Thông tin tài liệu
IN VITRO AND IN VIVO EVALUATION OF CUSTOMIZED
POLYCAPROLACTONE TRICALCIUM PHOSPHATE SCAFFOLDS
FOR BONE TISSUE ENGINEERING
ERVI SJU
(B.Eng.(Hons.), NUS)
A THESIS SUBMITTED
FOR THE DEGREE OF MASTER OF ENGINEERING
DEPARTMENT OF MECHANICAL ENGINEERING
NATIONAL UNIVERSITY OF SINGAPORE
2010
PREFACE
The thesis is submitted for the degree of Master of Engineering in the Department of
Mechanical Engineering at the National University of Singapore under the
supervision of Professor Teoh Swee Hin and Dr Alvin Yeo. No part of this thesis has
been submitted for other degree at other university or institution. Parts of this thesis
have been published or presented in the following:
INTERNATIONAL JOURNAL PUBLICATION
A. Yeo, E. Sju, B. Rai, S.H. Teoh. Customizing the degradation and load-bearing
profile of 3D polycaprolactone-tricalcium phosphate scaffolds under enzymatic and
hydrolytic conditions. Journal of Biomedical Materials Research Part B: Applied
Biomaterials. (Published online: 10 June 2008).
CONFERENCE PAPERS
E. Sju, A. Yeo, B. Rai, S.H. Teoh. In vitro and in vivo degradation profile of
untreated, sodium hydroxide- and lipase-treated PCL-TCP scaffolds. International
Conference on Advances in Bioresorbable Biomaterials for Tissue Engineering,
Singapore, 2008.
E. Sju, A. Yeo, B. Rai, S.H. Teoh. Enzymatic and hydrolytic degradation of poly(εcaprolactone)
tricalcium
phosphate
composite
scaffolds.
4th
International
Conference on Materials for Advanced Technologies (ICMAT), Singapore, 2007.
i
ACKNOWLEDGEMENTS
The author wishes to express her sincere gratitude and heartfelt appreciation to the
following people who have rendered generous support and technical assistance
leading toward the accomplishment of this project:
Professor Teoh Swee Hin (Department of Mechanical Engineering), supervisor,
for offering the privileged opportunity to work on this project and allowing the
author to join his team, for his expertise, kindness, and most of all, his patience.
His enthusiasm in research and continuous support have truly been a source of
inspiration and motivation for this project throughout.
Dr. Alvin Yeo (Department of Mechanical Engineering and National Dental
Centre), co-supervisor, for his patience and guidance on supervising the author
throughout the whole process. He has been an immense driving force behind this
project. One simply could not wish for a better or friendlier supervisor.
Dr. Bina Rai, mentor, for graciously sharing her knowledge and encouragement
in this project. Her kind assistance and time spent are greatly appreciated.
Dr. Zhang Zhiyong, Ms. Erin Teo Yi Ling and Mr. Mark Chong Seow Khoon,
PhD students, for their constructive feedbacks and for being excellent mentors.
They have gone out of their way to render assistance on many occasions.
Mr. Cheong Jia Jian, NUS alumnus, whom was unreserved in sharing his
knowledge and experience in this research field.
Mdm. Zhong Xiang Li (Materials Science Lab) for the use of the SEM (JEOL
JSM 5600LV) and the gold-sputtering machine.
ii
Dr. Zhang Yanzhong (Biomechanics Lab) and Ms. Eunice Tan Phay Sing
(NanoBiomechanics Lab) for the use of the Instron 3345 compressive
mechanical tester machine.
Dr. Jeremy Teo Choon Meng (DSO National Lab) and Ms. Lei Yang (Biosignal
and Instrumentation Lab) for the use of the Skyscan 1076 Micro-CT.
Dr. Lin Jian Hua and Ms. Juline Sim Siew Hong (PSB corporation) for their
assistance in Gel Permeation Chromatography.
Ms. Irene Kee (SingHealth Experimental Medicine Centre, Singapore General
Hospital) for her assistance in the animal handling and maintenance.
Ms. Han Tok Lin (Faculty of Dentistry, NUS) for her assistance in Histology.
Mr. Jackson Ong Sing Kiat and Dr. Chui Chee Kong (BIOMAT), Mr. Zhang
Jing (Biosignal and Instrumentation Lab), and fellow students at BIOMAT for
their support and encouragement throughout the fulfilling years.
To all, who have given contribution in one way or another.
And to all close friends, for being understanding during this challenging period.
Thank you for always being there during both good and bad times.
Last but not least, the author would like to thank her parents Mr. Sju Tjing
Kwang and Mrs. Tea Giok Tjian, and younger sister Ms. Lydia Sju, for their
constant love and support, without which this study would not have been
possible. Their undaunting confidence gave the author the strength to overcome
any difficulties. To them the author dedicates this thesis.
The author acknowledged the financial support by the following grants:
No. 016/06 from National Dental Centre (SingHealth), Singapore.
TDF/CD003/2006 from SingHealth (Talent Development Fund), Singapore.
iii
TABLE OF CONTENTS
PREFACE
i
ACKNOWLEDGEMENTS
ii
TABLE OF CONTENTS
iv
SUMMARY
ix
LIST OF TABLES
xi
LIST OF FIGURES
xii
LIST OF SYMBOLS
xviii
LIST OF ABBREVIATIONS
xx
CHAPTER 1: INTRODUCTION
1.1 BACKGROUND
1
1.1.1 Bone tissue engineering
1
1.1.2 Application in dentoalveolar defects
3
1.1.3 PCL-TCP scaffolds: Current drawbacks
4
1.2 RESEARCH OBJECTIVES
5
1.3 RESEARCH SCOPE
6
1.3.1 Part 1: Selective modification of PCL-TCP scaffolds
targeted for dentoalveolar reconstruction application
6
1.3.2 Part 2: Optimization of native and customized scaffolds
in vitro and their effects in initial bone healing
6
1.3.3 Part 3: Evaluation of PCL-TCP scaffolds in a clinically
relevant defect model
7
iv
CHAPTER 2: LITERATURE REVIEW
2.1 BONE PHYSIOLOGY
8
2.1.1 Composition
8
2.1.2 Morphology
11
2.2 POLY(ε-CAPROLACTONE)
13
2.2.1 Degradation of PCL polymer
14
2.2.1.1 Hydrolysis mechanism
16
2.2.1.2 Enzymatic degradation
17
2.3 TRICALCIUM PHOSPHATE (TCP)
2.3.1 Degradation mechanisms of calcium phosphate ceramics
18
19
2.3.1.1 Physicochemical degradation
19
2.3.1.2 Cellular degradation
21
2.3.1.3 Mechanical degradation
21
2.4 PCL-TCP SCAFFOLDS
22
2.4.1 Fabrication method of PCL-TCP scaffolds
24
CHAPTER 3: SELECTIVE MODIFICATION OF PCL-TCP SCAFFOLDS TARGETED
FOR DENTOALVEOLAR RECONSTRUCTION APPLICATION
3.1 INTRODUCTION
29
3.2 MATERIALS AND METHODS
30
3.2.1 Scaffold design and fabrication
30
3.2.2 Sterilization of scaffolds
30
3.2.3 Scaffold characterizations
31
3.2.3.1 Micro-computed tomography analysis
31
3.2.3.2 Gravimetric analysis
31
3.2.3.3 Compressive mechanical testing
32
3.2.3.4 Electron microscopy preparation and analysis
32
v
3.2.3.5 Molecular weight testing
3.2.4 Statistical analysis
33
33
3.3 RESULTS
34
3.3.1 Porosity measurements and 3D model analysis
34
3.3.2 Weight loss analysis
36
3.3.3 Compressive mechanical properties
37
3.3.4 Surface morphology analysis
38
3.3.5 Molecular weight analysis
40
3.4 DISCUSSION
40
3.5 CONCLUSION
43
CHAPTER 4: OPTIMIZATION OF NATIVE AND CUSTOMIZED SCAFFOLDS
IN VITRO AND THEIR EFFECTS IN INITIAL BONE HEALING
4.1 INTRODUCTION
44
4.1.1 In vitro degradation study
44
4.1.2 In vivo degradation study
45
4.2 MATERIALS AND METHODS
46
4.2.1 Scaffold design and fabrication
46
4.2.2 Sterilization of scaffolds
47
4.2.3 Animal husbandry
47
4.2.4 Scaffold implantation
48
4.2.5 Scaffold characterizations
49
4.2.5.1 Micro-computed tomography analysis
50
4.2.5.2 Gravimetric analysis
50
4.2.5.3 Compressive mechanical testing
50
4.2.5.4 Electron microscopy preparation and analysis
50
4.2.5.5 Molecular weight testing
50
vi
4.2.5.6 Histology preparation and analysis
50
4.2.6 Statistical analysis
51
4.3 RESULTS - In vitro degradation study
51
4.3.1 Porosity measurements and 3D model analysis
51
4.3.2 Weight loss analysis
57
4.3.3 Compressive mechanical properties
58
4.3.4 Surface morphology analysis
60
4.3.5 Molecular weight analysis
66
4.4 RESULTS - In vivo degradation study
67
4.4.1 Porosity measurements and 3D model analysis
67
4.4.2 Weight loss analysis
70
4.4.3 Compressive mechanical properties
71
4.4.4 Surface morphology analysis
72
4.4.5 Molecular weight analysis
75
4.4.6 Histology analysis
76
4.5 DISCUSSION
79
4.5.1 Comparison between in vitro and in vivo studies
4.6 CONCLUSION
82
83
CHAPTER 5: EVALUATION OF PCL-TCP SCAFFOLDS IN A CLINICALLY
RELEVANT DEFECT MODEL
5.1 INTRODUCTION
85
5.2 MATERIALS AND METHODS
88
5.2.1 Implant design and fabrication
88
5.2.2 Animal husbandry
89
5.2.3 Pre- and postoperative medication
90
5.2.4 Surgery 1 (Extraction and defect creation)
91
vii
5.2.5 Surgery 2 (Ridge augmentation)
93
5.2.6 Sacrifice
95
5.2.7 Micro-computed tomography analysis
96
5.3 RESULTS
96
5.3.1 Gross examinations
96
5.3.2 New bone formation
98
5.3.3 Ratio of bone volume fraction for PCL-TCP scaffolds
with respect to autografts
100
5.3.4 3D model analysis
101
5.3.5 Two-dimensional x-ray radiographs evaluation
102
5.4 DISCUSSION
104
5.5 CONCLUSION
109
CHAPTER 6: FINAL CONCLUSIONS AND RECOMMENDATIONS
6.1 FINAL CONCLUSIONS
110
6.2 RECOMMENDATIONS FOR FUTURE WORK
112
REFERENCES
114
APPENDICES
122
APPENDIX A – PART 1 STUDY
122
APPENDIX B – PART 2 STUDY
134
APPENDIX C – PART 3 STUDY
148
viii
SUMMARY
The research scope encompasses the degradation and load-bearing profile of 3D
bioresorable polycaprolactone-20% tricalcium phosphate (PCL-TCP) scaffolds under
enzymatic and hydrolytic conditions and subsequently to evaluate the efficacy of the
scaffolds in both small and large animal models. The purpose was to develop
scaffolds with desirable customized properties and increased degradation rates
suitable for application in dentoalveolar defects treatment. The scope of this thesis
ended with a large animal study, a stage just before preclinical trials.
