical characteristic of the yarn collected in the water

tank and the yarn collected on the mandrel as shown in Figure 3.3, it is apparent that

the yarn collected in the water tank was made out of aligned but wavy fibers.

However, the yarn collected by the mandrel was made out of highly aligned and

straight fibers. The straight fibers collected on the mandrel is probably due to the

mechanical drawing of the fibers and the surface tension exerted on the fibers as the

yarn was pulled through the air and collected on the mandrel. Observation of the

SEM images revealed the presence of ‘kinks’ on the yarn where the fiber was bent

backwards. These ‘kinks’ were probably the result of the elongation and

consolidation of the nonwoven mesh initially deposited on the water surface as it

passess through the hole.

Figure 3.3 SEM images of the collected yarn. a) Without going through the drawing process in the air. b) After going through the drawing process in the air.

19

Research on the effect of polymer solution concentration on fiber diameter for

electrospun fibers deposited on solid collector has shown that reduction in polymer

solution concentration would result in reduced fiber diameter (Tan et al. 2005;Mit-

uppatham et al. 2004). Similarly, by reducing polymer concentration, the diameter of

electrospun fibers collected in water also decreased. For a feed-rate of 10 ml/hr,

PVDF-co-HFP solution at the lowest concentration of 0.08 g/ml, beaded fibers were

formed and continuous collection of yarn was not possible. At a higher concentration

of 0.1 g/ml, fewer beads were observed and the collection of yarn was continuous.

The collection of yarn was also continuous for higher polymer concentration of 0.12

g/ml. Similarly, for PLLA/col (50:50 w/w), a higher concentration gives a higher

fiber diameter. Table 3.1 gives a summary polymer solutions and their corresponding

fiber diameter. Parameters favoring the formation of beaded fibers have been covered

various researchers (Fong et al. 1999;Shenoy et al. 2005;Yu et al. 2006). In brief,

beads are formed when the surface tension of the solution is greater than the ability to

stretch the solution without the solution breaking out into droplets. Thus, with a

higher concentration which corresponds to a greater viscosity, the solution can be

stretched further resulting in smoother fibers. Presence of beads along the fiber is

likely to reduce the mechanical strength of the yarn due to stress concentration thus

continuous yarn collection was not possible.

Table 3.1 Polymer solution and average fiber diameter. Polymer Concentration

(g/ml)

Average fiber

diameter(nm)

Remarks

20

PVDF-co-HFP 0.08 463 + 157 Beaded fibers

PVDF-co-HFP 0.1 723 + 226 Beaded fibers

PVDF-co-HFP 0.12 1224 + 393 Smooth fibers

PLLA/col (50:50 w/w) 0.03 351 + 71 Smooth fibers

PLLA/col (50:50 w/w) 0.05 520 + 125 Smooth fibers

PLLA/col (50:50 w/w) 0.08 690 + 154 Smooth fibers

PLLA/col (70:30 w/w) 0.08 790 + 157 Smooth fibers

PCL 0.08 511 + 151 Smooth fibers

PCL/col (50:50 w/w) 0.08 492 + 137 Smooth fibers

The feed-rate of the polymer solution was found to have a significant effect on the

diameter of the fibers, diameter of the yarn and the take-up speed of the yarn. Table

3.2 shows that with increasing feed-rate, the corresponding fiber diameter also

increases. Generally, with an increased volume of solution dispensed while

maintaining all other parameters, the same amount of solution stretching will result in

an increased fiber diameter. The variance in the fiber diameter also increases with

increased feed-rate. This increment could be due to the differences in the coagulation

rate of the fibers in contact with the water upon deposition. Due to the hydrophobicity

of PVDF-co-HFP, later fibers may be deposited on the earlier fibers collected on the

water surface. Earlier fibers in contact with the water will instantaneously coagulate,

as water is a non-solvent. Later fibers deposited on the earlier fibers will not

immediately come into contact with water and thus it has more time for the solvent to

evaporate resulting in a smaller fiber diameter. Theron et al (2004) has shown that

21

increasing feed-rate will decrease the fiber deposition area (Theron et al. 2004). Thus

at a higher feed-rate, more fibers will be stacked on top of one another thus the

differences in solidification rate will result in higher variance in fiber diameter.

Between feed-rate of 2 ml/hr and 5 ml/hr, there was a significant increase in the fiber

diameter. Formation of nanofibers using electrospinning has been attributed to

secondary bending instabilities due to its high charge density which led to increased

stretching of the fiber (Reneker et al. 2000;Mitchell and Sanders 2006). With a higher

feed-rate, the charge density on the electrospinning jet will be reduced. At a lower

feed-rate of 2 ml/hr, the charge density may be sufficiently large for secondary

bending instability to occur. However, at a feed-rate of 5 ml/hr, the reduced charge

density on the electrospinning jet may be insufficient for secondary bending

instability to take place thus giving a significantly larger fiber diameter.

Table 3.2 Fiber diameter with respect to feed rate for PVDF-co-HFP. Feed rate (ml/hr) Average fiber diameter (nm)

1 740 + 150

2 770 + 154

5 1190 + 288

10 1220 + 393

15 1300 + 429

The take-up speed of the rotating mandrel for solution feed-rate of 1 ml/hr and 2

ml/hr was 58 m/min while the take-up speeds for solution feed-rate of 5 ml/hr and

22

higher was 63 m/min. The take-up speed is defined as the minimum speed for the

yarn to be drawn continuously onto the rotating mandrel without it being washed in

the tank below. Since the speed of water flow was constant, the factors that influence

the take-up speed at different feed-rate could be due to the different surface area in

contact with the water between fibers of different diameter or the higher mass of the

yarn spun at higher feed-rate. However, more tests need to be carried out to determine

the dominant factor in influencing the take-up speed.

For continuous drawing of the yarn, a feed-rate of more than 5 ml/hr is preferred.

