Kehinde Yusuf Thesis (PDF 9MB)

Institute of Health and Biomedical Innovation (IHBI)

School of Chemistry, Physics and Mechanical Engineering

Science and Engineering Faculty

Queensland University of Technology

AN EXPLORATORY STUDY OF THE POTENTIAL

OF RESURFACING ARTICULAR CARTILAGE

WITH SYNTHETIC PHOSPHOLIPIDS

Kehinde Quasim Yusuf

Bachelor of Science (Chemical Engineering) (Hons.), University of Lagos, Nigeria – 2004

Thesis submitted in fulfilment of the requirements for the degree of Doctor of Philosophy in

Science and Engineering Faculty, School of Chemistry, Physics and Mechanical, Queensland

University of Technology.

January 2013

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Supervisors: Professor Kunle Oloyede

Associate Supervisors: Professor Ross Crawford

Associate Supervisors: Associate Professor Nunzio Motta

Keywords

Apparent diffusion coefficient, Articular cartilage, Atomic force microscopy, Complementary

energy, Confocal microscopy, Delipidization, Fick’s law of diffusion, Magnetic resonance

imaging, Mechanical Compression tests, nanosurface characterization, Osteoarthritis,

Relipidization, Residual energy, Resurfacing cartilage, Semipermeability, Strain energy,

Surface-active phospholipids, Surface amorphous layer.

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Abstract

This thesis is aimed at further understanding the uppermost lipid-filled membranous layer

(i.e. surface amorphous layer (SAL)) of articular cartilage and to develop a scientific

framework for re-introducing lipids onto the surface of lipid-depleted articular cartilage (i.e.

“resurfacing”). The outcome will potentially contribute to knowledge that will facilitate the

repair of the articular surface of cartilage where degradation is limited to the loss of the lipids

of the SAL only. The surface amorphous layer is of utmost importance to the effective load-

spreading, lubrication, and semipermeability (which controls its fluid management, nutrient

transport and waste removal) of articular cartilage in the mammalian joints. However,

because this uppermost layer of cartilage is often in contact during physiological function, it

is prone to wear and tear, and thus, is the site for damage initiation that can lead to the early

stages of joint condition like osteoarthritis, and related conditions that cause pain and

discomfort leading to low quality of life in patients. It is therefore imperative to conduct a

study which offers insight into remedying this problem.

It is hypothesized that restoration (resurfacing) of the surface amorphous layer can be

achieved by re-introducing synthetic surface-active phospholipids (SAPL) into the joint

space. This hypothesis was tested in this thesis by exposing cartilage samples whose surface

lipids had been depleted to individual and mixtures of synthetic saturated and unsaturated

phospholipids. The surfaces of normal, delipidized, and relipidized samples of cartilage were

characterized for their structural integrity and functionality using atomic force microscope

(AFM), confocal microscope (COFM), Raman spectroscopy, magnetic resonance imaging

(MRI) with image processing in the MATLAB® environment and mechanical loading

experiments. The results from AFM imaging, confocal microscopy, and Raman spectroscopy

revealed a successful deposition of new surface layer on delipidized cartilage when incubated

in synthetic phospholipids. The relipidization resulted in a significant improvement in the

surface nanostructure of the artificially degraded cartilage, with the complete SAPL mixture

providing better outcomes in comparison to those created with the single SAPL components

(palmitoyl-oleoyl-phosphatidylcholine, POPC and dipalmitoyl-phosphatidylcholine, DPPC).

MRI analysis revealed that the surface created with the complete mixture of synthetic lipids

was capable of providing semipermeability to the surface layer of the treated cartilage

samples relative to the normal intact surface. Furthermore, deformation energy analysis

revealed that the treated samples were capable of delivering the elastic properties required for

load bearing and recovery of the tissue relative to the normal intact samples, with this

capability closer between the normal and the samples incubated in the complete lipid mixture.

In conclusion, this thesis has established that it is possible to deposit/create a potentially

viable layer on the surface of cartilage following degradation/lipid loss through incubation in

synthetic lipid solutions. However, further studies will be required to advance the ideas

developed in this thesis, for the development of synthetic lipid-based injections/drugs for

treatment of osteoarthritis and other related joint conditions.

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Table of Contents

Keywords.............................................................................................................................................(ii)

Abstract...............................................................................................................................................(iii)

Table of contents..................................................................................................................................(v)

List of figures......................................................................................................................................(ix)

List of Tables......................................................................................................................................(xix)

List of Abbreviations .........................................................................................................................(xx)

Statement of authorship.....................................................................................................................(xxiii)

Abstract ........................................................................................................................................... iv

Table of Contents ............................................................................................................................. vi

List of Figures ................................................................................................................................... x

List of Tables .................................................................................................................................. xx

List of Abbreviations ...................................................................................................................... xxi

Statement of Original Authorship ................................................................................................. xxiv

CHAPTER 1: INTRODUCTION ................................................................................................... 1

CHAPTER 2: CRITICIAL REVIEW/APPRAISAL OF THE LITERATURE ............................ 7

2.1 Introduction ............................................................................................................................ 7

2.2 motivation for this thesis - Treatment of Osteoarthritis ............................................................ 9

2.3 Articular Cartilage Structure and Architecture ....................................................................... 11

2.4 Bio-mechano-chemical basis of the functional failure of articular cartilage ............................ 18

2.5 Components of Articular Cartilage and their functions........................................................... 20

2.5.1 Collagen fibres........................................................................................................... 23

2.5.2 Proteoglycans ............................................................................................................ 26

2.5.3 Water (Bound and Unbound), Ions and Chondrocytes ................................................. 34

2.6 Articular Cartilage Lipids ..................................................................................................... 37

2.6.1 Biochemistry of Lipids: Nomenclature and Structure .................................................. 37

2.6.2 Phospholipids: Properties and Functions ..................................................................... 42

CHAPTER 3: EXPLORATORY STUDY OF THE APPROACH AND METHODOLOGY ..... 51

3.1 Atomic force microscope (AFM) imaging of the surface of articular cartilage ........................ 51

3.1.1 Choice of cantilever for AFM imaging ....................................................................... 57

3.1.2 Optimization of Set-point and Scanning Parameters .................................................... 59

3.1.3 Real-time tracking of Trace and Retrace signals ......................................................... 61

3.2 Evaluation of the semipermeability of resurfaced lipid layer – diffusion study ....................... 68

3.3 Mechanical loading tests ....................................................................................................... 74

3.4 Removal of surface lipids - Delipidization ............................................................................. 80

3.5 lipid resurfacing - Relipidization ........................................................................................... 84

CHAPTER 4: APPROACH AND METHODOLOGY ................................................................ 86

4.1 Background .......................................................................................................................... 86

4.2 Critical Arguments and Testing of Hypothesis ....................................................................... 87

4.3 Experimental Study .............................................................................................................. 93

4.3.1 Sample Preparation .................................................................................................... 93

4.3.2 Delipidization Process - Surface Lipid Removal ......................................................... 93

4.3.3 Relipidization Process (Incubation in lipid-filled environment) ................................... 95

4.3.3.1 Case 1 ....................................................................................................................... 96

4.3.3.2 Case 2 ....................................................................................................................... 96

4.3.3.3 Case 3 ....................................................................................................................... 97

4.3.4 Atomic Force Microscopy (AFM) .............................................................................. 99

4.3.4.1 Principles of Operation .......................................................................................... 101

4.3.4.2 Mode of Operation .................................................................................................. 104

4.3.4.3 Atomic Force Microscopy (AFM) Tip/Stylus ........................................................... 107

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4.3.4.4 Imaging and Force Spectroscopy ............................................................................. 114

4.3.5 Confocal Microscopy (COFM) ................................................................................. 123

4.3.5.1 Nile Red Staining .................................................................................................... 126

4.3.6 Raman Spectroscopy ................................................................................................ 126

4.3.7 Magnetic Resonance Imaging ................................................................................... 130

4.3.8 Computational Analysis ........................................................................................... 135

4.3.9 Mechanical Compression Test .................................................................................. 138

CHAPTER 5: MICROSCOPIC AND CHEMICAL CHARACTERIZATION OF THE SURFACES OF

NORMAL, DELIPIDIZED, AND RELIPIDIZED ARTICULAR CARTILAGE ..................... 147

5.1 Introduction ........................................................................................................................ 147

5.2 Materials and Methods ........................................................................................................ 148

5.2.1 Atomic Force Microscopy Samples .......................................................................... 148

5.2.2 Confocal Microscopy Samples ................................................................................. 149

5.2.3 Raman Spectroscopy Samples .................................................................................. 149

5.2.4 AFM Imaging and Force spectroscopy ..................................................................... 150

5.2.5 Surface Lipid Removal (Delipidization) ................................................................... 151

5.2.6 Relipidization Process (Incubation in lipid-filled environment) ................................. 151

5.2.7 Confocal Microscopy (COFM) ................................................................................. 152

5.3 Analyses of AFM Imaging Results ...................................................................................... 152

5.4 Analyses of Nano-indentation Results (Force Curves) ......................................................... 153

5.5 Statistical analysis .............................................................................................................. 154

5.6 Results and Observations .................................................................................................... 155

5.6.1 Confocal Microscopy and AFM Imaging .................................................................. 155

5.6.2 Raman Spectrocopy ................................................................................................. 165

5.6.3 AFM Analysis and Force Spectroscopy .................................................................... 168

5.7 Conclusion ......................................................................................................................... 174

CHAPTER 6: EVALUATION OF THE FUNCTIONALITY OF ARTIFICIALLY LAID LIPID

MEMBRANE FOR ARTICULAR CARTILAGE RESURFACING ......................................... 175

6.1 Introduction ........................................................................................................................ 175

6.2 Pertinent Theory ................................................................................................................. 177


6.3.1 Articular cartilage samples ....................................................................................... 182

6.3.2 Deuterium oxide-Phosphate Buffered Saline (D2O-PBS) Solution ............................. 183

6.3.3 Atomic Force Microscope (AFM) Imaging ............................................................... 183

6.3.4 Magnetic Resonance Imaging (MRI) ........................................................................ 184

6.3.5 Determination of Apparent Diffusion Coefficients .................................................... 186

6.3.6 Statistical Analysis ................................................................................................... 187


6.5 Conclusion ......................................................................................................................... 198

CHAPTER 7: ASSESSMENT OF THE MECHANICAL INTEGRITY/ PHYSIOLOGICAL FUNCTION

OF RESURFACED ARTICULAR CARTILAGE ..................................................................... 199

7.1 Introduction ........................................................................................................................ 199


7.2.1 Articulr cartilage samples ......................................................................................... 201

7.2.2 Derivation of the energy components ........................................................................ 202

7.2.3 Statistical Analyses .................................................................................................. 205


7.4 Conclusion ......................................................................................................................... 218

CHAPTER 8: DISCUSSION AND CONCLUSIONS ................................................................ 219

REFERENCES ........................................................................................................................... .237

Appendix A …………………………………………………………………………………………….270

Appendix B …………………………………………………………………………………………...2709

Appendix C …... ……………………………………………………………………………………297

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List of Figures

Figure 2.1 A physical model of representing the physical interactions and structural coupling between the

components of the load bearing units of articular cartilage before loading (Balloons and

strings, (Broom and Marra, 1985). ................................................................................... 15

Figure 2.2 The balloon-string analogue suggested by Broom and Marra (1985). The 3-D string meshwork

and the air-filled balloons represent the collagen fibre network and the fluid swollen

proteoglycans of articular cartilage after loading. The flat plank sits on the articular surface in

contact with the load........................................................................................................ 16

Figure 2.3 Full picture showing an osteoarthritic articular cartilage (Garg, 2012). ............................. 20

Figure 2.4 Representation of the zonal variation and distribution of articular cartilage matrix constituents

from the surface and subchondral bone (Jadin et al., 2007). .............................................. 21

Figure 2.5 Schematic view of the articulating joint, the expansion shows the zonal architecture of

articular cartilage (Crouch, 1985)..................................................................................... 24

Figure 2.6 (A) Schematic diagram showing the chondrocyte distribution and (B) the structure of collagen

fibre network in the distinct zones (Buckwalter et al., 1994). ............................................ 25

Figure 2.7 Zonal architecture of articular cartilage (Jeffrey and Watt, 2003). .................................... 25

Figure 2.8 Schematic representation of cartilage extracellular matrix showing the proteoglycan aggregate

and aggrecan molecule (Pearle, et al., 2005). ................................................................... 26

Figure 2.9 Molecular structure of chondroitin sulphate monomer chain (Muir, 1978). ....................... 28

Figure 2.10 Molecular structure of keratan sulphate monomer chain (Muir, 1978). ........................... 28

Figure 2.11 Hyaluronic acid, a heteropolysaccharide with several thousand monomer units of N-acetyl

glucosamine and glucuronic acid formed (Stern, 2004). ................................................... 29

Figure 2.12 Electron micrograph of the oligolamella layer of SAPL adsorbed to the pleural epithelium,

which is similar to the surface of cartilage in vivo (Hills, 2000). ....................................... 32

Figure 2.13 Electrostatic bonding of the quaternary positive ions from the SAPL molecules with excess

negative charges from proteoglycan molecules on the articular surface (Hills, 2000). ....... 33

Figure 2.14 (a) and (b) Chemical structure of triacylglycerol R, R1, R2, and R3 denote aliphatic chain

hydrocarbons (Lehninger, et al., 2005). ............................................................................ 38

Figure 2.15 Chemical structure of Palmitic acid, a saturated and unbranched fatty acid (Lehninger, et al.,

2005). ............................................................................................................................. 39

Figure 2.16 Chemical structure of Oleic acid; an unsaturated and unbranched fatty acid (Lehninger, et al.,

2005). ............................................................................................................................. 39

Figure 2.17 Chemical structure of Cholesterol; an unsaturated and branched fatty acid (Lehninger, et al.,

2005). ............................................................................................................................. 40

Figure 2.18 A lipid bilayer structure, showing the hydrophilic head and hydrophobic tails (Inex

Pharmaceutical Corporation). .......................................................................................... 41

Figure 2.19 General chemical structure of phosphatidylcholines (Jump, 2002). ................................. 43

Figure 2.20 General structure of glycerophospholipids (Lehninger, et al., 2005). .............................. 43

Figure 2.21 A schematic representation of liposome structure (Britannica, 2007). ............................. 45

Figure 3.1 2-D AFM images of the surface of bovine humeral head articular cartilage (A) and (B) are

height images (Scale bars, 2 μm; full gray ranges: 1000 nm (A) and (B) 600 nm) (Jurvelin, et

al., 1996). ........................................................................................................................ 53

Figure 3.2 2-D topographical AFM image of the surface normal healthy adult pig articular cartilage. Full

scan size 30 x 30 µm; full grey range 1700 nm (Kumar et al., 2001). ................................ 54

Figure 3.3 AFM height images of the surfaces of bovine cartilage in synovial fluid (a) before and (b)

after washing with PBS. (Image size: 5 x 5 µm area) (Crockett et al., 2005). .................... 54

Figure 3.4 2-D topographical AFM images of the surface of fresh bovine articular cartilage: (a) 40 µm

scan with 1 µm height scale, (b) 20 µm scan with 2 µm height scale (Grant et al., 2006)... 55

Figure 3.5 2-D AFM images of the surface of fresh bovine articular cartilage (a) Topographical and (b)

Deflection images (scan size: 8 x 8 µm) acquired with a rectangular cantilever ................. 56

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Figure 3.6 Schematic view of a set of triangular and rectangular AFM cantilever carrying silicon nitride

tips. These cantilevers have extremely low spring constants, thus suitable for imaging in air

and liquid environments both with contact and tapping mode (Bruker AFM Probes, Madison,

WI, USA). ....................................................................................................................... 58

Figure 3.7 (a) and (b) Screen shots of the approach profiles of the AFM tip on the surface of cartilage

during two landing processes. .......................................................................................... 60

Figure 3.8 Schematic representation of the imaging process of normal cartilage with the AFM, the

similarity between the trace and retrace signals shows that the AFM tip is producing a good/

high resolution image of cartilage (a) beginning and (b) end of scan. ................................ 63

Figure 3.9 2-D (a) Height or topographical and (b) Deflection AFM images of the surface of normal

intact articular cartilage (frame size: 8 µm by 8 µm) acquired for the Forward scan; (c) Height

or topographical and (d) Deflection images for the backward scan. The images (a) and (c); (b)

and (d) look similar as expected from the oscillograph shown in Figures 3.8 (a) and (b). Note:

the above images were obtained with V-shaped cantilevers. ............................................. 64

Figure 3.10 High resolution (5 µm by 5 µm) 2-D topographical images of the surface of fresh bovine

cartilage obtained with V-shaped cantilevers using the optimized scanning parameters (a)

forward scan and (b) backward scan. The forward and backward scans are almost identical,

proving the accuracy of the scanning process. .................................................................. 65

Figure 3.11 2-D (a) Topographical and (b) Deflection images of the surface of normal intact articular

cartilage obtained with AFM (frame size: 8 µm by 8 µm). This is compared with the images

previously presented in the Figure 3.4, which was assumed to be acquired with triangular

cantilevers. This further supports the argument that triangular cantilevers are more suitable for

imaging cartilage surface. ................................................................................................ 67

Figure 3.12 (a) and (b) The articular surface is not parallel to the horizontal X-axis. The images would

have to realigned parallel to the X-axis using the custom-built MATLAB® code before the

ROI is selected. ............................................................................................................... 72

Figure 3.13 Screen capture of the GUI for realigning the inclined images shown in Figure 3.12. After

rotating the image with the AS parallel to X-axis, the concentration of H2O at any given

position and time in the tissue is calculated from the MR image. ...................................... 72

Figure 3.14 (a) and (b) Well-aligned MRI images of normal intact cartilage, where the articular surface

approximately parallel to the horizontal X-axis. ............................................................... 73

Figure 3.15 Screen capture of the GUI for calculating the concentration of H2O at any given position and

time in the tissue and the apparent diffusion coefficient of H2O in the matrix from the MR

image in Figure 3.13. ....................................................................................................... 73

Figure 3.16 Load-displacement curve for a well-placed indenter sitting parallel to the articular surface.

The curve is smooth showing uniform distribution of load................................................ 76

Figure 3.17 Energy diagram derived from a typical load-displacement curve, where SE and RE represent

strain energy and residual energy respectively. ................................................................. 77

Figure 3.18 (a) and (b) Load-displacement curves for a badly-placed indenter not sitting parallel to the

articular surface. The curve is not smooth with uniform distribution of load. .................... 78

Figure 3.19 Topographical (a, c, e) and deflection (b, d, f) 2-D Images of articular cartilage surface

(Frame size: 8 by 8µm). Normal articular surface (a, b); after 3min delipidization in

chloroform/methanol (c, d); and after 21min delipidization in chloroform/methanol (e, f). 81

Figure 3.20 Variation of surface lipid lost (height of SAL, nm) with time following delipidization with

chloroform:methanol (2:1). Normal intact (group 1); 3 min delipidization (group 2); 15 min

delipidization (group 3); and 21 min delipidization (group 4). .......................................... 82

Figure 4.1 A conceptualized flowchart for the research, showing the different steps followed to achieving

the objectives of this thesis. ............................................................................................. 92

Figure 4.2 The NT-MDT atomic force microscope and video camera placed in a sound proof

compartment to minimize external vibration .................................................................. 100

Figure 4.3 A schematic of AFM operation (Peter, Atomic Force Microscopy). ............................... 102

Figure 4.4 2-D topographical image of the surface of Teflon (Frame size: 8 by 8µm) obtained with the

AFM. ............................................................................................................................ 103

Figure 4.5 2-D deflection image of the surface of Teflon (Frame size: 8 by 8µm) obtained with the AFM.

..................................................................................................................................... 103

Figure 4.6 The SMENA head of the NT-MDT SPM for scanning in liquid environment. ................ 106

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Figure 4.7 SMENA head for measurements in a liquid environment (NT-MDT). ............................ 106

Figure 4.8 Schematic view of scanning in a drop of liquid with SMENA head (NT-MDT). ............. 107

Figure 4.9 Schematic view of an AFM tip captured with a focused ion beam (FIB). ........................ 108

Figure 4.10 Schematic view of an AFM tip that is carried by a flat cantilever captured with a scanning

electron microscopy (SEM). .......................................................................................... 109

Figure 4.11 Schematic top view of an AFM triangular cantilever (Butt, et al., 2005). ...................... 111

Figure 4.12 Schematic view of an AFM rectangular cantilever for contact and tapping modes (NT-MDT,

Moscow, Russia). .......................................................................................................... 112

Figure 4.13 Schematic view of an AFM triangular AFM cantilever carrying silicon nitride tip suitable for

contact and tapping modes (Advanced Integrated Scanning Tools for Nanotechnology). . 113

Figure 4.14 NT-MDT P47 Solver Pro atomic force microscope (AFM) with specimen mounted before

measurements with the SMENA® head. ......................................................................... 115

Figure 4.15 Articular cartilage sample mounted on the scanner head of the NT-MDT P47 Solver Pro

before measurements. .................................................................................................... 116

Figure 4.16 The versatile SMENA® head of the NT-MDT P47 Solver Pro for imaging biological samples

in liquid medium. .......................................................................................................... 117

Figure 4.17 Schematic representation of the imaging process of normal cartilage with the AFM. .... 118

Figure 4.18 Single point force-distance curve obtained with an AFM tip in contact mode. .............. 120

Figure 4.19 Schematic representation of a single point force-distance curve showing several stages

involved in force measurement with an AFM tip. The probe is brought into and out of contact

by a piezoelectric translator (carrying the chip to where the cantilever is attached) with the

specimen fixed to a point (Green, et al., 2002). .............................................................. 122

Figure 4.20 Schematic diagram illustrating the principal light pathways in a basic confocal microscope

configuration (Nikkon Microscopy). .............................................................................. 124

Figure 4.21 Schematic of a Leica SP5 confocal microscope (Leica Microsystems, Germany) available at

the cell imaging facility of the Institute of Health and Biomedical Innovation (IHBI),

Queensland University of Technology (QUT). ............................................................... 125

Figure 4.22 Flowchart of Raman spectroscopic measurement of a sample. ...................................... 127

Figure 4.23 Raman microscope (InVia Renishaw) that is available at QUT (University of Nebraska -

Lincoln, USA). .............................................................................................................. 129

Figure 4.24 A 4.7 Tesla Magnetic Resonance Imaging (Bruker Avance 200 MHz NMR micro

imaging/spectrometer, Germany) facility at Queensland University of Technology (QUT).134

Figure 4.25 A graphical user interface (GUI) developed with MATLAB® for computing the apparent

diffusion coefficient of cartilage matrix from magnetic resonance imaging data .............. 137

Figure 4.26 Purpose-built 1-D consolidometer used for quasi-static compression tests and its parts. 141

Figure 4.27 High sensitivity material testing machine (Instron), with the consolidometer carrying the

cartilage specimen mounted on. The quasi-static compression test was conducted on this rig.

..................................................................................................................................... 143

Figure 4.28 Computer set up with Bluehill® software installed for real-time data collection of data to the

compression tests. ......................................................................................................... 144

Figure 4.29 Typical load-displacement curve obtained for normal intact cartilage sample from the Instron

machine. ....................................................................................................................... 145

Figure 5.1 Schematic flowchart of AFM imaging, confocal microscopy, and Raman spectroscopy for

normal intact, delipidized, and relipidized cartilage specimens. ...................................... 150

Figure 5.2 Screen capture of the WSxM® software used to generate a 3D image from a 2D image

obtained with the Nova® program. ................................................................................. 154

Figure 5.3 (a) Cross-sectional view of a normal intact AS obtained with a confocal microscope, (b) 2-D

topographical image of the surface of normal intact articular cartilage (5 µm by 5µm), (c) 3-D

topographical image of normal articular cartilage after image processing (length (X) and

breadth (Y) of the scanned area, and the average peak height of SAL (Z)), (Yusuf, et al.,

2012). ........................................................................................................................... 156

Figure 5.4 (a) Cross-sectional view of an articular surface following the partial removal of a lipid layer

obtained with a COFM, (b) 2-D topographical image of the surface of delipidized articular

cartilage (5 µm by 5µm), (c) 3-D topographical image of the surface of delipidized articular

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cartilage after image processing (length (X) and breadth (Y) of the scanned area, and the

average peak height of SAL (Z)), (Yusuf, et al., 2012). .................................................. 157

Figure 5.5 (a) Cross-sectional view of AS following incubation in POPC for 24 hours at 37oC obtained

with a COFM, (b) 2-D topographical image of the surface of relipidized articular cartilage in

POPC for 24 hours at 37oC (5 µm by 5µm), (c) 3-D topographical image of the surface of

relipidized articular cartilage in POPC for 24 hours at 37oC after image processing (length (X)

and breadth (Y) of the scanned area, and the average peak height of SAL (Z)), (Yusuf, et al.,

2012). ........................................................................................................................... 159

Figure 5.6 (a) Cross-sectional view of articular cartilage following incubation in DPPC for 24 hours at

43oC obtained with a COFM, (b) 2-D topographical image of the surface of relipidized

articular cartilage in DPPC for 24 hours at 43oC (5 µm by 5µm), (c) 3-D topographical image

of the surface of relipidized articular cartilage in DPPC for 24 hours at 43oC after image

processing (length (X) and breadth (Y) of the scanned area, and the average peak height of

SAL (Z)), (Yusuf, et al., 2012). ..................................................................................... 160

Figure 5.7 Schematic representation of POPC-bilayers and DPPC molecules formed on the surface of

degraded cartilage after relipidization, (a) showing wavelike structure deposits of POPC on

the articular surface, and (b) showing particle-like deposits of DPPC on the articular surface

(Yusuf, et al., 2012)....................................................................................................... 162

Figure 5.8 (a) 2-D topographical image of the surface of relipidized articular cartilage in a mixture

containing all five SAPL for 24 hours at 37oC (5 µm by 5µm), (b) 3-D topographical image of

the surface of relipidized articular cartilage in a mixture containing all five SAPLs for 24

hours at 37oC after image processing (length (X) and breadth (Y) of the scanned area, and the

average peak height of SAL (Z)). ................................................................................... 164

Figure 5.9 Raman spectrum of bovine cartilage collected on a Raman microscopy system ( = 785 nm).

Band assignments for the spectra are presented in Table C1 in the appendix section (Appendix

C). ................................................................................................................................ 165

Figure 5.10 Comparison of the Raman spectral in the C-H stretching mode region acquired for cartilage

specimens with normal intact, delipidized, and relipidized in POPC, DPPC, and full SAPL

mix. The figure reveals that there is a measurable difference between the chemical properties

of all the sample groups tested. ...................................................................................... 166

Figure 5.11 Comparison of the C-H stretching mode region peak area for cartilage specimens with

normal intact, delipidized, and relipidized in POPC, DPPC, full SAPL mix, saline solution

(control). ....................................................................................................................... 167

Figure 5.12 Averaged force-indentation curve for normal intact, delipidized and relipidized cartilage in

POPC, DPPC, and full SAPL mix, showing the variation of the mechanical properties of the

articular surface under the three surface conditions. ....................................................... 172

Figure 5.13 A typical force-indentation curve for normal intact cartilage used for estimating the average

elastic strain energy of the surface amorphous layer, a measure of the layer’s resistance to

AFM tip penetration (Yusuf, et al., 2012). ..................................................................... 173

Figure 6.1 Schematic flowchart of specimen grouping, AFM imaging, magnetic resonance

measurements, and computational analysis for normal intact, delipidized, and relipidized

cartilage. ....................................................................................................................... 181

Figure 6.2 3D topographical image of normal healthy articular cartilage after image processing, showing

the nanostructural arrangement of the surface amorphous layer with several peaks and troughs

(length (X) and breadth (Y) of the scanned area, and average peak height of SAL (Z)).... 189

Figure 6.3 3D topographical image of the surface of delipidized articular cartilage after image

processing, showing the loss of the membranous overlay (surface amorphous layer) of the

articular surface (length (X) and breadth (Y) of the scanned area, and average peak height of

SAL (Z)). ...................................................................................................................... 190

Figure 6.4 3-D topographical image of the surface of relipidized articular cartilage in POPC after image

processing, showing partially restored lamella layer of lipids slightly similar to normal

articular surface (length (X) and breadth (Y) of the scanned area, and average peak height of

SAL (Z)). ...................................................................................................................... 191

Figure 6.5 3-D topographical image of the surface of relipidized articular cartilage in DPPC after image

processing, showing almost featureless structure of the articular surface when compared with

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normal intact articular surface (length (X) and breadth (Y) of the scanned area, and average

peak height of SAL (Z)). ............................................................................................... 192

Figure 6.6 3-D topographical image of the surface of relipidized articular cartilage in complete SAPL

mixture after image processing (length (X) and breadth (Y) of the scanned area, and average

peak height of SAL (Z)). ............................................................................................... 193

Figure 6.7 Typical 2D multi-spin multi-echo (MSME) images of cartilage specimens immersed in D2O –

PBS solution acquired at different times......................................................................... 196

Figure 6.8 Representative depth-wise concentration profiles for the MSME images shown in Figure 6.8

at different time steps obtained using the analysis with a purpose-built computational scheme

developed in MATLAB®. .............................................................................................. 197

Figure 7.1 Energy diagram derived from a typical load-displacement curve obtained for normal intact

cartilage sample subjected to loading and unloading test on the Instron machine, where SE and

RE represent strain energy and residual energy respectively. .......................................... 203

Figure 7.2 Schematic flow chart of the loading sequence followed for the normal intact, delipidized, and

relipidized articular cartilage samples. ........................................................................... 204

Figure 7.3 Screenshot of the G*Power statistical power analysis software used to determine the influence

of the relatively small sample size used in this study. ..................................................... 206

Figure 7.4 Strain energy plotted for articular cartilage specimens with normal intact, delipidized, and

relipidized (POPC, DPPC and complete SAPL mix) surfaces. ........................................ 210

Figure 7.5 Complementary energy plotted for articular cartilage specimens with normal intact,

delipidized, and relipidized (POPC, DPPC and complete SAPL mix) surfaces. ............... 211

Figure 7.6 Released energy plotted for articular cartilage specimens with normal intact, delipidized, and


Figure 7.7 Residual energy plotted for articular cartilage specimens with normal intact, delipidized, and


Figure 7.8 Energy ratio plotted for articular cartilage specimens with normal intact, delipidized, POPC,

DPPC and complete SAPL mix surfaces. ....................................................................... 214

Figure 7.9 Complementary energy versus strain energy for articular cartilage samples with normal intact,

delipidized, relipidized surfaces. .................................................................................... 215

Figure 7.10 Released energy versus strain energy for articular cartilage samples with normal intact,


Figure 7.11 Residual energy versus strain energy for articular cartilage samples with normal intact,


Figure 7.12 Residual energy versus released energy for articular cartilage samples with normal intact,


Figure 8.1 Reconstructed 3-D AFM images of articular cartilage with normal intact, delipidized, POPC-

treated, DPPC-treated, and complete SAPL mix treated surfaces. ................................... 225

Figure 8.2 Cross-linked phospholipid network structure present in cartilage specimens with normal intact

and complete SAPL mixture-treated surfaces. ................................................................ 226

Figure 8.3 Wave-like phospholipid structure present in cartilage samples with DPPC and POPC treated

surfaces. ........................................................................................................................ 227

Figure 8.4 A schematic scale showing the change in permeability of articular cartilage with different

surface conditions. Relipdization in synthetic DPPC, POPC and full SAPL mix resulted in the

transformation of the delipidized cartilage sample surfaces from a highly undesirable

permeable condition to a more effective surface with lower permeabililty and better

semipermeabilty characteristics. .................................................................................... 230

Figure 8.5 Load-displacement diagram for linear elastic material subjected to compressive loading test.

The complementary energy and the elastic strain energy are equal; therefore the energy ratio

for linear elastic materials is equal to one. ...................................................................... 234

xx

List of Tables

Table 4.1Surfactant species in bovine joint (Chen, et al., 2007b). ..................................................... 98

Table 4.2 The molecular weights (MW) and diffusion coefficients (D) for several solutes in cartilage

matrix, where IGF-1 is Insulin-like growth factor 1 (IGF-1), PFG is patella femoral groove

and FH is femoral head (Mauck, et al., 2003). ................................................................ 132

Table 5.1 Variation of height and elastic strain energy of the surface amorphous layer for normal,

delipidized, and relipidized articular cartilage. ............................................................... 170

Table 6.1 Average apparent diffusion coefficients for cartilage samples with normal intact, delipidized

and relipidized surfaces. ................................................................................................ 195

Table 7.1 Variation of total strain energy, elastic energy released, and residual energy of cartilage

matrices with normal intact, delipidized, and relipidized surfaces. .................................. 209

List of Abbreviations

Acronym Meaning

AC Articular cartilage

ADC Apparent diffusion coefficient

AI Artificial intelligence

ANOVA Analysis of variance

AS Articular surface

AFM Atomic force microscopy

AIHW Australian Institute of Health and Welfare

CCD Charge-coupled device

CE Complementary energy

COFM Confocal microscopy

CT Computer tomography

dGEMRIC delayed Gadolinium Enhanced MR imaging of

Cartilage

DLPC Dilinoleoyl-phosphatidylcholine

DPPC Dipalmitoyl-phosphatidylcholine

FCD Fixed charge density

FOV Field of view

FT-IR Fourier Transform Infrared spectroscopy

GAG Glycosaminoglycan

GUI Graphical user interface

HA Hyaluronic acid

HE Hysteresis energy

xxii

Acronym Meaning

HEPP

Hydrostatic pore pressure

HPLC High-performance liquid chromatography

IR Infrared

LSF Least square fit

MRI Magnetic resonance imaging

MSME Multi-spin multi-echo

NIR Near infrared

NMR Nuclear magnetic resonance

NSAIDS Non-steroidal ant-inflammatory drugs

OA Osteoarthritis

PBS Phosphate buffered saline

PC Phosphatidylcholine

PG Proteoglycan

PLPC Palmitoyl-linoleoylphosphatidylcholine

POPC

Palmitoyl-oleoyl-phosphatidylcholine

QA Quaternary ammonium

RE Residual energy

RF Radiofrequency

ROI Region of interest

RS Raman spectroscopy

RTM Radioactive tracer method

SAL Surface amorphous layer

SAPL Surface-active phospholipids

Acronym

Meaning

SE Strain energy

SEM Scanning electron microscopy

SPM Scanning probe microscope

SPC Saturated phosphatidylcholine

STEM Scanning transmission electron microscopy

TEM Transmission electron microscopy

TR Repetition time

USPC Unsaturated phosphatidylcholines

xxiv

Statement of Original Authorship

The work contained in this thesis has not been previously submitted to meet requirements for

an award at this or any other higher education institution. To the best of my knowledge and

belief, the thesis contains no material previously published or written by another person

except where due reference is made.

Signature: _________________________

23-01-2013

Date: _________________________

1

Chapter 1: INTRODUCTION

This thesis focuses on the degradation that is initiated from the articular surface,

which results mostly in the loss of the surface lipids, also, known surface-active

phospholipids (SAPL), a condition that is often encountered in the early stages of

tissue degeneration. The attrition of the SAPL layer impairs lubrication in the joint,

resulting in detrimental consequences on joint movement and human activities. The

capacity to replace the SAPL artificially might improve the quality of life of people

suffering from this condition. Articular cartilage is a complex structured fluid-

saturated biological gel which performs the physiological functions of load

bearing/processing, semipermeability and lubrication in the mammalian joints. Its

degradation leads to debilitating joint conditions such as osteoarthritis, which is

characterized by progressive loss of cartilage matrix constituents, namely, depletion

of the osmotically active proteoglycans, disruption of collagen fibre architecture, and

more important to this research its lipid content (both surface and intramatrix lipids).