Initially, the PCL-TCP scaffolds were degraded in either sodium hydroxide or lipase
solution for 0, 12, 24, 36, 48, 60, 72, 84, 96, and 108 hours. Samples were
recovered at each time point and the following properties of the scaffolds were
measured: porosity, 3D structure, weight loss, compressive strength and modulus,
surface morphology, polymer molecular weight, and histology. In the second part of
the study, in vitro and in vivo degradation behaviours of these treated scaffolds were
investigated. PCL-TCP scaffolds were monitored after immersion in standard culture
medium for 0, 6, 12, 18 and 24 weeks in vitro. In vivo degradation of the scaffolds
was performed by implanting these scaffolds subcutaneously at the back of rats for
12 and 24 weeks. Upon retrieval, analyses similar to those described above were
performed. Lastly, another in vivo study was conducted whereby PCL-TCP scaffolds
and sheets were evaluated as defect fillers and barrier membranes respectively for
novel guided bone regeneration technique in the reconstruction of localized
ix
dentoalveolar defects in a micropig model for up to 6 months. The possibility of the
PCL-TCP scaffold for use as a bone substitute was compared to the current gold
standard of using autogenous bone.
The first objective of the study was achieved with scaffolds of approximately 85%
porosity obtained after 96 hours of treatment in 3M NaOH and 12 hours in 0.1%
lipase. These pre-treated scaffolds demonstrated favourable mechanical strength,
structure, and surface morphology. Secondly, the in vivo degradation profile of
porous PCL-TCP scaffolds are comparable with the obtained in vitro profile. Further,
the degradation rate of the lipase-treated scaffolds was noted to be the highest. This
is followed by NaOH-treated scaffolds and native untreated scaffolds. Overall, the
data suggest that NaOH-treated scaffolds demonstrate the best degradation profile
and physical properties for dentoalveolar reconstruction applications. They possess
the potential to degrade in a controlled and predictable fashion and still display
favourable mechanical strength within a desired time period for new bone formation
to occur. Lastly, healing was uneventful in all micropigs showed that the PCL-TCP
scaffolds exhibited good biocompatibility. Across the tested treatment options, defect
sites augmented with autografts and collagen membranes showed the most
promising results with greater bone formation detected as compared to PCL-TCP
scaffolds and collagen membranes which were about 64% efficient. The collagen
membranes were found to offer the advantage of a reduced frequency of soft tissue
dehiscence in comparison to PCL-TCP sheets. More improvements are needed to
increase the efficiency of the PCL-TCP scaffolds in bone healing as they could ruled
out the need for harvesting grafts.
x
LIST OF TABLES
Table 3.1
Mw, Mn, and PDI of NaOH-treated and lipase-treated PCL-TCP
Scaffolds.
40
Table 4.1
Mw, Mn, and PDI of native, NaOH-treated, and lipase-treated
PCL-TCP Scaffolds in vitro.
66
Table 4.2
Mw, Mn, and PDI of native, NaOH-treated, and lipase-treated
PCL-TCP Scaffolds in vivo.
75
Table 5.1
Number of sites with soft tissue dehiscence for the implanted
autograft, collagen membranes, PCL-TCP scaffolds, and PCLTCP sheets.
98
xi
LIST OF FIGURES
Figure 1.1
Schematic diagram of research scope.
6
Figure 1.2
Schematic diagram of part 1 and part 2 study.
7
Figure 2.1
Composition of bone.
9
Figure 2.2
The assembly of collagen fibrils and fibers and bone mineral
crystals.
9
Figure 2.3
Microscopic structure of cortical and cancellous bone.
11
Figure 2.4
The hierarchical structure of bone from macrostructure to subnanostructure.
12
Figure 2.5
Chemical structure of PCL (as circled).
13
Figure 2.6
The degradation rate of PGA, PLA, and PCL.
16
Figure 2.7
The chemical structure of TCP.
19
Figure 2.8
Schematic diagram of the FDM process.
25
Figure 2.9
Sequence of the data preparation for FDM model fabrication.
26
Figure 3.1
Centrifuge tubes.
29
Figure 3.2
Illustration of scaffold with 0/60/120º lay-down pattern.
30
Figure 3.3
5x5x3mm PCL-TCP scaffold.
31
Figure 3.4
Porosity measurements of NaOH-treated and lipase-treated
PCL-TCP Scaffolds over time.
34
Figure 3.5
3D model of original scaffold (of 75% porosity) at 0 hour:
(L) top view, (R) tilted view.
35
Figure 3.6
3D model of scaffolds after 96 hours immersion in 3M NaOH:
(L) top view, and (R) tilted view.
35
Figure 3.7
3D model of scaffolds after 12 hours immersion in 0.1%
lipase: (L) top view, and (R) tilted view.
36
xii
Figure 3.8
Comparison of weight loss between NaOH-treated and
lipase-treated PCL-TCP scaffolds.
36
Figure 3.9
Compressive strength of PCL-TCP scaffolds when treated
with NaOH and lipase at pre-determined time intervals.
37
Figure 3.10
Compressive modulus of PCL-TCP scaffolds when treated
with NaOH and lipase at pre-determined time intervals.
Electron micrographs of original scaffold (of porosity 75%) at 0
hour: (L) overall view, and (R) higher-magnification view.
38
Figure 3.12
Electron micrographs of scaffold after 96 hours immersion in
3M NaOH: (L) overall view, and (R) higher-magnification
view.
39
Figure 3.13
Electron micrographs of scaffold after 12 hours immersion in
0.1% lipase: (L) overall view, and (R) higher-magnification
view.
39
Figure 4.1
Native (left), NaOH-treated (middle), and lipase-treated (right)
scaffolds.
45
Figure 4.2
Rat at the start of experiment (left) and at the end after 6
months (right).
46
Figure 4.3
50x50x3mm PCL-TCP scaffold.
46
Figure 4.4
5x5x3mm PCL-TCP scaffold.
47
Figure 4.5
Rat cages.
47
Figure 4.6
Rat shaved and scrubbed with iodine.
48
Figure 4.7
Scaffolds’ positioning.
48
Figure 4.8
Incision made (left), implanted scaffold (left, inset), scaffold
positions (right).
49
Figure 4.9
Sacrifice of rats.
49
Figure 4.10
Removal of scaffolds.
49
Figure 4.11
Porosity measurements of native, NaOH-treated, and lipasetreated PCL-TCP scaffolds after immersion in DMEM for 6,
12, and 18 weeks.
52
Figure 4.12
3D model of native scaffold (of 85% porosity) at week 0:
(L) top view, and (R) tilted view.
52
Figure 3.11
xiii
38
Figure 4.13
3D model of native scaffold after 6 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
53
Figure 4.14
3D model of NaOH-treated scaffold after 6 weeks immersion
in DMEM: (L) top view, and (R) tilted view.
53
Figure 4.15
3D model of lipase-treated scaffold after 6 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
53
Figure 4.16
3D model of native scaffold after 12 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
54
Figure 4.17
3D model of NaOH-treated scaffold after 12 weeks immersion
in DMEM: (L) top view, and (R) tilted view.
