Post on 04-Jan-2017
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
ii
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 Materials and Methods ........................................................................................................ 181
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
6.4 Results and Observations .................................................................................................... 188
6.5 Conclusion ......................................................................................................................... 198
CHAPTER 7: ASSESSMENT OF THE MECHANICAL INTEGRITY/ PHYSIOLOGICAL FUNCTION
OF RESURFACED ARTICULAR CARTILAGE ..................................................................... 199
7.1 Introduction ........................................................................................................................ 199
7.2 Materials and Methods ........................................................................................................ 201
7.2.1 Articulr cartilage samples ......................................................................................... 201
7.2.2 Derivation of the energy components ........................................................................ 202
7.2.3 Statistical Analyses .................................................................................................. 205
7.3 Results and Observations .................................................................................................... 207
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
xvi
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
relipidized (POPC, DPPC and complete SAPL mix) surfaces. ........................................ 212
Figure 7.7 Residual energy plotted for articular cartilage specimens with normal intact, delipidized, and
relipidized (POPC, DPPC and complete SAPL mix) surfaces. ........................................ 213
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,
delipidized, relipidized surfaces. .................................................................................... 216
Figure 7.11 Residual energy versus strain energy for articular cartilage samples with normal intact,
delipidized, relipidized surfaces. .................................................................................... 216
Figure 7.12 Residual energy versus released energy for articular cartilage samples with normal intact,
delipidized, relipidized surfaces. .................................................................................... 217
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
89
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:
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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
)
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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
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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
133
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.
6.3 MATERIALS AND METHODS
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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6.4 RESULTS AND OBSERVATIONS
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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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 normal intact articular surface (length (X) and breadth
(Y) of the scanned area, and average peak height of SAL (Z)).
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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)).
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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 MATERIALS AND METHODS
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
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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
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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).
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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.
7.3 RESULTS AND OBSERVATIONS
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
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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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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
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
% 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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
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
% handles structure with handles and user data (see GUIDATA)
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
284
% handles structure with handles and user data (see GUIDATA)
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);
axes(handles.axes1);
[image2 rect] = imcrop();
if isempty(image2)
set(handles.status,'String','User Cancelled');
else
[r,c,d]=size(image2);
axes(handles.axes7);
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
% handles structure with handles and user data (see GUIDATA)
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);
axes(handles.axes9);
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
(16bit)','FontName','Arial','FontSize',8)
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
% 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)
% eventdata reserved - to be defined in a future version of MATLAB
% 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)
% eventdata reserved - to be defined in a future version of MATLAB
287
% handles structure with handles and user data (see GUIDATA)
% 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')
index_selected = get(handles.dir_list,'Value');
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)
% eventdata reserved - to be defined in a future version of MATLAB
% 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
global interface_width;
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
289
% Derived from Mathematica code written by Konstantin Momot
global filename;
global rect;
global interface_width;
global f_surface;
global pxscale;
global tsteps;
global boolrot;
global deg;
%clear f_surface;
list_entries = get(handles.dir_list,'String');
index_selected = get(handles.dir_list,'Value');
ind = numel(index_selected);
axes(handles.axes3);
axis auto
for i=1:ind
iter_file = strcat(pwd, '/', list_entries{index_selected(i)},...
'/', filename);
if exist(iter_file,'file') == 2
y = dicominfo(iter_file);
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
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
pxscale = (1:1:r) .*
str2num(get(handles.px_width,'String'));
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
(16bit)','FontName','Arial','FontSize',8)
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
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)
% eventdata reserved - to be defined in a future version of MATLAB
% handles structure with handles and user data (see GUIDATA)
global filename;
global rect;
global boolrot;
global deg;
list_entries = get(handles.dir_list,'String');
index_selected = get(handles.dir_list,'Value');
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);
if exist(iter_file,'file') == 2
y = dicominfo(iter_file);
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
I = imrotate(I,deg,'bilinear','crop');
end
I2 = imcrop(I, rect);
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) .*
str2num(get(handles.px_width,'String'));
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