Cartilage degeneration can initiate from the surface, within the matrix, or from the

bone end. The scope this thesis does not include studying the collagen, proteoglycans

or fluid-related consequences, instead this research focus on the solid surfactant,

namely surface-active phospholipids (SAPL) that covers the articular surface to

facilitate tribological function.

Currently, the major commercially available non-surgical/non-invasive remedies for

osteoarthritis are: visco-supplements or chondroprotective agents such as hyaluronic

2

acid (HA) injection, and orally administered glucosamine and chondroitin sulphate.

It has been reported in the literature that these agents are generally incapable of

providing cartilage with the lost SAPL. For example, glucosamine and chondroitin

sulphates replenish the intramatrix constituents (proteoglycans and collagen), while

hyaluronic acid was initially believed to provide lubrication for the synovial joint.

More recent studies have established that HA is not the boundary lubricant in the

joint, rather the SAPL coating on the articular surface, which posses highly desirable

lubricating properties for efficient joint function. HA injection is costly, ineffective,

and painful, thereby discouraging medical practitioners from applying it despite its

approval for treatment and management of osteoarthritis. Furthermore, these non-

surgical remedies are palliative with short term potency.

It is argued that this research will create a framework that can be used to develop

lipid-based formula which will deliver a new functional contact layer that will

provide more effective lubrication for joints. This will ultimately improve the quality

of life of patients by enabling them to participate in exercises and physical activities

that would improve their health conditions, thereby preventing obesity and other risk

factors. This thesis extends the study of joint lipids associated with the articular

surface of cartilage, with emphasis on their structural and functional properties in

both the intact (healthy) and degraded (osteoarthritic) conditions. The expected

outcomes are potentially significant in advancing knowledge on the characteristics of

the surface amorphous lipid layer (SAL), thereby contributing to the potential

benefits of relaying this layer following osteoarthritic degeneration. More

specifically, it is envisaged that the knowledge created would provide the scientific

3

3

framework that could be adapted for customizing synthetic saturated and unsaturated

lipids for joint management and treatment.

The basic hypothesis of this research is that the surface-based degradation of

articular cartilage can be corrected, namely, that the surface layer of lipids lost as a

result of degeneration can be “relaid” or “resurfaced” following an appropriate

scientific framework. A major objective of this thesis was to evaluate this hypothesis.

Several experimental analyses were conducted, including nano-, micro-, and

macroscopic and chemical characterization of natural and artificially engineered lipid

layers. Atomic force microscopy (AFM), confocal microscopy (COFM), Raman

spectroscopy, magnetic resonance imaging (MRI), and analyses involving image

processing and applied mechanics, were employed in the experiments, where the

outcomes of these were used to validate the proposition and the feasibility of the

hypothesis.

The resurfacing process involved the deposition of single lipid components

(palmitoyl-oleoyl-phosphatidylcholine, POPC and dipalmitoyl-phosphatidylcholine,

DPPC) and complete joint SAPL mixture on the surfaces of lipid-depleted cartilage.

The single lipid species (DPPC and POPC) used in this exploratory study were

specifically chosen to simulate the role/contribution of each component in the joint

lipid mixture. This was based on the argument that the potential application of

synthetic phospholipids as a remedy for degenerated joint conditions may not

necessarily require all the lipid components found in the joint. Since DPPC is the

most economical of all the joint lipid species, a feasible and cost effective alternative

4

may be the application of lipid mixtures containing fewer species than the complete

joint lipid mixture in order to achieve a significant level of resurfacing.

Based on comparative analyses of the experimental results, it has been demonstrated

in thesis that the newly laid surfaces of synthetic lipids can provide delipidized

cartilage with the following characteristics:

surface structure reorganization,

semipermeability modification and

load processing improvement.

This research has produced outputs in both journal and conference papers, namely:

Journal papers

1. Yusuf K. Q., Momot K. I., Wellard R. M., and A. Oloyede A. (2013). A

magnetic resonance imaging study of diffusion through the surface of

normal, delipidized, and relipidized articular cartilage. Journal of

Materials Science Materials in Medicine, (accepted for publication).

2. Zenon Pawlak, Wieslaw Urbaniak, Adam Gadomski, Kehinde Q. Yusuf,

Isaac O. Afara and Adekunle Oloyede (2012). The role of lamellate

phospholipid bilayers in lubrication of joints. Acta of Bioengineering and

Biomechanics, (accepted for publication).

5

5

3. Yusuf K. Q., Pawlak Z., Motta N., and Oloyede A. (2011). A

microanalytical study of the surfaces of normal, delipidized, and

artificially “resurfaced” articular cartilage. Connective Tissue Research,

53(3):236-245. doi:doi:10.3109/03008207.2011.630764.

4. Pawlak Z., Petelska A. D, Urbaniak W., Yusuf K. Q., and Oloyede A.

(2012). Relationship between Wettability and Lubrication

Characteristics of the Surfaces of Contacting Phospholipids-Based

Membranes. Cell Biochemistry and Biophysics, 1-11. DOI 10.1007/s12013-

012-9437-z

5. Duong Q.T., Yusuf K.Q., Oloyede, A. (2011). 3D rendering of

proteoglycan distribution in articular cartilage: preliminary study for

computational modeling of cartilage structure. Vietnam Journal of Science

and Technology (under review).

Refereed full length conference paper

1. Zenon Pawlak, Wieslaw Urbaniak, Adam Gadomski, Kehinde Q. Yusuf,

Isaac O. Afara and Adekunle Oloyede (2012). The role of lamellate

phospholipid bilayers in lubrication of joints. The 32th All-Polish

Tribology conference, Autumnal school of Tribology 2012. “Tribology

Nearer Practice”. Department of Mechanical Engineering, Institute of

Machine Design and Operation, Wroclaw University of Technology, Poland;

6

Polish Tribological Society; and Section of the Fundamentals of Operation

and Maintenance, Polish Academy of Science, Poland (under review).

2. Yusuf K.Q., Gudimetla P., Pawlak Z., Oloyede, A. (2011). Preliminary

Characterisation of the Surface of Cartilage Following Exposure to

Saturated and Unsaturated Synthetic Lipids. In The First International

Postgraduate Conference on Engineering, Designing and developing the

Built Environment for Sustainable Wellbeing, Queensland University of

Technology, Brisbane, Qld, pp. 347-351.

7

Chapter 2: CRITICIAL REVIEW/APPRAISAL

OF THE LITERATURE

2.1 INTRODUCTION

Mobility plays a significant role in the quality of life of humans and animals as their

survival depends on this mechanism. We need to walk, sit, run and move from one

place to the other to carry out our day to day activities in order to earn a living, to

maintain our quality of life and contribute to the gross domestic product of the

country. This movement is made possible by the mammalian joint system, which

relies on articular cartilage and bone to perform its function. In the joints, articular

cartilage prevents bone-to-bone contact, playing a key role in load bearing and joint

lubrication. Due to these specialized functions of cartilage, it is inevitable that its

malfunction or degeneration would deleteriously affect the mammalian mobility and

hence quality of life.

Articular cartilage is susceptible to wear and tear due to ageing, traumatic load and

disease. Unlike other connective tissues such as kidney and liver, articular cartilage

has limited ability to repair itself if damaged because it does not have nerve supply

(aneural) and blood vessels (avascular). The most common degenerative condition of

articular cartilage is osteoarthritis. This disease is often characterised by progressive

loss of cartilage matrix constituents, namely, disruption of the collagen fibre

meshwork (Broom, 1986a), depletion of the swelling components or proteoglycans

(Mow et al., 1992a; Oloyede and Broom, 1993), and more importantly to this

research its surface amorphous layer that is rich in surface-active phospholipids

8

(SAPL) (Ballantine and Stachowiak, 2002; Hills and Monds, 1998; Oloyede et al.,

2004a, 2004b; Sarma et al., 2001). While lipids are present both on the surface and

within the cartilage matrix, the scope of this thesis is limited to those in the surface

layer. Articular cartilage degeneration, as is manifested in osteoarthritis and other

related joint conditions is a common cause of disability, which is a major

impediment to daily living and mobility especially during aging (Aigner et al., 2007;

Aigner et al., 2004; Hollander et al., 1995; Inerot et al., 1978; Lane et al., 1993). It

affects more than 16 % of Australians mainly the over 55 year olds, leading to

disability in almost 5 % of the sufferers (AIHW, 2010). The cost of treatment and

management (direct and indirect cost) is close to AUD $23.9 billion per annum

(Access Economics, 2007).

The current treatment and management options for osteoarthritis include anti-

inflammatory drugs, orally administered or injected glucosamine and chondroitin

sulphate, hyaluronic acid injections, balms, chondrocyte culture, cartilage

transplantation, partial and total joint replacements. Apart from joint replacement

which last for about nine to fifteen years, before revision surgery is required for

affected patients, the other treatment options are palliative, having short term

effectiveness, usually not more than 3 months. The use of synthetic lipid-based

treatment might offer a more lasting solution for the treatment of this debilitating

condition. This thesis aims to explore the possibility of resurfacing degraded

cartilage with synthetic surface-active phospholipids by creating a new surface

membrane, which is structurally and functionally viable and thus able to restore

the functions of this important membrane. It is strongly believed that the knowledge

gained in this research will provide a potential treatment and prevention of

osteoarthritis, thus reducing the medical cost of treatment and management of joint

conditions in general.

9

2.2 MOTIVATION FOR THIS THESIS - TREATMENT

OF OSTEOARTHRITIS

Generally the aims of treating joint conditions such as osteoarthritis are to reduce

pain, recover and/or optimize joint function and achieve an overall improvement in

patients’ quality of life. There are many treatment options for osteoarthritis. These

can be broadly classified into two: surgical and non-surgical treatments (Brandt et

al., 2003b).

The non-surgical treatment option offers non-invasive procedures for the treatment

of osteoarthritis. It can be divided into non-pharmacological, and local and systemic

drug therapies (Brandt, et al., 2003b). The non-pharmacological interventions

include patient education, exercise, diet, avoidance of adverse mechanical factors

(weight loss, appropriate footwear, walking aids and appliances etc.). While the local

and systemic drug therapies include creams/gels, balms, non-steroidal ant-

inflammatory drugs (NSAIDS), capsaicin, orally administered drugs such as

analgesics, chondroitin sulphate, glucosamine, minerals (Boron), vitamins, Coxibs,

local injections therapies such as intra-articular and peri-articular corticosteroids, and

hyaluronan (Brandt, et al., 2003b).

The surgical treatment option however involves invasive interventions often

recommended in severe osteoarthritis when other treatment options have failed.

These include arthroscopic lavage, cartilage transplantation, osteotomy, partial and

total joint replacement (Brandt, et al., 2003b). It has been observed that joint

replacement (total or uni-compartmental knee replacement) is the most effective

remedy. The replacement prostheses last between nine and fifteen years on average

before revision surgery is required. Most of the other treatment options mentioned

10

above are palliatives with short term potency, usually limited to 3 months. Also

noting that the surfaces of osteoarthritic hip and knee cartilage in the early stages of

joint degeneration are deficient in SAPL relative to a normal healthy tissue and that

the SAPL injected into the joints of patients with osteoarthritis (OA) resulted in an

increase in mobility and relief for up to 14 weeks of post injection monitoring

(Vecchio et al., 1999). It is envisaged that the result of this study will serve as a

contributory platform for the research on creation of new generation

chondroprotective remedies and injection, which is currently on-going on at

Queensland University of Technology (QUT).

The synthetic SAPL used in the trial study by Vecchio et al (1999) was a non-native

joint SAPL. Previous study by Beldiman et al (2008) found Dipalmitoyl-

phosphatidylcholine, (DPPC) to kill the cells, is it because it is a saturated lipid with

low solubility? This was not investigated in this thesis. Currently, none of the

pharmaceutical products that are being sold in the markets today contain the native

surfactant found in the joints (i.e. saturated and unsaturated SAPL mixture). For

example, Hyalgan® injection, which contains sodium hyaluronate, a chemical found

in the body offers short term relief for patients with osteoarthritis. More so,

Hyalgan® is expensive, ineffective and painful, discouraging practitioners from

applying it despite the general view that such non-surgical intervention is desirable

(Weis et al., 1981; Dahlberg et al., 1994; Lohmander et al., 1996; Marshall, 1998). It

is believed that the outcome of this study will provide relevant data for medical

engineers and pharmaceutical companies to develop potential products that can offer

effective and non-invasive/non-surgical treatment of the disease. A systematic

scientific study is required if we were to realize the goal of this approach to treatment

of osteoarthritis. As mentioned earlier, the focus of this research is to explore the

potential of resurfacing cartilage with synthetic phospholipids using a scientifically

11

based approach. To achieve this aim, it is imperative to understand the physical,

chemical and biological properties of articular cartilage lipids and its relationship

with the other matrix components.

2.3 ARTICULAR CARTILAGE STRUCTURE AND

ARCHITECTURE

Articular cartilage is a dense connective tissue that covers the articulating surfaces of

moving joints in mammals such as fingers, elbow, rib cage, shoulder, knee and hip. It

plays a crucial role in biomechanical mobility of humans by spreading static contact

load thereby reducing the stress/pressure in the joint within the physiological loading

range and serving as a wear-resistant protective material for bones, thus acting as an

extremely low coefficient of friction material protecting the ends of articulating

bones (Mow et al., 1992b). Therefore articular cartilage allows for easy and pain free

movement of joints, which is needed by humans and animals for their daily activities.

The tissue is a heterogeneous and anisotropic fluid-saturated poro-elastic material

with a highly complex structure and architecture (Mow et al., 1984; Rieppo et al.,

2009). Healthy adult human cartilage is 2-4 mm thick and is composed of

chondrocyte cells, a three dimensional meshwork of collagen fibres, proteoglycans

(PGs), lipids and a large volume of water (Meachim and Stockwell, 1974). The

cartilage can therefore be conceptualized as an unpartitionable fluid saturated

poroelastic stiff gel formed by the combination of these components in a specific

composition by mass. The special interactions between the components result in the

complexity of the structure and functions of the tissue, with each component making

12

unique contribution to the physiological characteristics and function of the entire

tissue (Buckwalter and Mankin, 1998; Stok and Oloyede, 2007).

Generally, articular cartilage consists of relatively small number of cells between 3 -

10% of the total volume (Wong and Hunziker, 1998), and an abundant extracellular

matrix with both functioning interdependently (Buckwalter et al., 1990; Van der

Kraan et al., 2002). These cells are called chondrocytes. The chondrocytes are

responsible for the synthesis of the matrix and probably its degradation (Kuettner et

al., 1982; Mankin et al., 1971), while the solid matrix plays an important part in

regulating the cells’ environment (Guilak, 2000; Meachim and Stockwell, 1974). The

chondrocytes also contain trace quantities of neutral internal lipids in their lacunae

(Collins et al., 1965; Ghadially et al., 1965; Stockwell, 1967; Stockwell, 1979).

These are known as intra-matrix lipids. The intra-matrix lipids have been shown to

positively influence the load-bearing properties of cartilage (Oloyede, et al., 2004a,

2004b).

In addition to the intra-matrix lipids, articular cartilage is covered by a thin layer of

surface-active phospholipids (SAPL) membrane, known as surface amorphous layer

(SAL) of nanoscopic thickness, between 500 - 850 nm (Yusuf et al., 2012), which is

believed to play a key role in the lubrication of the joint as solid lubricant (Gale et

al., 2007; Hills, 2000; Hills, 1989; Saikko and Ahlroos, 1997). The SAL layer has

been shown to be lost as a result of cartilage degeneration (Hills and Monds, 1998)

leading to non-physiological lubrication that can be argued to increase the wear of

the tissue. Also, this lipid layer is known to impart highly desirable physico-

biochemical function to the cartilage such as semipermeability (Chen et al., 2007a;

13

Chen et al., 2007b). The capacity of the SAL to act as a semipermeable membrane

allows it to control the diffusional transport of fluid and polar solutes into and out of

the avascular cartilage tissue and the removal of unwanted waste materials (Burstein

et al., 1993; Chen, et al., 2007a; Mauck et al., 2003). The focus of this research is to

further understand this layer, its role in influencing the diffusion characteristics of

cartilage matrix, its influence on the fluid flow and deformation behaviour of the

tissue matrix; and to test the hypothesis that this surface lipid layer can be resurfaced

with full functionality using synthetic phospholipids following the degradation. This

will be achieved by creating models of articular cartilage with normal intact,

delipidized, and relipidized surfaces to study how the surface lipids influence the

structural and functional behaviour of articular cartilage.

Conversely, the dense extracellular matrix of cartilage can be likened to a three-

component gel system consisting primarily of water (65 – 80%), collagen fibres (10

– 20%) and proteoglycans (10 – 15%) (Pearle et al., 2005). Since the cells occupy a

small fraction of the total volume of human articular cartilage, the physico-chemical

properties of the cartilage is governed mainly by the properties of the matrix

(Meachim and Stockwell, 1974). The collagen meshwork being a tensile element

preserves the structural cohesivity of the gel system, while the proteoglycan-water

subgel is responsible for both the fluid and solute transport across the tissue. The

proteoglycan (PG) molecules contain excess fixed negatively charged groups with

typical properties of a polyelectrolyte solution that leads to a high swelling pressure

(Maroudas, 1979; Oloyede et al., 1992; Olsen and Oloyede, 2002; Olsen et al.,

2004). So it is able to draw fluid in when the tissue is unloaded, because the PGs can

swell forever if allowed, they form part of the load carrying structural unit by being

14

entrapped within the three-dimensional network created by the fibrous collagen

component (Oloyede and Broom, 1996).

The physical interactions and coupling between the extracellular matrix components

of cartilage can be conceptualized macroscopically with the balloon-string model

suggested previously by (Broom and Marra, 1985) (Figures 2.1 and 2.2). The model

represents the structural coupling between the entrapping collagen fibres and the

highly deformable proteoglycans before and after loading. The collagen fibrils are

represented by nylon braids and the proteoglycans by inflated balloons. The repeated

crosslinking between these vital matrix constituents allows the tissue to carry out its

load bearing function as demonstrated in the model. It is observed that this model

only accounts for the major components of articular cartilage matrix (collagen,

proteoglycans and water). It does not incorporate the highly important surface

amorphous layer, which has been established to contribute significantly to load

bearing and joint lubrication (Hills 2000; Oloyede et al., 2008). This model can be

further extended to include the articular surface. In Figures 2.1 and 2.2, an analogy

was made on how the articular surface (represented by parallel surface braids) is

supported or anchored by the radial arrangement of collagen fibrils (represented by

the intertwined nylon braids) underlying the tissue surface. Figure 2.2 shows how an

external applied load (the bags of cement) is carried by the inflated balloon-spring.

This similar to what is experienced in the joint, where the articular cartilage covered

by phospholipid-rich surface amorphous layer (surface braids) provides cushioning

for the joint system.

15

Figure 2.1 A physical model of representing the physical interactions and structural

coupling between the components of the load bearing units of articular cartilage

before loading (Balloons and strings, (Broom and Marra, 1985).

The flat plank rest of the parallel braids which is

analogous to the articular surface

16

Figure 2.2 The balloon-string analogue suggested by Broom and Marra (1985). The

3-D string meshwork and the air-filled balloons represent the collagen fibre network

and the fluid swollen proteoglycans of articular cartilage after loading. The flat plank

sits on the articular surface in contact with the load.

It is also important to note that articular cartilage requires other components of the

joint such as the synovial membrane, lubricin, hyaluronic acid (HA) and viscoelastic

synovial fluid to function efficiently (Hills, 1996; Hills and Butler, 1984; Ropes

MW, 1953; Schmidt, 2007). The combined structure of collagen, proteoglycans and

other cartilage matrix components in the joint govern the deformation that can result

The flat plank rest of the parallel braids which is

analogous to the articular surface in contact with the load

17

from an applied load, and through this deformation distributes the load over a wider

contact area (with minimum stress), so that its resultant effect on the bone is

minimized within a physiological bearable limit (Freeman and Kempson, 1974;

Oloyede and Broom, 1996; Weightman and Kempson, 1979). The collagen and

proteoglycans make up the major components of the cartilage extracellular matrix;

however, when cartilage is traumatized or degenerated, its surface is often the first

victim of attack, thereby resulting in lipid depletion (Hills and Monds, 1998). This

loss consequently affects the smooth physiological function of the joint, such as

lubrication, load spreading and semipermeability, thereby hampering mobility and

human activities.

Furthermore, the surface of loaded normal intact cartilage has been shown to be able

to sustain a high pressurized fluid film while the contrast applies to degraded

cartilage (Oloyede and Broom, 1994). Also, the results established that there is a

close relationship between the SAL structure and the fluid pressure generated during

loading which is argued to facilitate weeping lubrication (Mow and Ling, 1969), and

the effective physiological function of the joint system. This thesis focuses on the

role of SAPL in load bearing/processing and semipermeability behaviour of articular

cartilage, and whether the surface of degraded cartilage layered with synthetic SAPL

is able to carry out these important functions of the tissue. Further studies will be

required to test capacity of the newly laid SAL to perform lubrication function

exhibited by normal intact cartilage surface as this is not reported in this thesis. In

order to conduct any study of articular cartilage, it is imperative to understand its

structural, mechanical and physico-chemical properties and how these properties are

influenced by joint conditions such as osteoarthritis.

18

2.4 BIO-MECHANO-CHEMICAL BASIS OF THE

FUNCTIONAL FAILURE OF ARTICULAR

CARTILAGE

Connective tissues in the body undergo changes due to ageing, disuse, wear and tear.

Articular cartilage is no exception to this rule. Unlike other tissues, the cartilage is

aneural, alymphatic and avascular. When the joint is traumatized, the tissue has a

limited chance to repair itself depending on the degree of the injury. There are

instances where the cartilage is unable to function effectively; in such cases it is said

to be diseased/degraded or arthritic. The most common form of arthritis in the joint is

osteoarthritis (OA) (Brandt et al., 2003a). Osteoarthritis is a degenerative joint

disease characterized by progressive erosion of articular cartilage, which results in

eventual loss of entire joint function. Changes in cartilage during the early stages of

osteoarthritis include surface fissuring, which may lead to loss of surface lipids,

mechanical softening, decrease in PG content, increase in water content, changes in

thickness, and increased permeability (Buckwalter and Brown, 2004; Guilak et al.,

1994; Wu et al., 2000; Yusuf, et al., 2012).

Additionally, the degradation of cartilage is often accompanied by joint

inflammation, sometimes episodically but without systemic effects and is related to

the introduction of breakdown products into the synovial fluid and their consequent

phagocytosis (Brandt, et al., 2003a). Osteoarthritis is also accompanied by changes

in the composition of hyaluronic acid (HA) contained in the synovial fluid: the

typical molecular weight of HA reduces from 1.0 107 Da in healthy joints to as low

as 2.0 105 Da in diseased joints (Ghosh, 1994; Tehranzadeh et al., 2005), and the

overall concentration of hyaluronic acid is diminished from the normal 3 mg/mL

19

(Ghosh, 1994). The ultimate result of this process is loss of cartilage elasticity,

smooth articular function, and joint lubrication.

Furthermore, it has been demonstrated that this painful and enervating disease is

often characterised by progressive loss of cartilage, the matrix constituents, namely,

depletion of the osmotically active proteoglycans (PGs) (Oloyede and Broom, 1993;

Zhu et al., 1993), disruption of the collagen fibre architecture (Broom, 1986a, 1986b;

LeRoux et al., 2000), and more relevant to this study, the lipid or surfactant content

(Guerra et al., 1996; Oloyede, et al., 2004a, 2004b; Sarma, et al., 2001) (Figure 2.3).

The depletion of the SAPL layer impairs lubrication in the joint with unfavourable

effects on joint movement and human activities (Gale, et al., 2007; Hills, 2000).

Oloyede et al (2004b) established in their consolidation experiment that the loss of

lipids from cartilage surface and matrix resulted in a non-physiological stiffening of

the matrix at loading rates that are commonly applied to the tissue during normal

function. With increased stiffness, the cartilage is embrittled resulting in an increased

tendency to premature fracture under normal physiological loads (Oloyede, et al.,

2004b). Osteoarthritis leaves affected patients with pain, discomfort and

consequently the inability to work freely. Therefore, making it critical for people

suffering from this devastating condition seek medical intervention.

20

Figure 2.3 Full picture showing an osteoarthritic articular cartilage (Garg, 2012).

2.5 COMPONENTS OF ARTICULAR CARTILAGE

AND THEIR FUNCTIONS

Hyaline cartilage is the most commonly found cartilage in articulating or diarthrodial

joints. It is divided into four major distinct zones; namely: superficial or tangential,

middle or transitional, deep or radial, and the calcified zone (Figure 2.4), with its

components distributed anisotropically across these zones (Glenister, 1976). The

relative thickness of the zones is dependent on the maturity of the skeleton, age and

species examined. It also varies accordingly from joint to joints, and from different

21

regions within the same joint compartments (Ateshian et al., 1991; Kiviranta et al.,

1987; Simon, 1970). The articular cartilage is covered by a mechanically strong

semi-translucent layer called the articular surface. This articular surface covers the

underlying matrix and is integrated structurally with it and has been characterized

microscopically as a distinct layer (Kamalanathan and Broom, 1993). The articular

surface is overlaid by the surface amorphous layer (SAL) or superficial lipid layer

(SPL) (Graindorge and Stachowiak, 2000; Guerra, et al., 1996; Sarma, et al., 2001).

Figure 2.4 Representation of the zonal variation and distribution of articular cartilage

matrix constituents from the surface and subchondral bone (Jadin et al., 2007).

The surface amorphous layer is comprised of lamellar bodies, charged

macromolecules (lubricin and hyaluronan), and negatively charged phospholipid

micelles (Hills, 1990, 1992) that create desirable conditions which, in the presence of

Calcified cartilage

Deep/ Radial

Middle / Transitional

Articular surface

Superficial / Tangential

Zones

22

water, provide a most astonishing low sliding friction (Gadomski et al., 2008; Hills,

2000; Pawlak et al., 2008; Pawlak and Oloyede, 2008). This in turn provides the

articular cartilage of mammalian joints with an almost frictionless lubrication created

and controlled by the superficial phospholipids layer that are chemically attached to

the articular surface. One of the objectives of this thesis is to characterize

microscopically and nanoscopically the nature of the layer that is formed by each

components of this phospholipid layer when cartilage with degraded surface is

exposed to them. A further objective is to study the differences at this microscopic

level between the layers formed by these components and the complete lipid mixture

found on the cartilage surface. This will provide an understanding of the effects of

degradative lipid loss on cartilage’s transport or material exchange property during

degradation and establish the relative effects of each lipid type on this transport and

load bearing of the intact and degraded cartilage relative to lipid loss.

At this junction, it is important to explain the role of the other matrix components,

such as collagen, proteoglycans, water, and ions, found in the several zones

underlying the articular surface and how they combine efficiently to keep the joint in

perfect condition. It should be noted that aim of the work is not to test the

lubrication behaviour of the surface amorphous layer; however, this research will

focus on determining how synthetic lipids behave when in contact with cartilage

surface, the role they play in load bearing and diffusion characteristics of articular

cartilage. The knowledge gained from this study will contribute to the existing

research on the potential application of synthetic surface-active phospholipid-based

treatment of degenerating articular cartilage.

23

2.5.1 COLLAGEN FIBRES

The collagen fibres account for about two-third of the entire dry weight of the

cartilage matrix. The matrix consists majorly of Type II collagen, which accounts for

about 10 – 20% of its wet weight. Collagen is a fibrous protein that is primarily

responsible for the shear stress and tensile stiffness of the tissue. The orientation of

its fibres changes from the surface to the subchondral bone. The fibres are able to

form an intra- and inter-molecular cross-linked network structure, which stabilizes

the matrix (Pearle, et al., 2005) and through this network, three zones of alignment

can be distinguished (shown schematically in Figures 2.5, 2.6 and 2.7 below). The

fibres close to the articular surface are aligned parallel to the surface forming the

superficial or tangential zone and become increasingly aligned normal to the surface

at greater depth from the transitional zone down to the calcified zone (Jeffery et al.,

1991; Meachim and Stockwell, 1974). This zonal difference in fibre structure results

in the variation of biomechanical properties of cartilage such as thickness and

density. Also, the collagen fibre orientation plays a significant role in fluid flow-

dependent response of articular cartilage during physiological loading. The very low

permeability of cartilage is attributed to the large frictional resistance of the collagen

fibre-proteoglycan interaction to fluid flow (Pearle, et al., 2005).

Previous studies by Jurvelin et al. (1996), Kumar et al. (2001), and Grant et al.

(2006) have established, through atomic force microscopy imaging, that the surface

of normal intact cartilage is featureless and when the articular surface is degraded, a

fibrous structure of collagen from the subsurface or underlying matrix is revealed.

Their hypothesis was further tested in this study because of the contrasting

24

observation by Hills et al. (1990), which showed that the surface amorphous layer is

arranged in lamellar-like nature. In addition, since the articular surface contains

mostly phospholipids and protein materials not collagen, therefore, the features

observed by earlier researchers could not have been collagen. This problem can only

be resolved through a high resolution microscopic imaging and combined with

rigorous image analysis, as demonstrated by this thesis.

Figure 2.5 Schematic view of the articulating joint, the expansion shows the zonal

architecture of articular cartilage (Crouch, 1985).

25

Figure 2.6 (A) Schematic diagram showing the chondrocyte distribution and (B) the

structure of collagen fibre network in the distinct zones (Buckwalter et al., 1994).

Figure 2.7 Zonal architecture of articular cartilage (Jeffrey and Watt, 2003).

26

2.5.2 PROTEOGLYCANS

Proteoglycans make up approximately 30% of the dry weight of articular cartilage.

Unlike the neutral collagen fibres, the proteoglycans are highly charged molecules

entrapped and immobilised within a 3-D meshwork of collagen fibrils (Chen and

Broom, 1998; Karvonen et al., 1992; Oloyede, et al., 1992). Their molecules are

composed of a central protein core that forms a backbone, to which many

glycosaminoglycan (GAG) chains are attached via covalent bonding. These chains

are able to extend perpendicularly from the backbone of the protein core in a

“bottlebrush-like” structure that allows trapping of large amounts of water (Figure

2.8).

Figure 2.8 Schematic representation of cartilage extracellular matrix showing the

proteoglycan aggregate and aggrecan molecule (Pearle, et al., 2005).

27

Glycosaminoglycans (GAGs), also known as mucopolysaccahrides consist of long

unbranched polysaccharide chains, which are made up of a repeating disaccharide

unit (Hardingham, 1981; Roughley and Lee, 1994). This repeating disaccharide unit

consists of glucoronic acid (a hexuronic acid or six-carbon sugar acid moiety)

attached to N-acetylglucosamine (a hexosamine or six-carbon acylated amino sugar

moiety) (Roughley and Lee, 1994). There are three major types of

glycosaminoglycans (GAGs) in cartilage; chondroitin sulphate (M.W., 20-50 kDa),

keratan sulfate (M.W., 20-50 kDa) and hyaluronan (M.W. of up to 5 Mega Daltons

with over 100 GAG chains). Chondroitin sulphate is the most abundant followed by

keratin sulphate and lastly by hyaluronan (Lehninger et al., 2005).

The hyaluronan, also called hyaluronic acid (HA) or hyaluronate are covalently

linked to aggrecan monomers through the presence of link proteins, which stabilize

the HA – Aggrecan molecules (Kiani et al., 2002). The aggrecan monomers consist

of core proteins attached to several chondroitin sulphate and keratan sulphate chains,

which constitute the abundant proteoglycans in cartilage (Figure 2.8 above). The

GAGs contain excess fixed negatively charged groups because the amino sugar

moieties in their structure are sulphated, thus they are able to maintain osmotic

equilibrium when entrapped by the elastic collagen fibre network within the tissue

during physiological conditions (Oloyede and Broom, 1994a; Olsen and Oloyede,

2002; Olsen, et al., 2004).

28

Figure 2.9 Molecular structure of chondroitin sulphate monomer chain (Muir, 1978).

Figure 2.10 Molecular structure of keratan sulphate monomer chain (Muir, 1978).

29

Figure 2.11 Hyaluronic acid, a heteropolysaccharide with several thousand monomer

units of N-acetyl glucosamine and glucuronic acid formed (Stern, 2004).

It is also important to note that articular cartilage is not a composite material but a

specialized biomechanically and physicochemically active three-component

biological hydrogel (Broom and Oloyede, 1998). In this gel, a distended 3-D network

of collagen fibre meshwork entraps the fluid swollen proteoglycans, and the

interstitial water molecules (containing H+ and OH

- ions) are attracted by the highly

negatively charged side chains of the proteoglycans (Figures 2.9, 2.10 and 2.11 )

(Ateshian et al., 2003; Broom and Oloyede, 1998; Olsen and Oloyede, 2002). The

unique interaction between cartilage matrix components, such as the collagen fibre

meshwork, proteoglycans, and interstitial fluid plays a crucial role in its extremely

low anisotropic permeability and hyperelastic stiffness, thereby resulting in a low

rate of water exudation from the matrix during physiological function (Ateshian and

Hung, 2003; Oloyede et al., 1998; Oloyede and Broom, 1994a; Oloyede and Broom,

30

1994b). This unique matrix arrangement governs the mechanism of articular cartilage

consolidation developed by Oloyede and Broom (1991).

The variation of the concentration of proteoglycans across the depth of cartilage from

the articular surface to the subchondral bone follows a “bell-like” shape that differs

from the zones as defined by the collagen fibres (O'Connor et al., 1988). The

concentration of proteoglycans is low close to the articular surface and increases to a

maximum at about 50% to 80% depth. It then decreases when approaching the

tidemark (O'Connor, et al., 1988). As earlier mentioned, the articular surface

comprises multi-bilayered phospholipid membranes, proteoglycans, glycoproteins,

cholesterol, hyaluronan and water molecules (Figure 2.11). The exact composition of

these components still remains unknown resulting in several arguments amongst

researchers (Jurvelin et al., 1996; Kobayashi et al., 1995; Kumar et al., 2001). It has

been argued that each of the components of articular cartilage surface amorphous

layer contributes significantly to lubrication in mammalian joints. For example,

Swann et al (1985) and Radin et al (1970) proposed that lubricin; a glycoprotein

(also known as proteoglycans 4, PRG4) unique to the synovial fluid, is the active

ingredient for joint lubrication. However, Schwartz and Hills (1998) proved that

lubricin being water soluble, acts only as a carrier for the highly insoluble surface-

active phospholipids that are deposited on the articular surface. The SAPLs are

deposited as oligolamella layer of phospholipids which, in the presence of water,

posses the desirable lubricating properties for effective joint function (Pawlak and

Oloyede, 2008).

The phospholipid molecules in the SAPL exhibit unique amphiphilic behaviour,

possessing positive quaternary ammonium ions (R4N+ or QA) and negative

phosphate ions. The quaternary ammonium (QA) ions has strong electrostatic bond

31

strength and thus able to bind to surfaces with excess negative charges. Since the

proteoglycan molecules in the articular surface have excess carboxyl and sulphate

ions (Figures 2.9 and 2.10), the tissue surface is attractive to the QA ions, thereby

leaving the excess phosphate ions accessible for the positive mobile ions (Na+, Ca

2+,

H+) present in the synovial fluid (Hills, 2000). The effect of the ions interactions

keeps the articular surface electrically neutral, and excellent boundary lubricant. This

mechanism is presented in Figure 2.13. It is hypothesized in thesis that this unique

surface chemical property/structure possessed by the articular surface due its surface

amorphous phospholipid layer is disrupted or lost when the surface lipids are eroded

following early stages of cartilage degeneration. In order to simulate the loss of

cartilage surface lipids, an artificial lipid extraction process was used

(delipidization). The removal of the SAPL, which will inevitably change the tissue

surface chemistry, will be evaluated using Raman spectroscopy. This

characterization method was chosen because of its ability to detect changes in

chemical bonding between the molecules of SAPL and the articular surface, and the

influence of these changes on the overall chemical properties of the cartilage surface

(Lim et al., 2011).