Lower feed-rate would often results in yarn breakage. Since the tensile strength of a

single nanofiber is weak, there must be sufficient nanofibers in a yarn for it to be

strong enough to withstand the drawing and winding process. For the yarn to be

sufficiently strong either larger diameter fibers are used or more fibers are used to

form the yarn. Thus, higher feed-rate would favor both criteria. Figure 3.4 shows that

with higher feed-rate, the yarn diameter increases, however, the scatter of the yarn

diameter also increases. With an increased feed-rate and assuming that all the fibers

deposited on the water surface was collected as yarn, conservation of mass will mean

that increased feed-rate will bring about a corresponding increase in yarn diameter at

the same take-up speed. Increment in the scatter with increased feed-rate may be due

to a few factors. As seen from table 3.2, a higher feed-rate will result in a higher

variance in fiber diameter. Thus, intrinsically, this will lead to greater scatter in yarn

diameter. Another factor is the deposition of the nanofiber on the water surface from

the vortex. Water at the edge of the vortex flows quickly down the hole, however,

23

away from the vortex, the water tends to flow in a circular motion around the vortex.

At a lower feed-rate, the fiber deposition area is larger thus there is a higher

probability that the fibers deposits over the vortex and drawn into a yarn. Conversely,

at a higher feed-rate, a smaller fiber deposition area (Theron et al. 2004) may result in

the fibers being deposited away from the vortex and not drawn through hole

consistently.

3.4 Biomedical applications

Electrospun nanofibrous yarn has numerous applications in biomedical applications.

PLLA/col and PCL/col are biodegradable and they have different degradation rates.

One immediate application of this yarn is as sutures. Nanofibrous topography may

0.0

20.0

40.0

60.0

80.0

100.0

120.0

140.0

160.0

4 6 8 10 12 14 16

Feedrate (ml/hr)

Ave

rag

e ya

rn d

iam

eter

(m

icro

ns)

Figure 3.4 Graph of average PVDF-co-HFP yarn diameter against feed-rate. The vertical line for each point depicts the scatter in the yarn diameter for each feed rate.

24

encourage better cell proliferation and adhesion at the suturing site thus promoting

faster recovery and less scar tissue formation. Bundles of nanofibrous yarns may be

used in ligament or tendon reconstruction. Presence of collagen in the yarn will also

encourage better cellular integration to the scaffold and have the potential to

accelerate integration with native ligament or tendon tissues once the scaffold has

completely resorbed. Recently, nanofibrous yarns have been tested as nerve guidance

channel for bridging of nerve gaps as shown in Figure 3.5. In vivo study base on rat

sciatic nerve model have been carried out and the result showed that nanofibrous

yarns were able to promote better functional recovery of the severed nerve.

Figure 3.5 Nanofibrous yarn used as intra-luminal guidance channel. [A] Overview of nerve guidance channel consisting of a nanofibrous conduit and nanofibrous yarn in the lumen. [B] SEM image of conduit extracted from rat sciatic nerve after 3 months. [Picture courtesy of H. S. Koh]

25

3.5 Conclusion

It has been shown that by using flowing water as a working substrate, nanofibers

deposited on it can be arranged to form yarn assembly. This process for yarn

fabrication is versatile and can be used for different polymers as demonstrated in this

chapter. Increasing the concentration and feed-rate will increase both the fiber

diameter and the corresponding yarn diameter. Long strands of yarn may be used to

form other structures using techniques such as weaving, knitting and braiding. It can

also be used as reinforcement for composite structures. Using the setup described in

this chapter, shorter strands of yarns can also be used to form three-dimensional

assembly in situ. This will be covered in details in the next chapter.

26

Chapter 4: Fabrication of 3D Assemblies using fluid

properties

4.1 Introduction

Increasing evidence have shown that cells behaves differently in scaffold that is two-

dimensional and three-dimensional especially stem cells (Battista et al. 2005). Thus

there is a need to assemble nanofibrous structure that has a distinct three-dimensional

form. Water and its interaction with materials offer novel techniques of structure

fabrication especially in the sub-micron to nano-dimension level. Fluid properties and

interactions such as surface tension (Syms et al. 2003), capillary force and physical

confinement (Maury et al. 2008) have been used to assemble ultra-small components.

From the previous chapter, it has been shown that a fluid such as water can be used to

arrange the nanofibers into other structures by modifying its flow pattern. Zhang et al

(Zhang et al. 2005) and Deville et al (Deville et al. 2006;Deville et al. 2006) have

used freezing ice to control the distribution of particles in suspension to form

complex structures. In this chapter, water flow and the interactions between materials

and water properties will be used to construct three-dimensional composite nanofibers

structure.

4.2 Experimental Procedures

Polycaprolactone (PCL) (Aldrich, Mn ca. 80,000), collagen type I (col) (Koken) and

poly-L-lactide (PLLA) (Polyscience Inc, Mw 300,000) was used. PCL and a blend of

27

PCL/collagen (70:30 w/w) were dissolved in HFP to give a concentration of 0.08

g/ml for each solution. PLLA/col (50:50 w/w) was dissolved in HFP to give a

concentration of 0.03 g/ml. A voltage of 12 kV was supplied to the needle using a

Gamma High Voltage Research HV Power Supplies and a constant feed-rate of 1

ml/hr was used. The distance between the tip of the needle and the water surface was

set at 14 cm. The setup is the same as that used to fabricate nanofibrous yarn as

shown in Figure 3.1 except that the yarn was allowed to accumulate in the tank below

the basin.

Once sufficient yarns were deposited in the tank, they were collected and either

freeze-dried or dried under room conditions. Yarns to be freeze-dried were packed in

three different molds, cylinders of diameter 8 mm and 15 mm, and a hemispherical

container of diameter 15 mm. Freeze-dried samples were either pre-frozen at –86 oC

for 24 hr or at –196 oC in liquid nitrogen for 15 min before transferring to a freeze

drier (Heto Lyolab 3000). Freeze-drying was carried out for 48 hrs before further

characterization.