54
Figure 4.18
3D model of lipase-treated scaffold after 12 weeks immersion
in DMEM: (L) top view, and (R) tilted view.
54
Figure 4.19
3D model of native scaffold after 18 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
55
Figure 4.20
3D model of NaOH-treated scaffold after 18 weeks immersion
in DMEM: (L) top view, and (R) tilted view.
55
Figure 4.21
3D model of lipase-treated scaffold after 18 weeks immersion
in DMEM: (L) top view, and (R) tilted view.
55
Figure 4.22
3D model of native scaffold after 24 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
56
Figure 4.23
3D model of NaOH-treated scaffold after 24 weeks immersion
in DMEM: (L) top view, and (R) tilted view.
56
Figure 4.24
3D model of lipase-treated scaffold after 24 weeks immersion
in DMEM: (L) top view, and (R) tilted view.
56
Figure 4.25
Weight loss of PCL-TCP Scaffolds In vitro.
58
Figure 4.26
Relative compressive strength of PCL-TCP Scaffolds In vitro.
59
Figure 4.27
Relative compressive modulus of PCL-TCP Scaffolds In vitro.
59
Figure 4.28
Electron micrographs taken after 6 weeks immersion in
DMEM for: (a,b) native, (c,d) NaOH-treated, and (e,f) lipasetreated scaffolds. (L) overall view, and (R) highermagnification view.
62
xiv
Figure 4.29
Electron micrographs taken after 12 weeks immersion in
DMEM for: (a,b) native, (c,d) NaOH-treated, and (e,f) lipasetreated scaffolds. (L) overall view, and (R) highermagnification view.
63
Figure 4.30
Electron micrographs taken after 18 weeks immersion in
DMEM for: (a,b) native, (c,d) NaOH-treated, and (e,f) lipasetreated scaffolds. (L) overall view, and (R) highermagnification view.
64
Figure 4.31
Electron micrographs taken after 24 weeks immersion in
DMEM for: (a,b) native, (c,d) NaOH-treated, and (e,f) lipasetreated scaffolds. (L) overall view, and (R) highermagnification view.
65
Figure 4.32
Electron micrographs of native scaffold (of 85% porosity) at
week 0: (L) overall view, and (R) higher-magnification view.
66
Figure 4.33
Porosity of PCL-TCP Scaffolds In vivo.
67
Figure 4.34
3D model of native scaffold after 3 months implantation:
(L) top view, and (R) tilted view.
68
Figure 4.35
3D model of NaOH-treated scaffold after 3 months
implantation: (L) top view, and (R) tilted view.
68
Figure 4.36
3D model of lipase-treated scaffold after 3 months
implantation: (L) top view, and (R) tilted view.
68
Figure 4.37
3D model of native scaffold after 6 months implantation:
(L) top view, and (R) tilted view.
69
Figure 4.38
3D model of NaOH-treated scaffold after 6 months
implantation: (L) top view, and (R) tilted view.
69
Figure 4.39
3D model of lipase-treated scaffold after 6 months
implantation: (L) top view, and (R) tilted view.
69
Figure 4.40
Weight loss of PCL-TCP Scaffolds In vivo.
70
Figure 4.41
Relative compressive strength of PCL-TCP Scaffolds In vivo.
71
Figure 4.42
Relative compressive modulus of PCL-TCP Scaffolds In vivo.
72
Figure 4.43
Electron micrographs taken after 3 months implantation:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated
scaffolds. (L) overall view, and (R) higher-magnification view.
73
xv
Figure 4.44
Electron micrographs taken after 6 months implantation:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated
scaffolds. (L) overall view, and (R) higher-magnification view.
74
Figure 4.45
H&E stain of native scaffolds after 3 months implantation.
76
Figure 4.46
H&E stain of native scaffolds after 6 months implantation.
76
Figure 4.47
H&E stain of NaOH-treated scaffolds after 3 months
implantation.
77
Figure 4.48
H&E stain of NaOH-treated scaffolds after 6 months
implantation.
77
Figure 4.49
H&E stain of lipase-treated scaffolds after 3 months
implantation.
78
Figure 4.50
H&E stain of lipase-treated scaffolds after 6 months
implantation.
78
Figure 5.1
Timeline for the complete micropig study.
87
Figure 5.2
15x10x8mm PCL-TCP scaffold (left) and 25x25x1mm PCLTCP sheet (right).
88
Figure 5.3
Bioresorbable collagen membrane from BioGide (left) and
temperature-controlled hot water bath (right).
89
Figure 5.4
Micropig housing facility at SEMC, SGH (left) and weighing of
micropig prior to the experiment (right).
90
Figure 5.5
Removal of all premolars and first molar (left), and the
extraction sites (right).
92
Figure 5.6
The flaps were re-approximated with Vicryl sutures (left), and
the defect sites were closed (right).
92
Figure 5.7
Schematic illustrations of the four tested grafting procedures.
93
Figure 5.8
Placement of PCL-TCP scaffolds and autografts (left),
followed by PCL-TCP sheets and collagen membranes (right).
94
Figure 5.9
Micropig under euthanasia (left), and the mandible was block
resected using an oscillating autopsy saw (right).
95
Figure 5.10
The recovered segment of mandible (left), the site after
removal (right).
95
Figure 5.11
The recovered segment of the mandible of a micropig.
97
xvi
Figure 5.12
Soft tissue dehiscence observed for the majority of grafts
covered with PCL-TCP sheets.
97
Figure 5.13
Bone volume fraction detected after 6 months of implantation
of autografts and PCL-TCP scaffolds for individual micropigs.
99
Figure 5.14
The average values of bone volume fraction detected after 6
months of implantation of autografts and PCL-TCP scaffolds.
100
Figure 5.15
The ratio of bone volume fraction for PCL-TCP scaffolds with
respect to autografts for individual micropigs.
101
Figure 5.16
PCL-TCP scaffold treated site: overview (left) and crosssection (right).
102
Figure 5.17
Autograft-treated site: overview (left) and cross-section (right).
102
Figure 5.18
X-ray image of a micropig’s left mandible treated with
autograft (posterior) and PCL-TCP scaffold (anterior), and
covered with collagen membrane.
103
Figure 5.19
X-ray image of a micropig’s right mandible treated with PCLTCP scaffold (posterior) and autograft (anterior), and covered
with collagen membrane.
103
Figure 5.20
X-ray image of a micropig’s left mandible treated with
autograft (posterior) and PCL-TCP scaffold (anterior), and
covered with collagen membrane.
104
xvii
LIST OF SYMBOLS
ºC
Celcius
CaCl2
Calcium Chloride
CO2
Carbondioxide
H&E
Hematoxylin & Eosin
H2O
Water
KCl
Potassium Chloride
KH2PO4
Potassium Dihydrogen Phosphate
kN
Kilonewton
kV
Kilovolt
mm
Milimeter
Mn
Number-average Molecular weight
MPa
Mega-pascal
Mw
Weight-average Molecular weight
NaCl
Sodium Chloride
Na2HPO4
Sodium Hydrogen Phosphate
NaOH
Sodium Hydroxide
O2
Oxygen
P
Probability
rpm
Revolution per minute
Tg
Glass transition temperature
xviii
Tm
Melting point
W0
Initial dry weight
Wdry
Dry weight at time t
μA
Microampere
μm
Micrometer
xix
LIST OF ABBREVIATIONS
3D
Three Dimensional
ABG
Autogenous Bone Graft
BMP
Bone Morphogenetic Protein
BV
Bone Volume
BVF
Bone Volume Fraction
CAD
Computer-aided design
CT
Computed Tomography
DMEM
Dulbecco’s modified Eagle’s medium
ECM
Extracellular matrix
FDA
US Food and Drug Administration
FDM
Fused Deposition Modeling
GA
Gravimetric Analysis
GBR
Guided Bone Regeneration
GPC
Gel Permeation Chromatography
IM
Intramuscular
ISO
International Standards Organization
IV
Intravenous
Lipase PS
Pseudomonas Lipase
Micro-CT
Micro-computed Tomography
PBS
Phosphate Buffered Saline
xx
PCL
Poly(ε-caprolactone)
PDI
Polydispersity Index
PGA
Poly(glycolic acid)
PLA
Poly(lactic acid)
QS
QuickSlice
rhBMP-2
Recombinant human Bone Morphogenetic Protein-2
RP
Rapid Prototyping
SD
Standard Deviation
SEM
Scanning Electron Microscope
SEMC
SingHealth Experimental Medicine Centre
SFF
Solid Free-form fabrication
SGH
Singapore General Hospital
STL
Stereolithography
TCP
Tricalcium Phosphate
THF
Tetrahydrofuran
TV
Tissue Volume
xxi
CHAPTER 1: INTRODUCTION
1.1
BACKGROUND
This section aims to provide background information regarding bone tissue
engineering strategy and the application in implant dentistry, as well as the current
drawback of PCL-TCP scaffolds in dentoalveolar defects treatment that lead the
author to pursue this research. Detailed research objectives and research scope are
discussed in the next and last sections respectively.
1.1.1 Bone tissue engineering
Loss of human tissues or organs is a devastating problem that can affect individuals
of all ages. Bone, a complex living tissue that provides internal support for all higher
vertebrates, is currently heralded as the most commonly replaced organ of the body.