32

Figure 2.12 Electron micrograph of the oligolamella layer of SAPL adsorbed to the

pleural epithelium, which is similar to the surface of cartilage in vivo (Hills, 2000).

Lamella layers of SAPL

33

Figure 2.13 Electrostatic bonding of the quaternary positive ions from the SAPL

molecules with excess negative charges from proteoglycan molecules on the articular

surface (Hills, 2000).

34

2.5.3 WATER (BOUND AND UNBOUND), IONS AND

CHONDROCYTES

Water is the major component of articular cartilage. It makes up approximately 70-

85% of the entire weight of the tissue. There are two types of water in cartilage;

bound and unbound (free). The amount of bound water in the tissue is negligible

when compared to the unbound water. Bound water makes up only about 1% of the

total weight of the matrix (Buckwalter et al., 1988; Maroudas et al., 1973). The

unbound water is freely exchangeable between the matrix components; therefore it is

available for the transport of the micro solutes and ions, which are accounted for the

osmotic swelling pressure generated by the negative fixed charge of the

proteoglycans (Maroudas, 1970, 1979; Maroudas and Venn, 1977; Stockwell, 1979).

The concentration of water like proteoglycans varies across the depth of cartilage. It

varies almost inversely as the concentration of proteoglycans across the depth of

cartilage matrix, with highest value near the articular surface containing the

superficial lipid layer (approximately 80%), thus facilitating interstitial fluid

controlled lubrication (Maroudas, 1968; Venn, 1978; Venn and Maroudas, 1977).

However, the concentration of water is lowest in the deeper regions of the cartilage,

near the subchondral bone (approximately 65%) (Huber et al., 2000; Pearle, et al.,

2005). When dissolved in ionic species, the water becomes active electrolytes in the

matrix, consisting of Na+, Ca

2+, H

+, OH

-, Cl

- and other micro ions. Altogether, the

total ionic content of cartilage is less than 1% of its wet weight. To maintain a

balanced osmotic condition in cartilage, these ions are required to be neutralized by

the fixed charges provided by the proteoglycans (Hardingham and Fosang, 1992;

Lerner and Torchia, 1986; Maroudas, 1968).

35

Unlike the fixed or immobile charges that exist in proteoglycans (the sulphate and

carboxylate groups of chondroitin and keratan sulphates attached to hyaluronan

protein core, Figure 2.8), the ion species that make up the active water of cartilage

display astonishing behaviour due to their ability to move freely in the physiological

aqueous joint environment (mainly controlled by water). Hence, they are referred to

as mobile ions. The movement of fluid and ions across the articular surface to the

joint cavity (synovial fluid) also controls the nutrition and removal of waste materials

from the tissue (Burstein, et al., 1993; Maroudas, 1979; Mauck, et al., 2003). This is

important to maintain the survival of the chondrocytes, and consequently the entire

matrix. One of the objectives of this research is to determine whether or not the

surface amorphous layer, through its semipermeability property has any significant

effect on the diffusion of fluid into and out of the matrix and also, test whether or not

it is possible to regenerate the functional semipermeability characteristics of normal

intact cartilage by incubating lipid depleted cartilage in solutions of synthetic

phospholipids. This was achieved using magnetic resonance imaging and

computational analysis.

Magnetic resonance imaging was used to track in real-time, the diffusion of water

through the articular surface from the matrix of cartilage samples immersed in a

deuterium oxide (D2O) environment. The apparent diffusion coefficients of water in

cartilage matrices with three surface conditions; normal, delipidized, and relipidized,

were measured using a purpose-built computational scheme designed with

MATLAB®. The outcome of this study was used to further characterize the surface

amorphous layer and also, evaluate the functionality of the new layer deposited on

degraded cartilage following relipidization.

36

Furthermore, for articular cartilage to function effectively as a load spreading

material, there is a need for constant and continuous lubrication of its surface.

Experiments have revealed that the coefficients of kinetic friction measured in

healthy joints in vitro range from 0.002 – 0.006 compared with a value of 0.04 for

Teflon, which is presently known to be one of the best boundary lubricants in

mechanical systems (Charnley, 1959; Jones, 1934; Little et al., 1969). This

extremely low coefficient of friction in diarthrodial joints has been investigated by

researchers for many years leading to development of several theories of joint

lubrication ranging from boundary to hydrostatic and hydrodynamic lubrication to

mention but a few.

Earlier studies have suggested that lipids could exert lubricating effects in the joints

by adsorbing to the surface of cartilage. To further support this, Little et al. (1969)

observed that rinsing articular surfaces with lipid solvents increased the joint friction

more than two fold. The outcome of their study and other recent studies established

the presence of layer of lipids on the surface of cartilage. These lipids were later

called surface-active phospholipids (SAPL) or surfactants. The focus of this thesis

in not to study the influence of the SAPL on articular cartilage lubrication, but to

understand the surface structural configuration and functional characteristics of

normal healthy cartilage and the consequence lipid loss to these properties, thereby

establishing the basis of comparison for the newly resurfaced cartilage following

incubation in synthetic surface-active phospholipids. A comprehensive description of

cartilage lipids, the structure and functions are presented in the next section.

37

2.6 ARTICULAR CARTILAGE LIPIDS

Lipids are large chemically heterogeneous group of compounds found in living

organisms. Their biological functions are as diverse as their chemistry. Lipids are

naturally-occurring molecules, such as fats, oils, waxes, cholesterol, steroids,

phospholipids and many more. The most common and defining feature of lipids is

their insolubility in polar solvents (e.g. water) and relative solubility in non-polar

solvents and solvents of low polarity (e.g. ether, chloroform, acetone & benzene)

(Lehninger, et al., 2005). Unlike other organic macromolecules such as

carbohydrates and proteins, lipids are defined by physical property (solubility) rather

than by structure. They are generally insoluble in water. The main biological

functions of lipids include energy storage, acting as structural components of cell

membranes, and important signalling molecules (Jump, 2002).

2.6.1 BIOCHEMISTRY OF LIPIDS: NOMENCLATURE AND

STRUCTURE

Biological lipids can be broadly divided into two groups based on the functions they

perform in living organisms, these are: storage (neutral) lipids and structural

(membrane) lipids (Lehninger, et al., 2005). Storage lipids such as fats and oils are

the universal stored forms of energy in living organisms. They are derivatives of

fatty acids, which are carboxylic acids with hydrocarbon chains, ranging from 4 to

36 carbons long (C4 to C36). Triacylglycerols, which are also referred to as

triglycerides, fats or neutral fats, are the simplest member of the fatty acids. They

are composed of three fatty acids each in ester linkage with a single glycerol (Figure

38

(a)

2.14 (a) and (b)). Triacylglycerols form a separate phase of microscopic oily droplets

within the cells, which serve as depots of fuel for metabolic activities (Lehninger, et

al., 2005).

Figure 2.14 (a) and (b) Chemical structure of triacylglycerol R, R1, R2, and R3 denote

aliphatic chain hydrocarbons (Lehninger, et al., 2005).

The hydrocarbon chains of triglycerides can either be branched or unbranched with

saturated or unsaturated carbon atoms depending on the type of fatty acids

(Figures 2.15, 2.16 and 2.17). The degree of unsaturation and length of the

(b)

39

hydrocarbon chain largely determines the physical properties of fatty acids and the

compounds that contain them. Fatty acids have poor solubility in water due to their

non-polar hydrocarbon chain. Their solubility in water decreases as the length of the

fatty acyl chain increases and the number of double bonds decreases. For example

palmitic acid (16:0, Molecular weight 256 g/mol) has a solubility of 0.0083 mg/g in

water, which is far much less than glucose (Molecular weight 256 g/mol) with a

solubility of 1100 mg/g in water (Lehninger, et al., 2005).

Figure 2.15 Chemical structure of Palmitic acid, a saturated and unbranched fatty

acid (Lehninger, et al., 2005).

Figure 2.16 Chemical structure of Oleic acid; an unsaturated and unbranched

fatty acid (Lehninger, et al., 2005).

40

Figure 2.17 Chemical structure of Cholesterol; an unsaturated and branched fatty

acid (Lehninger, et al., 2005).

Structural lipids, on the other hand, form a central architectural feature in biological

membranes. This is called the lipid bilayer, which acts as a barrier for the passage of

polar molecules and ions (see Figure 2.18 below). Structural lipids, unlike storage

lipids, are amphipathic; with one end of the molecule hydrophilic and the other end

hydrophobic. They form bilayers when their hydrophobic ends interact with each

other and their hydrophilic ends interact with water. Structural lipids include

phospholipids, glycolipids and archaeal ether lipids (Lehninger, et al., 2005). The

synthetic phospholipids used in this study belong to this class of lipids.

41

Figure 2.18 A lipid bilayer structure, showing the hydrophilic head and hydrophobic

tails (Inex Pharmaceutical Corporation).

In human adult cartilage, lipids form 1-5% of the total weight and are found both

within and outside the cells of the matrix. The intracellular lipids are neutral

(storage) lipids located in the lacunae of the chondrocyte cells (Stockwell, 1967),

while the extracellular lipids spread more diffusively throughout the matrix (Efskind,

1941; Schallock, 1942). In addition to these, phospholipids, which are the major

components of structural or membrane lipids, form a microscopic layer surface-

active phospholipids (SAPL) in conjunction with water, glycoproteins, cholesterol,

hyaluronic acid and other constituents on the articular surface (Hills, 1990; Sarma, et

al., 2001; Schwarz and Hills, 1998). These components combine to form a

membrane on articular cartilage called the surface amorphous layer (SAL), in which

the SAPL is the major component (Hills, 1990; Schwarz and Hills, 1998).

42

2.6.2 PHOSPHOLIPIDS: PROPERTIES AND FUNCTIONS

Phospholipids are fat derivatives, in which one fatty acid has been replaced by a

phosphate group and one of several nitrogen-containing molecules with the

hydrocarbon chains being hydrophobic (as in all fats) (Hills and Cotton, 1986). A

phospholipid molecule is said to be amphiphilic/amphipathic when it contains

phosphate and amino groups, which make up the hydrophilic polar head group and

a hydrophobic tail, respectively. The hydrophobic tail is made up of two fatty acid

chains, which may be saturated (i.e. carbon atoms are all connected by single

bonds) or unsaturated (i.e. some carbon atoms are connected by double

bonds). Each of the fatty acid chains has an even number of carbon atoms, which

result from the mode of their synthesis through the condensation of two-carbon

(acetate) units (Jump, 2002). The ability for SAPL to switch between hydrophilic and

hydrophobic forms plays major role in articular cartilage lubrication (Pawlak and

Oloyede, 2008).

There are two main types of phospholipids: phosphoglycerides and sphingolipids.

Most phospholipids belong to the phosphoglycerides, also called

glycerophospholipids, which are membrane lipids in which two fatty acids are

attached in ester linkage to the first and second carbon atoms of a glycerol backbone

and a highly polar/charged group is attached through a phosphodiester linkage to the

third carbon. Examples include phosphatidylcholine (PC),

phosphatidylethanolamine, phosphatidylglycerol, phosphatidylserine, and

phosphatidylinositol (Jump, 2002; Lehninger, et al., 2005).

43

Figure 2.19 General chemical structure of phosphatidylcholines (Jump, 2002).

Figure 2.20 General structure of glycerophospholipids (Lehninger, et al., 2005).

Saturated fatty acids

(e.g., palmitic acid)

Unsaturated fatty acids

(e.g., oleic acid)

X Head-group

substituent

44

The properties of phospholipids such as structure, molecular weight, melting point,

boiling point, solubility, and pH are characterized by the properties of the fatty acid

chains and the phosphate/amino group. The large hydrocarbon moiety (a long chain

of the form CH3(CH2)n, with n > 4) of the fatty acids, for example is non-polar.

However, the phosphate (PO43−

) group has negatively charged oxygen and positively

charged nitrogen that make up the polar (ionic) group. Figure 2.19 represents the

chemical structure of a saturated phospholipid, while Figure 2.20 represents the

general structure of glycerophospholipids, in which two fatty acids (with saturated

and unsaturated straight chains) are attached in ester linkage to the first and second

carbons of glycerol, and a highly polar or charged group (represented as - X above)

is attached through a phosphodiester linkage to the third carbon (Lehninger, et al.,

2005). Phospholipids such as ethanolamine and choline are quite soluble in aqueous

solutions. When mixed with water, they spontaneously form microscopic lipid

aggregates in a phase separate from the aqueous surroundings, with the hydrophobic

moieties in contact with each other and the hydrophilic-head groups interacting with

the surrounding water. Three lipid aggregates (a micelle, a lipid bilayer or a

liposome) can be formed depending on the solution conditions such as concentration,

temperature, ionic strength and pH and the nature of the lipids (Lehninger, et al.,

2005) (Figure 2.21).

45

Figure 2.21 A schematic representation of liposome structure (Britannica, 2007).

Recent studies (Hills, 1990) have shown that the articular surface is overlaid by a

thin layer of phospholipids (later called surface-active phospholipids, (SAPL)) of

macroscopic thickness, which is believed to contribute immensely to the lubrication

(Hills, 1989; Schwarz and Hills, 1998) and load processing in joints (Oloyede, 2004).

Hills (1996) further argued that the waxy appearance of the cartilage is closely

related to the presence of these surface lipids. In the lungs, SAPL is more commonly

referred to as “surfactant”, where it is produced by alveolar Type II cells in form of

lamella bodies, which is secreted onto the alveolar surface (Stratton, 1984). SAPL is

46

also synthesized and secreted in other parts of the body such as pleural (Hills, 1992),

pericardial (Hills and Butler, 1985) and peritoneal cavities (Grahame et al., 1985;

Ziegler et al., 1989) and joints (Gale, et al., 2007; Hills and Butler, 1984; Schwarz

and Hills, 1996), where its adsorption onto the surface of tissues at these sites has

been demonstrated using techniques such as electron microscopy (Ueda et al., 1985),

epifluorescence microscopy (Hills, 1992), flow cytofluoremetry (Hayem et al., 1994)

and autoradiography (Chen and Hills, 2000).

Also, SAPL imparts highly desirable physical and physiological properties, which

include: surface tension reduction (Clements, 1957), boundary lubrication (Gale, et

al., 2007), release (anti-stick) (Hills et al., 1998), semipermeability (Chen et al.,

2002) and physical barrier formation (Hills, 1991). These functional properties of

SAPL in several parts of the body, including the cartilage, have made it imperative

for further studies of the mechanism of SAPL and articular cartilage interaction.

Based on the fatty acid chains in phospholipids, SAPL can be classified into two

species namely saturated and unsaturated species. It has been established that the

composition and type of SAPL varies amongst organs. However, for a long time,

most research on human SAPL has focused on the saturated surfactant (dipalmitoyl-

phosphatidylcholine (DPPC) (Chen et al., 2005; Vecchio, et al., 1999) for reasons to

do with their role in sudden infant death syndrome and its associated neonatal

respiratory distress as earlier mentioned. Analysis of SAPL from bovine articular

cartilage revealed that this type of surfactant (SAPL) is not the one found in the joint

of mammals (Hills, 1996; Hills and Butler, 1984; Schwarz and Hills, 1996) and that

the unsaturated species are most dominant with phosphatidylcholine (41%),

phosphatidylethanolamine (27%) and sphingomyelin (32%) being the major

components (Chen, et al., 2007b; Sarma, et al., 2001). Dipalmitoyl-

47

phosphatidylcholine (DPPC), a di-saturated phosphatidylcholine (SPC), is the

main component of surfactant/SAPL found in the alveolar lining of the lung

(Stratton, 1984), leading to the now outdated assumption that this saturated

phosphatidylcholine (SPC) also makes up the main component of the

phosphatidylcholines (PCs) in non-alveolar sites of the body (Chen and Hills, 2000;

Hills, 1991; Paananen et al., 2002).

Contrary to this long held belief, recent studies have shown that the dominant SAPL

in non-lung sites such as the eustachian tube (Paananen, et al., 2002), the gastric wall

(Bernhard et al., 2001), and articular joints such as the mammalian knee (Chen, et

al., 2007b) are unsaturated phosphatidylcholines (USPC). DPPC, a saturated

SAPL, has a phase transition temperature of 41.3°C; it is unable to liquify at body

temperature and is neither absorbable nor adsorbable with consequences for its

contribution to joint function. On the other hand, the naturally unsaturated

surfactants that exist in articular joints have a phase transition temperature that is

below body temperature and are readily adsorbed, thereby indicating their potential

for providing longer term effectiveness.

The results of Chen et al (Chen, et al., 2007b) obtained from HPLC analysis of

bovine knee cartilage lipid content also provide more recent evidence confirming that

the results from the earlier investigation of Chen and Hills (2005), who discovered

that the dominant PC species on the surface of articular cartilage are the unsaturated

phosphatidylcholines (USPCs), namely Dilinoleoyl-phosphatidylcholine (DLPC),

Palmitoyl-linoleoylphosphatidylcholine (PLPC), Palmitoyl-oleoyl-

phosphatidylcholine (POPC), Dioleoyl-phosphatidylcholine (DOPC) and Stearoyl-

linoleoylphosphatidylcholine (SLPC) coexisting in mixture with a small quantity of

DPPC (8%) . It has been observed that researchers have established the composition

48

of SAPL in the joints. However, there is no study in the literature that gives insights

into determining the effect of either injecting the right composition or blend of

synthetic SAPL into degraded joint or incubating degenerated articular cartilage in

solutions of synthetic SAPL using compositions found in health joints. One of the

key objectives of this research is to close this gap.

Furthermore, it is observed that the compositions and types of SAPL present on the

surface cartilage has been established. However, there is no study in the literature

that offers insight into determining the mechanism(s) of interaction of synthetic

lipids with articular cartilage. It is, therefore, important to carry out a study, which is

targeted towards understanding how synthetic lipids behave when in contact with

articular cartilage within and around the joint environment. The outcome of this

informed how the lipids (surfactant) behave when in contact with cartilage, their

nature of interaction with cartilage, the manner/mode in which they are transported in

the joint, and overall, reveals the potential of repairing the surface of a degraded

cartilage through relipidization in synthetic lipid-rich environment. It is believed that

the outcome of this study will advance knowledge in the area of developing lipid-

based intervention in the management and treatment of dysfunctional joints and also

contribute to the work on chondroprotective injections currently being carried out

within the group at Queensland University of Technology (QUT). The specific

degradation addressed in this work is osteoarthritic degeneration.

In order to achieve the aims and objective of this research, the mechanism of

interaction between synthetic lipids and cartilage was studied in vitro for the first

time by incubating normal and degraded cartilage specimens in solutions of

synthesized saturated and unsaturated Surface-active Phospholipids (SAPLs),

which biomimic the natural lipid species and quantities in the human knee joint.

49

Additionally, microscopic analysis, nano-mechanical indentation tests, diffusion

studies and quasi-static compression tests were conducted to determine whether or

not relipidization with synthetic phospholipids has the potential to reverse the

functional properties of delipidized or degraded cartilage.

The microscopic examination involved the characterization of the structural

configuration of articular cartilage specimens with the surface amorphous layer

intact, and specimens with altered surface lipid layer following gradual removal with

lipid rinsing agents (delipidization). The results were then compared with those of

resurfaced cartilage samples obtained following a controlled deposition of synthetic

surface-active phospholipids (relipidization) on the surfaces of lipid depleted samples

to replace the lost surface amorphous layer.

Furthermore, nano-indentation tests, diffusion study, and quasi-static compression

tests were conducted to evaluate the functional viability of the resurfaced cartilage

relative to normal healthy tissue. The nano-indentation tests were conducted using

the atomic force microscope (AFM) to assess the resistance or rigidity of the new

surface layer created following relipidization. The diffusion studies appraised the

semipermeability characteristics, using a combination of magnetic resonance

imaging (MRI) and computational techniques, while the compression tests measured

mechanical properties, which are related to the fluid exudation behaviour of the

matrix such as: average total strain energy, average elastic energy lost in matrix

recovery, average complementary energy, average residual energy, and average

energy ratio of the resurfaced articular cartilage samples. The results of this work

will be fundamental to the resolution of the question of whether or not the

hypothesized potential repair role of phospholipids in joints has any real significance.

The detailed experimental protocol for the microscopic analysis, nano-mechanical

50

indentation tests, diffusion measurements and computational analysis, and

mechanical compression tests conducted to test the hypotheses of this thesis are

described in the approach and methodology chapter (Chapter three).

51

Chapter 3: EXPLORATORY STUDY OF THE

APPROACH AND

METHODOLOGY

This chapter presents the rationale for the methodologies used in the thesis and the

preliminary experiments conducted to develop the methods used for testing the

hypotheses of this research. Several modifications of existing methods for

characterizing the structural and functional properties of articular cartilage were

undertaken to meet the objectives of this research. The modifications conducted to

adapt the established experimental protocols are discussed.

3.1 ATOMIC FORCE MICROSCOPE (AFM) IMAGING

OF THE SURFACE OF ARTICULAR CARTILAGE

AFM is a versatile nano-characterization instrument that is used for imaging both

soft and hard materials. It can be operated in air, vacuum, or liquid environment.

Imaging of soft materials (biological tissues such as articular cartilage) is often

conducted in liquid environment, where the specimen is submerged in a

physiological saline medium (either 0.15 M saline (NaCl) or phosphate buffered

saline (PBS)). This experimental setup/arrangement is necessary to preserve the

integrity of the tissue during measurements and to keep specimen intact for further

testing, thereby improving the credibility of the results. Generally, AFM

measurement of articular cartilage, which comprises of imaging and force

spectroscopy, is performed in liquid medium (Jurvelin, et al., 1996; Kumar et al.,

2001; Park et al., 2004; Crockett et al., 2005, Grant et al., 2006). Therefore, all the

52

AFM imaging and force spectroscopy experiments reported in this thesis were

conducted in liquid environment with the cartilage specimens fully immersed in

PBS.

Generally, a number of factors are important for obtaining high resolution and

quality images with the AFM. These include:

selection of suitable cantilever (rectangular or triangular)

optimization of set-points and scanning/imaging parameters (scanning

speed/frequency)

mode of operation of the AFM (contact or semi contact/tapping mode)

real-time monitoring of trace and retrace signals with the oscillograph during

scanning, were found to be important.

It is worth noting that the AFM images of the surface of cartilage, which have been

published in the literature until today, generally have low resolution and quality as

illustrated in Figures 3.1 – 3.4 (a collection of 2-D AFM images of the surfaces of

normal articular cartilage).

53

Figure 3.1 2-D AFM images of the surface of bovine humeral head articular cartilage

(A) and (B) are height images (Scale bars, 2 μm; full gray ranges: 1000 nm (A) and

(B) 600 nm) (Jurvelin, et al., 1996).

54

Figure 3.2 2-D topographical AFM image of the surface normal healthy adult pig

articular cartilage. Full scan size 30 x 30 µm; full grey range 1700 nm (Kumar et al.,

2001).

Figure 3.3 AFM height images of the surfaces of bovine cartilage in synovial fluid

(a) before and (b) after washing with PBS. (Image size: 5 x 5 µm area) (Crockett et

al., 2005).

55

Figure 3.4 2-D topographical AFM images of the surface of fresh bovine articular

cartilage: (a) 40 µm scan with 1 µm height scale, (b) 20 µm scan with 2 µm height

scale (Grant et al., 2006).

The type of cantilevers used to obtain the above AFM images (Figures 3.1 – 3.3)

were not specified by the researchers in these studies. There is a high probability that

rectangular cantilevers were used. On the other hand, the images in Figure 3.4 are

likely to have been acquired with triangular cantilevers. Since, all the images shown

above (Figures 3.1 – 3.3) have low quality; with those obtained with triangular

cantilevers showing better outcome (Figure 3.4). More specifically, the low

resolution images in Figure 3.1, 3.2 and 3.4 with average frame size of 20 µm have

poor resolution and small depth of focus. These images were expected to be clear

enough to reveal the surface morphology of the articular surface at such low

resolutions. Although, the images in Figure 3.3 were acquired at high resolution (5 x

5 µm), the features are not clearly revealed. Therefore, there is a need for further

56

research into improving the resolution and quality of the AFM images of the articular

surface. In this study, preliminary experiment was conducted to image the surface of

normal cartilage with a rectangular cantilever. The experiment produced the images

shown in Figure 3.5. Even at a low resolution (8 by 8 µm), the AFM images are

clear; but do not reflect the nano-structural features of a normal intact articular

surface.

Figure 3.5 2-D AFM images of the surface of fresh bovine articular cartilage (a)

Topographical and (b) Deflection images (scan size: 8 x 8 µm) acquired with a

rectangular cantilever

Based on these observations and gaps outlined in the literature (Chapters two, three,

and four), it can be argued that high resolution images with good quality will provide

more information regarding the nano-structural properties of the surface amorphous

layer covering cartilage surface, thus creating more insight into the understanding of

a b

57

the role of this important membranous layer to articular cartilage function. This

thesis seeks to develop the protocol for acquiring high resolution images of the

articular surface with good quality using the AFM, and then apply this method for

the characterization of cartilage samples with normal intact, delipidized and

relipidized surfaces. The factors already highlighted above for obtaining high

resolution and quality images with the AFM will be considered and optimized to

achieve the objectives this study. The factors are: selection of suitable cantilever,

optimization of set-points and scanning/imaging parameters, and real-time

monitoring of trace and retrace signals with the oscillograph during scanning. These

factors are explained in the following sections.

3.1.1 CHOICE OF CANTILEVER FOR AFM IMAGING

Generally, cantilevers for AFM measurements can either be triangular or rectangular.

AFM cantilevers are usually triangular or V-shaped because of their high lateral

stiffness (stability) (Butt, et al., 2005). Triangular cantilevers are expensive, thus

making the rectangular-shaped cantilevers more cost effective alternative for AFM

applications. One of the key challenges in this research is to determine the most

suitable cantilever for imaging the surface of cartilage in a liquid environment.

Several preliminary experiments were conducted using AFM tips with rectangular

cantilevers; the results showed that these cantilevers were not suitable for imaging

the articular surface. On the other hand, V-shaped cantilevers (with low

stiffness/force constant) were found more suitable for imaging cartilage surface in

liquid medium (Yusuf, et al., 2011; Yusuf, et al., 2012).

58

Figure 3.6 Schematic view of a set of triangular and rectangular AFM cantilever

carrying silicon nitride tips. These cantilevers have extremely low spring constants,

thus suitable for imaging in air and liquid environments both with contact and

tapping mode (Bruker AFM Probes, Madison, WI, USA).

Triangular

cantilever

Rectangular

cantilever

59

3.1.2 OPTIMIZATION OF SET-POINT AND SCANNING

PARAMETERS

After selecting the right cantilever, it is important to determine the appropriate or

safe set-point for approaching the AFM tip on the sample surface during the landing

of the cantilever. A careful approach of the tip with low set-point within the range of

0.6 - 1.2 nA was found suitable in the preliminary experiments for AFM imaging and

measuring force curves (nano-indentation) on the articular surface. This low set-point

also prevented the tip from bumping into the soft surface of articular cartilage in the

liquid medium (PBS). Uncontrolled approach can lead to bumping of the tip on the

articular surface; this could damage the tip completely or adversely affect the results

and analyses of the force curves. The feedback system, which controls the erratic

deflection of the cantilever, was switched on and the feedback gain was kept at

constant value of 1.0 during the AFM experiment.

60

Figure 3.7 (a) and (b) Screen shots of the approach profiles of the AFM tip on the

surface of cartilage during two landing processes.

After landing (a)

Before

landing

Deflection

of cantilever

Cantilever deflecting

away from the AS

(b)

Before landing

After landing Deflection of

cantilever

61

In Figure 3.7 (a), the cantilever deflects away from the articular surface (AS) after

landing. This problem was resolved by retracting the tip away (backward) from the

AS. The AFM was given 30 minutes to stabilize/equilibrate, after which the set-point

was slightly increased and the landing was repeated. The AFM settings were

optimized until a good approached was achieved as shown in Figure 3.7 (b). In this

figure, at first, as the AFM tip approaches the articular surface (AS), it deflects away

due to surrounding external forces. The feedback system immediately returns the tip

back in contact with the AS, and this contact is maintained throughout the course of

the measurement in the region of approach on the AS. This is referred to as a good

approach. The settings used to achieve this landing were recorded and used for

subsequent measurements.

Furthermore, the scanning speed/frequency was also optimized to improve the

quality of the image and to prevent the tip from damage during scanning. The

scanning frequency was set to approximately 0.3 - 0.7 Hz.

3.1.3 REAL-TIME TRACKING OF TRACE AND RETRACE

SIGNALS

Another important factor that is worth noting during imaging with the AFM is the

trace and retrace signals. Although these signals have been used as input parameters

for measuring the microscale friction coefficients of the articular surface elsewhere

in the literature (Park, et al., 2004), they can also be used for real-time monitoring of

the progress of a scanning process. To obtain a high quality image, the trace and

retrace signals must track/match each other at every position during scanning,

62

thereby yielding an almost similar forward and backward images (Figures 3.3 (a) and

(b)).

In this study, the NOVA® program (NT-MDT, Moscow, Russia), which controls the

P47-Pro Solver scanning probe microscope (SPM) (NT-MDT, Moscow, Russia), was

used. This program allows continuous/ real-time monitoring of trace and retrace

signals using an inbuilt oscillograph installed with the program. The oscillgraph is

constantly monitored to ensure that the signals are tracking each other at every

position during the scanning process (Figure 3.8 (a) and (b)). The figures show the

sequence of imaging of normal intact cartilage surface with the AFM from the

beginning of the scan to the end. The imaging was conducted with a soft triangular

cantilever, using the set-point and scanning parameters described in Sections 3.1.1

and 3.1.2. The trace and retrace signals look similar, proving that the cantilever is

reasonably stable and the results of the scan reflect the surface configuration

expected for a normal intact articular cartilage surface.

63

Figure 3.8 Schematic representation of the imaging process of normal cartilage with

the AFM, the similarity between the trace and retrace signals shows that the AFM tip

is producing a good/ high resolution image of cartilage (a) beginning and (b) end of

scan.

Trace & retrace signals

(a)

Trace & retrace

signals

(b)

64

The resulting 2-D AFM images acquired from the above scans (using triangular

cantilevers) are shown Figures 3.9 and 3.10 below:

(a) (b)

(c) (d)

Figure 3.9 2-D (a) Height or topographical and (b) Deflection AFM images of the

surface of normal intact articular cartilage (frame size: 8 µm by 8 µm) acquired for

the Forward scan; (c) Height or topographical and (d) Deflection images for the

65

backward scan. The images (a) and (c); (b) and (d) look similar as expected from the

oscillograph shown in Figures 3.8 (a) and (b). Note: the above images were obtained

with V-shaped cantilevers.

Figure 3.10 High resolution (5 µm by 5 µm) 2-D topographical images of the surface

of fresh bovine cartilage obtained with V-shaped cantilevers using the optimized

scanning parameters (a) forward scan and (b) backward scan. The forward and

backward scans are almost identical, proving the accuracy of the scanning process.

a b

66

The images presented in Figure 3.10 were acquired at a high resolution, similar to

those in Figure 3.3. However, the former images appear to have better quality when

compared to the latter despite their similarity in resolution (5 µm by 5 µm). Also, the

images in Figure 3.10 clearly reveal that the nano-structural details of the

intact/unaltered articular surface.

Contrary to the results obtained for cartilage samples imaged with triangular

cantilevers, the images presented below were obtained with a rectangular cantilever

after several failed attempts with this cantilever to obtain a high resolution image of

good quality. The set-point and scanning parameters used for the scanning are the

same as described for the triangular cantilevers. Unlike the images obtained with the

triangular cantilevers (Figures 3.9 and 3.10), the 2-D AFM images below have very

low resolution and quality. The images do not resolve the structural details of

articular surface of normal intact cartilage, which was described by Hills et al. (1990)

as a lamella-like layer of surface-active phospholipids (SAPL). From the above

preliminary experimental results, it can be argued intuitively that triangular

cantilevers are preferable for AFM imaging of the surface of articular cartilage using

the set-up, and scanning procedures, and parameters outlined in this chapter.

67

(a) (b)

Figure 3.11 2-D (a) Topographical and (b) Deflection images of the surface of

normal intact articular cartilage obtained with AFM (frame size: 8 µm by 8 µm).

This is compared with the images previously presented in the Figure 3.4, which was

assumed to be acquired with triangular cantilevers. This further supports the

argument that triangular cantilevers are more suitable for imaging cartilage surface.

68

In summary, the following steps/procedure will be used for the AFM study in thesis:

Selection of triangular cantilevers for AFM measurements (both for imaging

and force spectroscopy)

Measurement will be conducted in liquid medium (PBS)

Selection of low set-point within the range of 0.6 - 1.2 nA for landing tip on

the sample surface

Feedback gain will be maintained at 1.0 during measurements

The scanning frequency will be set to approximately 0.3 - 0.7 Hz

The trace and retrace signals will be carefully monitored during imaging.

3.2 EVALUATION OF THE SEMIPERMEABILITY OF

RESURFACED LIPID LAYER – DIFFUSION STUDY

The following methods have been used to study the diffusion /semipermeability

characteristics of articular cartilage:

osmosis-type experiment using a saline filtration chamber (osmometer) by

Chen et al. (2007)

contrast enhanced computer tomography (CT) to study the diffusion of

contrast agents in articular cartilage (Kokkonen et al., 2011)

69

Nuclear magnetic resonance (NMR) spectroscopy to study the diffusion of

small solute in cartilage (Burstein, et al., 1993)

radioactive tracer method (RTM) using of solutes labelled with radioactive

isotopes (Burstein, et al., 1993; Maroudas, 1968; Torzilli et al., 1987)

magnetic resonance imaging (MRI) spectroscopy to study diffusion/bulk

transport of deuterium oxide (D2O) or heavy water in articular cartilage

(Burstein, et al., 1993).

MRI was chosen in this research to determine semipermeability properties of

cartilage matrices with normal intact, delipidized and relipidized surfaces because of

the numerous advantages it has over other existing methods highlighted above. It is

non-destructive and non-invasive, and unlike the radioactive tracer method (RTM), it

can be used for in vivo diagnosis of joint conditions such as osteoarthritis. The full

details of the experiment are described in Chapters four and six.

More relevant to this research is the work of Chen et al. (2007), which evaluated the

capacity of SAPL to provide semipermeability to articular cartilage using an

osmosis-type experiment by measuring the selectivity of ions (Na+, Cl

-, H

+, OH

-)

across SAPL membranes in a saline filtration environment (osmometer). In their

study, an artificial membrane was created with the SAPL extracted from surfaces of

bovine cartilage. The outcome of the study, which was only restricted to the surface

of cartilage, revealed that the SAPL possesses some level of semipermeability, by

selectively allowing the passage of specific ions through it (a physicochemical

process). However, this research did not investigate how fluid (water) is transported

70

through the articular surface into the matrix. Also, noting that the cartilage matrix is

made up of several interconnected/ interrelated layers, which are arranged in distinct

zones (Glenister, 1976), it is important to understand how the articular surface

(through its semipermeability) influences the diffusion/exudation of fluid into and

out of the matrix to fully understand this tissue both in healthy and degenerated

conditions. This will be addressed in this thesis by combining MRI and

computational techniques to measure the apparent diffusion coefficients of water

through the matrices of cartilage with normal, delipidized, and relipidized surfaces.

