nature inspired composite nanofibers teo wee eong · teo we, he w, ramakrishna s (2006) electrospun...
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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.
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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
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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.
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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
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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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