In fact, with over 1.3 million bone repair procedures performed per year in the United
States alone [Chim, 2006], the ability to come up with an innovative and effective
defects treatment to satisfy the major clinical need has indeed been a great
challenge for many researchers.
Historically, autogenous or allogenic bone grafts have been used for treatment in
bone defects. Often, the bone repair mechanism fails as a result of magnitude,
infection or other causes. Autogenous bone grafts are those made of tissue obtained
from the patient who receives the graft, while allogenic bone grafts are those made
of tissue from a human donor, usually post-mortem. However, these techniques
1
have some drawbacks. Harvesting of autogenous bone grafts induces additional
trauma and morbidity, increase operation times, and are often limited in supply. At
the site of bone transplantation, the risks of wound infection, necrosis, and resorption,
representing up to 30% of transplanted material have also been experienced [Betz,
2002; Horch, 2006]. Allogenic bone grafts present risks of possible disease
transmission and problems of religious implications [Hutmacher, 2005; Celil, 2006].
These limitations have then instigated new research aiming to provide a bone graft
engineered in the laboratory and readily available. The ultimate goal of this approach
was the regeneration rather than just the repair of skeletal tissue, and this treatment
strategy was later coined as “bone tissue engineering”.
A key component in tissue engineering for bone regeneration is the scaffold that
serves as a 3D template for initial cell interactions and the formation of boneextracellular matrix to provide structural support to the newly formed tissue. The
porous scaffold provides the necessary support for cells to attach, proliferate, and
maintain their differentiated function. The ability of the scaffold to be metabolized by
the body allows it to be gradually replaced by cells to form functional tissues [Pollok,
1996]. A well-designed scaffold for bone tissue engineering then plays an important
role in facilitating bone healing. To do so effectively, several qualities of an effective
scaffold material must be satisfied. Ideally, a scaffold should possess the following
properties: (1) a 3D structure with an increased porosity and a highly interconnected
pore network for cellular or vascular ingrowth and transport of nutrients and
metabolic waste; (2) biocompatibility and bioresorbability with controlled degradation
and resorption rates to match tissue replacement; (3) suitable surface properties for
cell adhesion, proliferation, and differentiation; and (4) sufficient mechanical
2
properties to match those of the tissues at the site of implantation [Hutmacher, 2001].
The latter is extremely crucial in skeletal tissue such as bone and cartilage where
certain mechanical properties are required. These scaffolds serve as temporary
load-bearing devices that provide adequate strength and help maintain space for
new bone formation to occur [Hutmacher, 2000; Rezwan, 2006; Zhou, 2007].
1.1.2 Application in dentoalveolar defects
In implant dentistry, clinical situations involving major defects or deformities as the
result of trauma or diseases are often faced. The outcome is a compromised and
deficient alveolar ridge, which is often extended and non-contained and frequently
requiring extensive guided bone regeneration (GBR) procedures. In the dentoalveolar skeleton, an inadequate bone volume always creates problems in the
prosthetic and esthetic reconstruction of partially and completely edentulous
situations. In an era where implant borne tooth restorations have became the
standard of care for the replacement of missing teeth, the quantity and quality of the
available bony ridge is critical in determining whether ridge augmentation is required
prior to dental implant placement [Adell, 1990; Jemt, 1993]. This will not only
determine the outcome of a favorable ridge shape and the contour of the overlying
soft tissue, but also the optimal three-dimensional placement of the dental implant.
This is where the role of scaffolds come into the picture as they may eliminate the
need for an extensive bone harvesting procedure from a donor site. However in
facing a complex biological system as the human body, the requirements of scaffold
materials for bone tissue engineering in dentoalveolar application can be extremely
challenging.
3
1.1.3 PCL-TCP scaffolds: Current drawback
The use of synthetic polymers in the field of tissue engineering has been widely
investigated in recent years, with advances in the scaffold technology extending their
usage to clinical applications such as bone regeneration. In particular of such
interest is poly(ε-caprolactone)-tricalcium phosphate (PCL-TCP) composite scaffold,
a synthetic biodegradable polymer frequently investigated for bone tissue
engineering applications. Recent studies on PCL-TCP scaffolds have demonstrated
favourable biocompatibility, bioactivity, and mechanical characteristics [Rai, 2004;
Schantz, 2003; Rai, 2005]. However due to their high molecular weight and
hydrophobicity, they degrade at a slow rate [Jeong, 2003; Ha, 1997]. This is a
disadvantage for bone tissue engineering purposes in dentoalveolar application, as
the new bone replacing the scaffold are inserted with dental implants for prosthetic
rehabilitation [Wu, 2004; Lei, 2006]. Degradation behaviors of porous scaffolds play
an important role in the engineering of new tissue, since the degradation rate is
intrinsically linked to cell vitality, growth, as well as host response. In order for a
biodegradable scaffold to be successful over the long term, the material must have a
rate of degradation that acts in concert with the ingrowth of new bone. Ideally, the
degradation and resorption kinetics of composite scaffolds should be designed such
that the cells are allowed to adhere, proliferate, and secrete their own extracellular
matrix (ECM) as the scaffolds gradually resorbs, creating space for new cell and
tissue growth. The physical support provided by the three-dimensional (3D) scaffold
should also be maintained until the regenerated tissue has sufficient mechanical
integrity to support itself [Putnam, 1996]. Thus, it would be desirable to control the
degradation of the PCL-TCP scaffolds to be in sync with the formation of new bone
4
targeted for dentoalveolar defects treatment (which takes approximately 5-6
months).
1.2 RESEARCH OBJECTIVES
The interest of this study was to investigate the degradation and load-bearing profile
of 3D bioresorable PCL-TCP scaffolds under enzymatic and hydrolytic conditions
and subsequently to evaluate the efficacy of the scaffolds in both small and large
animal models. The purpose was to develop scaffolds with desirable customized
properties and increased degradation rates suitable for application in dentoalveolar
defects treatment.
In this research, specific aims have been identified:
1. To obtain PCL-TCP scaffolds with the desired higher porosity of 85% by treating
them with 3M NaOH or 0.1% lipase-PBS medium under physiological conditions
for up to 108 hours.
2. To compare the degradation profile of treated and untreated PCL-TCP scaffolds
in vitro when immersed in standard culture medium for up to 24 weeks, and in
vivo when implanted in the subcutaneous back of rats for 24 weeks (6 months).
3. To evaluate the rate and extent of bone formation of PCL-TCP scaffolds in vivo
when implanted in a larger, clinically relevant defect model for up to 6 months.
Micropigs were chosen as the animal models.
5
1.3 RESEARCH SCOPE
In order to meet the objectives stated in the previous section, the research scope
(Figure 1.1) has been divided into three parts as follows:
In vitro
Small
animal
model
Large
animal
model
Figure 1.1: Schematic diagram of research scope.
1.3.1
Part 1: Selective modification of PCL-TCP scaffolds targeted for
dentoalveolar reconstruction application (in Chapter 3)
PCL-TCP scaffolds (75% porosity) were treated using 3M NaOH or 0.1% lipase for 0,
12, 24, 36, 48, 60, 72, 84, 96, and 108 hours. Samples were recovered at each time
intervals and properties such as porosity, mechanical strength, surface degradation
and surface characteristics of the scaffolds were evaluated. This part serves as an
initial stage of a larger project, in order to develop a scaffold of a higher porosity that
allows for a more rapid degradation whilst maintaining favourable mechanical
properties. A final porosity of about 85% was targeted for.
1.3.2
Part 2: Optimization of native and customized scaffolds in vitro
and their effects in initial bone healing (in Chapter 4)
In the second part of the study, PCL-TCP scaffolds of a higher porosity (85%) were
tested. The scaffolds were divided into 3 experimental groups: NaOH-treated, lipasetreated and untreated. They were (a) implanted subcutaneously into the back of rats,
6
or (b) immersed in DMEM growth media, for various time periods of up to 6 months.
Analysis similar to those described in part 1 were performed.
Pre-degradation Study
0, 12, 24, 36, 48, 60, 72, 84, 96, 108 hours
In Vitro Degradation Study
In Vivo Degradation Study
0, 6, 12, 18, 24 weeks
Micro-CT
Gravimetric
→ Porosity
→ Structure
→ Weight Loss
0, 12, 24 weeks
Mechanical
Testing
→ Strength
→ Stiffness
SEM
GPC
Histology
→ Surface → Molecular → Inflammation
Morphology Weight → Vascularisation
Figure 1.2: Schematic diagram of part 1 and part 2 study.
1.3.3
Part 3: Evaluation of PCL-TCP scaffolds in a clinically relevant
defect model (in Chapter 5)
In the third and last part of the study, PCL-TCP scaffolds and thin sheets were
implanted in the posterior mandible of micropigs, after two lateral ridge defects were
initially created in each side of the mandible. Following a healing period of 6 months,
the micropigs were sacrificed and the harvested specimens were characterized. The
scope of this thesis ended at the preclinical stage, which was this large animal study.