Macroscopic image was captured on a digital camera for visual examination.

Microscopic examination was carried out using an SEM. Samples for SEM were gold

coated for 120 s and 10 mA using a fine coater (Jeol JFC-1600 Autofine coater).

Diameters of the fibers were measured from the SEM images using Image J software

(National Institute of Health, USA). Hydrophobicity of the scaffold was determined

28

by a sessile drop contact angle measurement using a VCA Optima Surface Analysis

System (AST products, Billerica, MA).

4.3 Results and Discussions

With the onset of electrospinning above the water after a few minutes, a clump of

nanofiber yarns was observed in the water tank. The nanofiber assembly collected

was soft and easily conformed to the shape of the mold.

4.3.1 Physical structure observation

The drying condition was shown to affect the physical structure of the PCL scaffold.

Drying under room condition resulted in scaffold that was compact and shrunken

without any observable macro-pores as shown in figure 4.1a. In contrast, freeze-dried

scaffold was distinctly fluffy with numerous macro and micro-pores figure 4.1 b, c

and d.

29

Figure 4.1 [a] Polycaprolactone (PCL) 3D mesh dried in room condition with no visible pores. [b] PCL 3D mesh pre-frozen at -86 oC and freeze-dried in a cylinder with visible pores [c] PCL/col 3D mesh pre-frozen at -86 oC and freeze-dried in a hemispherical container showing visible pores [d] PCL/collagen 3D mesh pre-frozen at -86 oC and freeze-dried in a cylinder with visible pores (Teo et al. Current Nanoscience 2008; 4: 361-369 © Bentham Science Publishers).

Observation under the SEM revealed scaffolds dried either in room condition or

freeze-dried condition was made out of nanofibrous yarn as shown in Figure 4.2a and

b. This is in agreement with our previous study where the vortex formed by the water

exiting the basin hole caused the deposited nanofibers on the water surface to be

drawn into a yarn assembly. Closer examination revealed the tightly compacted yarns

of the scaffold dried in room condition while yarns were further apart in the freeze-

dried scaffold.

30

Figure 4.2 [a] PCL mesh dried under room condition showing yarn stacked closely on top of one another. [b] Freeze-dried PCL mesh pre-frozen at -86 oC showing distinct and isolated yarns made out of aligned nanofibres. [c] PCL/collagen mesh pre-frozen at -86 oC before freeze-drying showing ridge-like structures. [d] Disordered nanofibres that formed the ridges. All samples were packed in 15 mm diameter cylinder (Teo et al. Current Nanoscience 2008; 4: 361-369 © Bentham Science Publishers).

The influence of the drying condition on the PCL scaffold may be caused by capillary

pressure (Scherer 1990). When a wet sample is dried in a condition where water in

the liquid phase is converted to the gaseous phase, capillary pressure will be exerted

on the walls of the pore where the water resides. The tension (P) on the evaporating

water is related to the radius of curvature of the meniscus by r

P2

, where is the

surface tension, r is the radius of curvature of the meniscus. When the center of

curvature is in the vapor phase, the radius of curvature is a negative value resulting in

31

tension in the water (P > 0). Water evaporating from the surface of the scaffold will

encourage water from its interior to migrate to the surface. In the process, as the

volume of water decreases, the radius of curvature in the meniscus reduces resulting

in an increased tension in the water within the pore. If the walls of the pore are not

stiff or strong enough, the walls will converge causing the scaffold to shrink. To

maintain a porous structure in a mechanically weak scaffold, it is therefore necessary

to remove condition for capillary pressure to exist. This can be achieved by first

converting the water from liquid phase to solid phase and subsequently removing the

water in its solid state from the scaffold. By pre-freezing the scaffold followed by

freeze-drying, the dried PCL scaffold was able to maintain its porous structure.

4.3.2 Effect of hydrophobicity

In PCL/collagen (70:30 w/w) freeze-dried scaffold pre-frozen at –84 oC, the scaffold

microstructure was distinctly different from pure PCL freeze-dried scaffold. At the

macroscopic level, the texture was spiky compared to the fluffy PCL scaffold.

Observation under the SEM showed disordered nanofibers forming ridges radiating

from the axis of the scaffold as shown in Figure 4.2c and d. PLLA/col (50:50 w/w)

scaffold pre-frozen at –84 oC also showed similar physical characteristic.

The difference in the PCL/collagen (70:30 w/w) and PLLA/col (50:50 w/w) scaffold

freeze-dried scaffold from PCL freeze-dried scaffold may be due to their difference in

hydrophobicity. Ice is known to adhere better to hydrophilic materials (Raraty and

Tabor 1958) and the effect of cooling rate and ice crystal formation on freeze-dried

32

biological tissues has been well established (Schwabe and Terracio 1980;Meryman

1956;Ishiguro and Rubinsky 1994). Ice crystal formation and growth was known to

cause mechanical distortion in cells with larger ice crystals causing greater damages.

Heterogeneous nucleation will certainly occur during freezing of the scaffold due to

the presence of numerous surfaces acting as nucleation sites (Meryman 1956).

To determine the hydrophobicity of a material, surface water contact angle of the

scaffold was used. However, this widely used method requires the material to have

flat surface so that a baseline can be established. Thus to perform this test on the

scaffold, the 3D mesh was first compressed to give a flat surface before the

measurement was taken. As expected, the PCL scaffold was found to be relatively

more hydrophobic with a surface contact angle of 105 + 20o while the water droplet

immediately absorbed into the PCL/col scaffold upon contact.

When a container containing water is cooled below freezing temperature, ice

nucleation followed by crystal growth in the form of knife-edge ice front (Harrison

and Tiller 1963) proceeds from the wall (Langer 1980) to the interior due to faster

heat lost. This freezing pattern caused the distinctive ridge-like microstructure on the

surface of PCL/collagen (70:30 w/w) and PLLA/col (50:50 w/w) scaffolds. Figure 4.3

shows the schematic of the process.