The results will then be used to evaluate the semipermeability characteristics of the

surface membranous layer (SAL) of resurfaced/repaired cartilage relative to normal

intact cartilage surface.

MRI was used to acquire a time series of multi-spin multi-echo (MSME) images of

cartilage submerged in heavy water (D2O). The change in intensity of the MSME

images acquired with the MRI provided information that was used to track the

concentration of H2O at any given position and time in the tissue by using a

numerical iteration scheme developed with MATLAB®. The apparent diffusion

coefficient (ADC) was computed from a fit of the depth- and time-dependent signal

intensities, which contained the H2O signal intensities for every depth, every time

point in the time series obtained from the MRI data, to the solution of the 1-D

diffusion equation developed by Fick (1855) using appropriate boundary conditions

derived from the experimental set up (Crank, 1975; Burstein, et al., 1993; Kokkonen

et al., 2011). The ADC is a composite parameter, which accounts for the distribution

of depth-dependent intra and interlayer diffusion coefficient and the surface

interlayer permeability (Glenister, 1976). Also, since the SAL membrane is very thin,

the measured ADC can be approximated as a measure of its semipermeability. This

is further explained in Chapters of six and eight.

71

It also is important to note that some problems were encountered during processing

of the MR images, where some of the acquired images were not parallel to the X-

axis. The problem was resolved by writing a code in the MATLAB® program to

realign the specimen parallel to the X-axis, before the region of interest (ROI), which

is the section of the image containing only the cartilage, is selected. The selected

ROI in the sample image is the region, where the articular surface (AS) is

approximately parallel to the bone-cartilage interface. The complete MATLAB®

code and step-by-step procedure for applying the associated-graphical user interface

(GUI) developed are presented in Appendix B. The figures below are the sets of

MSME images of cartilage-on-bone acquired with the MRI spectrometer.

72

(a) (b)

Figure 3.12 (a) and (b) The articular surface is not parallel to the horizontal X-axis.

The images would have to realigned parallel to the X-axis using the custom-built

MATLAB® code before the ROI is selected.

Figure 3.13 Screen capture of the GUI for realigning the inclined images shown in

Figure 3.12. After rotating the image with the AS parallel to X-axis, the

concentration of H2O at any given position and time in the tissue is calculated from

the MR image.

73

(a) (b)

Figure 3.14 (a) and (b) Well-aligned MRI images of normal intact cartilage, where

the articular surface approximately parallel to the horizontal X-axis.

Figure 3.15 Screen capture of the GUI for calculating the concentration of H2O at

any given position and time in the tissue and the apparent diffusion coefficient of

H2O in the matrix from the MR image in Figure 3.13.

74

The summary of the procedure to be used the MR image and numerical iteration

scheme is highlighted below:

normal intact bone-cartilage plug is submerged in a NMR tube containing

D2O-PBS solution and quickly placed in the spectrometer

time course MSME images were acquired during 2.5 hr period as D2O-H2O

exchange takes place

the MR images are converted into an array of depth- and time-dependent

H2O signal intensities for every depth, every time point in the time series

the array is used as input for the least square fit (LSF) procedure for

estimating the ADC of H2O in cartilage

the procedure is repeated for the delipidized and relipidized samples

average values of the ADC is calculated and analyzed to determine its

statistical significance

the results are compared to assess the semipermeability characteristics of

the repaired cartilage relative to the normal intact samples

3.3 MECHANICAL LOADING TESTS

The load-processing capacity of articular cartilage, a direct function of its structural

integrity, is often evaluated using mechanical loading tests like any other highly used

engineering material. It is arguably the oldest and most common means of assessing

articular cartilage functional viability. Mechanical loading schemes have been

proposed and adopted for evaluating the “compliance/integrity” property of articular

cartilage to physiological loading conditions. These tests include compressive,

75

tensile, and tribological testing protocols, with the most common being those based

on compressive loading. Common compressive loading procedures include, but not

limited to, time independent static loading, cyclic loading test, quasi-static loading,

and dynamic loading test, with parameters such as stiffness, stress, strain, and strain

energy extracted from the loading profile for analysis of the material property of the

test sample. Although, tribological test, which involves the measurement of the

lubrication characteristics of cartilage surface, is an important test for the evaluation

of the resistance of repaired cartilage surface to shearing, it is not included in this

thesis and has been recommended for future studies in this research area.

Mechanical compression test was chosen in this thesis because it allows for the

measurement of the energy stored in the tissue, which is known as the strain energy

(SE). Strain energy is an all-encompassing parameter that can be used to measure the

mechanical integrity of cartilage. It is also a good indicator of the tissue’s load-

bearing capacity or overall functionality within the physiological loading range. In a

fluid-saturated material such as cartilage, strain energy is strongly related to the fluid

management by the tissue during loading or physiological function. Furthermore,

since the transport of fluid into and out matrix has been established to be influenced

by the permeability of the articular surface, it can be argued that the strain energy is

also dependent on the state or condition of the articular surface. This argument was

tested in this thesis by measuring the strain energy of cartilage samples with normal

intact, delipidized, and relipidized surfaces. The results were compared to determine

whether there is a close relationship between cartilage surface condition and the

energy stored or released by the tissue during loading, also, whether relipidization

with synthetic lipids can create a functional surface that is comparable to a normal

intact cartilage surface.

76

The mechanical compression test was conducted using a consolidometer type set-up

mounted on a material testing machine (Instron). The complete experimental set-up

is presented in Section 4.3.9 of Chapter four. The load-displacement data for each

cartilage specimen was logged during the experiment. The data for all the samples

were analysed using energy methods described in Section 7.2.4 of Chapter seven.

Prior to loading, the cartilage specimen was carefully placed such that the articular

surface was parallel to the indenter surface, and maintained in this position during

measurement, thereby avoiding any uneven load distribution that may affect the

outcome of the experiment. A typical load-displacement curve for a well-aligned

articular surface and indenter is shown in Figure 3.16.

Figure 3.16 Load-displacement curve for a well-placed indenter sitting parallel to the

articular surface. The curve is smooth showing uniform distribution of load.

77

The energy parameters derived from the load-displacement curves are presented in

Figure 3.17. The parameters provided further information on the load processing

capacity of resurfaced cartilage relative to the normal intact cartilage. The parameters

are calculated as the areas under the regions defined in the load-displacement curve

shown in Figure 3.17. Full details of the energy method and definitions of the

associated-energy parameters are presented in Section 7.2.4 of Chapter seven.

Figure 3.17 Energy diagram derived from a typical load-displacement curve, where

SE and RE represent strain energy and residual energy respectively.

During the compression tests, there were instances, where the contact between the

articular sample and the indenter was not maintained throughout loading-unloading

process. This resulted in a non-uniform distribution of load in the sample, thereby

affecting the results of the experiment. The load-displacement curves obtained for

78

these experiments are shown in Figure 3.18 (a) and (b). The problem was resolved by

removing the sample from the consolidometer and allowed to recover for 4 hrs in

saline, after which the experiment is repeated.

(a)

(b)

Figure 3.18 (a) and (b) Load-displacement curves for a badly-placed indenter not

sitting parallel to the articular surface. The curve is not smooth with uniform

distribution of load.

79

The summary of the steps/procedure to be used for the mechanical compression test

and strain energy analysis is highlighted below:

full thickness normal intact cartilage-bone laminate is placed in the

consolidometer filled 0.15 M saline and then transferred to the Instron

machine

before loading, the specimen is placed carefully such that the articular

surface is parallel to the indenter surface to avoid uneven load

distribution

sample is subjected to quasi-static loading-unloading process and the

load-displacement data for the sample is logged throughout the

measurement

the test is repeated for the corresponding delipidized and relipidized

samples

the load-displacement data is converted to energy parameters using

energy methods

the energy parameters are analysed using statistic methods

the results are compared to assess the mechanical integrity of the

resurfaced cartilage relative to their corresponding normal intact

samples

80

3.4 REMOVAL OF SURFACE LIPIDS -

DELIPIDIZATION

In this research, lipids were selectively removed from the articular surface in

accordance with the delipidization procedure described elsewhere in the literature

(Gudimetla et al., 2007) using Folch reagent (i.e. a mixture of chloroform/methanol

(2:1) v/v) (Folch et al., 1957). Briefly, the cartilage samples held on a retort stand

were carefully immersed in Folch solution with only the articular surface touching

the lipid rinsing solvent for 1, 3 and 21 min, and taking care to maintain the same

meniscus for all the samples.

The effectiveness of the delipidization process for the removal of the lipid-rich

surface amorphous layer (SAL) was examined using the AFM. Surface imaging was

conducted with the AFM; the images obtained were used to quantify the average

SAL height of the delipidized samples relative to the normal intact samples. The

experimental protocol for the AFM imaging of samples is presented in Section 3.1.

The normal and delipidized surface characterization was conducted on the same

samples, where the lipid removal was done subsequent to the imaging of the normal

intact specimen. The results of the preliminary AFM characterization are presented

below:

81

(a) (b)

(c) (d)

(e) (f)

Figure 3.19 Topographical (a, c, e) and deflection (b, d, f) 2-D Images of articular

cartilage surface (Frame size: 8 by 8µm). Normal articular surface (a, b); after 3min

delipidization in chloroform/methanol (c, d); and after 21min delipidization in

chloroform/methanol (e, f).

(a)

82

Figure 3.20 Variation of surface lipid lost (height of SAL, nm) with time following

delipidization with chloroform:methanol (2:1). Normal intact (group 1); 3 min

delipidization (group 2); 15 min delipidization (group 3); and 21 min delipidization

(group 4).

83

Figures 3.19 shows the 2-D topographical (a) and deflection (b) images acquired

simultaneously for a normal intact cartilage surface. The figures reveal that a normal

cartilage is covered by a non-fibrous 1ayer of organized surface structure. Figure

3.19 shows the 2-D topographical (c) and deflection (d) images of the surface of

articular cartilage exposed to chloroform:methanol (2:1) for 3 min. On the other

hand, Figures 3.19 (e) and (f) show the 2-D topographical (c) and deflection (d)

images of the articular surface after 21 min exposure in the lipid rinsing reagent.

Exposure of the surface of normal intact samples to Folch solution almost completely

removed the organized surface amorphous layer observed in the normal in the

normal intact samples.

Additionally, the box plot of the height of the surface amorphous layer (SAL) of

normal intact cartilage and cartilage, the surface of which has been subjected to

different delipidization times in chloroform:methanol (2:1) is shown in Figure 3.20.

In each of the delipidization groups, a decrease in the heights of SAL with time of

exposure in lipid rinsing solvent was observed. After establishing that the lipid

rinsing agent did remove SAPL from the articular surface using the protocol

described above, the delipidization process used throughout this thesis involved

soaking of kimwipes in Folch solution, and gradual wiping of the normal intact

cartilage surfaces with this soaked kimwipes. This allowed for better control of the

delipidization process, and prevents the direct soaking of the specimens inside the

aggressive lipid rinsing solvent (chloroform-methanol mixture), thereby minimizing

the possible disruption/depletion of other vital matrix components such as collagens

and proteoglycans.

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Further characterization was conducted using confocal microscopy and Raman

spectroscopy to examine the efficiency of the delipidization process. This is fully

explained in Chapters four and five of this thesis.

3.5 LIPID RESURFACING - RELIPIDIZATION

Relipidization is the process of resurfacing degraded cartilage surface with synthetic

lipids. The surfaces of delipidized cartilage were incubated to aqueous solutions

containing single SAPL species (DPPC and POPC) and complete SAPL mix, using

compositions found in human joints (Chen, et al., 2007b). The incubation was done

in a controlled environment using radial agitating chamber, which was set at a

physiological body temperature of 37oC for 24 hours (Oloyede, et al., 2008). This

incubation time has been established to be sufficient for the effective deposition of

the synthetic lipids on a lipid depleted cartilage surface (Oloyede, et al., 2008; Yusuf,

et al., 2011; Yusuf, et al., 2012). Also, the test tube containing the cartilage-lipid

solution was continuously stirred in the agitating chamber in order to simulate

physiological joint conditions (motion) and increase the rate of mass transfer or

deposition of the synthetic lipids onto the articular surface.

It is also important to note that the incubation of DPPC, a saturated SAPL, was

conducted at 43oC, in order to increase the solubility of DPPC, because it does not

dissolve readily in aqueous solution at the normal body temperature of 37oC

(Oloyede, et al., 2008). Preliminary experiments were conducted by dissolving

DPPC in deionized water at room temperature (25oC), body temperature (37

oC),

43oC and 50

oC. DPPC did not dissolve at temperatures below 43

oC, while

temperatures above 43oC were found to be high for the human body. Also, higher

85

temperatures (> 43oC) might decompose or breakdown the phospholipid chains in the

DPPC. It can be argued that for future clinical applications of synthetic SAPL as

intra-articular injection, the DPPC should be preheated to 43oC before mixing with

the other unsaturated SAPL species (POPC, SPLC, DLPC, and PLPC), and the

injection applied immediately after mixing. More information about the incubation

process is discussed in the following chapters of this thesis.

After relipidization, the treated samples will be characterized using the methods

described in this chapter. The outcome of the characterization will provide relevant

information for testing the hypothesis of this research i.e. whether or not the surface

of a delipidized articular cartilage can be replaced with a new functional surface lipid

layer using synthetic surface-active phospholipids (SAPL).

In conclusion, the AFM measurements (imaging and force spectroscopy), diffusion

study and mechanical loading tests described above will be conducted for all the

sample groups (normal intact, delipidized, and relipidized) examined in this research.

Furthermore, the methods developed in this chapter were applied in the following

chapters to obtain data for validating the hypotheses of this thesis.

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Chapter 4: APPROACH AND

METHODOLOGY

4.1 BACKGROUND

The fact that articular cartilage contains lipids has been well documented by several

researchers in the field of cartilage research (Bonner et al., 1975; Collins, et al.,

1965; Ghadially, et al., 1965; Hills and Butler, 1984; Oloyede, et al., 2004a, 2004b;

Saikko and Ahlroos, 1997; Stockwell, 1967), where it has been established that

articular cartilage contains lipids at two sites; within the cells (as intra-cellular lipids)

(Collins, et al., 1965) and in the matrix outside the cells (as extra-cellular lipids)

(Ghadially, et al., 1965); and on the outmost layer of the surface in nano-

constitutients, namely the surface-active phospholipids (SAPL), that form a layer of

microscopic thickness (Guerra, et al., 1996; Hills and Butler, 1984; Sarma, et al.,

2001). This research will focus only on the surface lipids. A large amount of research

has been conducted to investigate the role of lipids in healthy joints and equally on

the consequence of their depletion in diseased cartilage. In spite of the numerous

applications of synthetic phospholipids for the treatment of diseases as explained in

the previous chapters, there are few studies in the literature on the potential of lipid-

based treatment of joint diseases (Oloyede et al., 2008; Vecchio, et al., 1999; Yusuf

et al., 2011; Yusuf, et al., 2012). The thesis bridges this gap by exploring the

possibility of resurfacing degraded articular cartilage with synthetic phospholipids to

87

restore/repair the lost surface structural characteristics, and more generally, the

biomechanical function of the tissue.

Following the aim of this research, which is to scientifically explore the

characteristics of artificially laid synthetic phospholipid layer on the surface of

degraded articular cartilage, experimental and computational studies were conducted

to test the hypothesis that the structural and functional characteristics of a

dysfunctional articular surface can be restored or remodelled successfully. The

analyses addressed several fundamental questions which provide insight that

contribute to the understanding of the nature of the interaction(s) between synthetic

phospholipids and the surface of articular cartilage.

4.2 CRITICAL ARGUMENTS AND TESTING OF

HYPOTHESIS

To understand the nature/mechanism of the interaction of cartilage with synthetic

phospholipids, normal and artificially degraded cartilage specimens were incubated

in aqueous solutions of synthetic phospholipids with different concentrations and

combinations (saturated and unsaturated SAPL species), and under different

environmental conditions such as temperature and incubation time. These lipid

combinations were guided by compositions and quantities of natural lipid species

found in the human knee joint (Chen, et al., 2007b). All the lipids were mixed in

deionized water at room temperature.

Furthermore, a set of experiments was conducted in which single components of the

synthetic phospholipids found in the joints were exposed to artificially degraded

88

tissue samples. By using the different single components of the synthetic SAPL, the

conditions approximating lipid loss was simulated/modelled. This will be required if

we are to understand the capacity of synthetic phospholipids in any joint treatment

application. If the individual phospholipids were incapable of adhering on their own

to a delipidized cartilage surface, there may not be a reason to continue the

investigation for entire lipid mixtures as well as widen the scope to include agents

such as lubricin and hyaluronic acid. Therefore, it can be argued that assessment of

joint cartilage lipids at the individual component level is fundamental to the

resolution of the question of whether or not the hypothesized repair role of

phospholipids in joints has any real basis.

Additionally, since the mammalian joint system comprises of saturated and

unsaturated SAPLs, the use of representative species from both groups, allowed us to

test whether the lipids of both types exhibit similar/different characteristics when

cartilage samples is exposed to them i.e. either the lipids adhere to the articular

surface, diffuse into the matrix or combine the two mechanisms simultaneously or in

succession. It is hypothesized that the resurfacing process which is dependent on the

nature of interaction between the synthetic phospholipids and the articular surface

will involve adsorption, diffusion or a combination of these two mechanisms. To

resolve this, it is important to address the following fundamental research questions:

Whether or not the molecules of individual phospholipids species (saturated

and unsaturated) will disperse randomly, diffuse into the matrix or organize

themselves and then adsorb/deposit directly on to the articular surface over a

period of time, forming a bi-layered structure which creates and restores the

characteristics that are exhibited by normal intact cartilage. If the lipids

adsorb, what is the nature of adsorption? (Is it physical or chemical?) What is

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the strength of adhesion or cohesion of the adsorbed phospholipids to the

articular surface, and will this bonding be strong enough keep the SAPL

attached to the articular surface during physiological function.

Whether or not the newly laid surface will have similar or close enough

semipermeability characteristics relative to that of normal intact cartilage

surface.

Whether or not surface lipids will influence the mechanical properties of the

matrix, such as: fluid flow pattern, matrix deformation and energy dissipation

under physiological loading conditions.

Whether or not the incubation conditions such as temperature, time,

concentrations, and mix or combinations of synthetic lipid species (saturated

and unsaturated) will influence the outcome of the resurfacing process.

Whether or not the surrounding joint nutrient molecules such as hyaluronic

acid, lubricin, and synovial fluid will influence the outcome of the resurfacing

process.

The answers to these questions will be critical to the understanding of the manner or

mode in which synthetic lipids behave when in contact with articular cartilage, and

more importantly whether or not the newly laid cartilage surface will exhibit similar

structural and functional characteristics when compared to normal healthy cartilage

surface. The questions raised above will be addressed when testing hypothesis

developed in this research. Also, based on the established facts, gaps identified in the

literature, and research questions raised, this thesis hypothesizes that the:

90

Exposure of degraded articular cartilage surface to solutions of individual species

or mixtures of synthetic phospholipids will restore or recondition it structurally

and functionally to a level close the normal healthy articular cartilage condition.

While this hypothesis might appear simple, to prove that the properties of degraded

articular cartilage can be restored or remodelled, it is a huge task because of the

complex nature of articular cartilage. Additionally, it is important to note that the

first challenge in testing the above hypothesis was to determine whether or not it is

possible to replace or resurface the lost surface amorphous layer (SAL) membrane on

a degraded cartilage through relipidization or incubation in solutions containing

synthetic phospholipids. After a successful relipidization, the structural and

functional characteristics of the newly laid surface were assessed to determine its

mechanical strength and degree of functionality relative to normal intact cartilage

surface.

The surface structural properties were measured using microanalytic characterization

techniques which involved the combination of the following methods: confocal

microscopy (COFM), atomic force microscopy (AFM) and rigorous image

processing to compare the images and nanostructural surface characteristics of

cartilage specimens with the three surface conditions (normal, delipidized and

relipidized). More importantly, the properties of the resurfaced matrices, which were

obtained after relipidization or incubation of artificially delipidized cartilage samples

in aqueous synthetic phospholipids, were compared to their corresponding normal

intact specimens. In order to completely resolve the topographical features or surface

configuration of samples with the three surface conditions, the images acquired with

the atomic force microscope (AFM) were subjected to further image processing

(using WSxM, v4.0 Beta 1.3, an image processing and analysis software by Nanotec

91

Electronica, Spain) (Horcas et al., 2007). These two techniques have never been

combined before to analyse surface of relipidized articular cartilage.

The second challenge in this thesis is to evaluate the viability/functionality of the

repaired cartilage samples by comparing the nano-biomechanical characteristics

(such as force distribution and average elastic strain energy), the diffusion or

semipermeability characteristics (such as apparent diffusion coefficients) and the

macro-mechanical properties (such matrix deformation, total strain energy, elastic

energy lost in recovery and residual elastic energy) of the resurfaced cartilage

specimens relative to their normal counterparts. The nano-biomechanical

characterization was conducted through nano-indentations of the cartilage matrices

using the atomic force microscope (AFM), the diffusion study was conducted by

combining magnetic resonance imaging and computational analysis, and the macro-

mechanical properties were measured using mechanical loading tests. Magnetic

resonance imaging (MRI) was used to measure the diffusion of water across the

articular surface of cartilage matrices with normal intact, delipidized, and relipidized

surfaces. A purpose-built computational scheme developed with MATLAB® was

then used to fit the appropriate solution of the generalized diffusion equation to the

experimental data (Burstein, et al., 1993).

In general, it is strongly believed that the outcome of this research will enhance

understanding of the role of synthetic phospholipids, the mechanisms involved in

their interaction with articular cartilage, as well as provide significant information for

research involving the manufacture of drugs and injections for the treatment and

management of joint diseases, more importantly osteoarthritis. The protocols and

experiments needed to achieve the objectives of this research are described in detail

in the next sections. Figure 4.1 shows the idealized flow chart of the thesis from the

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conceptual stage through the development of the hypothesis to its testing, and

measurement of key parameters for the assessment of the functionality of the

resurfaced/repaired cartilage.

Figure 4.1 A conceptualized flowchart for the research, showing the different steps

followed to achieving the objectives of this thesis.

93

4.3 EXPERIMENTAL STUDY

4.3.1 SAMPLE PREPARATION

Articular cartilage samples used in this study were visually normal and intact. The

samples were prepared from the patellae of 3-4 year old healthy joints of bovine

animals (n = 40) harvested from a local abattoir within 24 hrs of slaughter and stored

at -20oC until required for testing. The samples were thawed in continuous running

water at room temperature and kept in a saline (0.15 M NaCl)/phosphate buffered

saline (PBS) solution prior to testing. Osteochondral plugs, full thickness articular

cartilage-bone laminate, were taken from the thawed joints and trimmed into

specimens of specific dimensions depending on the type of experiment they are to be

used for. During the entire sample preparation procedure, care was taken to maintain

the integrity of the samples.

4.3.2 DELIPIDIZATION PROCESS - SURFACE LIPID

REMOVAL

Delipidization was applied as a model for lipid loss in this research because the

method has already been established and utilised previously to study cartilage

surfaces in the degenerative state (Gudimetla et al., 2007; Oloyede, et al., 2008;

Oloyede, et al., 2004a, 2004b; Yusuf, et al., 2011; Yusuf, et al., 2012). Although

cartilage contains both intramatrix and surface lipids, delipidization can be confined

to the articular surface. Since the objective of this thesis is to selectively remove only

the surface lipids, delipidization was confined to the surface of cartilage. This was

MRI &

CONSOLIDATION

TEST

FINITE ELEMENT

ANALYSIS

94

achieved by gradual wiping of the cartilage surface with lipid rinsing agents to

selectively remove the surface lipids.

Delipidization is a method used for artificially extracting/removing lipids from

biological materials such as cartilage. There are various methods of delipidization,

which include; mechanical, enzymatic and chemical delipidization. Mechanical

delipidization involves the use emery cloth or sandpaper or glasspaper of different

grit sizes to carefully wipe out the surface amorphous or phospholipid layer (SAL) of

articular cartilage. The choice of Sandpaper or the grit size of Sandpaper to be used

for delipidization depends on the level of SAL removal required, but it is often

difficult to control the amount SAL that is removed. The enzymatic delipidization

uses an enzyme, known as phospholipase (for example, A1 and A2) to hydrolyse or

breakdown the phospholipid chains present in the SAL in order to achieve the lipid

removal. Although the actions of enzymes are specific, and are able to target the

component or compound of interest, the use of enzymes for cartilage delipidization is

not common.

A more common, well established, and more often used method for extracting lipids

in tissues and membranes is chemical delipidization. The most popular chemical

lipid extraction method was developed by Folch (1957). This method uses the Folch

reagent/solution which is made up of chloroform and methanol, usually in the ratio

2:1 (Folch et al., 1957). It is important to note that other reagents such as ethanol,

propylene glycol, have been used for the removal of lipids inside the matrix

(Gudimetla, et al., 2007; Oloyede, et al., 2004b) and on the surface of cartilage

(Yusuf, et al., 2011; Yusuf, et al., 2012), however, in thesis, chloroform-methanol

(2:1) solution was used. This is because, in our opinion it is more aggressive, and can

achieve a much quicker lipid extraction, thereby reducing the cartilage sample

95

exposure time in the lipid rinsing agent. In doing so, the integrity of the samples is

preserved after the delipidization process, thus, the sample is good enough for

subsequent experiments (Gudimetla, et al., 2007).

4.3.3 RELIPIDIZATION PROCESS (INCUBATION IN LIPID-

FILLED ENVIRONMENT)

Relipidization is the process of reintroducing synthetic lipids either by intra-articular

injection into the joint to repair a degraded cartilage, or by in vitro incubation of

lipid-depleted cartilage in solutions of synthetic lipids in a controlled environment

(Oloyede, et al., 2008). The in vitro relipidization process was used in this study. As

mentioned in the previous chapters, the nano-thick uppermost membranous layer of

articular cartilage consists of saturated and unsaturated phospholipids in different

compositions. It is not known at this stage whether the entire mixture is needed to

effectively resurface a degenerating tissue. It is therefore logical to study the nature

of interactions of the individual components with delipidized cartilage, before

extending the study to mixtures containing all of the components in the right joint

compositions.

This thesis considered three case scenarios for testing the potential of resurfacing

artificially degraded cartilage using aqueous solutions of synthetic surface-active

phospholipids. The first and second scenarios are pilot studies in which two single

components of the SAPL mix found in the mammalian joints are used to establish the

fundamental basis of articular cartilage-synthetic lipid interactions upon which other

experiments were based. The two representative synthetic lipid species used in these

experiments are palmitoyl-oleoyl-phosphatidylcholine (POPC) and dipalmitoyl-

phosphatidylcholine (DPPC), for case one and case two respectively. They both

96

represent the saturated SAPL (DPPC) which is the least in composition (8 wt. %),

and unsaturated SAPL (POPC) which is twice the composition of DPPC (17.5 wt. %)

of joint surfactant, which were selected as representative components in this thesis.

In the third case scenario, the entire joint SAPL species was used. A detailed

description is provided in the subsequent sections of this thesis.

4.3.3.1 CASE 1

In this case, the effect unsaturated lipids was tested using POPC component, one of

the most abundant (17.5 wt. %) unsaturated phospholipids. The test specimens were

placed in labelled test tubes containing 5ml of 1 wt. % of POPC in aqueous solution

(Avanti Lipids, Alabama, USA). The test tubes containing the specimens were

placed in a radial agitating incubator which was maintained at 37oC, and the samples

were incubated for 24 hours as required. Preliminary experiments revealed that this

time interval was enough for the effective deposition of the synthetic lipids on the

cartilage surfaces (Oloyede, et al., 2008; Yusuf, et al., 2011; Yusuf, et al., 2012).

However, it is believed that further analyses to determine the bond strength could

reveal a need for optimization.

4.3.3.2 CASE 2

This second scenarios involved testing the effect of the only saturated phospholipids

(DPPC) present in the joint. The test samples were placed in labelled test tubes

containing DPPC (Avanti Lipids, Alabama, USA), with the same concentration as

POPC (1 wt. %). The test tubes were placed in a radial agitating incubation chamber

which was set at 43oC incubated for 24 hours. This was done to increase the

97

solubility of DPPC, because it does not dissolve in an aqueous solution at the normal

body temperature of 37oC (Oloyede, et al., 2008).

4.3.3.3 CASE 3

The analyses of the composition of the SAPLs in the mammalian knee joints reveal

that they mostly contain unsaturated phospholipids; 30% palmitoyl-

linoleoylphosphatidylcholine (PLPC), 23% dilinoleoyl-phosphatidylcholine (DLPC),

17.5% palmitoyl-oleoyl-phosphatidylcholine (POPC) and 16% stearoyl-

linoleoylphosphatidylcholine (SLPC), 8% saturated dipalmitoyl-phosphatidylcholine

(DPPC) (Chen, et al., 2007b) (Table 4.1). In this study, the relipidization process was

extended to include all components of the SAPL species found in the joint, in their

right compositions. The specimens were placed in labelled test tubes containing 5ml

of 1 wt. % of the complete SAPL mix in aqueous solution (Avanti Lipids, Alabama,

United States of America). The test tubes were placed in a radial agitating incubator

and maintained at 37°

C. The samples were incubated for 24 hrs as required.

Preliminary studies (Yusuf, et al., 2011; Yusuf, et al., 2012) demonstrated that this

time interval was enough for effective deposition of the synthetic lipids on the

cartilage surface.

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Table 4.1Surfactant species in bovine joint (Chen, et al., 2007b).

Surfactant

Shorthand

description

Percentage in the knee

joint

Palmitoyl-

linoleoylphosphatidylcholine

(PLPC)

16:0/18:2

30%

Dilinoleoyl-phosphatidylcholine

(DLPC)

18:2/18:2

23%

Palmitoyl-oleoyl-

phosphatidylcholine

(POPC)

16:0/18:1

17.5%

Stearoyl-

linoleoylphosphatidylcholine

(SLPC)

18:0/18:2

16.0%

Dipalmitoyl-phosphatidylcholine

(DPPC)

16:0/16:0

8%

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4.3.4 ATOMIC FORCE MICROSCOPY (AFM)

Atomic force microscopy (AFM) is the most widely used form of scanning probe

microscopy (SPM) with applications in surface, material and biological sciences

(Fotiadis et al., 2002; Hörber and Miles, 2003; Humphris et al., 2005; Müller et al.,

2000). AFM is chosen for this work because of its several advantages over other

conventional electron microscopy such as scanning electron microscopy (SEM),

transmission electron microscopy (TEM), scanning transmission electron microscopy

(STEM), low-voltage electron microscopy and reflection electron microscopy.

Firstly, AFM has the capacity to provide very high resolution images of test

specimens which allows for the observation of surface topographic features with

nanoscale resolution that may not be seen with other imaging tools (Jurvelin, et al.,

1996; Radhakrishnan and Mao, 2004). Secondly, with very easy image processing,

AFM produces true 3-D surface profiles of samples while other imaging tools only

give 2-D projection images.

Furthermore, and more importantly to this study, is the fact that the AFM does not

require any special sample treatments such as metal/carbon coatings (applied in SEM

measurements) that would irreversibly change or damage the sample. In particular,

keeping the articular surface as intact as possible during measurements on a

nanoscale level is crucial to this study, because any modifications due to external

influence during sample preparation will alter the experimental results. However,

scanning electron microscopy (SEM) and transmission electron microscopy (TEM)

require an expensive vacuum environment for proper operation. Most AFM modes

can work perfectly in ambient air or even a liquid (phosphate buffered solution)

environment. This makes it possible to use the AFM to study biological

macromolecules and even living organisms (Bowen et al., 2000) (Figure 4.2 below).

100

Figure 4.2 The NT-MDT atomic force microscope and video camera placed in a

sound proof compartment to minimize external vibration

In spite of the numerous benefits of AFM, it also has some setbacks. It is only able to

scan very small image sizes. Unlike the SEM and TEM that can image sample areas

on the order of millimetres by millimetres (mm x mm) and a field depth on the order

of millimetres (mm), the AFM can only image a maximum height on the order of

micrometres (µm) and a maximum scanning area of around 150 by 150 µm. Also,

the scanning speed of the AFM is slow compared to the other electron microscopes.

101

It takes several minutes for a typical scan with the AFM, while an SEM is capable of

scanning images at near real-time, but overall, the entire experimental procedure,

which comprises of equipment set-up, sample preparation and imaging with the

AFM is much quicker than the SEM and TEM. Generally, AFM produces a better

scan quality/resolution than the SEM and TEM.

4.3.4.1 PRINCIPLES OF OPERATION

The atomic force microscope comprises a cantilever with a tip (probe) at the free end

of the cantilever. The tip is usually made of silicon or silicon nitride with an

extremely small radius of curvature usually in the order of nanometres. The probe,

which is mounted to a cantilever spring (arm), is used to scan the sample surface by

relative movement between the tip and the sample via the piezoelectric ceramics

attached to the scanning element. When the tip is brought very close to the sample,

the Van der Waals forces between the tip and sample leads to a deflection of the

cantilever which can be measured with sensitive optical methods (using the

photodiode detector). During scanning, the force between the probe and the sample is

determined by observing the deflection of the cantilever. If the stiffness (spring

constant) of the cantilever is known, then Hooke’s law can be applied to calculate the

interactive forces between the tip and the sample using the cantilever deflection (Butt

et al., 2005; Heinz and Hoh, 1999).

102

Topographical images of specimens can be acquired with the AFM using two

methods. Firstly, the images can be acquired by plotting the tip-sample distance that

is measured with the piezoelectric translator (height position of the piezoscanner).

This height position (tip-sample separation) is controlled by a feedback electronics

system, and the feedback loop maintains a constant force between the probe and

sample (Figure 4.3). On the other hand, a topographic image can be obtained by

plotting the deflection of the cantilever against the sample position. This is also

known as a deflection image (Figures 4.4 and 4.5).

Figure 4.3 A schematic of AFM operation (Peter, Atomic Force Microscopy).

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Figure 4.4 2-D topographical image of the surface of Teflon (Frame size: 8 by 8µm)

obtained with the AFM.

Figure 4.5 2-D deflection image of the surface of Teflon (Frame size: 8 by 8µm)

obtained with the AFM.

Partially smooth surface pattern

Deflection image surface pattern

104

4.3.4.2 MODE OF OPERATION

Primarily, atomic force microscopes are designed to operate in two modes; static and

dynamic. Based on the separation between the tip (stylus or probe) and the sample

during scanning, AFMs can be classified further into three modes namely; contact

mode, semi-contact mode and non-contact mode. The contact mode is commonly

known as the static mode, while semi-contact (tapping mode) and non-contact are

referred to as a dynamic mode. In the static mode, the tip drags across the specimen

surface while in contact. However, in the dynamic mode, the cantilever is oscillated

by an external piezoelectric element at or about the resonant or fundamental

frequency which is usually between 5 - 400 kHz.

In addition, the choice of mode of operation depends on two factors, namely: the

nature of the substrate to be analysed and the parameters to be evaluated from the

AFM measurements. For example, in the contact mode the probe is perpetually in

contact with the sample surface during scanning and the force between the tip and

the surface is kept constant during scanning by maintaining a constant deflection.