7
CHAPTER 2: LITERATURE REVIEW
2.1 BONE PHYSIOLOGY
2.1.1 Composition
Bone, a subset of a large and diverse group of tissues collectively referred to as
connective tissue, is the main building block of the human skeletal system. Bone is
made up of organic and inorganic (mineral) matter, cells, and water (Figure 2.1). The
organic matter is concentrated in the bone matrix, which amounts to about 35% of
the dry weight of bone. It consists of 90% collagen, which is thus by far the most
abundant bone protein. Collagen assembles in an organised pattern within the bone
microstructure and modulates bone calcification sites (Figure 2.2). Its complex threedimensional structure, comparable to that of a rope, gives bone tissue its tensile
strength. The remainder of the bone matrix is made up of various noncollagenous
proteins such as cytokines, osteonectin, osteopontin, osteocalcin, growth factors,
bone sialoprotein, hyaluronan, thrombospondin, proteoglycans, phospholipids, and
phosphoproteins [Rho, 1998; Wang, 2001; Glimcher, 1989; Fleisch, 2000]. Together
they play an important role in bone remodelling and in osteogenesis. The mineral
matter of bone consists mainly of mineral salts known as hydroxyapatites, which are
largely made up of calcium phosphates. Tiny crystals of these salts lie within and
between the collagen fibers in the extracellular matrix, producing the compressive
strength and stiffness that is so characteristic of bone [van Gaalen, 2008]. The
8
proper combination of the fibers and salts then allows bones to be both strong and
durable without being brittle [Glimcher, 1998; Baron, 1996].
Figure 2.1 (above): Composition of bone
[Fleisch, 2000].
Figure 2.2 (right): The assembly of
collagen fibrils and fibers and bone
mineral crystals [Rho, 1998].
Bone’s function is both biomechanical and metabolic. Biomechanically, bone acts to:
(1) maintain the shape of the skeleton, (2) protect soft tissues in the cranial, thoracic
and pelvic cavities, (3) transmit the forces of muscular contraction during movement,
and (4) supply a framework for bone marrow. Metabolically, bone (1) serves as a
reservoir for ions, especially calcium ions, and (2) contributes to the regulation of the
extracellular matrix composition [Ferrer, 2007].
Bone is a self-repairing structural material; it is capable of adapting its mass, shape
and properties to the changes in mechanical requirements and endures voluntary
physical activity for life without breaking. This capacity stems from the fact that bone
is in fact alive, and contains cells which work continuously to regenerate and repair it
9
[Bronner, 1999; Ferrer, 2007]. Bone tissue contains five basic types of bone cells:
osteogenic cells, osteoblasts, osteocytes, osteoclasts, and bone-lining cells.
Osteogenic cells respond to traumas, such as fractures, and begin the healing
process immediately by giving rise to bone-forming cells (osteoblasts) and bonedestroying cells (osteoclasts). They can be found in the bone tissue which contacts
the endosteum and the periosteum. Osteoblasts are cell which synthesize and
secrete basic un-mineralized compound to help in the process of bone repair, bone
growth, or bone regrowth. Osteoblast-secreted extracellular matrix may initially be
amorphous and noncrystalline, but it gradually transforms into more crystalline forms
[Boskey, 2003]. Mineralization is a process of bone formation promoted by
osteoblasts and is thought to be initiated by the matrix vesicles that bud from the
plasma membrane of osteoblasts to create an environment for the concentration of
calcium and phosphate, allowing crystallization [Barckhaus, 1978; Celil, 2006].
Where the bone tissue has higher metabolism, the osteoblast cells are more plentiful,
this includes the border of the medullary cavity and under the periosteum. A mature
osteoblast is known as an osteocyte. While osteocytes are technically a different
bone cell altogether, the osteoblast changes into an osteocyte over time.
Osteoblasts have the unique ability to secrete bone tissue and form the tissue
around itself like a protective wall of bone tissue. They are responsible for the
maintenance of healthy bone by secreting enzymes and directing the bone mineral
content. They also control the calcium release from the bone tissue to the blood. The
cells which are responsible for the breakdown of bone tissue, which releases
calcium, are known as osteoclasts. Osteoclasts are vital to the process of bone
growth, remodeling, and healing. The last type of cells are bone-lining cells. They
are made from osteoblasts along the surface of most bones in an adult, and are
10
thought to regulate the movement of calcium and phosphate into and out of the bone
[Chenu, 1998].
2.1.2 Morphology
Macroscopically, bone can be divided into an outer part called cortical or compact
bone, which makes about 80% of the total skeleton, and an inner part named
cancellous, trabecular, or spongy bone. Cortical bone is very dense and contains
only microscopic channels. Forming the outer wall of bones, it bears most of the
supportive and protective function of the skeleton. Cancellous bone, on the other
hand, makes up the remaining 20% of bone mass in the body. It consists of
trabeculae which form an interconnected lattice. Cancellous bone can be found in
vertebrae, fracture joints, ends of long bones and in foetuses. The whole structure,
an outer cortical sheath and an inner three-dimensional trabecular network, allows
optimal mechanical function under customary loads [van Gaalen, 2008; Brickley,
2008; Rho, 1998; Ferrer 2007].
Figure 2.3: Microscopic structure of cortical and cancellous bone [US National
Cancer Institute’s SEER Program, 2009].
11
Microscopically, woven and lamellar bone can be distinguished. Woven bone is the
type formed initially in the embryo and during growth, and is characterized by an
irregular array of loosely packed collagen fibrils. It is then replaced by lamellar bone,
so that it is practically absent from the adult skeleton, except under pathological
conditions of rapid bone formation, such as occur in Paget's disease, fluorosis, or
fracture healing. In contrast, lamellar bone is the form present in the adult, both in
cortical and in cancellous bone. It is made of well-ordered parallel collagen fibers,
organized in a lamellar pattern called osteons or haversian systems. The osteon
consists of a central canal called the osteonic (haversian) canal, which is surrounded
by concentric rings (lamellae) of matrix. Between the rings of matrix, the osteocytes
are located in the lacunae. The osteonic canals contain blood vessels that are
parallel to the long axis of the bone. These blood vessels interconnect, by way of the
canaliculi, with vessels on the surface of the bone [van Gaalen, 2008; Rho, 1998;
Ferrer, 2007].
Figure 2.4: The hierarchical structure of bone from macrostructure to subnanostructure [Rho, 1998].
12
2.2 POLY(ε-CAPROLACTONE)
Poly(ε-caprolactone) (PCL) is a semi-crystalline, biodegradable, and bioresorbable
polymer widely used in tissue engineering recently [Teoh, 2004]. It has a melting
point (Tm) of 60ºC and a low glass transition temperature (Tg) of -60ºC that gives it
rubbery characteristics and be relatively ductile at room temperature [Gan, 1999]. It
is synthesized by ring-opening polymerization of ε-caprolactone monomers. As a
homopolymer belonging to the aliphatic polyester family, the repeating molecular
structure of PCL consists of a 5 non-polar methylene group and a single relatively
polar ester group. The presence of this hydrolytically unstable aliphatic-ester linkage
along the polymer backbone attributed to the biodegradability of the polymer [Perrin,
1997]. When the polymer is implanted in the body, hydrolysis of polymer backbone
reduces the molecular weight of polymer and the degraded products can be
metabolized in the body. The presence of methylene groups on PCL also renders it
non-polar; hence, PCL is hydrophobic and its resistance to a number of medium
such as water, oil and solvent gives it a slow degradation rate.
Figure 2.5: Chemical structure of PCL (as circled) [Wikimedia, 2007].
The biocompatibility of PCL has been confirmed through extensive in vitro and in
vivo studies and approved by US Food and Drug Administration (FDA) for its usage
13
in various medical applications namely sutures and drug delivery systems [Zein,
2002; Pitt, 1981]. The in vitro biocompatibility of PCL scaffolds was investigated by
Hutmacher et al. It was found that both human fibroblasts and osteoblasts colonized
the struts and bars and formed a cell-to-cell and cell-to-extracellular matrix
interconnective network throughout the entire 3D honeycomb-like architecture
[Hutmacher, 2000]. In an in vivo study, intramedullary pins made of PCL were
implanted into a rat humerus osteotomy model. Gross post mortem examination
revealed normal soft tissue and callus formation. Nonunion, lymhadenopathy,
infection and sinus drainage were not seen in any of the PCL specimens. Histology
verified the absence of osteolytic regression around the implant site and foreign
body giant cell reactions. Decalcified humeri demonstrated osteoblastic and
osteoclastic activity [Lowry, 1997]. Hence based on a large number of tests, the
polymer PCL is currently regarded as non-toxic and tissue compatible materials.
Besides being bioresorbable and biocompatible, the polymer can also be processed
with ease into many shapes and forms [Rezwan, 2006]. All the abovementioned
qualities make PCL an ideal candidate for biomedical applications including
controlled drug releases and resorbable matrices as scaffolds for tissue engineering.
2.2.1 Degradation of PCL polymer
Degradation behaviours of scaffolds play an essential role in the engineering of new
tissue, as the rate of degradation is intrinsically linked to many cellular processes
including cell viability, tissue growth, as well as the host response [Lei, 2006]. Once
implanted in the body, a porous scaffold should maintain its mechanical properties
and structural integrity until the ingrowth of new tissue could adapt to the
environment and excrete sufficient amount of extracellular matrix. During this time, it
14
is expected of the scaffold to be largely degraded and absorbed by the body,
enabling the space occupied by porous scaffolds to be replaced by the newly formed
tissue [Alsberg, 2003]. Ideally, the degradation rate should match to or be slightly
slower than the rate of tissue formation [Hedberg, 2005; Rai, 2006].