33

Figure 4.3 Schematic of the ice crystals pushing the entangled yarn to form disordered fibres and ridges on the PCL/col scaffold. [a] Stage 1. Nanofibrous yarns were suspended in the water. [b] Stage 2. As the water cools, ice nucleates on the wall of the cylinder and on the nanofibres. [c] Stage 3. Growing ice front from the cylinder wall pushes against the nanofibres resulting in [d] ridges made out of random nanofibres (Teo et al. Current Nanoscience 2008; 4: 361-369 © Bentham Science Publishers).

At temperatures below freezing, ice nucleation will occur on any surfaces with

impurities. This would include the spaces between the hydrophilic nanofibers where

water molecules are present. Crystal growth amongst in the yarn would push the

nanofibers apart disrupting the yarn structure. Since water cools faster at the wall of

the mold, ice crystal growth will be faster and the ice front would soon hit the

nanofibers before they were frozen in place. As the growing ice front pushes against

regions of entangled nanofibers and sections of nanofibers held by ice, ridge-like

microstructure consisting of disordered nanofibers will be formed. As ice nucleation

and growth also concurrently proceed at the core of the scaffold freezing the

nanofibers in place, a boundary consisting of frozen inner nanofibr core and the

displaced nanofibers at the surface of the scaffold was formed. This boundary layer is

shown in Figure 4.4.

34

Figure 4.4 Cross section of the PCL/col scaffold cooled at -86 oC showing inhomogeneous pores and a boundary layer between the displaced nanofibers on the surface and compacted fibers at the inner core (Teo et al. Current Nanoscience 2008; 4: 361-369 © Bentham Science Publishers).

In contrast with PCL/collagen (70:30 w/w) and PLLA/col (50:50 w/w) scaffolds,

nanofibers in the PCL scaffold was able to maintain its yarn assembly at the surface.

The hydrophobic nature of PCL accounts for this differences in microstructure

assembly after freezing. In water, hydrophobic surfaces are known to exert attractive

forces (Israelachvili and McGuiggan 1988;Christenson and Claesson 2001;Yaminsky

1999) thus PCL nanofibers would adhere to one another in water driving out any

water molecules between them. Christenson and Claesson (1998) have shown that

many vapor cavities would bridge two hydrophobic surfaces at the point of contact

(Christenson and Claesson 1988). Without any water molecules between the

nanofibers, ice nucleation and growth cannot proceed thus the nanofibers were not

pushed apart and the yarn microstructure remained after freezing.

35

4.3.3 Effect of cooling rate

Microstructure of PCL/collagen (70:30 w/w) and PLLA/col (50:50 w/w) scaffolds has

been shown in the previous section to be formed by ice crystal growth. One of the

factors influencing ice nucleation and crystal growth is cooling rate. When the

cooling rate increases, ice crystal size decreases but the number of ice crystals

increases. Nevertheless, reduction in crystal growth would cause less mechanical

distortion in water-filled structures. Therefore, hypothetically, reduction in crystal

size would reduce the amount of disorder in original yarn microstructure in

hydrophilic scaffolds. To test this hypothesis, PCL/collagen (70:30 w/w) and

PLLA/col (50:50 w/w) scaffolds placed in 8 mm diameter cylinder were pre-frozen in

liquid nitrogen at –196 oC. A comparison of PCL/collagen (70:30 w/w) physical

structures pre-frozen at –86 oC and at –196 oC is shown in Figure 4.5.

36

Figure 4.5 Freeze-dried PCL/col [a] prefrozen at -86 oC showing “spikes”, [b] mesh pre-frozen in liquid nitrogen has a relatively smoother surface. [c] Cross-section of the mesh pre-frozen at -86 oC showing the ridges are only found on the surface while [d] the mesh pre-frozen in liquid nitrogen did not exhibit any apparent differences in the microstructure from the surface to the interior.[e] Higher magnification showing microstructure of scaffold pre-frozen at -86 oC where the “spikes” were ridges formed by disordered nanofibres and [f] mesh pre-frozen in liquid nitrogen with distinct yarns (Teo et al. Current Nanoscience 2008; 4: 361-369 © Bentham Science Publishers).

Both PCL/collagen (70:30 w/w) and PLLA/col (50:50 w/w) pre-frozen at –196 oC

retained their original yarn microstructure. At –196 oC, numerous ice nucleations

37

occur on the surface of the nanofibers and these ice crystals quickly freeze the yarn in

place and any ice crystal growth that may disrupt the nanofibrous assembly was

retarded. However, the nanofibrous yarns in the hydrophilic scaffolds were not as

tightly bundled as the hydrophobic PCL yarn. Although ice crystal growth is reduced,

ice crystal forming on the nanofibers will still cause the nanofiber to separate during

freezing. Figure 4.6 shows a schematic of the ice crystal formation on the scaffold at

–196 oC.

Figure 4.6 Schematic of the ice nucleation and growth on PCL/col mesh cooled at -196 oC. [a] Stage 1, nanofibrous mesh is suspended in water. [b] Numerous and rapid ice nucleation on the surface of the nanofibers while ice nucleation and growth commence from the cylinder wall. [c] Complete freezing of mesh before the ice growing from the cylinder wall reaches it (Teo et al. Current Nanoscience 2008; 4: 361-369 © Bentham Science Publishers).

4.4 Application in Tissue Engineering

Despite numerous in vitro studies demonstrating favorable proliferation, adhesion and

gene expression of cells cultured on nanofiber mesh, however organs such as bone

and cartilage require a three-dimensional environment rather than a two-dimensional.