While in the non-contact mode, the probe is far enough and does not touch the

surface. In the semi-contact (tapping mode), which is intermediate between the

contact and non-contact mode, intermittent contact occurs. It is observed that when

analysing soft samples such as cells, cartilage and biopolymers, the cantilever drags

across the surface. This may result in the tip and the surface of the tissue getting

damaged because of the direct contact, but with the use of cantilevers with low

stiffness (soft cantilevers), this problem can be minimized. However, the dragging

effect is completely eliminated in the non-contact mode.

105

Furthermore, in the tapping mode, the dragging effect is better controlled because,

during imaging, the probe makes oscillatory contacts on the sample surface. Thus

making it more suitable for imaging cartilage surface, but results from preliminary

experiments have shown that we can effectively image cartilage surface in contact

mode with very soft triangular cantilevers. Also, to image cartilage in tapping mode,

there is need to determine the resonant frequency. This parameter is very difficult to

measure for triangular cantilevers, especially in a liquid environment. Therefore with

soft triangular cantilevers, the contact mode offers an easy and faster method for

imaging cartilage (Jurvelin, et al., 1996).

In addition, imaging of cartilage samples in this work was done in a liquid

environment (phosphate buffered saline, PBS) to preserve the integrity of the tissue

during measurements and also, make the samples suitable for further testing. This

was achieved using the SMENA head, which was specifically designed by NT-MDT

for liquid and biological applications. Figures 4.6, 4.7 and 4.8 below show the

SMENA head and its set-up for scanning experiments.

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Figure 4.6 The SMENA head of the NT-MDT SPM for scanning in liquid

environment.

Figure 4.7 SMENA head for measurements in a liquid environment (NT-MDT).

.

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Figure 4.8 Schematic view of scanning in a drop of liquid with SMENA head (NT-

MDT).

4.3.4.3 ATOMIC FORCE MICROSCOPY (AFM) TIP/STYLUS

The AFM tip or stylus is the most important element of the AFM, as both imaging

and force spectroscopy are conducted with the tip. It is a high precision micro-

fabricated material, with a very sharp point made of hard material such as silicon,

silicon oxide or silicon carbide. The tip is mounted on the cantilever which allows for

its vertical movements (up and down in the z-direction). It has been established that

resolution of an AFM depends on the shape and size of the tip (Dreyer and

Wiesendanger, 1995) (Figures 4.8 and 4.9).

108

Figure 4.9 Schematic view of an AFM tip captured with a focused ion beam (FIB).

109

Figure 4.10 Schematic view of an AFM tip that is carried by a flat cantilever

captured with a scanning electron microscopy (SEM).

The AFM tip is often carried by a long (usually between 10-200 µm), specially-

designed cantilever. The performance of the AFM relies largely on the mechanical

properties of the cantilever, which is characterized by its spring constant and the

Tip/Stylus

110

resonant frequency (Butt, et al., 2005). These parameters largely depend on the

geometry of the cantilever such as length, size (width and thickness), shape

(rectangular or triangular), and the properties of the material from which it is

manufactured. In principle, the spring constant and the resonant frequency can be

calculated from the material properties and geometry/dimensions of the cantilever,

and this process is known as cantilever calibration. For example, the spring constant

and resonance frequency for a rectangular cantilever (with a constant rectangular

cross sectional arm) can be calculated from the equations (Butt, et al., 2005):

(4.1)

(4.2)

Where, is the spring constant of the cantilever (Nm-1

), F is the applied force (N),

is the deflection of the cantilever at its end (m), E is the Young’s modulus of the

material of construction of the cantilever (Pa), is the cantilever width (m), t is the

cantilever thickness (m), L is the cantilever length (m), is the resonance frequency

of the cantilever (Hz), and is the density of material of the construction of the

cantilever (kgm-3

).

For non-rectangular cantilevers, equations (4.1) and (4.2) would have to be modified.

For triangular or V-shaped cantilevers, the spring constant at first

approximation using the parallel beam approximation can be expressed as

(4.3)

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Equation (4.3) above is similar to equation (4.1) multiplied by a factor of 2, this

accounts for the width of the two arms of the rectangular beam (2 ) that makes up

of the V-shaped cantilevers (Figure 4.10) (Butt, et al., 2005). A more accurate

expression for calculating the spring constant of triangular cantilever derived by

Sader (1995) which is more generally used is given as

(4.4)

Where is the opening angle of the cantilevers as shown in Figure 4.11. In this

thesis, the method developed by Sader (1999) was used to calibrate the triangular

cantilevers (Butt, et al., 2005; Sader et al., 1999; Sader et al., 1995).

Figure 4.11 Schematic top view of an AFM triangular cantilever (Butt, et al., 2005).

112

Cantilevers are usually made of silicon and silicon oxide coated with a pure native

oxide layer of 1-2 nm thickness (Butt, et al., 2005). The top and bottom sides of

cantilevers are often coated with a layer of gold in order to increase its reflectivity.

The most critical factor for any cantilever is its sensitivity; therefore a good

cantilever must have a high sensitivity (Butt, et al., 2005). Also, a good cantilever

should have a high lateral stiffness. In our opinion, since triangular cantilevers have

two arms upon which the tip is mounted on, they tend to have a higher lateral

stiffness than rectangular cantilevers (Figures 4.12 and 4.13). One of the key

challenges and breakthrough in this thesis was to determine the most suitable

cantilevers for imaging cartilage in a physiological saline/liquid environment.

Through rigorous preliminary experiments, it was discovered that a soft triangular

cantilever was suitable for imaging articular cartilage surfaces (Yusuf, et al., 2011;

Yusuf, et al., 2012).

Figure 4.12 Schematic view of an AFM rectangular cantilever for contact and

tapping modes (NT-MDT, Moscow, Russia).

Rectangular cantilever arm

Tip/Stylus

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Figure 4.13 Schematic view of an AFM triangular AFM cantilever carrying silicon

nitride tip suitable for contact and tapping modes (Advanced Integrated Scanning

Tools for Nanotechnology).

Apart from application for sample imaging, the AFM can be used to accurately

measure forces in nano to pico scale. This has made it possible to measure and detect

intra- and inter-molecular interaction forces between atoms and mechanical

properties of molecules. Generally, the AFM possesses several properties that make

it ideal for measuring forces (also known as force spectroscopy), including high

displacement sensitivity (of the order of 0.01 nm), small tip-sample contact area (in

the range of 10 nm2), and the ability to operate in a physiological liquid environment

(Green and Sader, 2002; Green et al., 2002). Working in a liquid medium eliminates

capillary forces that can interfere with the experimental results (Weisenhorn et al.,

Tip/Stylus

114

1989) and also preserves the integrity of the sample. A detailed protocol for AFM

imaging and force measurements will be discussed in the next section.

4.3.4.4 IMAGING AND FORCE SPECTROSCOPY

In this study, surface topographical images and force curves were obtained for

cartilage samples with different surface conditions. Each sample was secured to a

sample holder, and then submerged in PBS solution ready for AFM measurements

using the SMENA® head of the NT-MDT P47 Solver scanning probe microscope

(SPM) (NT-MDT, Russia). The imaging was performed with very soft triangular

cantilever (spring constants of between 0.05 – 0.10 kN/m) carrying appropriate

contact tips (Veeco probes, California, USA) (Yusuf, et al., 2011; Yusuf, et al.,

2012). The triangular cantilevers were calibrated to determine the force constants,

which were used for the conversion of the cantilever deflection from nanoAmperes

(nA) to nanometres (nm) and hence, nanoNewtons (nN) using a published method

(Butt, et al., 2005; Sader, et al., 1999). In addition, preliminary experiments and

previous studies (Yusuf, et al., 2011; Yusuf, et al., 2012) showed that rectangular

cantilevers were unsuitable for imaging cartilage (Section 3.1 of Chapter three).

After mounting the specimen and setting up the AFM, and to ensure that the drift of

the cantilever deflection angle was minimized before imaging, the instrument was

allowed to undergo thermal relaxation for 30 minutes (Jurvelin, et al., 1996). Figures

4.14, 4.15, and 4.16 show the procedures involved in mounting the specimen onto

the sample holder of the NT-MDT P47 Solver atomic force microscope. The surface

topography was measured by scanning through a cartilage surface using triangular

cantilevers operating in contact mode as explained earlier. In order to minimize any

115

disruption of the articular surface caused by the tip during approach/landing and

imaging, very soft triangular cantilevers.

Figure 4.14 NT-MDT P47 Solver Pro atomic force microscope (AFM) with

specimen mounted before measurements with the SMENA® head.

116

Figure 4.15 Articular cartilage sample mounted on the scanner head of the NT-MDT

P47 Solver Pro before measurements.

117

Figure 4.16 The versatile SMENA® head of the NT-MDT P47 Solver Pro for

imaging biological samples in liquid medium.

As previously explained in chapter 3, the trace and retrace signals were continuously

monitored with the oscillograph to ensure that they were tracking each other. Figure

4.17 shows the schematic diagram of a real-time imaging process of normal cartilage

with the AFM. The trace and retrace images look similar, thereby making the results

of the scan accurate, reflecting the surface architecture normal intact cartilage SAL.

118

Figure 4.17 Schematic representation of the imaging process of normal cartilage with

the AFM.

AFM imaging was conducted in the contact mode, and images were obtained along

the 2D planes of the articular surfaces of over 150 samples randomly selected from

the 320 normal intact specimens prepared for this study. It was ensured that each

sample set was taken from the same joint. In order to obtain high resolution images,

the deflection signal was minimized by optimizing the scanning parameters such as

feedback gain, set-point and scanning speed/frequency using the methodology

developed in Section 3.1 of Chapter three. The images and force curves were used as

standards for normal intact cartilage surfaces during the characterization process.

119

Specimens were then randomly selected from already imaged normal samples to be

subjected to delipidization.

The delipidization was conducted to artificially remove the surface lipids, thereby

simulating lipid loss in early articular cartilage degradation. The delipidization

process involved progressively wiping the surface of unaltered normal cartilage with

kimwipes that had been soaked in Folch reagent (i.e. a mixture of

chloroform:methanol (2:1)) (Folch, et al., 1957). After surface lipid removal, the

delipidized samples were immersed in phosphate buffered saline solution to recover

before AFM imaging and force spectroscopy.

Following delipidization and AFM measurements, the delipidized samples were

subjected to relipidization, which involves incubation in aqueous solutions

containing different concentrations and combinations of synthetic SAPL (both

saturated and unsaturated species) in a radial agitating incubator which was

maintained at 37oC for 24 to 48hrs. AFM imaging and force spectroscopy were

repeated for the relipidized samples. The results obtained were then compared with

those of normal and delipidized cartilage to determine the effect of the synthetic

phospholipids on the surface configuration and functional behaviour of degraded

cartilage. The outcome of the experiments will either prove or disprove the

hypothesis of this thesis.

Furthermore, it is important to note that several chemical and biomechanical

parameters can be deduced from the force curves. These are stiffness (Loparic et al.,

2010; Park et al., 2004; Radmacher, 1997; Stolz et al., 2004), binding/adhesion

forces (Heinz and Hoh, 1999; Hsieh et al., 2008), and strain energy (Yusuf, et al.,

2012). The strength of adhesion also known as adhesion/binding force can be

120

estimated from the force of attraction between the AFM probe (tip) and the

surfactant-cartilage surface. This is because the intermolecular interactions (caused

by Van der Waals forces or dipole-dipole interactions) between the surfactant

molecules and articular surface cause the probe to be retracted into the sample

surface, and this retraction force is equivalent to the adhesion force. The adhesion

force is then calculated from the force-distance curve by taking the most negative

force detected during the retraction curve (Heinz and Hoh, 1999; Park, et al., 2004),

as shown in Figure 4.18.

Figure 4.18 Single point force-distance curve obtained with an AFM tip in contact

mode.

Tip-sample separation distance (m)

Adhesion force

Forc

e (N

)

121

For a better appreciation of force curves, it is imperative to understand how a single-

point force-distance curve is obtained (Figure 4.19). In Figure 4.19 below, two

important regions can be identified. The first region is the non-contact region (I),

which corresponds to positions A to B. This region is the zero force line. The second

important region is the contact region (II) which corresponds positions B to C, was

used in this thesis to evaluate the nano-mechanical and possibly the functional

characteristics of the newly laid lipid layers relative to the normal intact cartilage

sample surfaces. Generally, for soft tissues and highly deformable materials such as

articular cartilage, the exact position of contact of the tip on the sample surface

during nano-indentation is difficult to determine accurately. Once, contact is

established, there is approximately no separation between the tip and the sample

surface, in AFM terminology, the “distance” parameter becomes “indentation” (Butt,

et al., 2005).

At the beginning of the cycle (during the approach in the non-contact region), the

distance between the tip and the sample is large, and no deflection in cantilever is

observed (I); the zero force line. As the AFM tip is gradually brought close into

contact with the sample surface at constant speed, it experiences an attractive force

from the sample and then “jumps into contact” on its surface. At this point, the force

acting on the tip is greater than the cantilever stiffness (Green, et al., 2002). The

cantilever continues to move further in towards the sample until a pre-set maximum

force (known as the set point) is reached (III). At this point, the direction of motion is

reversed, the AFM probe and sample are separated and the cycle is reversed. The

molecular interactions or adhesive force between the tip and sample keep the tip still

in contact with the sample surface as the cantilever is retracted, thus deflecting or

bending the cantilever (IV). Once the adhesion force is overcome, the tip and sample

are separated, and the tip returns back to its original position. In this thesis, we

122

measured the elastic strain energy generated as a result of the resistance of the

surface amorphous layer to tip penetration during nano-indentation. This was

calculated as the area under the contact region in the force-distance curve (region II),

the procedure for obtaining this parameter is explained in detail in Chapter five.

Figure 4.19 Schematic representation of a single point force-distance curve showing

several stages involved in force measurement with an AFM tip. The probe is brought

into and out of contact by a piezoelectric translator (carrying the chip to where the

cantilever is attached) with the specimen fixed to a point (Green, et al., 2002).

Non-contact region

A

B

C

D

123

4.3.5 CONFOCAL MICROSCOPY (COFM)

In this thesis, confocal microscopy was used to assess the outcome of the

delipidization and relipidization processes. The microstructural characteristics of the

surface of cartilage were studied with the confocal microscope while the surface

amorphous layer or superficial phospholipid layer was intact, and then compared

with the results obtained following delipidization and relipidization. The results were

used to establish the presence of SAPL on the surface of normal cartilage, determine

whether the surface lipids have been removed or degraded following delipidization

with lipid rinsing agents, and whether synthetic lipids are deposited on the surface of

the delipidized tissue following incubation in aqueous solutions of synthetic

phospholipids. The detailed microscopic protocol is explained in the following

section.

Confocal microscopy (COFM) is an optical imaging technique used for the

optimization of micrograph optical resolution and contrast, by using point source

illumination and a spatial pinhole designed to eliminate any out-of-focus light in

samples with higher thickness than the focal plane. It was discovered in 1955 by

Marvin Lee Minsky, an American cognitive scientist in the field of artificial

intelligence (AI). Confocal microscopy has numerous advantages over other

conventional optical microscopy techniques, such as: shallow depth field, elimination

of out-of-focus glare, and the ability to collect serial optical sections from thick

specimens (Nikkon Microscopy). It has several applications in biomedical sciences

and bioengineering including imaging of living fixed and living cells and tissues that

have been labelled with fluorescent dyes such as Nile red, Fluorescein, Rhodamine,

and Texas red (Figure 4.20). Figure 4.21 below shows a Leica SP5 confocal

microscope (Leica Microsystems, Germany). One major setback of confocal

124

microscopy is that the sample to be tested or imaged has to be labelled with

fluorescent probes. COFM was used in thesis because it allows for selective probing

of cartilage lipids. The staining protocol is explained in following section.

Figure 4.20 Schematic diagram illustrating the principal light pathways in a basic

confocal microscope configuration (Nikkon Microscopy).

125

Figure 4.21 Schematic of a Leica SP5 confocal microscope (Leica Microsystems,

Germany) available at the cell imaging facility of the Institute of Health and

Biomedical Innovation (IHBI), Queensland University of Technology (QUT).

126

4.3.5.1 NILE RED STAINING

The histological staining method adopted in this research has been previously

published (Fowler and Greenspan, 1985). A stock solution of Nile red was prepared

by dissolving 0.5 mg of Nile red (Sigma-Aldrich, Australia) in 1ml of acetone, and

was stored in dry ice and protected from light sources. When ready for staining, a

working solution of the dye was prepared by adding 10 µl of the stock solution to

1ml of 75 vol. % glycerol. The resulting working solution was shaken vigorously to

obtain a homogenous mixture, and a drop was added to each frozen section on the

microscopic slide and the entire slide covered with cover slips. After about 5-10

minutes of staining, images of the stained sections were captured under a Leica SP5

confocal microscope (Leica Microsystems, Germany).

4.3.6 RAMAN SPECTROSCOPY

For more rigorous characterization of the surface properties of cartilage samples used

in this thesis, chemical analyses of the surfaces of articular cartilage specimens with

normal intact, delipidized, and relipidized surfaces were conducted using Raman

spectrometer. While this method was used to establish the differences between the

chemical properties of the various samples used to test our hypothesis, it also served

a confirmatory test for the outcomes of the delipidization and relipidization

processes.

Raman spectroscopy is a photon-based spectroscopic technique that is based on the

inelastic scattering of monochromatic light, which is usually emitted from a laser

source in the visible, near infrared, and or near ultraviolet range. The frequency of

the incident monochromatic light (photons) from the laser changes upon interaction

127

with a sample (articular cartilage) thereby resulting in an inelastic scattering of the

light. As the photons pass through the sample, certain wavelengths are absorbed by

the sample molecules and then reemitted. The frequency of the reemitted photons by

the sample changes, shifting up or down relative to the incident light, this

phenomenon is called Raman effect (Raman, 1930). This frequency shift provides

important information about the vibrational and rotational transitions in molecules,

which can be used to study the biomolecular and biochemical structures and

conformations of samples in solid, liquid, and gaseous states. Figure 4.22 presents

the flowchart of Raman spectroscopic measurement of a sample.

Figure 4.22 Flowchart of Raman spectroscopic measurement of a sample.

It is worth noting that Raman spectroscopy is a complementary method to Fourier

Transform Infrared spectroscopy (FT-IR), i.e. most functional groups in biological

tissues and other materials that are IR inactive are Raman active, and vice-versa.

Sample Incident

monochromatic light for

excitation sample

molecules (E)

E (Rayleigh scattering)

The scattering pattern of the incident photons provides the

fingerprint of the chemical composition of the test sample

E3

E2

E1

128

Raman spectroscopy has several advantages over other optical spectroscopic

methods that make it suitable for chemical characterization of articular cartilage with

different surface conditions (normal intact, delipidized, and relipidized). It is non-

destructive with super fast spectra acquisition, it does not require special sample

preparation, the results are highly specific providing chemical fingerprint of the test

sample, it eliminates water interference in biological tissue, it also allows for remote

analysis of sample through light transmission by optical fibres over long distance. In

the research, Raman spectroscopy technique was used to probe changes in the

chemical properties of cartilage samples with altered surface amorphous layer (SAL)

configuration (both delipidized and relipidized samples) relative to the normal

unaltered surfaces.

Raman measurements were conducted on cartilage specimens with normal intact,

delipidized, and relipidized surfaces using the commercially available Raman

spectrometer (inVia, Renishaw, UK) (Figure 4.23). The spectrometer consists of

monochromators, a filter and a charge-coupled device (CCD) detector (Adebajo et

al., 2006). The spectra were excited by a Spectra-Physics Model 127 He-Ne laser

operating at 785 nm with a resolution of 2 cm-1

, laser power of 100 mW (normal

horizontally polarized), an exposure time of 30s and five repeated accumulations

(Adebajo et al., 2006).The spectra were obtained using Olympus BHSM microscope

with inbuilt 50X objective lens, recorded within spectral range of 800 – 3200 cm-1

using the synchro mode of the instrument software, WiRE 3.0 (Renishaw, UK)

(Adebayo et al., 2006). The spectra were calibrated using Silicon wafer. The spectra

analysis and processing were performed with both GRAMS®

software (Galactic

Industries Corporation, Salem, NH, USA) and Microsoft EXCEL®

2007

(Microsoft Corporation, Redmond, Washington, USA) spreadsheet.

129

Figure 4.23 Raman microscope (InVia Renishaw) that is available at QUT

(University of Nebraska - Lincoln, USA).

130

4.3.7 MAGNETIC RESONANCE IMAGING

Diffusion study was conducted using magnetic resonance imaging (MRI) and

computational analysis for the assessment of the effectiveness of relipidization in

restoring the altered semipermeability property of degraded articular cartilage. This

is based on the philosophy that the physiological function of articular cartilage is

primarily dependent on its fluid flow/exudation behavior. Fluid exudation is crucial

for the survival of the matrix because it controls the exchange of essential nutrients

(solutes) to the cells (chondrocytes) and carries away unwanted waste materials

(Burstein, et al., 1993; Mauck, et al., 2003). It is therefore hypothesized that any

modification or change in the articular surface membrane structure through lipid loss

will influence the tissue’s semipermeability, and hence, the effective exchange of

vital nutrients and waste materials in and out of the matrix.

The hypothesis was tested by studying the ingress of heavy water (deuterium oxide,

D2O), the diffusate through the surface of cartilage into the matrix. This was

performed for articular cartilage samples with normal intact, delipidized, and

relipidized surfaces, by combining magnetic resonance measurements and

computational techniques developed to solve the diffusion related problem (Burstein,

et al., 1993; Kokkonen et al., 2011). The apparent diffusion coefficients of water

were measured for cartilage matrices in the three testing conditions (normal,

delipidized, and relipidized). The results obtained were further used to deduce

significant information on how the surface amorphous lipid layer influences the

diffusion of water across the matrix of cartilage, and in general, the exchange of

important nutrients and waste removal. The full details of the procedures are

presented in Chapter six of the thesis.

131

It is important to note that various diffusates have been used to study role of

diffusion in the physiological state of articular cartilage (Burstein, et al., 1993;

Gardiner et al., 2007; Mauck, et al., 2003; Zhang et al., 2007). Table 4.2 presents a

list of diffusion agents/solutes, their molecular weights and measured diffusion

coefficients in cartilage matrix (Mauck, et al., 2003). Deuterium oxide or heavy

water was applied in this research, because it is denser than H2O; hence, it is able to

displace the H2 molecules of water (H2O). Also, D2O is insoluble in water; therefore

giving a higher image contrast with MRI. The MRI technique used to evaluate the

diffusion characteristics of cartilage matrices with different surface conditions has

several advantages over existing methods, such as radioactive tracer method (RTM)

(Burstein, et al., 1993; Maroudas, 1968; Torzilli et al., 1987), which involves the use

of solutes labelled with radioactive isotopes. MRI is non-invasive; thereby making it

suitable for in vivo applications. Additionally, the radioactive tracer method has a

restricted application for in vivo diagnosis or assessment of joint conditions, because

of the use of radioactive isotopes (Burstein, et al., 1993), thus making the magnetic

resonance imaging method a more viable alternative.

132

Table 4.2 The molecular weights (MW) and diffusion coefficients (D) for several

solutes in cartilage matrix, where IGF-1 is Insulin-like growth factor 1 (IGF-1), PFG

is patella femoral groove and FH is femoral head (Mauck, et al., 2003).

Solute

MW

Gel

D (m2/s)

Glucose (7 µM) 180 Mature bovine

cartilage

4.83E-10

Dextran (0.4 µM) 20000 Mature bovine

cartilage

1.58E-10

H-1H20 18 bovine cartilage 1.38E-09

Na-23NaCl (0.5 µM) 58.5 bovine cartilage 7.56E-10

F-19CF3CO2 (0.03 M) 113 bovine cartilage 5.92E-10

H-1glucose (0.05 M) 180 bovine cartilage 3.13E-10

Gd-DTPA 530 Calf FPG cartilage 1.40E-10

Gd-lysozyme 14300 Calf FPG cartilage 2.50E-11

Gd-trypsinogen 24000 Calf FPG cartilage 5.00E-12

Gd-ovalbumin 45000 Calf FPG cartilage 4.00E-12

Na+

23 Human adult cartilage 4.60E-10

Cl- 35.5 Human adult cartilage 7.20E-10

Inulin 5000 Human FH adult

cartilage

2.23E-11

IGF-1 7600 Calf bovine cartilage 6.30E-12

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Magnetic resonance imaging (MRI) is a relatively new technique for imaging

cartilage. It is largely used to visualize joint morphology from which relevant

information can be obtained for use in assessing articular cartilage health, such as:

joint shape, cartilage thickness and volume (Deborah and Martha, 2003; Eckstein et

al., 1999; Peterfy et al., 1995; Tieschky et al., 1997). MRI also offers a non-

destructive method for the biochemical analysis of cartilage. For example, it can be

used to quantify the GAG concentration in cartilage by using delayed Gadolinium

Enhanced MR imaging of Cartilage (dGEMRIC) (Bashir et al., 1997; Kurkijärvi et

al., 2004; Nissi et al., 2007; Trattnig et al., 2007).

Furthermore, MRI has several applications in tissue engineering research. It is a

powerful tool for the diagnosis, assessment and real-time monitoring of joint diseases

(Burstein et al., 2000). A major advantage of the MRI and other imaging techniques

such as atomic force microscopy (AFM), confocal microscopy (COFM), scanning

electron microscopy (SEM), and transmission electron microscopy (TEM) over

spectroscopic methods such as nuclear magnetic resonance (NMR), near infrared

(NIR), infrared (IR), and Raman spectroscopy (RS) is that the results obtained from

these imaging devices are easy to understand and interpret by scientists, surgeons and

non-spectroscopists (Chenery and Bowring, 2003). In this study, multi-spin multi-

echo (MSME) images acquired with the MRI spectrometer was used to track, in real-

time, the diffusion/egress of water through the articular surface from the matrix of

cartilage samples immersed in a deuterium oxide (D2O) environment. As earlier

mentioned, these measurements were conducted for articular cartilage specimens

under three different surface conditions (normal healthy, delipidized, and

relipidized).

134

Figure 4.24 A 4.7 Tesla Magnetic Resonance Imaging (Bruker Avance 200 MHz

NMR micro imaging/spectrometer, Germany) facility at Queensland University of

Technology (QUT).

135

Following the MRI experiments and image acquisitions, a computational scheme

developed with MATLAB®

(MathWorks, Natick, Massachusetts, United States of

America) was used to fit the experimental data to the solution of the 1-D diffusion

equation (Fick’s law) (Burstein, et al., 1993; Crank, 1975; Fick, 1855). Through this

method, the apparent diffusion coefficients of water in cartilage under three surface

conditions (normal, delipidized, and relipidized) were computed, and with these

results, the semipermeability characteristic of the resurfaced cartilage was evaluated.

It is believed that the outcome of this study can be used to further characterize and

determine the influence of the surface lipid membrane on articular cartilage function.

4.3.8 COMPUTATIONAL ANALYSIS

The change in intensity of the multi-spin multi-echo (MSME) images acquired with

the MRI spectrometer only provided information that can be used to track the

concentration of H2O at any given position in the tissue (i.e. a depth-wise

concentration profile) (Burstein, et al., 1993). However, further analysis is required

to obtain the relevant parameter for assessing the possible role of cartilage surface

lipid membrane (SAL) in the diffusion of fluid into and out of the matrix, and also,

the potential of resurfacing degraded cartilage with synthetic phospholipids. This was

achieved by using a numerical iteration scheme to convert the MSME images into

apparent diffusion coefficients (D). The apparent diffusion coefficient was computed

from a fit of the MRI data to the solution of the 1-D diffusion equation developed by

Fick (1855) using appropriate boundary conditions derived from the experimental set

up (Crank, 1975). A full description of the Fick’s law of diffusion, definition of its

associated variables and constants, development of boundary conditions for the

solution, the generalized solution of the diffusion equation, the underlying principles

136

and implementation of the experiments and analysis of the results are presented in

Chapter six.

A purpose-built computational scheme developed with MATLAB® was used to

determine the apparent diffusion coefficients (D) for cartilage specimens with normal

intact, delipidized and relipidized surfaces. This was achieved by fitting the MRI

data obtained from the experiments described in Section 6.3.6, to the diffusion

equation solution presented in Section 6.2 of Chapter six. The fitting was conducted

in accordance with the least squares method described in Appendix A. Figure 4.25

presents a snapshot of the MATLAB® GUI (graphical user interface) used for the

conversion of the MR data of a specimen whose apparent diffusion coefficients (D)

is to be determined. The full MATLAB® code for calculating the apparent diffusion

coefficient of the cartilage matrix is presented in the appendix section of this thesis

(Appendix B). A detailed step-by-step procedure for applying the GUI is also

presented in Appendix A.

137

Figure 4.25 A graphical user interface (GUI) developed with MATLAB® for

computing the apparent diffusion coefficient of cartilage matrix from magnetic

resonance imaging data

138

4.3.9 MECHANICAL COMPRESSION TEST

Fluid exudation though the articular surface of cartilage controls the load-carriage

mechanism and lubrication of the mammalian joints. The physiological function of

joints is largely dependent on its fluid flow behaviour of the fluid-saturated tissue.

The consolidation theory developed for fluid-saturated porous media such as clays

(Von Terzaghi, 1943), and later applied to cartilage by Oloyede (1991) provides an

explanation for the understanding of the underlying principle involved in load

management in the joint (Oloyede and Broom, 1991). The theory states that when a

porous, fluid saturated material, such as the cartilage is loaded; the applied stress is

initially carried by the fluid component only, after a period of time, the stress is

gradually transferred to the solid matrix as the fluid exudes the tissue, resulting in its

deformation. At the end of the fluid exudation process, a load-carriage stiffness is

developed in the solid component of the matrix, which ultimately stores part of the

energy transferred by the applied load (Oloyede, et al., 2004b).

The stored energy is referred to as the elastic strain or internal energy of the tissue.

This energy is a measure of the mechanical integrity of articular cartilage. Also, the

excess un-stored energy from the external load is the complementary energy. The

ratio of the complementary energy to the internal or strain energy is the energy ratio.

When cartilage is unloaded, a large portion of the elastic energy stored in the matrix

is released to allow for the deformed cartilage to recover; this process is

accompanied by loss of energy, known as hysteresis energy or elastic energy lost due

to matrix recovery. The unused energy left in the tissue is the residual energy. This

system of energy classification is based on the energy methods. It is noted that this

concept has never been extensively applied to articular cartilage biomechanics

139

Energy methods are used for obtaining solutions to elasticity problems, determine

deflections of structures, mechanical systems and machines (Boresi et al., 1993).

They are also known as scalar methods, because energy is scalar quantity, with a

wide range of applications both for linearly elastic material behaviour with small

displacement, and for non-linear elastic materials with large deformation behaviour

such as cartilage (McGibbon and Krebs, 2002). In this thesis, the energy method was

used to investigate the effect of surface conditions on the mechanical integrity of

articular cartilage in different surface conditions (normal intact, delipidized and

relipidized surfaces). The process involved the analysis of load-displacement data

obtained from a series of mechanical compressive load-unloading process using a

consolidometer type set-up mounted on a material testing device (Instron). A brief

description of the experimental set-up is presented below.

It is worth noting that the 1-D consolidometer arrangement with highly porous disc

placed in between the indenter and the articular surface used in the compression test

was done in order to confine the fluid exudation only through the articular surface

during loading, thereby ensuring one-dimensional conditions (Oloyede, et al.,

2004b). The consolidometer comprises of a circular frustum-like top made of

stainless steel containing an internal thick walled stainless steel cell at the lower end

with internal diameter of 14 mm which opens into a 2 mm bore, which was designed

to be connected to a Dynisco pressure transducer (USA), used for measuring

hydrostatic pore pressure (HEPP), if required. In this study, HEPP was not measured,

only the force-displacement values were obtained, these were further used to

compute strain energy related parameters for assessing the load bearing capacity of

140

the cartilage samples. The pressure transducer end was replaced with a screw to

prevent flow fluid in the horizontal direction.

The test sample, a 14 mm diameter cartilage-off-bone plug was mounted in the

consolidometer and clamped tightly using a circular clamping ring (R). Following

the mounting of the specimen, a 1.5 mm thick porous disc (P), with 10 mm diameter

was placed on the articular surface. This high porosity stainless steel disc was placed

between the articular surface and the steel indenter to allow the exuded fluid to flow

freely out of the tissue through its surface only during loading. This set-up provides a

means of studying the influence of articular surface condition on the

diffusion/percolation and exudation of interstitial fluid through cartilage surface, and

ultimately, the load bearing properties of the matrix.

The consolidometer parts and specimen are arranged such that only 10 mm of the

total 14 mm diameter area of the sample surface is available for indentation. Prior to

loading, the cartilage specimen was carefully placed such that the articular surface

was parallel to the indenter surface, thereby avoiding any uneven load distribution

that may affect the outcome of the experiment. The set up of a consolidometer and its

components is shown in Figures 4.26. During the sample mounting procedure, the

consolidometer was filled with physiological saline solution to preserve the integrity

of the tissue.

141

Figure 4.26 Purpose-built 1-D consolidometer used for quasi-static compression tests

and its parts.

The 0.15 M saline filled consolidometer carrying the cartilage specimen was

transferred on to a high sensitivity Instron material testing machine (Model 5944,

Instron Pty Ltd, Victoria, Australia), fitted with a 2 kN load cell of 0.0005 N

sensitivity (Figure 4.27). Each specimen was compressed to an equivalent

displacement of 20 % strain at a loading rate of 0.001 s-1

. After reaching the

nominated deformation, which is equivalent to 20 % strain, the load was immediately

relaxed and the tissue was allowed to return to equilibrium loading position (zero

Clamping ring

(R) with O ring

Porous disk (P)

Consolidometer

Screw replacing

the pressure

transducer

142

strain) within a time interval of 5 s. The load was applied on top the porous disc (P)

placed on the articular surface using a 3 mm plane-needed polished stainless steel

indenter.

It should be noted that indenter-to-articular surface contact was maintained during

the unloading (recovery) phase, and the 20 % strain chosen was sufficient to store

enough elastic energy in the cartilage during deformation, whilst preventing tissue

damage (Ficklin et al., 2007). The load-displacement data for each sample was

logged throughout the experiment. After completing the test for the normal intact

samples, the test was repeated for their corresponding delipidized and relipidized

counterparts. The full description of the experimental protocol, data acquisition and

result analysis are presented in Chapter seven.

143

Figure 4.27 High sensitivity material testing machine (Instron), with the

consolidometer carrying the cartilage specimen mounted on. The quasi-static

compression test was conducted on this rig.

Consolidometer carrying cartilage sample

mounted on a material testing machine

144

Figure 4.28 Computer set up with Bluehill® software installed for real-time data

collection of data to the compression tests.

Bluehill Software®

interface for data collection

145

Figure 4.29 Typical load-displacement curve obtained for normal intact cartilage

sample from the Instron machine.

Unload

Loading curve

Unloading curve

147

Chapter 5: MICROSCOPIC AND CHEMICAL

CHARACTERIZATION OF THE

SURFACES OF NORMAL,

DELIPIDIZED, AND RELIPIDIZED

ARTICULAR CARTILAGE

5.1 INTRODUCTION

The surface amorphous layer of articular cartilage is of primary importance to its

load bearing and lubrication function. This lipid-filled layer is degraded or

eliminated when cartilage degenerates due to diseases. This chapter examines the

characteristics of the surface overlay on articular cartilage using a combination of

optical microscopy (with a confocal microscope) and nanosurface characterization

(with an atomic force microscope) methods to evaluate the hypothesis that the

surface of articular cartilage can be repaired by exposing degraded cartilage to

aqueous synthetic lipid mixtures. Also, the tests were further extended to assess the

nano-structural, chemical, and functional characteristics of the newly laid surface

amorphous layer.