Different factors may affect the degradation kinetics of a scaffold. This include the
chemical composition and configurational structure, processing history, porosity,
polydispersity, environmental conditions, stress and strain, crystallinity, size, surface
morphology, chain orientation, distribution of chemically reactive compounds within
the matrix, additives, presence of original monomers and overall hydrophilicity.
[Rezwan, 2006]
In general, PCL, like other members of this family of aliphatic polyesters such as
poly(glycolic acid) (PGA) and poly(lactic acid) (PLA), is degraded by non-enzymatic
random hydrolytic scission of esters linkage [Coombes, 2004]. In the case of PCL,
several reports have shown that enzymes might play a role to some extents [Gan,
1999; Jeong, 2004]. Based on the hydrophilicity of monomeric units, PGA degrades
fast, PLA slow and PCL very slow. PLA is much more hydrophobic than PGA due to
the additional methyl group in the structure of PLA. Hence PGA degrades much
quicker in weeks time than PLA, which the latter can remain stable for over 1 year or
more depending on its degree of crystallinity [Mano, 2004]. It has been found that
the degradation of PCL with a high molecular weight (Mn of about 50,000) is
remarkably slow, requiring 3 years for complete removal from the host body [Rezwan,
2006].
15
Fast
PGA
Slow
PLA
PCL
Figure 2.6: The degradation rate of PGA, PLA, and PCL.
One of the main advantages of PCL is the non-toxic nature of the degradation
products, reported mainly to be CO2 and H2O [Pitt, 1981; Woodard, 1985], making it
safe for medical applications.
2.2.1.1 Hydrolysis mechanism
The degradation of poly(α-hydroxy esters) in the aqueous media generally proceeds
via a random, bulk hydrolysis of the ester bonds in the polymer chain. This process
was mainly due to the ends of the carboxylic chains that are produced during the
ester hydrolysis. During degradation, the soluble oligomers which are close to the
surface leach out towards the aqueous medium faster than the chains located inside
the matrix. This gradient of concentration in acidic groups then leads to the formation
of a layer composed of less degraded polymer [Mano, 2004]. Woodard et al. have
also extensively studied the intracellular degradation of PCL polymer [Woodard,
1985]. They reported that polymer encapsulation by collagen filaments containing
only occasional giant cells was observed during the first stage (non-enzymatic bulk
hydrolysis). Significant weight loss of the polymer was not observed during this stage
that lasted about 9 months. After this time period, the molecular weight decreased to
about 5000, followed by the onset of the second stage of degradation. The rate of
chain scission slowed, the hydrolytic process began to produce short chain
oligomers and weight loss was observed. Eventually the polymer was observed to
16
fragment into a powder that was observed inside the phagosomes of macrophages
and giant cells [Lei, 2006]. Inside these cells, the degradation was rapid, requiring
only 13 days for complete absorption in some cases. It was noted that PCL fibers
were susceptible to enzymatic degradation as well.
2.2.1.2 Enzymatic degradation
The studies of both in vivo and in vitro biodegradation of a given polymer are
important for biomedical applications. Special research interests have also been paid
to the enzymatic biodegradation [Gan, 1999]. One of the available model is the
classical Michael-Menten enzymatic model. However, this model is usually valid for
homogeneous systems in which both enzyme and substrate are water-soluble. Most
polymers are water-insoluble, so the enzymatic degradation is more likely a
heterogeneous kinetic process [Timmins, 1997]. It was proposed that those enzymes
soluble in water will first bind to the polymer substrate and then slowly catalyze the
hydrolytic scission of polymer chains [Mukai, 1993]. The surface area of polymer
materials will then have a greater influence on the enzymatic degradation. In the
case of an enzymatic biodegradation between PCL and lipase PS, the process
mainly involved two essential steps: (1) the adsorption of Lipase PS onto the PCL
and; (2) the interaction between Lipase PS and PCL. In principle, the second step is
dependent on the characteristics of Lipase PS and PCL, while the first step is related
to the total concentration of Lipase PS and PCL. It was reported that within the
Lipase PS-PCL system, the degradation rate was mainly dependent on the first step
[Gan, 1999]. In addition, the amount of degradation and the degradation rate of PCL
depended only on the concentration of Lipase PS and independent of the PCL
17
concentration. Several results also showed that enzymatic degradation is a rapid
method to study the degradation of PCL [Gan, 1999].
2.3 TRICALCIUM PHOSPHATE (TCP)
Calcium phosphates, or more accurately calcium orthophosphates, are salts of the
orthophosphoric acid. They were one of the first bioceramics that were specifically
developed for bone repair [Barrère, 2008]. The main driving force behind the
development of calcium orthophosphates as bone substitute materials is their
chemical similarity to the mineral component to mammalian bones and teeth. As a
result, in addition to being non-toxic, they are biocompatible, not recognized as
foreign materials in the body and, most importantly, both exhibit bioactive behavior
and integrate into living tissue by the same processes active in remodeling healthy
bone. They exhibit excellent bone-bonding properties that are related to the surface
reactivity, via dissolution/precipitation mechanisms. This leads to an intimate
physicochemical bond between the implants and bone, termed osteointegration. In
addition, their degradation products are entirely metabolized in a natural way by our
body [den Hollander, 1991; Lai, 2005]. These features are unique and contribute to
their potential in bone tissue engineering.
The first clinical attempt to use calcium phosphate compound was in the successful
repair of bony defect reported by Albee and Morrison in 1920 [LeGeros, 2002]. Since
then, several calcium phosphate biomaterials have been developed and used
successfully in clinics. One of them is tricalcium phosphate (TCP), which belongs to
18
the categories of bioresorbable and bioactive compounds.
Dental applications of tricalcium phosphate ceramics include
the filling of defects due to periodontal loss, as well as
repairing cleft palates. In orthopaedics, tricalcium phosphate
remains an implant material for defect filling where a
resorbable material is indicated [Barrère, 2008]. Tricalcium
phosphate is a white crystalline powder (hexagonal crystals)
Figure 2.7: The
chemical structure
of TCP [Chemical
land, 2007].
with a melting point of 1670ºC. It is insoluble in cold water, but decomposes in hot
water.
2.3.1 Degradation mechanisms of calcium phosphate ceramics
In artificial or natural aqueous environments calcium phosphates can degrade via:
1. Solution-mediated mechanisms leading to physicochemical dissolution of the
ceramic with possibly phase transformation
2. Cell-mediated mechanisms via macrophages and osteoclasts
3. Loss of mechanical integrity as a result of the aforementioned mechanisms.
In biological systems, degradation of calcium phosphates is a combination of nonequilibrium processes that occur simultaneously or in competition with each other
[Barrère, 2008].
2.3.1.1 Physicochemical degradation
The physicochemical degradation, or dissolution, of calcium phosphate ceramics can
be described as a dissolution-reprecipitation cascade which is the result of
exchanges at a solid-liquid interface. In artificial or natural aqueous environments
these ceramics dissolve, this physicochemical process is typical of inorganic
19
substrates, i.e. having dominant ionic features. It is the result of a multi-component
dynamic process that cannot be mimicked in vitro. However, in vitro experiments
simplifying the biological environment have led to conclusions that, in general, fit with
in vivo observations [Barrère, 2008].
From a thermodynamic point of view, most calcium phosphates are sparingly soluble
in water, while some are very insoluble, but all dissolve in acids. Their solubility,
defined as the amount of dissolved solute contained in a saturated solution when
particles of solute are continually passing into solution (dissolving) while other
particles are returning to the solid solute phase (growth) at exactly the same rate
[Wu, 1998], decreases with the increase of pH [de Groot, 1983].
From a surface reactivity viewpoint, physicochemical dissolution can be seen as
ionic transfer from the solid phase to the aqueous liquid via surface hydration of
calcium, phosphate species and possible impurities present in the biomaterial. As a
result of these ionic transfers, phase transformations occur at the ceramic surface. A
phase transformation occurs for calcium phosphate phases which are unstable
under physiological conditions, such as tricalcium phosphate [Barrère, 2008]. The
physicochemical dissolution behavior in vitro and in vivo can be affected by the
crystalline features, the thermodynamical solubility, the structure, and the presence
of additives [Barrère, 2000; Elliot, 1994; LeGeros, 1991; Radin and Ducheyne, 1994].
20
2.3.1.2 Cellular degradation
The typical cellular degradation of calcium phosphates is mediated by osteoclasts.