This has prompted efforts to create three-dimensional nanofibrous matrix. Srouji et al

38

(2007) stacked layers of flat mesh to give the scaffold sufficient depth. However, this

method of scaffold fabrication is very inefficient (Srouji et al. 2008). Ki et al (2007)

deposited electrospun nanofibers on a methanol coagulation bath until a 3D scaffold

was formed. However, the use of large amount of methanol which is harmful to

health and highly flammable poses a hazard if this method is to be used on a large

scale (Ki et al. 2007). Moreover, the structure is consists entirely of randomly

oriented nanofibers with no organization at the micro-structural level. Studies have

also shown that cells cultured on aligned nanofibers tend to elongate in the direction

of fiber alignment (Zhong et al. 2006). The dynamic flow method as described in this

chapter is able to fabricate three-dimensional matrices with micro-structural

organization. This allows further investigation of cell activity in 3D matrices with

different micro-structural organization. In vitro studies have shown that a composite

scaffold consisting of synthetic polymers and natural polymers showed better cell

activity compared to pure synthetic polymer (He et al. 2005). Potential applications

include cartilage, liver and breast regeneration since these organs require three-

dimensional matrix. The ability to fabricate a three-dimensional composite blend of

PCL/collagen (70:30 w/w) and PLLA/col (50:50 w/w) allows further studies into the

cell activity in a three-dimensional nanofibrous matrix environment.

4.5 Conclusion

The ability to construct block three-dimensional nanofibrous scaffold is a significant

advancement in the electrospinning technique. This opens up a wide range of organ

and tissue regeneration possibilities as many organs and tissues are three-dimensional

39

nanofibrous matrices. The ability to control the microstructure of the scaffold using

different freezing and drying conditions allows researchers the potential to better

tailor the synthetic regenerative scaffold for the particular tissue or organ.

40

Chapter 5: Mineralization of three-dimensional matrix

5.1 Introduction

In bone regeneration, incorporation of calcium phosphate is known to promote better

integration between implanted scaffold and the surrounding bone. In Chapter 3, a

dynamic fluid flow method has been shown assemble nanofibers into the form of

yarn. This method is then used to generate 3D nanofiber matrices and the effect of

drying condition on the matrix microstructure was investigated in Chapter 4. In this

chapter, mineralization of the 3D matrix is carried out to deposit calcium phosphate

nanoparticles on its surface.

5.2 Experimental Procedures

PLLA/col (50:50 w/w) and PLLA samples were prepared as described in the previous

chapter. In brief, flowing water with a vortex through a hole at the base of the basin

was used to collect the electrospun nanofibers. As the collected nanofibers flow down

the vortex, clumps of nanofibers were deposited on a tank below the basin. The

clumps of PLLA/col (50:50 w/w) or PLLA were placed in a cylindrical mold of inner

diameter 8 mm and freeze-dried. Cancellous bone cubes (CANCUBE30 S) of

dimension 3 mm x 3 mm x 3 mm were purchased from LifeNet Health.

Mineralization was first carried out on PLLA/col and PLLA samples. The samples

were first soaked in 0.5 M CaCl2 for 1 hr. The samples were then washed using de-

ionized (DI) water before soaking in 0.3 M Na2HPO4. Subsequently, the samples

41

were washed with DI water before repeating one more cycle of soaking in 0.5 M

CaCl2 and in 0.3 M Na2HPO4. The mineralized samples were then freeze dried.

To examine the effect of a flow system in mineralization of 3D scaffold, two methods

of mineralization technique were used on PLLA/col (50:50 w/w) 3D scaffold. First

was static mineralization which follows conventional alternate dipping mineralization

technique where the sample is dipped in CaCl2 followed by Na2HPO4. The other was

a flow mineralization technique where the alternate dipping mineralization has been

modified to encourage the solution to flow through the sample. In static

mineralization, the sample was kept in a mold and mineralization was carried out by

submerging the sample into 30 ml of 0.5 M CaCl2 for 1 hr. The solution was drained

and rinsed with 100 ml of de-ionized (DI) water. The sample was then rinsed with 10

ml of 0.3 M Na2HPO4 solution before submerging in 30 ml of 0.3 M Na2HPO4

solution for 1 hr. The solution was drained and rinsed with 100 ml of de-ionized (DI)

water. Subsequently, the sample was rinsed with 10 ml of 0.5 M CaCl2 solution

before submerging in 30 ml of 0.5 M CaCl2 solution for 1 hr. The last step of rinsing

and submerging of the sample in 30 ml of 0.3 M Na2HPO4 solution for 1 hr was

carried out as per described above before the mineralized sample was freeze-dried

and characterized. A schematic of the flow mineralization setup for PLLA/col sample

was shown in Figure 5.1. A peristaltic pump was used to circulate the solution

through the PLA/col 3D scaffold. Masterflex® 96400-16 tubing was used with the

pump at a flow rate of 40 ml/h. The mineralization procedure was as such, first, 30 ml

of 0.5 M CaCl2 was pumped through the scaffold for 1 h. Next, the solution was

42

drained except for the chamber containing the scaffold. 100 ml of DI water was used

to flush through the tube and the scaffold in the chamber. Without draining the water

in the chamber, 10 ml of 0.3 M Na2HPO4 was used to displace the water in the

chamber. After which 30 ml of 0.3 M Na2HPO4 was pumped through the tube and

scaffold for 1 h. Another round of draining, flushing and pumping of CaCl2 and

Na2HPO4 was carried out while ensuring that the chamber was filled with liquid at all

times. After mineralization, the sample was washed with DI water before freeze

drying.

Figure 5.1 Setup for mineralization of the 3D scaffold.