This chapter investigates whether or not it is possible to “resurface” a degraded

cartilage surface with synthetic lipids and attempts to determine the mechanism

governing cartilage-lipid interaction which facilitates such a process, if any. This

study examines the validity of this hypothesis by exposing cartilage specimens with

148

delipidized surfaces to aqueous solutions containing single components and mixtures

of synthetic phospholipids, and using optical (confocal) microscopy, chemical

characterization (Raman spectroscopy), and nanosurface characterization (atomic

force microscopy, (AFM)) to obtain information on the surface about biochemical

and biomechanical properties for the evaluation of the resurfaced tissue’s

functionality relative to the normal intact cartilage surface.

5.2 MATERIALS AND METHODS

The surfaces of cartilage samples under normal, delipidized and relipidized

conditions were characterized with atomic force microscopy (AFM) and confocal

microscopy (COFM), and Raman spectroscopy. The lipid overlay on the normal

specimens was removed chemically (delipidization), and lipids were re-introduced

via incubation (relipidization) in an aqueous solution of synthetic lipids at the

physiological temperature of 37oC, to restore to the artificially removed lipid layer.

5.2.1 ATOMIC FORCE MICROSCOPY SAMPLES

As stated in Section 4.3.1 of Chapter four, the cartilage specimens used in these

experiments were prepared from the patellae of 3-4 year old bovine animals (n = 40)

harvested from the local abattoir and stored at -20oC until required for testing. The

samples were thawed in continuous running water at room temperature and kept in a

phosphate buffered saline (PBS) solution prior to testing. Osteochondral plugs, full

thickness articular cartilage-bone laminate (320 specimens), were taken from the

149

thawed joints and trimmed into specimens of about 5mm by 5mm. The bony layer

underlying the cartilage was dabbed with a paper towel and immediately glued onto a

Petri dish (1.5 cm in diameter) using a fast-drying Loctite® 454 glue (Henkel

Australia PTY Ltd, Victoria, Australia). The Petri dish was mounted onto the AFM

sample holder, ready for AFM measurements. During gluing, the articular surface

was moistened repeatedly with drops of PBS to preserve its integrity.

5.2.2 CONFOCAL MICROSCOPY SAMPLES

Cryostat sections were cut from normal, delipidized and relipidized samples which

were prepared for this study, and the thickness of each section was 8µm. The

sections were carefully placed on standard microscopic slides immediately after

cutting, and then the cryostat sections were left to dry in a controlled humidity

environment. A total of 20 normal, 20 delipidized and 60 relipidized (from normal

and delipidized samples) specimens were examined under the confocal microscope

shown in Section 4.3.5 of Chapter four (Figure 4.17). The results of the experiment

are presented in the results and observation section of this chapter.

5.2.3 RAMAN SPECTROSCOPY SAMPLES

Articular cartilage samples used in this experiment were prepared from the patellae

of 3-4 year old joints of bovine animals (n = 3) harvested from a local abattoir within

24 hrs of slaughter and stored at -20oC until required for testing as described Section

4.3.1 of Chapter four. The samples were thawed in continuous running water at room

temperature and kept in a phosphate buffered saline (PBS) solution prior to testing.

150

Osteochondral plugs, full thickness articular cartilage-bone laminate, were taken

from the thawed joints and trimmed into specimens of about 12 mm diameter. The

sample preparations and measurements were conducted in PBS solution in order to

keep the sample intact. Raman measurements were conducted on cartilage specimens

(n = 18, 3 samples per group) with normal intact, delipidized, and relipidized

surfaces using the protocol described in Section 4.3.6 of Chapter four.

5.2.4 AFM IMAGING AND FORCE SPECTROSCOPY

The experimental protocol for the nanosurface characterization and microscopic

imaging and Raman measurements of samples is presented in the schematic flow

chart (Figure 5.1).

Figure 5.1 Schematic flowchart of AFM imaging, confocal microscopy, and Raman

spectroscopy for normal intact, delipidized, and relipidized cartilage specimens.

INCUBATION IN

FULL SAPL

MIXTURE

NORMAL

CARTILAGE

AFM

&

COFM

&

RAMAN

DELIPIDIZATION

INCUBATION IN

DPPC

(SAT. SAPL)

INCUBATION IN

POPC

(UNSAT. SAPL)

AFM

&

COFM

&

RAMAN

AFM

&

COFM

&

RAMAN

151

5.2.5 SURFACE LIPID REMOVAL (DELIPIDIZATION)

The surface lipids on the normal intact samples were artificially removed using the

delipidization method developed by Folch (1957). The rational for this method has

been explained in Section 4.3.2 of Chapter four. Additionally, it was hypothesized

that the removal of the surface lipids (SAPL), through delipidization will alter the

chemical characteristics of the articular surface. This hypothesis was tested using

Raman spectroscopy. The protocol for the delipidization process is explained details

in Section 4.3.2 of Chapter four.

Following the delipidization procedure, AFM imaging and force measurements were

repeated for the delipidized samples (n = 60) using the same scanning parameters for

the normal intact articular cartilage specimens, as earlier described in Section 4.3.4.4

of Chapter four. After the AFM experiments, the delipidized samples were divided

into three sets, ready for relipidization.

5.2.6 RELIPIDIZATION PROCESS (INCUBATION IN LIPID-

FILLED ENVIRONMENT)

After the AFM measurements on the delipidized samples, the specimens were

randomly divided into three sets containing 20 samples each, while ensuring that

each sample set was taken from the same joint. The three case scenarios described in

Sections 4.3.3.1, 4.3.3.2, and 4.3.3.3 of Chapter four were used for testing the

potential of resurfacing delipidized cartilage with synthetic surface-active

phospholipids. The detailed procedure is presented in Section 4.3.3 of Chapter four.

152

After the relipidization process, each sample was removed from its test tube

container and then rinsed in PBS solution. The bony layer underlying the cartilage

was dabbed with a paper towel and immediately glued onto a Petri dish using a fast-

drying Loctite®

454 glue. The Petri dish containing the relipidized specimen was

mounted back onto the AFM and filled with PBS using a pipette. The AFM

measurements were repeated for the relipidized specimens using the same scanning

parameters as previously described for the normal and delipidized cartilage samples

in Section 4.3.4.4 of Chapter four.

5.2.7 CONFOCAL MICROSCOPY (COFM)

This method was adopted to evaluate the outcome of the incubation process.

Specifically, confocal microscopy was used for the qualitative appraisal of the

characteristic condition of the cartilage surface relative to the lipid type to which it

was exposed. The protocol of this evaluation is described in Section 4.3.5 of Chapter

four.

5.3 ANALYSES OF AFM IMAGING RESULTS

In order to facilitate a better understanding of the topography of the surface

amorphous layer of cartilage obtained with the atomic force microscope, the 2-D

images acquired with the Nova® program (NT-MDT, Moscow, Russia) for all the

specimens tested were exported into the image processing and analysis software,

WSxM, v4.0 Beta 1.3 (Nanotec Electronica, Spain), and then converted into 3-D

images. This was achieved by rendering the 2D images into 3D using the three-

153

dimensional package integrated in a free Windows-based application (WSxM®,

Nanotec Electronica, Spain) for data acquisition and image processing in scanning

probe instruments such as the AFM (Horcas, et al., 2007). The 2D images obtained

directly from the AFM with the Nova®

program were post-processed by exporting to

the WSxM®

software to generate 3D images (see Figure 5.6). A full description of

this procedure is available in the literature (Horcas, et al., 2007).

5.4 ANALYSES OF NANO-INDENTATION RESULTS

(FORCE CURVES)

For complete characterization of the surface amorphous layer of cartilage, the force-

distance curves obtained for normal intact, delipidized, and relipidized (incubated in

POPC, DPPC and complete SAPL mixtures) cartilage specimens, were analyzed

with MATLAB® (MathWorks, Natick, MA, USA) to obtain the average force

(distributed along the x-y plane) and penetration depth for a given specimen. The

force curves were collected at different points along the sample surface, which were

then plotted against each other and presented in the results section of this chapter.

154

Figure 5.2 Screen capture of the WSxM® software used to generate a 3D image from

a 2D image obtained with the Nova® program.

5.5 STATISTICAL ANALYSIS

Statistical analyses were conducted to determine the variations and similarities

between the groups of samples used in this experiment. It is well established that

there are inherent differences in the properties of cartilage samples taken from the

155

same patellae, and hence from different patellae. Therefore, the statistical difference

between the normal intact specimens was evaluated using analysis of variance

(ANOVA) with repeated measures and multiple comparisons according to the

Bonferroni post hoc tests. The same analysis was applied to the set of delipidized

samples. A two-tailed, unpaired Student’s t-test was also performed to establish

whether or not there is a significant difference between the normal and delipidized

samples from across the same patella. All analyses were done with respect to a

significance level of p ≤ 0.05 or a 95% confidence interval.

Since it is practically impossible to use the same delipidized samples for

relipidization in DPPC, POPC and complete SAPL mixtures, two-tailed, unpaired

Student’s t-tests were applied to determine whether there is a significant difference

between the groups of samples created from the delipidized specimens for

relipidization. Also, the statistical significance of the results of relipidization in

DPPC, POPC and full SAPL mixture relative to the normal intact specimens was

determined through Student’s t-test.

5.6 RESULTS AND OBSERVATIONS

5.6.1 CONFOCAL MICROSCOPY AND AFM IMAGING

The surface topographical images, force curves and confocal micrographs for normal

intact, delipidized, and relipidized cartilage samples are presented in this section.

156

Figure 5.3 (a) Cross-sectional view of a normal intact AS obtained with a confocal

microscope, (b) 2-D topographical image of the surface of normal intact articular

cartilage (5 µm by 5µm), (c) 3-D topographical image of normal articular cartilage

after image processing (length (X) and breadth (Y) of the scanned area, and the

average peak height of SAL (Z)), (Yusuf, et al., 2012).

157

Figure 5.4 (a) Cross-sectional view of an articular surface following the partial

removal of a lipid layer obtained with a COFM, (b) 2-D topographical image of the

surface of delipidized articular cartilage (5 µm by 5µm), (c) 3-D topographical image

of the surface of delipidized articular cartilage after image processing (length (X) and

breadth (Y) of the scanned area, and the average peak height of SAL (Z)), (Yusuf, et

al., 2012).

158

The cross-sectional view of a normal intact articular cartilage acquired with a

confocal microscope after staining with Nile Red is shown in Figure 5.3a. The figure

reveals that the surface amorphous layer forms a distinct microscopic layer which

overlays the cartilage surface (Graindorge et al., 2006; Kobayashi, et al., 1995). A 2-

D AFM topographical image of the surface of a normal intact cartilage obtained with

the atomic force microscope is presented in Figure 5.3b, and it reveals that an

organized non-fibrous layer with several peaks and troughs covers the intact cartilage

surface. Hills et al. (1990) described this structure as an oligolamella layer formed by

the SAPL.

The confocal micrograph cross-sectional view of the cartilage surface following

partial lipid removal is depicted in Figure 5.4a, while Figure 5.3a shows that of a

normal unaltered articular surface layer. The figure demonstrates that the

microscopic lipid layer was removed after wiping the articular surface with Folch

reagent. The 2-D AFM topographical image of the cartilage surface obtained after

wiping with Folch reagent reveals that the wiping almost completely removed the

organized lipid surface layer, but no fibre structure was observed in the subsurface

layer in contrast to the work of (Jurvelin, et al., 1996) and (Grant et al., 2006)

(Figure 5.8b).

159

Figure 5.5 (a) Cross-sectional view of AS following incubation in POPC for 24

hours at 37oC obtained with a COFM, (b) 2-D topographical image of the surface of

relipidized articular cartilage in POPC for 24 hours at 37oC (5 µm by 5µm), (c) 3-D

topographical image of the surface of relipidized articular cartilage in POPC for 24

hours at 37oC after image processing (length (X) and breadth (Y) of the scanned area,

and the average peak height of SAL (Z)), (Yusuf, et al., 2012).

160

Figure 5.6 (a) Cross-sectional view of articular cartilage following incubation in

DPPC for 24 hours at 43oC obtained with a COFM, (b) 2-D topographical image of

the surface of relipidized articular cartilage in DPPC for 24 hours at 43oC (5 µm by

5µm), (c) 3-D topographical image of the surface of relipidized articular cartilage in

DPPC for 24 hours at 43oC after image processing (length (X) and breadth (Y) of the

scanned area, and the average peak height of SAL (Z)), (Yusuf, et al., 2012).

161

The cross-sectional view of articular cartilage following 24 hours of incubation in

unsaturated POPC at 37oC, obtained with a confocal microscope, is presented in

Figure 5.5a. The figure reveals a distinct layer of the POPC deposits on the surface of

the delipidized cartilage after the incubation process. A 2-D AFM topographical

image of the surface of delipidized cartilage after 24 hrs incubation in unsaturated

POPC at 37oC is shown Figure 5.5b. After relipidization in unsaturated POPC, there

was a considerable change in the surface structure of the relipidized tissue in contrast

to the delipidized condition, with seemingly partial restoration of the surface lipid

structure.

The cross-sectional view of articular cartilage following 24 hrs incubation in

synthetic DPPC (synthetic lipid) at 43oC obtained with a confocal microscope is

illustrated in Figure 5.6a. The DPPC deposit is not as distinct as the POPC layer.

The 2-D AFM topographical image of the surface of cartilage after 24 hrs incubation

in saturated DPPC at 43oC is presented in Figure 5.6b. In comparison to the effect of

POPC, incubation in saturated DPPC did not result in the deposit of a distinct surface

lipid layer (Figure 5.7b). However, the DPPC molecules settled in a particulate form

with several discontinuities between the lipid aggregates inside the void spaces on

the articular surface as against POPC. The POPC molecules were deposited as lipid

bi-layers in a continuous wavelike structure on the articular surface (Figure 5.7a).

162

Figure 5.7 Schematic representation of POPC-bilayers and DPPC molecules formed

on the surface of degraded cartilage after relipidization, (a) showing wavelike

structure deposits of POPC on the articular surface, and (b) showing particle-like

deposits of DPPC on the articular surface (Yusuf, et al., 2012).

163

After conversion to 3-D, Figures 5.3b and 5.4b yielded Figures 5.3c and 5.4c

respectively. Figures 5.5c and 5.6c represent the 3-D images of Figures 5.5b and

5.6b respectively. When visualized in 3-D, the articular surface of normal intact

cartilage consists of an organized lipid layer with several peaks and troughs which

are analogous to an aerial view of a city consisting of several buildings of different

heights and surface areas (Figure 5.3c). After delipidization, the surface topography

changed drastically with the loss of numerous peaks, indicating surface lipid

depletion (Figure 5.4c). The consequence of incubation of the delipidized articular

surface in unsaturated POPC seems to suggest a level of partial surface restoration

with a pattern of peaks similar to that revealed on the normal cartilage surface

(Figure 5.5c). This implies that the POPC was able to create a potential new

functional lipid layer, while incubation in saturated DPPC did not restore the lost

lipid surface structure (Figure 5.6c).

The 2-D topographical image of artificially delipidized cartilage surface exposed to

an aqueous solution containing the full SAPL mixture in a physiological condition

(37oC) for 24 hours (Figure 8a). The relipidization resulted in a much improved

surface configuration when compared with results obtained for surfaces layered with

POPC and DPPC. The newly resurfaced cartilage with SAPL mixture containing

both saturated and unsaturated phospholipid species has almost similar configuration

as normal intact cartilage surface (Figure 5.3a). The reconstructed 3-D image (Figure

8b) also demonstrated similar patterns of SAL peaks and troughs observed in the

normal intact cartilage surfaces shown in Figure 5.3c, with almost similar continuous

wavelike lipid bi-layer structure observed in cartilage surfaces treated with POPC

(Figure 5.7a).

164

Figure 5.8 (a) 2-D topographical image of the surface of relipidized articular

cartilage in a mixture containing all five SAPL for 24 hours at 37oC (5 µm by 5µm),

(b) 3-D topographical image of the surface of relipidized articular cartilage in a

mixture containing all five SAPLs for 24 hours at 37oC after image processing

(length (X) and breadth (Y) of the scanned area, and the average peak height of SAL

(Z)).

165

5.6.2 RAMAN SPECTROCOPY

Raman was used to examine the chemical composition of the surfaces of cartilage

samples. The concentration of lipids present on the articular surface of the cartilage

samples with different surface conditions was investigated using vibrational Raman

spectroscopy. The relative weights of lipids on the surfaces of the various samples

were extracted, and their relative amounts appear to vary with the surface condition

(normal unaltered, delipidized, and relipidized). Raman spectra of bovine cartilage

exhibit features that are predominantly from protein, collagen, and lipids.

Figure 5.9 Raman spectrum of bovine cartilage collected on a Raman microscopy

system ( = 785 nm). Band assignments for the spectra are presented in Table C1 in

the appendix section (Appendix C).

The region between 3100 and 2800 exhibits the C-H stretching vibrations of –CH2

and –CH3 functional groups which are dominated by the fatty acids of the various

membrane amphiphiles (phospholipids) and by some amino acid-side chain

vibrations. The region between 1800 and 1500 cm-1 is dominated by the

166

conformation-sensitive amide I and amide II bands, which are the most intense bands

in the spectra in all samples tested. Of importance to this study, is the region

dominated by the fatty acids. The integrated intensity in the C-H stretch region in the

fatty acids was used as an indicator of the total fatty acid concentration. The C-H

stretching and -CH2 deformation spectral regions were used to monitor changes in the

lipid-chain lateral packing characteristic in the bilayer assemblies. This is presented

in Figure 5.10. The figure reveals that there is a measurable difference between the

chemical properties of all the sample groups tested.

Figure 5.10 Comparison of the Raman spectral in the C-H stretching mode region

acquired for cartilage specimens with normal intact, delipidized, and relipidized in

POPC, DPPC, and full SAPL mix.

120

170

220

270

320

370

420

2800 2850 2900 2950 3000 3050 3100

Inte

nsi

ty (

arb

. un

its)

Wavenumber(cm-1)

167

Figure 5.11 Comparison of the C-H stretching mode region peak area for cartilage

specimens with normal intact, delipidized, and relipidized in POPC, DPPC, full

SAPL mix, saline solution (control).

168

The bar chart (Figure 5.11) shows that there are observable differences in the

quantity of lipids present on the articular surfaces of all the sample groups examined,

with the normal cartilage samples having the largest amount of lipids, closely

followed by the samples incubated in full SAPL mix and those treated with POPC.

The delipidized samples and those exposed to saline (placebo) have the least lipid

levels. These results further support the outcomes of the AFM and confocal

microscopy characterization.

All the characterization results prove/confirm the effectiveness of the delipidization

process in removing the surface lipids, and modifying the chemical characteristics of

the articular surface. It can also be argued that relipidization in synthetic

phospholipids led to the reversal of the chemical properties of the delipidized

samples, edging towards the normal intact surface. Additionally, samples incubated

in complete joint SAPL mixture gave the most promising resurfacing outcome,

followed by samples incubated in single unsaturated POPC solution. Similar to the

AFM characterization results, DPPC showed little promise of remodelling the

surface properties of the delipidized cartilage specimens to a level close to the

normal intact surface.

5.6.3 AFM ANALYSIS AND FORCE SPECTROSCOPY

Table 5.1 presents the average effective height of the surface amorphous layer and

the elastic strain energy of the surface amorphous layer for normal, delipidized and

relipidized articular cartilage. The effective height of the surface amorphous layer is

169

the difference between the highest and lowest scanned peak heights by the AFM,

since it is practically impossible to determine the absolute height without knowing

the absolute bottom layer of the surface amorphous layer. The average effective

height was computed from the 2-D atomic force microscopy images of the randomly

selected samples and the values of this parameter for the specimens in the delipidized

and relipidized conditions were compared separately to those of normal intact

samples using a two-tailed, unpaired Student’s t-test.

The results of the analysis reveal that there was a considerable reduction in the height

of the surface amorphous layer after delipidization, with the normal measured as

(858.80 ± 275.95 nm) and the delipidized as (716.90 ± 149.96 nm), with p < 0.0001.

After incubating the delipidized specimens in POPC and DPPC, there was a

significant increase in the surface amorphous layer height (1794.37 ± 218.18 nm, p <

0.0002) and (1541.53 ± 153.21 nm, p < 0.0001) respectively, relative to their normal

intact counterparts. Relipidization in complete SAPL mixture resulted in the creation

of new surface amorphous layer with an effective height of 1986.19 ± 175.01nm, and

p < 0.0001 in comparison with normal unaltered cartilage samples. These results

demonstrate that wiping the cartilage surface with chloroform/methanol (2:1) results

in reducing the size of the surface amorphous layer, and incubation in solutions of

synthetic lipids results in the deposition of new lipid layers to a thickness that is

comparable or higher than that of the normal intact surface amorphous layer.

170

Table 5.1 Variation of height and elastic strain energy of the surface amorphous layer

for normal, delipidized, and relipidized articular cartilage.

Sample Average effective

height of SAL

(nm)

(Mean ± SD)

Average elastic strain

energy (J x 10-15

)

(Mean ± SD)

Normal 856.80

±

275.95

15.92

±

1.05

Delipidized 716.90

±

149.96

2.38

±

0.27

Relipidized in POPC 1794.37

±

218.18

2.68

±

0.75

Relipidized in DPPC 1541.53

±

153.21

3.21

±

0.49

Relipidized in

complete SAPL

1986.19

±

175.01

5.91

±

0.32

171

Force-distance curves plotted for samples in the three testing conditions (normal

intact, delipidized, and relipidized) reveal that the mechanical properties, and

probably the functional capacity of cartilage, are different under the three surface

conditions (Figure 5.12). In addition, the elastic strain energy generated as a result of

the resistance of the surface amorphous layer to tip penetration during nano-

indentation was calculated as the area under the contact region in the force-distance

curve (Figure 5.13); the results are presented in Table 5.1. The average strain energy

of the surface amorphous layer for normal intact cartilage was significantly higher

than that of the delipidized samples, (15.92 x 10-15

± 4.62 J, p < 0.0001) and (2.38 x

10-15

± 0.75 J, p < 0.0001) respectively.

Additionally, relipidization of the artificially delipidized samples in solutions of

POPC and DPPC, the nanoscale elastic strain energy values measured were

significantly less than that of the normal samples and marginally lower than that of

the delipidized samples (Table 5.1). These differences were statistically significant

(POPC; 2.68 x 10-15

± 0.75 J, p < 0.0001) and (DPPC; 3.21 x 10-15

± 0.49 J, p <

0.0001) when compared to their corresponding normal samples, which therefore

reveals that the individual SAPL species (POPC and DPPC) were unable to restore

the functional characteristics of the surface of the lipid depleted cartilage specimens.

Conversely, incubation of the delipidized specimens in aqueous solutions containing

the complete SAPL mixture resulted in a higher average elastic strain energy values

in comparison to those of the POPC and DPPC treated samples (5.91 x 10-15

± 0.32

J).

172

Figure 5.12 Averaged force-indentation curve for normal intact, delipidized and

relipidized cartilage in POPC, DPPC, and full SAPL mix, showing the variation of

the mechanical properties of the articular surface under the three surface conditions.

173

Figure 5.13 A typical force-indentation curve for normal intact cartilage used for

estimating the average elastic strain energy of the surface amorphous layer, a

measure of the layer’s resistance to AFM tip penetration (Yusuf, et al., 2012).

174

5.7 CONCLUSION

This study has demonstrated that it is possible to resurface degraded cartilage with

commercially available synthetic phospholipids, and the potential to achieve some

degree of articular cartilage surface repair with phospholipids, following

degeneration or wear. However, further research would be necessary to determine the

most appropriate choice, quantity, mix of the phospholipids, and the optimum

incubation conditions such as time and temperature that would be required to restore

a damaged cartilage surface to its normal intact condition or at least sufficiently close

to this state in the treatment of osteoarthritis and other related conditions.

175

Chapter 6: EVALUATION OF THE

FUNCTIONALITY OF

ARTIFICIALLY LAID LIPID

MEMBRANE FOR ARTICULAR

CARTILAGE RESURFACING

6.1 INTRODUCTION

The surface lipid layer of articular cartilage, also known as surface-active

phospholipid (SAPL) have been observed to impart highly desirable physico-

biochemical characteristics to the tissue such as semipermeability (Chen, et al.,

2007a). The capacity of the SAPL to act as a semipermeable membrane allows it to

control the diffusion of fluid and solutes into and out of the avascular cartilage tissue

(Chen, et al., 2002; Mauck, et al., 2003). As explained in the previous chapters,

when cartilage is diseased or degenerated, its surface is often the first victim of

attack, thereby resulting in lipid depletion (Hills and Monds, 1998). This loss

consequently affects the smooth physiological function of the tissue and, more

relevant to this present work, its semipermeability characteristics, thereby hampering

mobility and human activities (Ballantine and Stachowiak, 2002; Oloyede, et al.,

2004a, 2004b; Sarma, et al., 2001; Vecchio, et al., 1999). Resurfacing of degraded

cartilage surface with synthetic phospholipids might be a possible remedy for

reversing this problem (Oloyede, et al., 2008; Vecchio, et al., 1999; Yusuf, et al.,

2011; Yusuf, et al., 2012).

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The surface layer of articular cartilage described above performs unique functions,

which facilitate the well-being of the physiological joints, namely, load-spreading,

lubrication, and semipermeability (Chen, et al., 2007a; Flachsmann et al., 2005;

Pawlak, et al., 2008; Pawlak and Oloyede, 2008). Consequently, the test for viability

of any engineered surface layer replacement must include the ability of such layers to

deliver these important physiological functions. This chapter addresses the

semipermeability of synthetic lipid-based replacement for the surface amorphous

layer of degraded articular cartilage. The semipermeability of the layers of saturated

Dipalmitoyl-phosphatidylcholine (DPPC), which had been previously used in a

clinical trial for the treatment of degraded joints is examined in this study (Vecchio,

et al., 1999), Palmitoyl-oleoyl-phosphatidylcholine (POPC) an abundant unsaturated

component of the lipid mixture of the articulation joints, and mixtures containing all

the surface-active phospholipids species found in the joints, which have not been

used for such purpose before, to provide semipermeability.

In the previous chapter and our already published works (Yusuf, et al., 2011; Yusuf,

et al., 2012), it was established that it is possible to recreate the lost surface

amorphous layer of degraded articular cartilage using synthetic phospholipids. Also,

we assessed the rigidity of the layers created nanoscopically using the atomic force

microscope. In this chapter, the work is further extended to the evaluation of these

lipid-based surface overlays created with synthetic surface-active phospholipids

relative to their ability to provide a diffusion barrier for the exchange of nutrients and

expression of wastes, namely semipermeability. To this end, magnetic resonance

imaging (MRI) was used with deuterium oxide (D2O) as the diffusate/diffusion

agent, and statistical comparisons between the apparent diffusion coefficients from

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the completely delipidized cartilage surface, surfaces carrying artificial/synthetic

lipid layers, and normal cartilage with intact surface layers, was conducted.

The atomic force microscope (AFM) was used to characterize the samples

nanoscopically under the three surface conditions (normal, delipidized, and

relipidized) before MRI measurements to ascertain the exact condition of the

surfaces used in these experiments and the effect of relipidization. Finally, the MRI

data was used to determine the apparent diffusion coefficients using a computational

scheme that was developed with MATLAB®

(MathWorks, Natick, Massachusetts,

United States of America).

6.2 PERTINENT THEORY

The diffusion of fluids or solutes within a medium can be modelled based on Fick’s

mathematical theory of diffusion, which is generally known as Fick’s law (Fick,

1855). For diffusion processes reducible to a single dimension, Fick’s second law of

diffusion has the form

(6.1)

where c is the concentration of the diffusing species, in the present case water (H2O),

D is the diffusion coefficient (diffusivity) of the species in the medium (m2/s), x is

the path length or thickness of the sample (m) and t is time (s) (Crank, 1975). In this

thesis, the 1D solution is appropriate because the sample was approximated as a

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constant-thickness layer of articular cartilage, and the diffusion of fluid is restricted

through one direction as explained in the subsequent section.

To determine the apparent diffusivity of water (H2O) in articular cartilage under

normal, delipidized, and relipidized conditions using magnetic resonance imaging

(MRI), the efflux/egress of H2O out of the tissue through the articular surface was

measured experimentally. The experiment was setup such that a bone-cartilage plug

was immersed in a deuterium oxide (D2O) environment, and the efflux of H2O was

monitored in real time via a series of proton (1H) magnetic resonance imaging

measurements. The intensity of the image in each volume element (voxel), at any

given time is proportional to concentration of H2O in that voxel (Burstein, et al.,

1993). The apparent diffusion coefficient was computed from a fit of the

experimental data to the solution of the diffusion equation (6.1) using appropriate

boundary conditions derived from the experimental set up.

The generalized analytic solution to Equation (6.1) can be obtained using the

separation of variables method (Crank, 1975).

(6.2)

where the coefficients Am, Bm are dependent on the initial condition (IC) and the

eigenvalues m are dependent on the boundary conditions.

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In order to adapt the general solution given by equation (6.2) to the experimental set

up employed in this present work, one initial condition and two boundary conditions

are required. The following initial condition and boundary conditions were used:

IC = C(x, t = 0) and BCs = C(t) or C (t) at the boundary

(i) Initial condition (IC):

(6.3)

where c0 is the initial concentration of H2O in the tissue and a is the thickness of

sample (measured from the bone-cartilage interface to the articular surface)

(ii) Boundary condition at the bone-cartilage interface (BC1):

Reflective boundary condition was imposed at the bone-cartilage interface ( ).

This is due to the impermeable nature of this interface (Crank, 1975). The H2O

molecules diffuse in the positive direction. Therefore, the diffusion flux across this

interface was taken as:

(6.4)

(iii) Boundary condition at the articular surface (BC2):

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Absorbing boundary condition was imposed at the articular surface ( ). The

concentrations of H2O at this position are kept at zero, but their first derivatives are

not zero, and there is a non-zero net flux through the articular surface (Crank, 1975).

This implies that once the water has diffused out through the articular surface, it was

assumed that it rapidly convect away so as not to establish a concentration profile

outside the articular surface. Therefore, the BC at the articular surface is:

(6.5)

Solving equation (6.2) subject to the initial and boundary conditions (equations (6.3-

6.5)) yields the solution:

(6.6)

where .

The parameters and D were adjusted during the least square fitting (LSF).

The parameter D has the meaning of the apparent diffusion coefficient because it is

dependent on the distribution of the local diffusivities at different locations as well as

the permeability of the tissue. D is therefore a lumped/composite parameter (Bihan,

1995). The apparent diffusion coefficient of water in cartilage, D was determined

from a fit of equation (6.6) to the results from the MRI measurements using a

specifically designed computational scheme written in MATLAB®

. A noise term was

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also introduced to equation (6.6) during the least square fitting to account for the

positive noise level in the magnetic resonance (MR) images. This allowed for the

optimum treatment of the experimental data.


The experimental protocol for the atomic force microscopy imaging, MRI

measurements, and computational analysis is presented in Figure 6.1.

Figure 6.1 Schematic flowchart of specimen grouping, AFM imaging, magnetic

resonance measurements, and computational analysis for normal intact, delipidized,

and relipidized cartilage.

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6.3.1 ARTICULAR CARTILAGE SAMPLES

Articular cartilage specimens used in these experiments were prepared from the

patellae of 3-4 year old bovine animals (n = 6) harvested from the local abattoir and

stored at -20oC until required for testing. The samples were thawed out in continuous

running water at room temperature and kept in phosphate buffered saline (PBS)

solution prior to testing. A 13 mm diameter stainless steel punch was used to cut

twenty osteochondral plugs (5 samples per patella), containing full thickness articular

cartilage-bone laminate from the thawed joints. The samples were divided into three

groups (A, B, and C) and placed in PBS solution for 1h for rehydration and

equilibration until ready for further processing, followed by AFM and MRI

measurements. The first group of samples (A), comprising of normal intact

specimens was used for the control experiments without surface modification, while

the second (B) and third (C) groups were set aside for delipidization and

relipidization respectively. Figure 6.1 shows the schematic flowchart of the

experimental process.

The normal intact samples from the second (B) and third (C) groups already imaged

with the AFM were subjected to delipidization using the protocol already described

in Section 4.3.2 of Chapter four. After the delipidization process, the specimens (n =

25) from the two groups (B and C) were placed in PBS for 30 min for rehydration

and to remove the lipid rinsing agent and any organic solvent left on the surface of

the tissue. After rehydration, the group B specimens (n = 5) were set aside for AFM

and MRI measurements while the group C samples (n = 15) were used for

relipidization.

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Furthermore, the three case scenarios described in Sections 4.3.3.1, 4.3.3.2, and

4.3.3.3 of Chapter four were considered for testing the effect of different synthetic

surface-active phospholipid combinations on the potential of remodelling the

semipermeability of artificially delipidized cartilage. Briefly, the group C samples

already delipidized (n = 15) were further divided into three sets and then subjected to

the relipidization. After the incubation, each sample was removed from its test tube

container and then rinsed in PBS solution, prior to AFM and MRI measurements.

6.3.2 DEUTERIUM OXIDE-PHOSPHATE BUFFERED SALINE

(D2O-PBS) SOLUTION

The D2O-PBS solution that was used for the MRI experiments was prepared by

dissolving 0.5 mg mL-1

of sodium azide, NaN3 (Sigma-Aldrich, New South Wales,

Australia), and 0.99g of PBS (Sigma-Aldrich, New South Wales, Australia) in 100ml

of 99.9% D2O (Merck Chemicals, Victoria, Australia) (Burstein, et al., 1993). The

resulting solution was tightly enclosed and stored in a cool area during the entire

period of the experiment.

6.3.3 ATOMIC FORCE MICROSCOPE (AFM) IMAGING

Surface imaging was performed to demonstrate the effect of delipidization on the

surface amorphous layer (SAL) of cartilage and to determine the effectiveness of the

relipidization process (i.e., to ensure that the replacement synthetic phospholipids did

adhere or settle onto the delipidized cartilage surface), before the samples were used

for the diffusion studies.

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AFM imaging was conducted on samples from the three groups (A, B and C), with

each sample immersed in PBS solution in accordance with the protocol previously

published (Yusuf, et al., 2011; Yusuf, et al., 2012). The detailed procedure for the

imaging process is explained in Section 4.3.3.4 of Chapter four. The surfaces of

osteochondral plugs (13 mm diameter) with full thickness articular cartilage, still

attached to the underlying bones, were imaged. The imaging was carried out on the

normal, delipidized, and relipidized specimens prior to the diffusion experiments.

6.3.4 MAGNETIC RESONANCE IMAGING (MRI)

Magnetic resonance imaging was conducted using a Bruker Avance nuclear

magnetic resonance (NMR) spectrometer (Bruker BioSpin, Ettlingen, Germany) with

a 7.0 T vertical bore magnet operating at room temperature. The system was

equipped with a 1.1 Tm-1

(110 G cm-1

) triple-axis gradient set and a Micro2.5

microimaging probe. The radiofrequency (RF) coil used in the imaging was a 15

mm birdcage proton (1H) resonator (Bruker BioSpin, Ettlingen, Germany). The test

specimen was immersed in PBS solution inside a 15 mm NMR tube with custom-

made Teflon plugs designed to orient the cartilage sample with the normal to the

articular surface aligned at the “magic angle” (54.7°) with respect to the static

magnetic field B0. This facilitated the interpretation of the data by maintaining

relatively uniform transverse relaxation rates across the depth of the cartilage in the

acquired multi-spin multi-echo (MSME) images (De Visser et al., 2008a; De Visser

et al., 2008b).