Osteoclasts are multinucleated bone cells derived from hematopoietic stem cells that
differentiate along the monocyte/macrophage lineage. They are responsible for bone
resorption by acidification of bone mineral leading to its dissolution and by enzymatic
degradation of demineralized extracellular bone matrix. The mature osteoclast is a
functionally polarized cell that attaches via its apical pole to the mineralized bone
matrix by forming a tight ring-like zone of adhesion, the sealing zone. This
attachment involves the specific interaction between the cell membrane and some
bone matrix proteins via integrins (transmembrane adhesion proteins mediating cellsubstratum and cell-cell interactions). In the resorbing compartment, situated under
the cell and delimited by the sealing zone, osteoclasts generate a milieu acidification
resulting in the dissolution of bone mineral. This osteoclastic acidification is mediated
by the action of carbonic anhydrase that produces carbon dioxide, water and protons
that are extruded across the cell membrane into the resorbing compartment
[Kartsogiannis, 2004]. The degree of osteoclastic activity and dissolution process of
a calcium phosphate ceramic depends on the nature of the calcium phosphate. In
the case of the degradation of highly soluble tricalcium phosphate ceramics in vivo,
Zerbo et al. have shown that the degree of physicochemical dissolution was higher
than osteoclastic resorption [Zerbo, 2005].
2.3.1.3 Mechanical degradation
The mechanical degradation of calcium phosphates is the result of both previous
degradation mechanisms. From a mechanical point of view, the calcium phosphate
ceramics are brittle polycrystalline materials for which mechanical properties are
21
governed by the grain size, grain boundaries and porosity [LeGeros, 2002]. Under
humid conditions, e.g. in liquids or physiological fluids, and as a consequence of the
physicochemical dissolution mechanisms, calcium phosphate ceramics undergo a
decrease of mechanical strength [de Groot, 1983; Mirtchi, 1989; Pilliar, 2001,
Raynaud, 1998] and of resistance to fatigue [de Groot, 1983; Raynaud, 1998]. The
mechanical strength of a material can be seen as its resistance to fracture formation
under specific and acute stress at a time point, while failure by fatigue includes an
additional parameter which is the long-duration strength. Normally, decrease of
strength of brittle ceramic materials is caused by slow or subcritical crack growth,
occurring under stress, sometimes assisted by environment factors [de Groot, 1983].
Parameters influencing the mechanical strength degradation in vitro and in vivo are
directly related to the parameters influencing the physicochemical dissolution
[Barrère, 2008].
2.4 PCL-TCP SCAFFOLDS
Many methods have been developed to enhance the properties of biodegradable
polymers in order to improve the rate of degradation and the mechanical properties.
One way to do so is by physical blending [Gan, 1999].
The incorporation of a tricalcium phosphate (TCP) into a poly(ε-caprolactone)
polymer matrix produces a hybrid or composite material. This bioceramic allows to
tailor the desired degradation and resorption kinetics of the polymer matrix
[Hutmacher, 2000]. Our team has hypothesized that the addition of TCP can
22
accelerate the degradation of the PCL polymer. It was shown that the TCP particles
were only physically blended into the polymer and they occupied random spaces in
the polymer. After the scaffold was immersed in solution, the TCP particles being
hydrophilic tend to fall off and interact with the surrounding medium. The falling of
TCP then created voids within the polymer, thus exposing their surfaces to hydrolytic
attack and weakening the overall structure of the PCL [Lei, 2006].
The composite approach can circumvent the limitations of pure ceramics for example
it offsets the problems of brittleness and the difficulty of shaping hard ceramic
materials to fit bone defects [Hu, 2007]. A composite material would also improve
biocompatibility and hard tissue integration in a way that ceramic particles, which are
embedded into the polymer matrix, allow for increased initial quick spread of serum
proteins compared to the more hydrophobic polymer surface [Hutmacher, 2000]. In
addition, the basic resorption products of TCP would buffer the acidic resorption byproducts of the aliphatic polyester like PCL and may thereby help to avoid the
formation of an unfavorable environment for the cells due to a decreased pH
[Hutmacher, 2000]. Finally, the addition of ceramics dispersed throughout the
polymeric matrix results in a superior compressive strength of the composite
compared to non-reinforced materials. All these qualities thus, render a promising
future for the application of PCL-TCP scaffolds in tissue engineering purposes. The
proposed mechanism of degradation manifested by PCL-TCP composite scaffolds
are expected to be a result of combining PCL polymer with TCP ceramic [Lei, 2006].
23
2.4.1 Fabrication method of PCL-TCP scaffolds
Along with the material selection, fabrication methods are also critical for designing
biological scaffolds. Numerous fabrication technologies have been applied to
process biodegradable and bioresorbable materials into scaffolds of high porosity
and surface area. One of them is solid free-form fabrication (SFF) or rapid
prototyping (RP) technique known as fused deposition modeling (FDM), which has
been applied to fabricate complex-shaped tissue engineering constructs. Unlike
conventional machining which involves constant removal of materials, RP is able to
build scaffolds by selectively adding materials, layer-by-layer, as specified by a
computer program. Each layer represents the shape of the cross-section of the
computer-aided design (CAD) model at a specific level. In addition, another potential
benefit offered by RP technology is the ability to create parts with highly reproducible
architecture and compositional variation across the entire matrix due to its computer
controlled fabrication [Hutmacher, 2004].
A traditional FDM machine consists of a head-heated-liquefier attached to a carriage
moving in the horizontal x-y plane. The function of the liquefier is to heat and pump
the filament material through a noozle to fabricate the scaffold following a
programmed path which is based on CAD model and the slice parameters. Figure
2.8 shows a schematic representation of the FDM process. The FDM method
involves the melt extrusion of filament materials through a heated nozzle and
deposition as thin solid layers on a platform. The nozzle is positioned on the surface
of a build platform at the start of fabrication. It is part of the extruder head (FDM
24
head), which also encloses a liquefier to melt the filament material fed through two
counter-rotating rollers.
Figure 2.8: Schematic diagram of the FDM process [Hutmacher, 2001].
Figure 2.9 shows how each layer is made of raster roads deposited in the x and y
directions. A fill gap can be programmed between the roads to provide horizontal
channels. Subsequent layers are deposited with the x-y direction of deposition, the
raster angle, programmed to provide different lay-down patterns. The sequence of
data preparation from step 1 to 4 is: importing of computer-aided design (CAD) data
in STL (stereolithography) format into QS (QuickSlice), slicing of the CAD model into
horizontal layers and conversion into SLC format, creation of a deposition path for
each layer and conversion into SML format for downloading to the FDM machine,
and FDM-fabrication process with a filament modeling material to build the actual
25
physical part in an additive manner layer by layer, respectively [Hutmacher, 2001;
Hutmacher, 2008].
Figure 2.9: Sequence of the data preparation for FDM model fabrication
[Hutmacher, 2001].
26
The method of synthesis for PCL-TCP scaffolds used in this study was described in
detail in recent literatures [Hutmacher, 2001; Rai, 2006]. The first step is filament
fabrication. Pellets of PCL (catalog no. 440744) from Aldrich Chemical Co., Inc.
(Milwaukee, WI) are used. The polymer has an average number-average molecular
weight of 80,000 with a melt index of 1.0 g/10 min. The polymer pellets are kept in a
desiccator prior to usage. PCL pellets are physically blended with TCP granules prior
to filament fabrication. Filament fabrication then is performed with a fiber-spinning
machine (Alex James & Associates Inc., Greensville, SC). The pellets are melted at
190°C in a cylinder with an external heating jacket. After a hold time of 15 min, the
temperature is lowered to 140°C and the polymer melt is extruded through
spinnerets with a die exit diameter of 0.064 in. (1.63 mm). Each batch of PCL pellets
weighs about 30 ± 1 g. The piston speed is set at 10 mm/min. The extrudate is
quenched in chilled water placed 40 mm below the die exit. The combination of
temperature, piston speed and height drop to water quenching settings produces a
filament diameter of 1.70 ± 0.10 mm. The PCL filaments are fabricated to have a
consistent diameter to fit the drive wheels of the FDM system. The filaments are
vacuum-dried and kept in a desiccator prior to usage.
The next step is scaffold design and fabrication [Zein, 2002; Hutmacher, 2000]. The
PCL filaments are fed into a FDM 3D Modeler RP system from Stratasys Inc. (Eden
Prairie, MN). Stratasys Quickslice software is manipulated to produce the desired
dimensions. The head speed, fill gap, and raster angle for every layer are
programmed through this software and saved in the Slice file format. Lay-down
patterns of 0/60/120° are used to give a honeycomb, fully interconnected matrix
architecture and mechanical properties suitable for rapid vascularization and
27
maintenance of the structural integrity of tissue engineered bone grafts in loadbearing applications [Schantz 2002; Schantz, 2003; Hutmacher, 2001]. The use of
the highly reproducible and computer-controlled FDM technique allows the
fabrication of bone grafts that can be designed based on computed tomography (CT)
scans of individual defect sites [Endres, 2003; Hutmacher, 2000].
28
CHAPTER 3: SELECTIVE MODIFICATION OF
PCL-TCP SCAFFOLDS TARGETED FOR
DENTOALVEOLAR RECONSTRUCTION APPLICATION
3.1 INTRODUCTION
This pre-degradation study served as an initial stage to develop a scaffold of a
higher porosity that allows for a more rapid degradation whilst maintaining
favourable mechanical properties. A final porosity of about 85% was targeted for.
The time frame for the action of lipase and NaOH required to achieve this desired
porosity on the PCL-TCP scaffold degradation was also determined.
180 small blocks of PCL-TCP scaffolds with porosity 75%
were equally divided into two groups. Each composite
were placed individually into clean centrifuge tubes and
completely immersed in either 3M NaOH or 0.1% lipasePBS solution (Amano Lipase PS, Sigma-Aldrich, USA).