Verification of the presence of collagen in the PLLA/col blend after freeze-drying

was carried out using Avatar 380 ATR-FTIR (Attenuated Total Reflection Fourier

Transform Infrared). To observe the distribution of calcium phosphate nanoparticles

in the sample, the sample was sectioned normal to its principal axis into at least 5

pieces before gold sputtering and observation under SEM. EDX (Energy-dispersive

X-ray spectroscopy, S-4300 FE SEM HITATCHI) was carried out to determine the

presence of calcium phosphate in the mineralized sample. To check the composition

43

and phase of the minerals, XRD (X-ray diffraction) was used. The scaffold was

crushed using pastel and mortar to form a solid “pancake” for XRD.

To find the amount of minerals in the mineralized scaffold, two methods were used.

The first method involves finding the weight increase in the scaffold following

mineralization and freeze drying. In the second method, the mineralized scaffold was

weighted followed by sintering at 500 oC, with a ramp rate of 10 oC/min and a dwell

time of 30 min. The weight of the ash remains will give us the percentage mass of the

minerals in the mineralized scaffold. The temperature for sintering was found using

TGA.

An Instron 3345 was used to measure the compressive strength and modulus of the

scaffolds. A strain rate of 0.0033 /s was applied and a 10 N load cell was used. The

compressive modulus was the peak stiffness taken from a plot of stiffness against

strain (n = 7). The compressive strength was defined by the stress at which the

stiffness had reduced by 3% from the maximum stiffness (n = 7) (Li and Aspden

1997).

5.3 Results and Discussions

Bone is known to be a hierarchically organized structure consisting of both organic

materials and inorganic nano-crystals. Since minerals are found on and within

collagen fibrils, numerous studies have suggested that collagen contains numerous

44

binding sites for their nucleation (Urry 1971;Glimcher and Muir 1984;Curley-Joseph

and Veis 1979). Rhee et al studied the mechanism where the carboxylate group of

collagen chelated with a calcium ion which may represent the first step of nucleation

towards hydroxyapatite (HA) crystal formation (Rhee and Lee 2000). Since collagen

is known to enhance cell adhesion and proliferation and is able to provide favorable

binding sites for HA formation, it has been incorporated into synthetic biodegradable

polymers for mineralization to construct bone graft. The spectra of PLLA and

PLLA/col blended 3D nanofiber scaffold were shown in Figure 5.2. The presence

amide II band at 1547cm-1 and amide I band at 1656 cm-1 in PLLA/col blend

indicates the presence of collagen in the scaffold. These peaks were distinctly absent

in PLLA scaffold.

Figure 5.2 Spectra of PLLA and PLLA/col blended 3D nanofiber scaffold.

Using the alternate soaking mineralization technique, numerous mineral nanoparticles

were deposited on the PLLA/col sample while much lesser mineral nanoparticles

45

were observed on the surface of PLLA samples as shown in Figure 5.3. Due to the 3D

nanofibrous architecture, some mineral nanoparticles were found in the inner core of

the PLLA sample. Gopal et al have shown that particles less than 1 µm were attracted

to nanofibers. Mechanisms such as direct interception, inertial impaction and

diffusion have been proposed. However, since this was a static mineralization, the

most probable mechanism would be diffusion (Brownian Motion) where the

precipitated nanoparticles flow into the scaffold and retained by adsorption force.

Rinsing was not sufficient to detach the nanoparticles which agree to the observation

made by Gopal et al (Gopal et al. 2007). In PLLA/col sample, mineral nanoparticles

were found both on the surface and at the inner core with greater density. A study by

Ngiam et al showed that PLLA nanofibers is a poor material for HA deposition.

However, with the addition of collagen into PLLA, the resultant scaffold showed

excellent mineral deposition (Ngiam et al. 2009).

Figure 5.3 SEM images of mineralized [A] PLLA sample and [B] PLLA/col sample.

46

3D nanofibrous scaffold presents a different environment for mineralization as

compared to a flat film or mesh. Both static mineralized scaffold and flow

mineralized scaffold resulted in relatively rigid without any visible compression.

Gross examination of static mineralized scaffold showed absence of macroscopic

pores. Observation of the cross section of static mineralized scaffold under SEM

showed uneven distribution of the minerals from the periphery of the scaffold to its

core. While numerous nanoparticles covered the nanofibers at the periphery of the

scaffold, the amount of mineral deposition was visibly less, closer to the core as

shown in Figure 5.4. The distribution of the nanoparticles was also not uniform along

the length of the nanofibers.

Figure 5.4 Distribution of minerals on across the cross-section of static mineralized scaffold.

47

Alternate dipping method has been successfully used to deposit mineral nanoparticles

on nanofibrous membrane in less than an hour (Ngiam et al. 2009), however, 3D

scaffold requires a much longer duration for the ions to diffuse into its core and

sufficient mineral deposition to stiffen the structure. Moreno and Varughese (1981)

used the following equation to describe the rate of hydroxyapatite precipitation in

seeded crystal growth experiment (Moreno and Varguhese 1981),

nkASR where R is the rate of hydroxyapatite precipitation k is proportionality constant A is the surface area of the seeding crystal S is the degree of supersaturation n is the order of the reaction

and v

spK

IPS

1

where IP is the ion activity product of hydroxyaptatite Ksp is the solubility product of hydroxyaptatite v is the number of ions per mole of the solid phase

During mineralization, heterogeneous chemical reaction where the calcium ions

compete for binding sites on the nanofibers takes place. Non-uniform deposition of

calcium phosphate on the nanofibers in static mineralization may be due to

discontinuous precipitate concentration on the nanofibers (Samson et al. 2000). As

given in the equation above, high surface area of the nanofibers and porous scaffold

encourages rapid precipitations when phosphate ions were introduced. The

corresponding competition for the phosphate ions gave rise to localized regions of

undersaturation where the calcium phosphate dissolves as diffusion of the phosphate

48

ions lags the rate of nucleation and crystal growth whereas regions of oversaturation

encourages calcium phosphate precipitation. Rapid precipitation also caused blockage

of the pores on the scaffold. Reduction of free phosphate ions with time and pore

blocking on the surface of the scaffold further reduces phosphate concentration

available for diffusion into the core of the scaffold. This results in little or no minerals

found in the core of the scaffold as the solution is undersaturated in calcium

phosphate.