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The NMR tube containing the specimen was placed inside the RF coil and carefully

mounted in the spectrometer. The position of the sample was marked on the tube and

the coil to ensure that the sample could be returned to the exact initial position for the

next experiment. This was done to ensure that the imaging plane was perpendicular

to the articular surface. After mounting the RF coil in the MRI spectrometer, multi-

spin multi-echo (MSME) 2D images of selected slices were acquired for normal

intact cartilage specimens (from group A) firstly in PBS solution. The imaging

parameters were as follows: repetition time (TR), 1000 ms, echo time, 7 ms, and one

average per slice, using a 1 mm slice thickness, a field of view (FOV) of 20 mm X

20 mm, and a 128 X 128 matrix size. Following the measurements, the sample was

removed from the tube and placed in PBS solution, while the NMR tube was allowed

to dry before the next experiment to avoid the alteration of the concentration of the

D2O–PBS solution that will be used for the diffusion study.

The sample already imaged was brought out of the PBS solution, dabbed carefully

with paper towel and its lateral sides (including the sub-layer bone) was covered with

fast-drying Loctite® 454 glue to prevent the D2O and H2O from diffusing laterally

during the diffusion experiment, thereby constraining the fluid flow to one direction

(i.e. only through the articular surface) (Kokkonen, et al., 2011). The glued sample

was placed back in the NMR tube and a measured volume of D2O–PBS solution was

added. The tube was inserted into the RF coil, mounted in the spectrometer and the

measurements previously conducted for the sample in pure PBS solution were

repeated for the specimen in D2O-PBS solution as a time course. The time taken

between the addition of the D2O-PBS solution and the acquisition of the first MSME

image was noted on the stopwatch. Subsequent MSME images were acquired during

the following 2.5 hr. Within this period, over 50 images were obtained and the time

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interval between each acquisition was approximately 150 s. Preliminary studies

showed that the 2.5 hr timeframe was sufficient for almost complete replacement of

H2O with D2O in the cartilage sample. The experiment was then repeated for the

delipidized and the relipidized samples from groups B and C respectively. After

measurements were completed, the acquired images were exported, ready for

computational analysis.

6.3.5 DETERMINATION OF APPARENT DIFFUSION

COEFFICIENTS

This section presents the numerical iteration scheme for determining the apparent

diffusion coefficients (D) from the time series of MR images. The first step of the

analysis involved converting each image in the time series (Figure 6.7) to the depth

profile of H2O signal intensity (Figure 6.8). Figure 6.8 was constituted into an array

of depth- and time-dependent signal intensities which contained the H2O

signal intensities for every depth, every time point in the time series. This array was

used as the input for the least square fit (LSF) procedure, where the entire constituted

data set was subjected to fitting over the time range (the duration of

the time series) and the range of from 0 to (estimated sample thickness) using

equation (6.6). The fit used the initial and boundary conditions in equations (6.3 -

6.5) and had five adjustable parameters namely, H2O signal intensity in the tissue

before D2O ingress ( ), sample thickness ( ), apparent diffusion coefficient (D),

exposure time of sample to the D2O-PBS solution ( ), and a noise term ( ).

The fitting was conducted in accordance with the least squares method and

incorporated a noise term with the other adjustable LSF parameters described in the

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section following equation (6.6). The iteration involved selecting a region of interest

(ROI), which is normally a flat region of the cartilage matrix from a relevant MR

image; and the sample thickness at this region determined by the software. During

this process, the concentration profiles which represent the distribution of H2O across

the depth of cartilage (Figure 6.8) for all time steps were fitted simultaneously. The

value of the apparent diffusion coefficient was determined as the value of

corresponding to the converged fit (Table 6.1). The LSF analysis was implemented

in a MATLAB®-based interface designed in-house. The detailed procedure for

calculating the apparent diffusion coefficient and a full description of the MATLAB®

code are presented in the appendix (Appendixes A and B).

6.3.6 STATISTICAL ANALYSIS

Statistical analyses were conducted to determine the significance of the measured

apparent diffusion coefficients. The mean and standard deviation of the apparent

diffusion coefficients for normal, delipidized, and relipidized cartilage samples are

presented. A two-tailed, unpaired Student’s t-test with Welch correction was used to

appraise the data. The analysis revealed a high level of statistical confidence (p <

0.05) at 95% in the differences between the apparent diffusion coefficients of the

specimens tested, namely, delipidized, relipidized (with POPC, DPPC and compete

SAPL mixture), and control (normal intact). The method of analysis of variance

(ANOVA) with repeated measures and corrections for multiple comparisons

according to the Bonferroni post hoc tests was also used to determine the statistical

differences between the different groups of samples, with the analyses conducted

using a significance level of p < 0.05 or a 95% confidence interval.

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The topographical images of articular cartilage specimens with normal intact,

delipidized, and relipidized (in aqueous solutions of POPC and DPPC) surfaces were

acquired with the AFM (Figures 6.3 - 6.5). An investigation of the 3D resolved AFM

image of the unaltered cartilage surface (Figure 6.2) showed a unique organization of

the SAL in a lamella-like arrangement as previously described by Hills et al. (1990).

Wiping of the intact surface with lipid rinsing reagent resulted in a noticeable change

in the surface topography (Figure 6.3). Thus, confirming the effectiveness of the

delipidization process. On the other hand, relipidization in POPC partially restored

the lost SAL nanostructural surface patterns (Figure 6.4); and the incubation in

DPPC did not yield any significant improvement in the surface configuration relative

to normal intact cartilage surface (Figure 6.5). Relipidization in aqueous solution

containing the complete SAPL species provided a much better resurfacing outcome,

with the newly laid surface exhibiting almost similar SAL pattern observed in normal

intact cartilage surface (Figure 6.6).

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Figure 6.2 3D topographical image of normal healthy articular cartilage after image

processing, showing the nanostructural arrangement of the surface amorphous layer

with several peaks and troughs (length (X) and breadth (Y) of the scanned area, and

average peak height of SAL (Z)).

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Figure 6.3 3D topographical image of the surface of delipidized articular cartilage

after image processing, showing the loss of the membranous overlay (surface

amorphous layer) of the articular surface (length (X) and breadth (Y) of the scanned

area, and average peak height of SAL (Z)).

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Figure 6.4 3-D topographical image of the surface of relipidized articular cartilage in

POPC after image processing, showing partially restored lamella layer of lipids

slightly similar to normal articular surface (length (X) and breadth (Y) of the scanned

area, and average peak height of SAL (Z)).

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DPPC after image processing, showing almost featureless structure of the articular

surface when compared with normal intact articular surface (length (X) and breadth

(Y) of the scanned area, and average peak height of SAL (Z)).

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complete SAPL mixture after image processing (length (X) and breadth (Y) of the

scanned area, and average peak height of SAL (Z)).

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The measured average apparent diffusion coefficients for cartilage samples with

different surface conditions are presented in Table 6.1. The average apparent

diffusion coefficient computed for the delipidized specimens was significantly higher

than those of their normal counterparts (p = 0.003, Table 6.1). Upon incubation of

the artificially delipidized tissue samples in three separate solutions containing

synthetic POPC, DPPC and complete SAPL mixture, the average apparent diffusion

coefficients were significantly lessoned with p = 0.0354 (for POPC), p = 0.0136 (for

DPPC), and p = 0.0017 (for complete SAPL mixture) respectively. When compared

with the normal untreated specimens, the measured apparent diffusion coefficients

were greater for the cartilage surfaces exposed to POPC, DPPC, and full SAPL mix

with statistical difference of p = 0.0065, p = 0.004, and p = 0084 respectively. The

average apparent diffusion coefficient of the POPC-treated samples was slightly

lower than that of the DPPC, there was no statistical difference between them (p =

0.7809). However, the values obtained for the samples exposed to the complete

SAPL mix were far lower than those of POPC and DPPC, with a value approaching

those of normal intact specimens (Table 6.1).

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Table 6.1 Average apparent diffusion coefficients for cartilage samples with normal

intact, delipidized and relipidized surfaces.

Sample condition

Apparent diffusion coefficient (µm2s

-1)

(mean ± SD)

Normal

463.1 ± 37.5

Delipidized

704.1 ± 52.2

Relipidized in POPC

600.5 ± 19.1

Relipidized in DPPC

609.0 ± 17.6

Relipidized in full SAPL mix

541.2 ± 26.3

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Figure 6.7 Typical 2D multi-spin multi-echo (MSME) images of cartilage specimens

immersed in D2O – PBS solution acquired at different times.

It is important to note that image A was acquired at the beginning of the diffusion

time course, while B, C and D were captured several minutes later. The change in

intensity of the images was used to track the D2O ingress into the cartilage matrix.

The bright colour corresponds to high H2O content and the dark colour corresponds

to low H2O content.

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Figure 6.8 Representative depth-wise concentration profiles for the MSME images

shown in Figure 6.8 at different time steps obtained using the analysis with a

purpose-built computational scheme developed in MATLAB®.

In the Figure 6.8, the curves represent the distribution of H2O across the cartilage

from the subchondral bone (x = 0) to the articular surface (x = xmax) at different

times. Curves A, B, C, and D correspond to the images A, B, C, and D presented in

Figure 6.7 respectively.

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6.5 CONCLUSION

It has been demonstrated, using individual components and complete mixture of

surface-active phospholipids found in the normal surface amorphous layer overlaying

the articular cartilage, that it is possible to modify the permeability of the delipidized

cartilage surface, reconstituting it to a form approaching the normal intact surface

behaviour. This result adds further support to the hypothetical position that it is

possible to artificially “resurface” a degenerating articular cartilage, especially in the

early stage condition such as osteoarthritis, by introducing phospholipid mixtures

into the joint space. It is acknowledged that this work is only at the proof-of-concept

stage and still requires significant further research, in which more case scenarios will

be considered with various combinations of phospholipid constituents such as SLPC,

PLPC, and DLPC, and other components of the joint fluid, such as lubricin and

hyaluronic acid.

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Chapter 7: ASSESSMENT OF THE

MECHANICAL INTEGRITY/

PHYSIOLOGICAL FUNCTION OF

RESURFACED ARTICULAR

CARTILAGE

7.1 INTRODUCTION

This study extends the existing knowledge on the surface characteristics of intact,

degraded and resurfaced cartilage presented in chapters four and five. It provides

further insight into the effect of surface condition, concomitant diffusion/percolation

and exudation of interstitial fluid, on load processing within the articular cartilage

matrix. This knowledge would contribute further to our understanding of:

(i) the effect lipid loss during early stage cartilage degeneration, on its

biomechanics in relation to its elastic deformation and osmotic recovery, with

the potential to contribute to a further understanding of cartilage behaviour

during function at low rates of loading,

(ii) the mechanical functionality of the cartilage matrix following relipidization,

especially with respect to answering the question of whether or not

relipidization can form the surface “seal” to restore cartilage to a condition

similar to that of the normal intact sample.

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The fluid component of cartilage plays a significant role in its stress-sharing

behaviour, and hence, the loading processing action of the joint (Oloyede and

Broom, 1993; Oloyede and Broom, 1991). In Chapter six, magnetic resonance

measurements and computational analysis were used to show that the alteration of

the surface amorphous layer of cartilage through delipidization and relipidization,

changes its semipermeability or diffusion characteristics. It is therefore hypothesized

that changes in the surface amorphous layer of articular cartilage will influence the

manner with which the solid matrix components process applied external load, stores

and releases energy in a load-unloading process. This would be similar to the type of

action accompanying short term low velocity or static activity of the joint.

The load bearing/processing characteristics and related energy parameters were

measured from the load-unloading behaviour of the tissue obtained from the

mechanical compression tests using the Instron material testing machine. Further

analyses were then conducted on the energy components associated with the stress-

strain curve of articular cartilage to determine the relationships between the energy

developed as a result of matrix deformation and that released to facilitate recovery.

The details of these experiments have already been described in Chapter four,

Section 4.3.9.

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7.2.1 ARTICULR CARTILAGE SAMPLES

The articular cartilage samples used in this study were obtained from the patellae of

3-4 year old bovine animals (n = 8) harvested from the local abattoir and stored at -

20oC until required for testing. The samples were thawed out in continuous running

water at room temperature and kept in saline solution (0.15M sodium chloride) prior

to testing. A 14 mm diameter stainless steel punch was used to cut osteochondral

plugs (n = 32), containing full thickness articular cartilage-bone laminate. Each

sample was mounted in the consolidometer and then transferred to the Instron

machine using the procedure presented in Section 4.3.9 of Chapter four. The

cartilage plug was subjected to the loading and unloading protocol described in the

approach and methodology chapter (Chapter three). After the loading process, the

specimen was removed from the consolidometer and submerged in 0.15 M saline

solution to recover. The sample was left in the saline solution for 4 hrs to recover

completely until ready for delipidization; previous studies have established that

optimum time for a loaded cartilage to recover is 2-3 hrs (Oloyede, et al., 2004a,

2004b).

The normal intact samples (n = 32) from Section 7.2.1 above were subjected to

delipidization after their recovery. A full description of the delipidization protocol is

presented in Section 4.3.2 of Chapter four. Following delipidization, specimens were

rinsed repeatedly with 0.15 M saline solution, and later placed in saline solution for

30 mins for rehydration. After rehydration, each sample was placed in the

202

consolidometer filled with 0.15 M saline solution, and then mounted on the Instron

machine as previously described for the normal cartilage samples. While on the

material testing device, the sample was subjected to the same loading and unloading

protocol described in Section 4.3.9 of Chapter four. After the loading, the sample

was removed from the consolidometer and placed in saline solution for 4 hrs to fully

recover. After the recovery, the samples were divided into three groups comprising

of eight samples (n = 8) per group, while ensuring that each sample set was taken

from the same joint. The samples were further subjected to relipidization.

The three case scenarios described in Sections 4.3.3.1, 4.3.3.2, and 4.3.3.3 of Chapter

four were used for relipidizing the delipidized cartilage specimens (the three sample

groups). Prior to the mechanical compression tests, each sample from the

incubation/relipidization process, was removed from its test tube container, and

gently rinsed in 0.15 M saline solution. The rinsed samples were sandwiched into the

consolidometer filled with 0.15 M saline solution, and then mounted on the material

testing device. Loading protocol and input parameters used for the normal intact and

delipidized specimens were also used for the relipidized and control test samples.

7.2.2 DERIVATION OF THE ENERGY COMPONENTS

The load-displacement curves obtained for the normal intact, delipidized and

relipidized specimens were analysed using energy methods described in Section

4.3.9 of Chapter four. The parameters derived from theses curves are strain energy,

elastic energy lost in matrix recovery or hysteresis energy, complementary energy,

residual energy, and energy ratio. These energies were calculated as the areas under

203

the regions defined in the load-displacement curve shown in Figure 7.1. Their

average values were statistically analysed and used to assess the mechanical integrity

of resurfaced cartilage relative to their corresponding normal intact counterparts.

Figure 7.1 Energy diagram derived from a typical load-displacement curve obtained

for normal intact cartilage sample subjected to loading and unloading test on the

Instron machine, where SE and RE represent strain energy and residual energy

respectively.

Figure 7.2 presents the protocol for the experiments conducted and the parameters

derived for accompanying articular cartilage’s functional response when carrying

normal, delipdized and relipidized surfaces.

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Figure 7.2 Schematic flow chart of the loading sequence followed for the normal

intact, delipidized, and relipidized articular cartilage samples.

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7.2.3 STATISTICAL ANALYSES

The data presented in this study represent the mean and standard deviation (SD) of

the strain energy, hysteresis energy, complementary energy, residual energy, and

energy ratio, for normal, delipidized, and relipidized cartilage samples. A two-tailed,

unpaired Student’s t-test was used to determine if there is any significant difference

between the normal and delipidized samples and, also, the statistical significance of

the results of relipidization of artificially delipidized specimens in DPPC, POPC and

full SAPL mixture relative to the normal intact specimens. The statistical differences

across the different groups of samples were evaluated using analysis of variance

(ANOVA) with repeated measures and multiple comparisons according to the

Bonferroni post hoc tests. All the analyses were conducted with a significance level

of p < 0.05 or a 95% confidence interval.

Furthermore, a statistical appraisal was conducted using power analysis, with

G*Power statistical power analysis software (Faul et al., 2007), to determine whether

or not the relatively small sample size (n = 24, 8 samples per group) used in the

analysis described above (Student’s t-test and ANOVA) is sufficient (Zhu, et al.,

1993).

206

Figure 7.3 Screenshot of the G*Power statistical power analysis software used to

determine the influence of the relatively small sample size used in this study.

207

The screenshot showing the power analysis result (Figure 7.3) reveals that effective

size, f = 0.954 for the samples tested. When compared with the effect size convention

proposed by Cohen (1988), (small f = 0.10, medium f = 0.25, large f = 0.40), the

measured effective size (f) in this study is greater than the value of the large f

proposed by Cohen (1988). The results of the power analysis established that the

sample size used for the experiments passed the criteria proposed by Cohen (1988),

and the specimen size is sufficient for the Student’s t-test and ANOVA.


The average and standard deviation of the strain energy, hysteresis or released

energy, complementary energy, residual energy, and energy ratio for cartilage

samples with normal intact, delipidized, and relipidized (in POPC, DPPC and

complete SAPL mix) surfaces are presented in Table 7.1. The average strain energy

calculated for the delipidized specimens was significantly lower than those of their

normal counterparts (p < 0.001, Table 7.1). Upon incubation of the lipid depleted

samples in separate solutions containing synthetic POPC, DPPC and complete joint

SAPL mixture, the average strain energies of POPC and DPPC treated samples were

significantly lower than those of the normal samples, for POPC, p < 0.001, and

DPPC, p < 0.001, respectively. On the other hand, incubation in full SAPL mix

resulted in a significant increase in the average strain energy (p < 0.001, relative to

the normal unaltered samples.

208

A comparison of the average strain (or internal) energy to the other energy-related

parameters revealed similar trend for the average released energy, residual energy,

and complementary energy for the delipidized samples exposed to POPC, DPPC, and

full SAPL mix when compared with their corresponding normal intact counterparts.

Interestingly, almost similar patterns were observed for the POPC and DPPC treated

samples when their average strain, hysteresis, residual, and complementary energies

were compared with each other. The POPC treated samples exhibited higher strain

energy, released energy and residual energy, while DPPC treated samples presented

slightly higher average complementary energy (Figures 7.4 - 7.7). Overall, the

calculated values of the energy parameters for these two sample groups were also

found to be lower than the normal and full SAPL treated samples (Figures 7.4 – 7.8).

As expected, the samples incubated in the complete SAPL mixture showed more

promising resurfacing outcomes, with their energy values approaching those of the

normal samples.

Furthermore, the energy ratio calculated as the ratio of the complementary energy to

the strain energy showed a different pattern compared to the other energy parameters.

The normal intact samples have an average energy ratio of (1.679 ± 0.255). The

samples exposed to POPC solution has the lowest average energy ratio (1.596 ±

0.230), while those incubated in synthetic DPPC solution has the highest value

(3.134 ± 0.503) (Figure 7.8). Delipidization resulted to an increase in the energy

ratio, with a measured averaged value of 1.776 ± 0.423, when compared to their

corresponding normal unaltered samples. However, there was no statistical

difference between these two sample groups. The delipidized specimens exposed to

aqueous solutions containing POPC and full SAPL mixture, with average energy

209

ratios of 1.596 ± 0.230 and 1.634 ± 0.170 respectively, are closest to the normal

intact cartilage samples (Figure 7.8).

Table 7.1 Variation of total strain energy, elastic energy released, and residual energy

of cartilage matrices with normal intact, delipidized, and relipidized surfaces.

Sample

condition

Average

strain

energy (J x

10-3

)

(Mean ±

SD)

Average

elastic

energy

released (J x

10-3

)

(Mean ±

SD)

Average

residual

energy

(J x 10-3

)

(Mean ±

SD)

Average

complementary

energy

(J x 10-3

)

(Mean ± SD)

Energy

ratio (CE/SE)

Normal 1.878

±

0.543

0.496

±

0.187

1.365

±

0.381

3.496

±

1.360

1.679

±

0.255

Delipidized 1.057

±

0.063

0.315

±

0.172

0.599

±

0.138

1.916

±

1.051

1.776

±

0.423

Relipidized

in POPC

0.969

±

0.164

0.224

±

0.050

0.744

±

0.132

1.559

±

0.419

1.596

±

0.230

Relipidized

in DPPC

0.698

±

0.303

0.152

±

0.051

0.546

±

0.252

1.829

±

0.308

3.134

±

0.503

Relipidized

in all SAPL

mixture

1.419

±

0.345

0.331

±

0.088

1.088

±

0.258

2.025

±

0.656

1.634

±

0.170

210

For the purpose clarity and better interpretation of the results presented in Table 7.1,

pictorial representations of the measured strain energy, complementary energy,

released energy, residual energy, and energy ratio are shown in Figures 7.4 - 7.8

respectively. The bar charts were plotted using the mean values of these parameters

with their 95% confidence interval (CI).

Figure 7.4 Strain energy plotted for articular cartilage specimens with normal intact,

delipidized, and relipidized (POPC, DPPC and complete SAPL mix) surfaces.

211

Figure 7.5 Complementary energy plotted for articular cartilage specimens with

normal intact, delipidized, and relipidized (POPC, DPPC and complete SAPL mix)

surfaces.

212

Figure 7.6 Released energy plotted for articular cartilage specimens with normal

intact, delipidized, and relipidized (POPC, DPPC and complete SAPL mix) surfaces.

213

Figure 7.7 Residual energy plotted for articular cartilage specimens with normal

intact, delipidized, and relipidized (POPC, DPPC and complete SAPL mix) surfaces.

214

Figure 7.8 Energy ratio plotted for articular cartilage specimens with normal intact,

delipidized, POPC, DPPC and complete SAPL mix surfaces.

215

It should also be noted that because the properties of articular cartilage vary with

position across the same joint and from joint to joint, the average values of the

energies presented in Table 7.1 and Figures 7.4 - 7.8 may not truly reflect the

differences in the load processing behaviour of the different samples tested.

Therefore, scatter plots (Figures 7.9 – 7.12) were used to show the relationship

between the different energy parameters extracted from the load-displacement data

obtained for the samples. The graphs reveal noticeable differences between the

parameters for the normal and delipidized samples. The energy parameters for the

relipidized/resurfaced samples suggest changes in their load-bearing (functional)

properties, with the values for the resurfaced samples approaching those of normal

unaltered specimens (Figures 7.9 – 7.12). The scatter plot of the complementary

energy versus strain energy shows a slight overlap between the delipidized and

resurfaced cartilage samples (Figure 7.9).

Figure 7.9 Complementary energy versus strain energy for articular cartilage samples

with normal intact, delipidized, relipidized surfaces.

216

Figure 7.10 Released energy versus strain energy for articular cartilage samples with

normal intact, delipidized, relipidized surfaces.

.

Figure 7.11 Residual energy versus strain energy for articular cartilage samples with

normal intact, delipidized, relipidized surfaces.

217

A linear relationship was observed between the released and strain energy (Figure

7.10), residual and elastic strain energy (Figure 7.11) for all the sample groups.

However, the complementary energy versus strain energy (Figure 7.9), and the

residual versus released energy (Figure 7.12) produced an almost non-linear

relationship. The elastic property seems to be maintained across the sample groups

tested, with the normal and relipidized samples having higher strain energies, while

the delipidized samples have lower values. Thus, when intact, the tissue is able to

store more energy when loaded. Removal of the SAL to delipidization consequently

reduces the elastic strain energy, thereby making the tissue stiffer.

Figure 7.12 Residual energy versus released energy for articular cartilage samples

with normal intact, delipidized, relipidized surfaces.

218

7.4 CONCLUSION

This study has demonstrated that the removal of surface lipids of articular cartilage

changes its load processing characteristics, and that incubation in synthetic

phospholipid solutions potentially restores the tissues’ mechanical behaviour

(function), pushing it towards those of normal intact samples. Overall, the complete

SAPL mixture provided better resurfacing outcomes than the individual components

(DPPC and POPC); thereby further supporting the results from our previous

experiments in Chapter five and six.

219

Chapter 8: DISCUSSION AND CONCLUSIONS

This thesis has developed scientific protocols, as well as demonstrated their

potentials, for resurfacing the articular cartilage following loss of its lipid-rich

surface amorphous layer based on a sound understanding of the chemical

composition of the phospholipids content of this vital surface layer of the tissue. The

newly created phospholipid layers were characterized nanoscopically,

microscopically, and macroscopically to establish the structural and functional

efficacy of the newly engineered layer. Numerical analysis of the magnetic

resonance imaging data provided information on the influence of the newly laid

surface on the semipermeability characteristics of the matrix, while energy analysis

from loading-unloading tests provided important assessment parameters on the load-

bearing/load-processing effectiveness as a result of the relipidization procedure.

This thesis has rigorously investigated the potential effect of synthetic lipid

resurfacing/relipidization of lipid-depleted articular cartilage on its structural and

functional integrity. An experimental procedure was developed to resurface

artificially degraded cartilage using single components and mixtures of

phospholipids found in the mammalian joints. In the course of this research, several

case studies were considered, including the conduction of a proof-of-concept study

involving individual phospholipids components (a saturated specie, DPPC; and

unsaturated specie, POPC) of the total lipid constituents found in the human joints. A

significant contribution of this study is the insight into the potential of reconditioning

degraded cartilage surfaces with synthetic phospholipids. It also established a step-

220

by-step approach for resurfacing cartilage with synthetic phospholipids, and the roles

of the individual lipid components in cartilage physiological function.

The relipidization experiment conducted with the individual SAPL species developed

a platform, which enhanced the understanding of the contributions of each of the

lipid components to cartilage surface function. It can be argued that if the single

phospholipid species were incapable of adhering on their own to delipidized cartilage

surfaces, there may not be a reason to continue the investigation and widen the scope

to include complete SAPL mixture, and extend the study to include the effect of

factors such as the influence of lubricin and other agents such as hyaluronic acid.

Therefore, it is strongly believed that this proof-of-concept study is fundamental to

resolving the question of whether or not the hypothesized repair role of

phospholipids in joints has any real basis. In addition, the inability of the single

phospholipid species to effectively adhere to the surface of delipidized cartilage and

it remodel its functional behaviour was the motivation for extending the investigation

to include complete joint SAPL mixture used in this research.

Using atomic force microscopy (AFM) to image the soft surface of articular cartilage

in a phosphate buffered saline (PBS), as well as confocal microscopy (COFM) and

Raman spectroscopy, the surface of normal, delipidized and relipidized cartilage

specimens have been characterized. The AFM imaging demonstrated that the

removal of the surface amorphous layer (SAL), which overlays the intact cartilage

surface, leads to the exposure of a disorganized structure in the same manner as

previously observed by Crockett et al. (2005) (Figures 5.3 and 5.4, Chapter five).

These results are further supported by the outcomes of the Raman measurements,

221

which established that delipidization led to the modification of the chemical

characteristics of the normal intact the articular surface.

Confocal microscopy of the incubated delipidized specimens demonstrates that

synthetic lipids can indeed settle and form layers on the surface of degraded cartilage

(Figures 5.5a and 5.6a). In fact, the incubation in unsaturated POPC, saturated DPPC

and complete SAPL mixture led to dissimilar structural overlays of average effective

thicknesses 1794 nm, 1541 nm and 1986 nm respectively, compared to the normal

sample which is 856 nm (Table 5.1, Chapter five). Effective thickness was used

because it is only possible to obtain a height that is relative to the lowest peak of the

surface amorphous layer.

When juxtaposed with the earlier work of Vecchio et al. (1999), these results could

be interpreted as suggesting the potential capacity of synthetic lipids to “resurface”

the tissue. While these initial experiments indicate the potential of resurfacing

cartilage with synthetic lipids, further characterization methods are required both in

terms of understanding the right lipid mix, optimum time of incubation and quantity

of lipids in solution which is required for the creation of a more viable surface, the

macro-functional/biomechanical compatibility of these new surface layers and the

diffusion or semipermeability assessment of the newly laid surface layer. The

semipermeability and macromechanical tests were conducted and presented in

Chapters six and seven respectively.

It is worth noting that there are inherent differences in articular cartilage samples

used in the study, and there is a possibility of errors due to the sensitivity of the

222

nano-characterization instrument (AFM). However, while there is an appreciable

overlap between the effective height of the surface amorphous layer for the normal

and delipidized samples, the average elastic strain energy demonstrates that the

delipidization resulted in a significant difference (p < 0.0001) in the surface

properties of the samples (Table 5.1, columns 2 and 3, rows 2 and 3). Also, whilst the

deposited lipid layer using POPC, DPPC and full SAPL mixture, are significantly

higher than the surface amorphous layer in the normal specimens, it can be seen that

the elastic surface strain energy values are much lower than those of the normal

specimens (Table 5.1, Columns 2 and 3, Rows 4 - 6). The values of the average

elastic strain energies of the surfaces suggest that, despite the encouraging new lipid

layer formation, the AFM results reveal that the newly laid lipid-filled surface is still

not fully capable of viable physiological load-spreading. This is unsurprising for the

surfaces laid with POPC and DPPC, which are only single components of the entire

lipid species found in the surface amorphous layer overlaying the articular cartilage.

However, results for the full lipid mix provides more encouraging outcomes for the

potential of using mixtures of synthetic-lipid based treatment for joint diseases.

The reconstituted 3-D images have served to resolve the current prevalent opinion on

the nature of the structure exposed by the delipidization process, namely whether or

not this is the dendritic/fibrous structure of collagen of the underlying matrix that is

revealed when the lipids are removed. The comparative analysis between the normal

and delipidized samples seems to suggest that it is not collagen fibres that are seen in

these images, and that the surface of normal cartilage is not featureless as previously

proposed by Kumar et al. (2001), Jurvelin et al. (1996), and Grant et al. (2006), but

is instead characterized by a series of ridges and valleys (Figure 5.3c) that appear like

223

a fibrous meshwork in 2-D (Figure 5.3b). This result is also supported by the work of

Orford and Gardner, (1985) and Crockett et al. (2005).

From the results obtained for cartilage in the three surface conditions, namely normal

intact, delipidized and relipidized (in DPPC, POPC and complete SAPL mix); it was

deduced that the elastic strain energy of the surface amorphous layer due to its

resistance to nano-indentation for normal intact cartilage was about five times larger

than that of delipidized and relipidized samples in POPC and DPPC, and three times

larger than that for samples relipidized in complete SAPL mixture. Since, the

average elastic strain energy of samples treated with full SAPL mixtures is

significantly higher than those exposed to POPC and DPPC, this demonstrates that

relipidization in complete SAPL mix provides a better resurfacing outcome than in

solutions containing POPC and DPPC alone, and that these single species were

insufficient to completely remodel and functionalize the compromised articular

surface layer. Also, the force-displacements results reveal that the relipidization of a

degraded articular surface in either DPPC or POPC did not restore its compliance to

nano-mechanical load-spreading when compared to normal intact surfaces (Figure

5.12), thereby, justifying the motive for extending the study to include full SAPL

mixture due to inability of the single SAPL species to deliver the desired outcome.

At this point, it is important to mention that there is a close relationship between

articular cartilage surface structure and its function (load-spreading and lubrication)

(Oloyede and Broom, 1996) (Broom and Oloyede, 1998). This is evident in the AFM

images and force curves obtained for the tissue in normal, delipidized and relipidized

conditions (Figures 8.1 and 5.12). As mentioned earlier, both delipidization and

224

relipidization have considerable effects on the structural configuration of the surface

of articular cartilage, due to differences in topography. An analysis of the force

curves reveal that the tissue’s surface resistance to penetration during nano-

compression is seemingly dependent on the type of lipids used. It is therefore

proposed that the mechanical efficiency of a regenerated cartilage surface layer

would depend significantly on the type, and combination of lipids employed. This

requires further testing where the methodology would include the study of the type,

composition and quantity of lipids that would be required to engineer the creation of

a new functional cartilage surface layer.

225

Figure 8.1 Reconstructed 3-D AFM images of articular cartilage with normal intact,

delipidized, POPC-treated, DPPC-treated, and complete SAPL mix treated surfaces.

226

A closer angular view of the reconstructed 3-D AFM images of articular cartilage

samples with different surface conditions (Figure 8.1) reveals that the normal intact

and complete SAPL-treated surfaces have similar surface structural network, with

several crosslinking between the phospholipid molecules (Figure 8.2). However, this

structure is not present in the delipidized articular surface, possibly due to the

disruption of the surface network following lipid removal. The POPC- and DPPC-

treated samples exhibited a wave-like structure, without crosslinking between the

lipid molecules (Figure 8.3). It is argued that this cross-linked phospholipid network

structure exhibited in cartilage with normal and complete joint SAPL-treated surface

is due to the presence and interactions of the various SAPL species on their surfaces

(saturated and unsaturated species).

Figure 8.2 Cross-linked phospholipid network structure present in cartilage

specimens with normal intact and complete SAPL mixture-treated surfaces.

227

Figure 8.3 Wave-like phospholipid structure present in cartilage samples with DPPC

and POPC treated surfaces.

Furthermore, since semipermeability is a major functional characteristic of the

articular cartilage surface and it is one of the significant factors determining its

physiological efficacy, it is necessary that any research on resurfacing or layering the

artificial surface of degraded cartilage with synthetic phospholipids, which is the

objective of this research, includes a test of the capacity of such surfaces to provide

semipermeability to both fluid and microconstituents. The diffusion experiments

described in chapter five was conducted using DPPC, POPC, and complete SAPL

mix to resurface delipidized cartilage samples provides contrasting surface

properties, i.e. saturated and unsaturated artificial phospholipid layers against which

semipermeability can be tested. The atomic force microscope (AFM) observations

(Figures 6.2 – 6.6, Chapter six), which are in complete agreement with our earlier

results (Yusuf, et al., 2012), demonstrate that the results presented in this thesis can

be considered with confidence, to represent the diffusion barriers or permeability

properties of the surfaces with different surface lipid configurations/structure.

228

It is worth noting that the diffusion of solutes through articular cartilage is influenced

by the local composition and structure of the matrix, due to its inherent anisotropic

nature. Previous studies have established that diffusion coefficient varies with depth

from the articular surface to the subchondral bone (Leddy and Guilak, 2003;

Maroudas et al., 1968). The results presented in Table 6.1 (Chapter six) is an

“apparent” diffusion coefficient, which is a lumped/composite parameter accounting

for the distribution of depth-dependent intra and interlayer diffusion coefficient as

well as the surface interlayer permeability (Glenister, 1976). Consequently, it can be

argued that the measured diffusion coefficients are averaged values of a highly

variable parameter which does not refer to any particular physical location within the

sample but depends on the weighted-average of the local diffusion coefficient

values. This is analogous to the methodology used in (Burstein, et al., 1993;

Kokkonen, et al., 2011).