Figure 5.5 shows the SEM images of the cross section of the flow mineralized

scaffold. Unlike the static mineralized scaffold, minerals can be found both on the

surface and the core of the mineralized scaffold and the mineral distribution on the

fibers were more uniform. However, there were still isolated pockets of nanofibers

where no minerals were observed.

49

Figure 5.5 Distribution of the minerals in the cross-section of the scaffolds using flow mineralization.

Unlike the static mineralization method, flow mineralization forces the ions into the

highly porous scaffold. This allows a more uniform distribution of free ions

throughout the scaffold. Mineral nucleation and growth is not restricted by the rate of

diffusion and there is more homogeneous solution saturation on the surface of the

nanofibers. However, due to the highly tortuous path in the scaffold for the fluid to

flow through, isolated pockets in the scaffold will experience different flow rate and

may result in varying minerals deposits throughout the scaffold.

50

Examination of a commercially available bone chip using EDX gave an average Ca/P

ratio of 1.06 + 0.23. The Ca/P ratio found in the static mineralized scaffold at

different points also varies with an average Ca/P ratio of 1.88 + 0.69 which is closer

to the stoichiometric ratio of HA than the commercial bone chip. Ca/P ratio of flow

mineralized scaffold was found to be 2.44 + 1.35. In natural bones, it is widely

accepted that the calcium phosphate minerals are mostly in the form of amorphous

hydroxyapatite. While the stoichiometric ratio of Ca/P ratio is about 1.67, natural

bone have Ca/P ratio varies from individuals about 1.9 to 2.2 (Akesson et al.

1994;Tzaphlidou and Zaichick 2004;Zaichick and Tzaphlidou 2003) and it can go as

high as 4.78 (Zaichick and Tzaphlidou 2003).

When the scaffold was crushed using a pastel and motar, the minerals were not

detached from the nanofibers suggesting that the adhesion between the nanoparticles

and the nanofibers were good. As the scaffold was not brittle, a consolidated solid flat

pan-cake was formed. No attempts were made to separate the minerals from the

polymer so as not to alter the composition and form of the minerals for

mineralization. As shown in figure 5.6, the XRD result showed numerous peaks with

a main peak observed close to 32 suggesting that the minerals formed on the scaffold

were amorphous calcium phosphate or possibly HA. However, due to the presence of

numerous peaks and the overlapping of peaks from PLA, it is not possible to verify

whether the majority of the minerals were hydroxyapatite.

51

Figure 5.6 XRD showing the presence of amorphous calcium phosphate crystals in the mineralized scaffold.

The presence of different calcium phosphate phases in the mineralized scaffold is

probably due to the tortuous paths which the calcium and phosphate ions took as they

flow towards the core of the scaffold. At the surface of the scaffold, ions would start

to adhere to the carboxylate groups found on the collagen. Therefore, going deeper

into the scaffold, the ion concentration would start to reduce. As the pore size varies

at different points on the scaffold, the flow rate would differ. This results in sub-

Mineralized scaffold

Unmineralized scaffold

52

optimum calcium and phosphate ion concentration and composition away from the

surface of the scaffold giving rise to amorphous calcium phosphate in different

phases.

Base on the weight increase after mineralization, the minerals accounted for 70.1 +

4.5% (n = 10) of the weight in the static mineralized scaffold and 50.6 + 8.3% (n =

10) of the weight in the flow mineralized scaffold. Through sintering, the static

mineralized scaffold was found to comprise of 62.3 + 4.0% (n = 5) minerals and 43.3

+ 6.9% (n = 5) minerals in flow mineralized scaffold. The difference in the weight

could be due to the presence of water in the sample.

Figure 5.7 shows the ash remains of the static mineralized scaffold and flow

mineralized scaffold. Sintered static mineralized scaffolds were either hollow or had a

black inner core while the surface remains relatively whiter. The black ash was likely

to be carbon remains of sintered PLLA and collagen and the white surface was the

calcium phosphate deposits. This provides further evidence that minerals were mainly

deposited on the surface of the scaffold. Unlike the scaffold mineralized using static

mineralization method, the ash remains of the flow mineralized scaffold had a

relatively uniform color distribution throughout and had a solid core. This provides

further evidence that calcium phosphate was more evenly distributed throughout the

scaffold in the flow mineralization method.

53

Figure 5.7 Ash remains of mineralized scaffold after sintering at 500 oC. [A] Static mineralized scaffold showing a hollow core. [B] Flow mineralized scaffold showing a solid core.

In native bone, the minerals are deposited on the collagen fibrils throughout the bone.

Studies have shown that presence of hydroxyapatite on nanofibers facilitate osteoblast

proliferation and secretion of minerals (Zhang et al. 2008). Therefore, it is preferred

that the minerals are deposited throughout the scaffold. As seen in Figure 5.4, the

minerals on the outer surface of the scaffold were not adhering to the nanofibers.

Thus it is likely that the minerals may be dislodged easily when it is subjected to

mechanical stresses. Migration of the mineral nanoparticles to other part of the body

in vivo may cause unnecessary inflammatory reaction.

The compressive strength and modulus of the scaffolds were significantly influenced

by the mineralization process as shown in Figure 5.8. Static mineralized scaffold

showed an increment of almost three times the compressive strength and almost twice

the modulus of unmineralized scaffold. In flow mineralized scaffold, the compressive

strength of 103 kPa showed even more significant increase over unmineralized

scaffold (17 kPa) and static mineralized scaffold (41 kPa). The compressive modulus

54

of flow mineralized scaffold at 782 kPa also showed significant increase over

unmineralized (148 kPa) and static mineralized scaffold (276 kPa).

Figure 5.8 Mechanical properties of scaffold mineralized under different condition [A] Compressive strength. [B] Compressive Modulus

Presence of minerals in the scaffold has been shown to improve compressive strength

and modulus. However, a more significant contribution is the arrangement of the

55

minerals nanoparticles on the fibers that made up the scaffold. Static mineralized

scaffold, despite having 20% more mineral content than flow mineralized scaffold,

had a compressive strength and modulus that was respectively six times weaker and

almost three times weaker than flow mineralized scaffold. From SEM images, it has

been shown that the minerals on static mineralized scaffold were found on the surface

while minerals were found throughout flow mineralized scaffold. Minerals were also

more homogeneously covering the surface of individual nanofibers in the flow

mineralized scaffold. This had an additional strengthening effect on the fibers and

thus the whole structure. However, the compressive strength and modulus of flow

mineralized scaffold pales in comparison with human cancellous bone (Li and

Aspden 1997). Figure 5.9 shows the SEM image of flow mineralized scaffold versus

cancellous bone. The minerals and nanofibers in cancellous bone were more

compacted than those of the mineralized scaffold. The c-axis of the bone mineral is

also in the direction of the nanofibers which the mineralized scaffold lacked. Further,

mineral nanoparticles are also arranged in an orderly manner in the collagen triple

helix structure. Certainly, such greater order and hierarchical organization of the

minerals and collagen in bone contributed significantly to its mechanical properties.

56

Figure 5.9 Comparison of mineral nanoparticles distribution in [A] mineralized nanofibrous scaffold and [B]cancellous bone.

5.4 Conclusion

The alternate dipping method has been used successfully to create a composite

scaffold consisting of PLLA/col scaffold and calcium phosphate minerals. Although

the Ca/P ratio of the deposited minerals is close to the stoichiometric ratio of HA,

XRD was not able to confirm that the majority of the minerals were consisted of HA.

Nevertheless, various researchers have shown that calcium phosphates of various

phases are biocompatible and are suitable for use as bone repair graft. SEM images

and gross observation of the sintered mineralized scaffold showed that most of the

calcium phosphate minerals were concentrated on the surface. Therefore,

modification of the alternate dipping method is required to encourage mineral

deposition in the core of the 3D scaffold.

57

Chapter 6: Research Summary and Future

Recommendations

The next generation of regenerative scaffold will increasingly base on mimicking

native tissue extracellular matrix (ECM). Electrospinning has provided an avenue for

researchers to fabricate nanofibrous structure which may potentially mimic ECM.

Studies by various researchers have shown that electrospun nanofibers provide a

favorable environment for adhesion and proliferation of cells in vitro. To translate in

vitro results into usable regenerative scaffold for various organ regeneration and

tissue engineering, a flat mesh produced by conventional electrospinning is

inadequate. In particular, an appropriate bone regenerative scaffold has to be a 3D

construction and ideally mimic the hierarchical organization of natural bone.

This thesis presents several breakthroughs in the electrospinning process which

allows the eventual fabrication of a composite 3D nanofibrous structure for

biomedical applications. Since nanofiber does not have the mechanical strength to be

handled to form 3D architectures using mechanical devices, a new method of

manipulating the nanofibers using water fluid properties has been developed. The

fluid nature of water allows movement and arrangement of the nanofibers without

breaking them. The diameter of the yarn and the nanofibers that made up the yarn can

be altered by varying the solution concentration. The composite biodegradable

nanofibrous yarn has various biomedical applications such as suture or ligament and

58

tendon repair. Research has also shown that the nanofibrous yarn can be used as intra-

luminal guidance channel for bridging nerve gap. Nanofiber manipulation using water

has been further developed to give a 3D nanofibrous scaffold. Adjusting the freezing

and drying condition allows some control on the microstructure of the 3D scaffold.

Since collagen can be incorporated into the 3D nanofibrous scaffold, mineralization

can be carried out to obtain high density mineral deposits thus mimicking the

architecture and composition of natural bone. In bone regenerative scaffold,

researchers have shown that various phases of calcium phosphates such as tricalcium

phosphates (McCullen et al. 2009) and dicalcium phosphate dehydrate (Li et al. 2008)

were able to promote cell differentiation and proliferation. Therefore, it is expected

that the composite 3D scaffold will provide a bioactive and biocompatible

environment for bone regeneration.

As shown in this thesis, mimicking nature's hierarchical structure is able to

significantly increase the mechanical strength of the composite. Nevertheless, the

nanofibrous composite strength is much weaker than bone. These differences may be

due to the much higher level of hierarchical organization in bone. The collagen

nanofiber molecules are helically arranged with nano-hydroxyapatite crystals

periodically situated within the nanofiber. Although it is possible to blend nano-

hydroxyapatite crystals into electrospun nanofibers, the distribution of the crystals

will be random. Surrounding the collagen nanofibers in bone are compacted nano-

hydroxyapatite crystals with their c-axis in the direction of the collagen alignment. In

man-made mineralized composite, it is difficult to replicate such level nano-crystals

59

density and controlled orientation. These attributes account for the superior strength

of bone.

With the ability to construct nanofibrous yarn, 3D nanofibrous scaffold and

mineralized 3D nanofibrous scaffold, a new generation of biomimicking regenerative

scaffold has been developed and have the potential to significantly improve the

quality of life for patients in need of implants. Given the optimism and favorable in

vitro studies using nanofibers, these constructs are ready for the next stage of in vivo

studies. Future research should also involve examining how the various hierarchical

organizations contribute to the strength of the material. Both the strength of collagen

and calcium phosphate are individually low but their combination in an organized

manner has a multiplication effect on the strength of their composite. While one way

of improving the strength of composite materials is to use higher strength

reinforcements, an alternative and more efficient method may be to look into

hierarchical organization of the reinforcements in the composite. This study has

shown that at the nanometer level, even simple level of hierarchical organization can

achieve significant strengthening of composite.

60

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