Additionally, the MRI results can be considered to represent the combined effect of

the surface permeability and subsurface diffusion characteristics of the various tissue

groups. Given that the SAL lipid membrane is quite thin, it can be argued that the

results obtained for the apparent diffusion coefficients are good indicators of the

permeability/semipermeability of the lipid membranes of the SAPLs. This suggests

that the measured average apparent diffusion coefficients (Table 6.1) that are

presented in the thesis should be treated with caution. Nevertheless, the data provide

a good comparison and estimate of the permeability properties of the normal,

delipidized, and relipidized cartilage samples studied, since it can be deduced from

Fick’s first law of diffusion that the rate of diffusion (diffusion flux) across a

membrane, such as the articular surface, is directly proportional to the permeability

229

of the membrane (Fick, 1855). More so, diffusion flux is proportional to diffusion

coefficient, providing supporting evidence for the assumption made in the

comparisons of the permeability of the different surfaces studied.

The results from the numerical analyses of the MRI data reveal a significant

difference between the diffusion characteristics of normal and delipidized samples,

thus establishing that the delipidization process resulted in a significant change in the

semipermeability property of the tissue (p = 0.003, Table 6.1, Colum 2, Rows 2 and

3) following the loss of the lipid layer. The measured average apparent diffusion

coefficient (ADC) for the delipidized specimens was significantly higher (about

52%) than that of the corresponding normal counterparts. Upon relipidization in

POPC and DPPC, there was a considerable decrease in the average apparent

diffusion coefficient of the delipidized specimens, about 14.7% and 13.4%

respectively. When compared with the normal intact sample surfaces, the POPC and

DPPC laid surfaces were significantly higher, about 29.7% and 31.5%, respectively.

The sample exposed to complete SAPL exhibited even lower ADC when compared

with delipidized, DPPC and POPC treated specimens.

Given that the diffusion coefficient is directly proportional to the permeability (Fick,

1855); these results provide evidence that the synthetic phospholipid layer laid

artificially to surface of delipidized cartilage is capable of modifying a highly

permeable delipidized cartilage surface to a more semipermeable state (Figure 8.4).

Although, the apparent diffusion coefficients of the cartilages samples exposed to

DPPC and POPC were higher than those of the normal intact samples, samples

incubated in complete SAPL mixture provided a relatively high semipermeable

230

surface that is close to natural unaltered articular surface, it is inferred that

relipidization with synthetic lipids could offers a potential remedy for managing

early stage joint degeneration. This will require more rigorous studies as the result

presented in this thesis has only established a scientific approach that can be

developed to realize this treatment option.

Figure 8.4 A schematic scale showing the change in permeability of articular

cartilage with different surface conditions. Relipdization in synthetic DPPC, POPC

and full SAPL mix resulted in the transformation of the delipidized cartilage sample

surfaces from a highly undesirable permeable condition to a more effective surface

with lower permeabililty and better semipermeabilty characteristics.

After successful relipidization, the newly laid lipid layers were characterized

structurally using nano- and microscopic methods with the AFM and confocal

231

microscope. The semipermeability properties were also evaluated using a

combination of magnetic resonance imaging and image processing techniques.

Additionally, the “resurfaced” cartilage samples were subjected to macro-mechanical

compression tests to assess their loading processing capacity/mechanical integrity

relative to their normal intact counterparts. Load-displacement data obtained from

this test were used to derive several energy parameters which were then related to the

load processing capability of the samples’ matrices. Detailed descriptions of this

procedure are fully documented in Section 7.2.2 of Chapter seven. The compression

test also provided relevant information on how the surface condition of cartilage

influences its elastic deformation and osmotic recovery.

Although the exudation and influx of fluid out and into the articular cartilage matrix

during loading and unloading has been shown to be strongly correlated to its fixed

charge density (FCD), a function of its proteoglycan content (Maroudas et al., 1969;

Maroudas and Venn, 1977; Muir, 1978), a number of studies (Oloyede, et al., 2008;

Oloyede, et al., 2004a, 2004b) have also demonstrated the effect of the surface of

cartilage, particularly its lipid layer or surface amorphous layer, on this characteristic

of its matrix. This thesis supports and extends the results of existing studies in the

literature by showing that a significant improvement in the functional viability of the

tissue can be achieved by relipidizing the surface of lipid-depleted cartilage samples

using synthetic phospholipids. This functional property of the tissue was assessed via

mechanical load-unloading tests. It has also been demonstrated that the load

processing capacity of cartilage is dependent on the energy parameters derived from

the load-displacement curves.

The mechanical tests results reveal that articular cartilage regardless of its surface

conditions (either normal, delipidized, or relipidized) is capable of storing energy

232

(elastic strain energy) when it is loaded and its matrix deformed. When unloaded, a

portion of this stored energy is released (hysteresis energy) as the deformed matrix

recovers. This magnitude of these energies can be used to evaluate the functional

capacity of the tissue. This thesis has used energy parameters derived from the

mechanical compression tests results to establish the relationship between cartilage

surface condition and its capacity to process load efficiently. The 1-dimensional

consolidation set-up adapted in the mechanical testing protocol was effective in

confining the fluid flow only through the surfaces of the samples, thus accentuating

the influence of cartilage surface on its overall functional viability.

The energy transformation during the loading and unloading of cartilage is analogous

to the phenomenon observed in the stretching and release of an elastic spring or

catapult. During the stretching phase, an amount of potential elastic energy is stored

in the spring by the external pulling (or tensile) force. A portion of this stored

potential elastic energy is immediately transformed into kinetic energy as the spring

is released, with the remaining converted to other forms of energy (for example, heat

and sound). The amount of energy stored and released in the spring system is related

to its stiffness (Hooke’s law). Applying this principle to the load-unloading

(compression test) results presented earlier in Table 7.1 of Chapter seven, the

capacity of normal cartilage samples (with intact SAL) to store and release more

energy than the delipidized samples (with altered surfaces) may be attributed to the

increase in matrix stiffness as a result of the effect of delipidization as suggested by

Oloyede et al (2004b) (Figures 7.4 and 7.6, Chapter seven). Relipidization in single

SAPL components (DPPC and POPC) did not have any significant influence on the

stored energy, surprisingly; their released energies were lower than the delipidized

233

samples. However, treatment in complete joint SAPL mixture resulted in an increase

in the elastic strain energy approaching those of normal intact samples, therefore

increasing the potential for applying SAPL mixtures as remedy for joint

degeneration. This might be due to the fact that the complete joint lipid mixture is

able to form a seal that restores the functional characteristics of the degraded

cartilage surface.

It is also important to note that the average complementary energy for all the sample

groups were higher than the corresponding average strain energies. As earlier started

in Section 7.2.2 of Chapter seven, the ratio of the complementary energy to the strain

energy is the energy ratio. For linear elastic materials, the load-displacement plot is

linear, with the complementary energy having the same value as the strain energy,

therefore, the energy ratio for such materials is equal to one (Boresi, Schmidt et al.

1993) (Figure 8.5). However, articular cartilage undergoes a large non-linear

deformation under compressive loading, resulting in a non-linear J-shaped load-

displacement curve (Nguyen and Oloyede 2001; Oloyede, Gudimetla et al. 2004).

The mechanical tests results reveal that the average complementary energies of all

the sample groups tested are greater than the strain energies; their energy ratios are

also greater than one. It is interesting to observe that the DPPC treated samples have

almost twice the energy ratio when compared with other sample groups, while POPC

and full SAPL mixture treated samples have almost similar energy ratios as the

normal intact samples (Figure 7.8). The large difference in the energy ratio of the

DPPC treated samples may be a likely explanation for joint inflammation (gout)

outcome of the clinical trial of Vecchio et al. (1999); however, further research is

required to validate this claim.

234

Figure 8.5 Load-displacement diagram for linear elastic material subjected to

compressive loading test. The complementary energy and the elastic strain energy

are equal; therefore the energy ratio for linear elastic materials is equal to one.

At this point in time, it should be noted that while the results from this thesis

demonstrate that the surface-active phospholipids are capable of adhering to the

delipidized articular surface, albeit with the complete joint SAPL showing more

235

promise than single unsaturated POPC and saturated DPPC, they do not yet provide a

complete picture of the resurfacing process, which is aimed at creating fully

functional surface that can mimic a natural intact articular surface. This is partly

because, in this exploratory study, only arbitrary concentrations of the synthetic

phospholipids have been used (1g per 100 ml of deionized water), which do not

strictly conform to the concentrations found in the joint. However, it is believed that

the concentrations applied in this research are adequate for testing our hypothesis,

while noting that the concentrations of lipids used in the experiments are not exactly

the same as the amounts found in the joint, this is explained in detail by Chen et al.

(2007) and Sarma et al. (2001).

Despite the limitations highlighted, the results of this study demonstrate that cartilage

can be potentially resurfaced, especially with synthetic lipid mixtures; while also

furthering creating a scientifically based knowledge in the advancement of the non-

surgical treatment of joint diseases. It is recommended in this thesis that further

research will be required before it can be fully concluded that the resurfacing of

articular cartilage with full functionality relative to the physiological cartilage

surface is achievable. This study would arguably include:

I. Addition of lubricin and hyaluronic acid (HA) to the SAPL mixture.

II. Assessment of the lubrication characteristics of the newly laid SAPL

membranes.

III. Evaluation of the time-dependence of the incubation process to understand

how long it might take for the lipids to fully attach to the articular surface, for

236

example if applied to the joint surface as an injection. This could include

animal model studies.

IV. The use of all components of the SAPL mixture may not be required for the

creation of an effective surface layer. This needs to be understood for

economic optimization in the event that the lipid treatment is adopted as non-

surgical intervention.

237

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270

APPENDIX A

Measurement of the Apparent Diffusion Coefficient from D2o

Ingress Experiment

271

A1.1 EXPERIMENT SETUP

1. A fully hydrated bone-cartilage plug that has been kept in H2O-PBS solution

was brought out for the MR measurements.

2. The URJ image of the sample oriented at the magic angle (54.70) with respect

to the magnetic field (B0) was obtained

The image should be weakly T2 – weighted (echo time, TE ~10ms or

shorter).

Check that the image intensity within the AC is more or less uniform -

i.e. that the T2’s vary only weakly with depth. This is important for

the interpretation of the subsequent D2O ingress experiment data.

3. Prepare D2O-PBS solution: Same as “normal” PBS but with D2O instead of

H2O

4. Fill the NMR tube with the magic-angle Teflon plug with D2O-PBS.

The bone-cartilage plug in the tube was quickly placed in the spectrometer

(so that it is aligned at the magic angle).

The approximate time the sample was placed in D2O-PBS was noted.

5. Obtain a time course of weakly T2 – weighted images.

The Images should be equidistantly spaced in time and separated by as short a

time as possible (ideally 1.5 – 2min):

272

At the end of step (5), we will have the primary data set describing the ingress

process: NT images (NR x NP) voxels each:

i.e., the raw dataset contains: NR x NP x NT voxels.

A1.2 IMAGE PROCESSING

Convert the raw data set into a data set to be used for least squares fitting (LSF)

1. Based on the image from step (2) on page 271 of the appendix, estimate the

thickness (ho) of the articular cartilage (AC) in the bone-cartilage plug. The

estimate should be conservative, in that it must not exceed the actual

thickness of AC in order to avoid including the saline points into the LSF (but

it is ok to under-estimate the thickness; this would result in some AC voxels

being dropped.

2. Identify the AC region of interest (ROI): Identify a region of the sample

where the articular surface (AS) is approximately parallel to the bone-

cartilage interface, draw a rectangle of thickness ho, this is the LSF ROI:

273

3. Construct the LSF dataset: Assuming that the ROI contains NROI voxels, we

now have (NROI x NT) data points. Each data point is characterized by the

time acquired, distance from bone and intensity

Data Point 1 = (t1, x1, s1)

Data Point 2 = (t2, x2, s2) *

.

.

.

Etc., total = (NROI x NT) points

Here, Tk = one of the ti’s (tk = t1, or t2, or t3, . . . )

0 ≤ xk ≤ ho

Sk ≥ 0

274

4. Optional: If the dataset (*) is too large for an LSF, the data points can be

sorted into a histogram with respect to X and only the average value for every

X bin can be used.

i.e.: Divide the X range into

NB bins:

0 x1 x2 h0 = XB

Place all points with a given t and Xi-1 ≤ X ≤ Xi into bin I and calculate the

average intensity for these points:

Note: Take only the Si’s corresponding to the same ti

This way we end up with a reduced data set of (NT X NB) points

Point 1 = ( ) NT values of t

Point 2 = ( ) NB points per t

.

.

Point NB +1 = ( )

.

. (**)

NB bins

275

Total NB X NT points in the reduced dataset

A1.3 LEAST SQUARES FITTING

The data set (*) or the reduced data set (**) is then subjected to LSF in order to

extract the apparent diffusion coefficient (D).

Assuming a perfectly reflective boundary condition (BC) at the bone cartilage

interface and a perfectly absorbing BC at the AS, the exact solution for the

diffusional egress of H2O from the AC is given by

Where =

a = exact thickness of AC

t = time from the start of diffusional transport

C0 initial H2O concentration within the AC

X = distance from the bone

Several assumptions/approximations are implicit in equation (1):

(A1.) The bone-cartilage interface is impermeable this is modeled by the reflective

boundary condition at x = ø;

276

(A2.) As soon as the H2O molecules diffuse out of AC, they are rapidly carried away

by convention or rapid diffusion in PBS. This is modeled by the absorbing boundary

condition at the AS:

(A3.) The cartilage layer covered by the ROI has a perfectly uniform thickness:

a = constant over ROI

(A4.) The proton spin density and T2 are perfectly uniform within the AC. This is an

approximation achieved experimentally by orienting the AC at the magic angle (MA)

with respect to . This is modeled by the uniform initial condition per H2O

distribution:

(A5.) The imaging plane is exactly perpendicular to the AS (and therefore to the

bone surface). This approximation allows the x for every voxel to be calculated from

the image of a single slice.

Furthermore, to make equation (1) is usable in a practical setting, the following

modifications must be made:

(A6.) The exact thickness of the cartilage is not known; therefore; a must be an

adjustable parameter of the LSF.

(A7.) Because each image is acquired over the time span of 1-2min, the values of the

tk are also poorly defined. For this reason, “t” in equation (1) needs to be replaced

with “t – t0” where t0 is an adjustable parameter of LSF.

(A8.) Because the UR image is acquired in magnitude mode, noise is strictly

positive. Therefore, equation (1) needs to include a “noise” term – also an adjustable

parameter.

Therefore, the actual LSF equation was as follows:

277

Where C1 is noise term

The last approximation used is that truncation is appropriate

(A9.) The terms after m=M in equation (2) make a negligible contribution and

therefore do not affect the outcome of the LSF.

Approximations (A1) to (A10) are all the approximations needed for the

interpretation of the MRI data.

Table A1The LSF five adjustable parameters

Parameter Initial Value Meaning/ purpose

Signal intensity at the

smallest x,

Smallest t.

MR signal intensity priot

to D20

A h0 Thickness of AC

D

Apparent D

Time D20 was put in

contact with AC

To fix the poorly define

“Acquisition time”

ø Noise in MR image

278

Once again, all (NROI x NT) [or NB x NT] data points must be fitted simultaneously.

This enables to fix all 5 adjustable parameters with an acceptable reliability.

The apparent diffusion coefficient, D, extracted from the LSF, is a complicated

function of two factors:

1) The actual diffusion coefficient of H20 within the cartilage

2) The permeability of the articulate surface i.e. the probability of H20

molecules escaping once they reached x=a

It also contains the last implicit assumption (or rather, approximation)

(A10.) The diffusivity of H20 within AC is perfectly uniform. This is known

experimentally not to be strictly the case, so the “D” has meaning only within this

approximation.

279

APPENDIX B

Computational Scheme/MATLAB® Code and GUI for Determining

Apparent Diffusion Coefficient from Magnetic Resonance Imaging

Data

280

% Authors - Tim Gurnett and Andrew Knuckey, QUT High Performance

Computing and

% Research Support

% Other authors are acknowledged within their respective code.

% DO NOT reproduce the following code without this statement or

without recognizing

% the original authors in a similar manner.

function varargout = diffusion_analysis(varargin)

% DIFFUSION_ANALYSIS MATLAB code for diffusion_analysis.fig

% DIFFUSION_ANALYSIS, by itself, creates a new

DIFFUSION_ANALYSIS or raises the existing

% singleton*.

%

% H = DIFFUSION_ANALYSIS returns the handle to a new

DIFFUSION_ANALYSIS or the handle to

% the existing singleton*.

%

% DIFFUSION_ANALYSIS('CALLBACK',hObject,eventData,handles,...)

calls the local

% function named CALLBACK in DIFFUSION_ANALYSIS.M with the

given input arguments.

%

% DIFFUSION_ANALYSIS('Property','Value',...) creates a new

DIFFUSION_ANALYSIS or raises the

% existing singleton*. Starting from the left, property value

pairs are

% applied to the GUI before diffusion_analysis_OpeningFcn gets

called. An

% unrecognized property name or invalid value makes property

application

% stop. All inputs are passed to diffusion_analysis_OpeningFcn

via varargin.

%

% *See GUI Options on GUIDE's Tools menu. Choose "GUI allows

only one

% instance to run (singleton)".

%

% See also: GUIDE, GUIDATA, GUIHANDLES

281

% Edit the above text to modify the response to help

diffusion_analysis

% Last Modified by GUIDE v2.5 17-Oct-2011 11:20:13

% Begin initialization code - DO NOT EDIT

gui_Singleton = 1;

gui_State = struct('gui_Name', mfilename, ...

'gui_Singleton', gui_Singleton, ...

'gui_OpeningFcn', @diffusion_analysis_OpeningFcn,

...

'gui_OutputFcn', @diffusion_analysis_OutputFcn,

...

'gui_LayoutFcn', [] , ...

'gui_Callback', []);

if nargin && ischar(varargin{1})

gui_State.gui_Callback = str2func(varargin{1});

end

if nargout

[varargout{1:nargout}] = gui_mainfcn(gui_State, varargin{:});

else

gui_mainfcn(gui_State, varargin{:});

end

% End initialization code - DO NOT EDIT

%%% Mouse location for D

% --- Executes just before diffusion_analysis is made visible.

function diffusion_analysis_OpeningFcn(hObject, eventdata, handles,

varargin)

%% This function has no results args, see OutputFcn.

% hObject handle to figure

% eventdata reserved - to be defined in a future version of MATLAB

% handles structure with handles and user data (see GUIDATA)

% varargin command line arguments to diffusion_analysis (see

VARARGIN)

% Choose default command line results for diffusion_analysis

handles.results = hObject;

% Update handles structure

282

guidata(hObject, handles);

if nargin == 3

initial_dir = pwd;

elseif nargin > 4

if strcmpi(varargin{1},'dir')

if exist(varargin{2},'dir')

initial_dir = varargin{2};

else

errordlg('Input argument must be a valid

directory','Input Argument Error!')

return

end

else

errordlg('Unrecognised input argument','Input Argument

Error!')

return;

end

end

% Populate the listbox

load_listbox(initial_dir,handles)

% UIWAIT makes diffusion_analysis wait for user response (see

UIRESUME)

% uiwait(handles.figure1);

% --- Outputs from this function are returned to the command line.

function varargout = diffusion_analysis_OutputFcn(hObject,

eventdata, handles)

%% varargout cell array for returning results args (see VARARGOUT);

% hObject handle to figure



% Get default command line results from handles structure

varargout{1} = handles.results;

283

% --- Executes on button press in openImage.

function openImage_Callback(hObject, eventdata, handles)

%% hObject handle to openImage (see GCBO)



global filename;

global image;

global boolrot;

[filename, pathname] = uigetfile({'*.*','All Files'},'Select Image

File','MultiSelect','off');

if filename ~= 0

y = dicominfo(strcat(pathname,filename));

image = dicomread(y);

axes(handles.axes1);

imshow(image,[]);

%text((c/2)-120,-.1*r,sprintf('Original Image'));

set(handles.axes1,'Visible','ON');

set(handles.crop,'Visible','ON');

set(handles.status,'String','New Image Loaded..');

set(handles.rotate,'Visible','ON');

boolrot = 0;

end

% ---

function clear_Callback(hObject, eventdata, handles)

%#ok<*INUSL,*DEFNU>

%% hObject handle to multiImage (see GCBO)

% eventdata reserved - to be defined in a future version of

MATLAB


axes(handles.axes3)

cla

axes(handles.axes9)

cla

% --- Executes on button press in crop.

function crop_Callback(hObject, eventdata, handles)

%% hObject handle to crop (see GCBO)



284


global image2;

global rect;

global filename;

global interface_width;

clear f_surface;

str1 = 'Create a rectangle to crop image';

str2 = 'Double click inside rectangle to finalise selection';

str3 = strvcat(str1,str2);

set(handles.status,'String',str3);


[image2 rect] = imcrop();

if isempty(image2)

set(handles.status,'String','User Cancelled');

else

[r,c,d]=size(image2);


imshow(image2,[]);

%axes(handles.axes9);

image2 = int16(image2);

for i=1:r

avg_intensity(i) = mean(image2(i,:));

end;

%pxscale = (1:1:length(avg_intensity)) .*

str2double(get(handles.px_width,'String'));

%interface_width = pxscale(length(pxscale));

%plot(pxscale,avg_intensity,'bx');

%title(sprintf('Pixel intensities in selected timesteps for

image %s',filename),'FontName','Arial')

%xlabel('Depth (mm)','FontName','Arial','FontSize',8)

%ylabel('Mean Pixel Value of row

(16bit)','FontName','Arial','FontSize',8)

set(handles.status,'String','Region Cropped');

set(handles.uipanel3,'Visible','ON');

set(handles.uipanel7,'Visible','ON')

end

%----Executes on button press in multiImage

285

function multiImage_Callback(hObject, eventdata, handles)

%% hObject handle to multiImage (see GCBO)

% eventdata reserved - to be defined in a future version of

MATLAB


global filename;

global rect;

global boolrot;

global deg;

list_entries = get(handles.dir_list,'String');

index_selected = get(handles.dir_list,'Value');

m = numel(index_selected);


hold on

axis auto

ColOrd =

'rgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrgbmkrg

bmkrgbmkrgbmkrgbmk';

for i=1:m

iter_file = strcat(pwd, '/', list_entries{index_selected(i)},...

'/', filename);

if exist(iter_file,'file') == 2

y = dicominfo(iter_file);

I = dicomread(y);

if boolrot == 1;

I = imrotate(I,deg,'bilinear','crop');

end

I2 = imcrop(I, rect);

I2 = int16(I2);

[r,c] = size(I2);

for j=1:r

avg_intensity(j)= mean(I2(j,:));

end

lngth = length(avg_intensity);

pxscale = (1:1:lngth) .*

str2num(get(handles.px_width,'String'));

plot(pxscale,avg_intensity,'.','Color',ColOrd(i));

286

title(sprintf('Pixel intensities in selected timesteps for

image %s',filename),'FontName','Arial','FontWeight','bold')

xlabel('Depth (mm)','FontName','Arial','FontSize',8)

ylabel('Mean Pixel Value of row


set(gca,'FontName','Arial','FontSize',8)

else

errordlg(sprintf('Select one or more directories containing

file %s',filename))

end

end

% --- Executes on button press in exit.

function exit_Callback(hObject, eventdata, handles)

%% hObject handle to exit (see GCBO)



% Clear all variables in memory

clear all;

% Close all figures created by the program and exit

close all;

% --- Executes during object creation, after setting all properties.

function px_width_CreateFcn(hObject, eventdata, handles)

%% hObject handle to px_width (see GCBO)


% handles empty - handles not created until after all CreateFcns

called

% Hint: edit controls usually have a white background on Windows.

% See ISPC and COMPUTER.

if ispc && isequal(get(hObject,'BackgroundColor'),

get(0,'defaultUicontrolBackgroundColor'))

set(hObject,'BackgroundColor','white');

end

% --- Executes on selection change in dir_list.

function dir_list_Callback(hObject, eventdata, handles)

%% hObject handle to dir_list (see GCBO)


287


% Hints: contents = cellstr(get(hObject,'String')) returns dir_list

contents as cell array

% contents{get(hObject,'Value')} returns selected item from

dir_list

get(handles.figure1,'SelectionType');

if strcmp(get(handles.figure1,'SelectionType'),'open')


file_list = get(handles.dir_list,'String');

dirname = file_list{index_selected};

if handles.is_dir(handles.sorted_index(index_selected))

cd(dirname)

load_listbox(pwd,handles)

else

errordlg('Please select valid directories, not files.',...

'Unable to open directory')

end

end

% --- Executes during object creation, after setting all properties.

function dir_list_CreateFcn(hObject, eventdata, handles)

% hObject handle to dir_list (see GCBO)


% handles empty - handles not created until after all CreateFcns

called

% Hint: listbox controls usually have a white background on Windows.

% See ISPC and COMPUTER.

if ispc && isequal(get(hObject,'BackgroundColor'),

get(0,'defaultUicontrolBackgroundColor'))

set(hObject,'BackgroundColor','white');

end

% ---

function load_listbox(dir_path,handles)

%%

cd (dir_path)

dir_struct = dir(dir_path);

[sorted_names, sorted_index] = sortrows({dir_struct.name}');

288

handles.filenames = sorted_names;

handles.is_dir = [dir_struct.isdir];

handles.sorted_index = sorted_index;

guidata(handles.figure1,handles)

set(handles.dir_list,'String',handles.filenames,'Value',1)

set(handles.text8,'String',pwd)

% ---

function f = pde_solution(b,x)

%% b is a 1x4 vector containg in order - D, t0, c0, c1

%%% x must be n*2, column 2 is t

%initialise vars


N = 30;

h0 = interface_width;

D = b(1); %1.0*10^-3;

t0 = b(2); % 40140 - 300

c0 = b(3); %16000;

c1 = b(4); %500;

[r,c] = size(x);

% each row of expr corresponds to a value of x

exprs = zeros(r,1);

% big sigma section of equation

for m = 0:N

exprs = exprs + ( ( ( 4 * (-1)^m ) / (pi + 2 * pi * m) ) * cos(

(pi*(m+ 0.5)*x(:,1)) / h0 ) .* ...

exp( -D * (x(:,2) - t0) * ((pi * (m + 0.5) / h0) * (pi*(m +

0.5) / h0)) ) );

end

f = c0 * exprs + c1;

% --- Executes on button press in nlin_push.

function nlin_push_Callback(hObject, eventdata, handles)

%% hObject handle to nlin_push (see GCBO)



289

% Derived from Mathematica code written by Konstantin Momot

global filename;

global rect;


global f_surface;

global pxscale;

global tsteps;

global boolrot;

global deg;

%clear f_surface;



ind = numel(index_selected);


axis auto

for i=1:ind

iter_file = strcat(pwd, '/', list_entries{index_selected(i)},...

'/', filename);



timestr = y.AcquisitionTime;

tsteps(i) = str2num(timestr(1:2))*3600 +

str2num(timestr(3:4))*60 + str2num(timestr(5:6));

I = dicomread(y);

if boolrot == 1


end


I2 = int16(I2);

[r,c] = size(I2);

for j=1:r

avg_intensity(j)= mean(I2(j,:));

end

pxscale = (1:1:r) .*


interface_width = pxscale(length(pxscale));

f_surface(i,1:r) = avg_intensity;

else

290

errordlg(sprintf('Select all directories containing file

%s',filename))

end

end

warning off all

options = statset('MaxIter',1000, 'TolX',1e-10);

coeff = [1.0e-4, 3.6e4, 3.0e4, 1.0e3];

[nplots,m] = size(f_surface);

t = ones(1,m);

yvec = f_surface(1,:);

xvec = pxscale;

time = t * tsteps(1);

for j = 2:nplots

yvec = cat(2,yvec,f_surface(j,:));

xvec = cat(2,xvec,pxscale);

time = cat(2, time, t * tsteps(j));

end

[coeff_fit, resid, J, COVB, mse] = nlinfit([xvec',time'], yvec',

@pde_solution, coeff, options);

ci = nlparci(coeff_fit,resid,'jacobian',J);

x_append = [0, pxscale];

f_fit = pde_solution(coeff_fit,[x_append', [1; t']*mean(tsteps)]);

%note that this will skew the curve if more images are selected

either side of the halfway point

plot(x_append,f_fit,'r-')

title(sprintf('Fitted diffusion curve for all timesteps of

%s',filename),'FontName','Arial','FontWeight','bold')

xlabel('Depth (mm)','FontName','Arial','FontSize',8)

ylabel('Mean Pixel Value of row


set(gca,'FontName','Arial','FontSize',8)

set(findobj(handles.axes3,'Type','line','-

and','Color','r'),'LineWidth',1.5);

set(handles.up_ci,'String',num2str(mean(ci(1,2))))

291

set(handles.low_ci,'String',num2str(mean(ci(1,1))))

set(handles.avgD,'String',num2str(coeff_fit(1)))

% --- Executes on button press in button_surf.

function button_surf_Callback(hObject, eventdata, handles)

% hObject handle to button_surf (see GCBO)



global tsteps;

global pxscale;

global f_surface;

figure;

reltime = tsteps - tsteps(1);

surf(pxscale,reltime,f_surface);

title('Mean pixel value over time as surface')

xlabel('Depth (mm)')

ylabel('Time (sec)')

zlabel('Mean Pixel Value (16bit)')

% --- Executes on button press in pushbutton12.

function pushbutton12_Callback(hObject, eventdata, handles)

% hObject handle to pushbutton12 (see GCBO)



fig2 = figure('visible','off');

h = copyobj(handles.axes3,fig2);

set(h,'position', [0.1 0.1 0.85 0.85],'FontName','Arial')

[filename, ext, user_canceled] = imputfile;

if (user_canceled == 0)

saveas(h, filename, ext)

end

close(fig2)

% --- Executes on button press in pushbutton13.

292

function pushbutton13_Callback(hObject, eventdata, handles)

% hObject handle to pushbutton13 (see GCBO)



fig2 = figure('visible','off');

h = copyobj(handles.axes9,fig2);

set(h,'position', [0.1 0.1 0.85 0.85],'FontName','Arial')

[filename, ext, user_canceled] = imputfile;

if (user_canceled == 0)

saveas(h, filename, ext)

end

close(fig2)

% --- Executes on button press in rotate.

function rotate_Callback(hObject, eventdata, handles)

% hObject handle to rotate (see GCBO)



global image;

global boolrot;

global deg;

axes(handles.axes1)

set(handles.status,'String',strvcat('Select bottom left', 'and right

pixels in sample'));

[x,y] = ginput(2);

xlen = x(2) - x(1);

ylen = y(2) - y(1);

deg = atand(ylen/xlen);

imtest = imrotate(image,deg,'bilinear','crop');

if isempty(imtest)

set(handles.status,'String','User cancelled rotate.');

else

image = imtest;

cla

imshow(image,[]);

set(handles.status,'String',strvcat('Image rotated.', 'Select

stripe to isolate','analysis region.'));

end

293

boolrot = 1;

% --- Executes on button press in nlin_layer.

function nlin_layer_Callback(hObject, eventdata, handles)

% hObject handle to nlin_layer (see GCBO)



global filename;

global rect;

global boolrot;

global deg;



ind = numel(index_selected);

warning off all

options = statset('MaxIter',1000, 'TolX',1e-10);

coeff = [1.0e-4, 3.6e4, 3.0e4, 1.0e3];

Da_all = zeros(ind,1);

Db_all = zeros(ind,1);

Dc_all = zeros(ind,1);

Dd_all = zeros(ind,1);

for j = 1:ind

iter_file = strcat(pwd, '/', list_entries{index_selected(j)},...

'/', filename);



timestr = y.AcquisitionTime;

tsteps(j) = str2num(timestr(1:2))*3600 +

str2num(timestr(3:4))*60 + str2num(timestr(5:6));

I = dicomread(y);

if boolrot == 1


end


I2 = int16(I2);

294

[r,c] = size(I2);

midpartition = round(r * 0.5);

tenthpartition = round(r * 0.1);

if tenthpartition == 0

tenthpartition = 1;

end

I2_d = I2(1:tenthpartition,:);

I2_c = I2(tenthpartition+1:midpartition,:);

I2_b = I2(midpartition+1:end-tenthpartition,:);

I2_a = I2(end-tenthpartition+1:end,:);

% figure()

% subplot(2,2,1), imshow(I2_a,[])

% subplot(2,2,2), imshow(I2_b,[])

% subplot(2,2,3), imshow(I2_c,[])

% subplot(2,2,4), imshow(I2_d,[])

avg_intensity_a = mean(I2_a,2)';

avg_intensity_b = mean(I2_b,2)';

avg_intensity_c = mean(I2_c,2)';

avg_intensity_d = mean(I2_d,2)';

pxscale = (1:1:r) .*


time = ones(1,r) .* tsteps(j);

coeff_fit_d =

nlinfit([pxscale(1:tenthpartition)',time(1:tenthpartition)'],avg_int

ensity_d', @pde_solution, coeff, options);

coeff_fit_c =

nlinfit([pxscale(tenthpartition+1:midpartition)',

time(tenthpartition+1:midpartition)'],avg_intensity_c',

@pde_solution, coeff, options);

coeff_fit_b = nlinfit([pxscale(midpartition+1:end-

tenthpartition)',time(midpartition+1:end-

tenthpartition)'],avg_intensity_b', @pde_solution, coeff, options);

coeff_fit_a = nlinfit([pxscale(end-

tenthpartition+1:end)',time(end-

tenthpartition+1:end)'],avg_intensity_a', @pde_solution, coeff,

options);

Da_all(j) = coeff_fit_a(1);

Db_all(j) = coeff_fit_b(1);

Dc_all(j) = coeff_fit_c(1);

295

Dd_all(j) = coeff_fit_d(1);

end

end

fig1 = figure();

time = tsteps - tsteps(1);

plot(time,Da_all,'r.',time,Db_all,'b.',time,Dc_all,'g.',time,Dd_all,

'm.')

xlabel('time (s)','FontName','Arial','FontSize',10)

ylabel('Estimated D','FontName','Arial','FontSize',10)

set(gca,'FontName','Arial','FontSize',10);

axis auto

legend('Layer "a"','Layer "b"', 'Layer "c"', 'Layer "d"')

296

B1 A graphical user interface (GUI) for computing the apparent diffusion coefficient

of cartilage matrix from the magnetic resonance data.

297

APPENDIX C

Raman Spectral Band Assignments

298

Table C1 Raman band assignments

Sample Assignment Component Tissue

Normal 2949 CH3 and CH2

str

lipid

2890 CH2 str lipid

1666 Amide I Collagen Cartilage and bone

1642 Amide I Collagen Cartilage and

bone

1455 CH2 deformation

Protein, lipid Cartilage and bone

1429

1347

1320

1251 Amide III Collagen Cartilage,

subchondral

bone

1168

1005 Phenylalanine collagen Cartilage,

subchondral

bone

939

923 Proline Collagen Cartilage,

subchondral

bone

857 Hydroxyproline Collagen Cartilage, subchondral

bone

817

6-2

POPC

b d

1454 1692

1398 1051

1361 1033

997 1005

877 969

940

858

818

6-3

DPPC

b e

1662 1666 Amide I, occurrence of

three of more

bands with max

299

intensity at

1636, 1656 ( -helical) and

above 1670 ( -sheets)

correspond to

different secondary

protein

structures.

1456 1455

1289 1249 Amide III

1036 1210

1005 1171

939 1005

815 940

920

853

814

Blue-characteristics of collagen

The Band assignments were obtained from: Esmonde-White, Analyst, 2011 136 (8)

1675-1685, and Naumann Applied Spectroscopy Reviews vol 36, 2-3 2001.

300

Spectra collected from different areas of the sample

Sample Assignment

1 POPC 2952

1670

1649

1455

1267

1246

3 DPPC

a

2950

1669

1457

1247

1005

940

857

817

2-SAPL 2949

2901

1661

1640

1455

1432

1255

1245

1006

939

878

857

817

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