The tubes were then sealed and put into an incubator of
constant temperature 37ºC. Ten scaffolds were removed
Figure 3.1:
Centrifuge tubes.
at each predetermined time intervals of 12, 24, 36, 48, 60, 72, 84, 96, and 108 hours.
These samples were gently rinsed with PBS before drying for 48 hours in an
incubator that maintained a temperature of 37ºC and controlled relative humidity of
29
30%. The scaffolds were then characterized in terms of their porosity, mechanical
strength, surface degradation and surface morphology.
3.2 MATERIALS AND METHODS
3.2.1 Scaffold design and fabrication
Scaffold specimens were fabricated with PCL-TCP
(80:20%) filaments by using a fused deposition
modeling (FDM) 3D Modeler RP system from
Stratasys Inc (Eden Prairie, MN). Blocks of 50 x 50
Figure 3.2: Illustration of
scaffold with 0/60/120º laydown pattern [Zein, 2002].
x 3 mm were purchased directly from Osteopore
International Pte Ltd, Singapore. Each composite
manifested a lay-down pattern of 0/60/120º with a typical honeycomb array of
interconnected equilateral triangle, and porosity of about 75%. TCP existed as nonuniformly distributed particles on the rods of PCL. The specimens were then cut into
smaller blocks of 5 x 5 x 3mm dimension by using a cutter under aseptic conditions.
3.2.2 Sterilization of scaffolds
The raw pieces of scaffold blocks were pre–treated before used in the degradation
experiment. After being rinsed 3x with phosphate buffered saline (PBS, 137mM NaCl,
2.7mM KCL, 10mM Na2HPO4, 1.8mM KH2PO4, pH7.4), the PCL-TCP scaffolds were
sterilized in 70% ethanol for 24h. This was followed by rinsing twice in PBS with
centrifugation at 1000 rpm for 10 min. The scaffolds were dried in humidified
30
atmosphere at 37oC and 5% CO2 for 1h and soaked for 3h
in PBS before loading. This latter process of pre–wetting the
porous scaffolds was to ensure that PBS solution had
permeated through all the pores of the scaffolds and to let
the scaffolds became more hydrophillic.
Figure 3.3:
5x5x3mm PCL-TCP
scaffold.
3.2.3 Scaffold characterizations
Upon retrieval from the respective mediums, the scaffold specimens were subjected
to characterizations for analysis of their porosity, structure, percentage weight loss,
compressive mechanical properties, surface morphology, and molecular weight.
Several methods employed to characterize the PCL-TCP scaffold samples were:
3.2.3.1 Micro-computed tomography analysis (n = 3)
Microcomputed tomography (SkyScan 1076 In Vivo X-Ray Microtomograph,
Belgium) was performed to monitor the surface and porosity changes of the
scaffolds before and after implantation. The parameters were set at 104 kV energy,
98 μA intensity, and 35 μm pixel size. The specimens were scanned through 180o,
and the image data from the scanned planes were subsequently thresholded and
reconstructed to create 3-D images for quantitative histomorphometric analyses.
3.2.3.2 Gravimetric analysis (n = 3)
Scaffolds’ weight losses during degradation were measured by the changes in dry
weight after incubation or implantation for specifed time periods. Weights were
normalized with respect to zero-time measurements. For such tests, three scaffolds
31
were removed, gently rinsed in PBS and dried in vacuum oven for 48 hours. Values
obtained for triplicate samples were averaged. Percent weight loss was computed
according to the following equations (3.1):
W0 − Wdry
W0
Weight loss (%) =
Where
×100
(3.1)
W0 is the initial dry weight as measured at time 0, and Wdry is the dry weight
at time t.
3.2.3.3 Compressive mechanical testing (n = 5)
Evaluation of mechanical properties of the specimens were performed with an
Instron 3345 uniaxial testing system (table-top tester 3345; Instron, Canton, MA),
with a 1kN load cell. Five specimens from each group were to be tested for each
concentration study at each time point. Each specimen was to be placed between 2
flat plates for compressional testing. The scaffolds were compressed at a crosshead
speed of 1mm/min up to 80% of the scaffolds total thickness. A stress-strain curve
was plotted using the experimental data obtained to determine the compressive
modulus and compressive strength.
3.2.3.4 Electron microscopy preparation and analysis (n = 2)
Surface morphological changes of the PCL-TCP scaffolds were characterized using
the JEOL JSM – 5600LV SEM operating at an accelerating voltage of 15kV under
high vacuum mode. Prior to the usage of the SEM, the PCL-TCP scaffolds have to
be pre-treated first. The scaffolds samples were immersed in 2.5% gluteraldehyde
(Sigma, Germany) at 4°C overnight. They were then rinsed in PBS for 10 minutes
32
and dehydrated in a graded ethanol series of 25% (5 min), 50% (10min), 75% (10
min), 95% (10 min) and 100% (10 min, 3 times). Following that, the scaffolds were
placed in an oven dessicator overnight to dry. As the scaffolds were viewed under
high vacuum condition in the SEM to attain high magnification, they have to be gold
coated first which is a destructive method. The gold splattering of the fracture
surfaces was conducted with JEOL JFC – 1200 fine coater in a high vacuum
chamber for 40 seconds at a current of 10mA.
3.2.3.5 Molecular weight testing (n = 3)
Each of the scaffold samples was selected and partially dissolved into
Tetrahydrofuran (THF). The THF solute was then analyzed for molecular weight
distribution by Gel Permeation Chromatography (GPC) using Waters 600_717_2414
System. The measurements were carried out at an elution rate of 1 ml/min using
THF as the mobile phase solvent through Styragel column refractors. A total of four
7.8 x 300mm column were used: Styragel HR 0.5, Styragel HR 1, Styragel HR 3 and
Styragel HR 4. Polystyrene Standards from Waters were used to obtain the
calibration curve.
3.2.4 Statistical analysis
All quantitative data (the mechanical strength and molecular weight loss) were
expressed as mean values ± the standard deviation (SD) of the mean. Data
analyses and comparisons were performed using Student’s paired t-test. A value of
p< 0.05 was considered to be statistically significant.
33
3.3 RESULTS
3.3.1 Porosity measurements and 3D model analysis
Results from the micro CT analysis demonstrated an increase in porosity values in
both treatment groups. Lipase-treated scaffolds showed a faster rate of degradation
than NaOH-treated ones. Pre-degraded scaffolds of the desired 85% porosity were
obtained when they were immersed in 3M NaOH for 96 hours and in 0.1% lipase for
12 hours (Figure 3.4). Refer to Appendix A1 – Table A.1 for the complete data.
140
3M NaOH
0.1% lipase
120
Porosity (%)
100
81.78 84.44
80
75.61 75.61
80.92
81.08
80.85
81.14
98.23
98.12
97.69
96.47
95.45
93.4
80.01
82.01
98.51
98.32
83.41
87.44
60
40
20
0
0
12
24
36
48
60
72
84
96
108
Degradation Time (Hours)
Figure 3.4: Porosity measurements of NaOH-treated and lipase-treated PCL-TCP
scaffolds over time.
The 3 dimensional images reconstructed from these two treatment groups similarly
showed a more significant degradation action by the lipase than the NaOH treatment.
It has been suggested that the chemical action of sodium hydroxide mainly causes
surface erosion, whereas the enzymatic action involves scission of the polymer
chain backbone and hence a more significant degradation activity [Wan, 2005].
34
Whilst attaining the desired 85% porosity, both the treated PCL-TCP scaffolds
maintained favourable 3D morphology with the usual interconnected pore network
(Figure 3.6 and 3.7). An untreated PCL-TCP scaffold of around 75% porosity was
shown in Figure 3.5.
Figure 3.5: 3D model of original scaffold (of 75% porosity) at 0 hour:
(L) top view, (R) tilted view.
Figure 3.6: 3D model of scaffolds after 96 hours immersion in 3M NaOH:
(L) top view, and (R) tilted view.
35
Figure 3.7: 3D model of scaffolds after 12 hours immersion in 0.1% lipase:
(L) top view, and (R) tilted view.
3.3.2 Weight loss analysis
There is an increasing trend for the weight-loss percentage as shown in Figure 3.8.
100
82.3
Weight loss (%)
80
92.77
91.28
90.36
90
92.8
90.61
85.66
73.71
70
60
52.8
50
39.48
45.13
40
30
15.76
20
10
22.33
20.32
21.94
24.59
26.18
17.06
3M NaOH
0.1% lipase
0
0
0
0
10
20
30
40
50
60
70
80
90
100
110
120
Degradation Time (Hours)
Figure 3.8: Comparison of weight loss between NaOH-treated and
lipase-treated PCL-TCP scaffolds.
The values for NaOH-treated at 96 hours and lipase-treated at 12 hours scaffolds
were found to increase logarithmically to 39.48±2.25 % and linearly to 45.13±7.41 %
36
respectively. These values demonstrated a significant loss of mass as the scaffolds
were degraded over the course of time. Refer to Appendix A4 – Table A.4 for the
complete data.
3.3.3 Compressive mechanical properties
The mechanical properties of the treated PCL-TCP scaffolds over time are shown in
Figures 3.9 and 3.10. For the NaOH-treated scaffolds at 96 hours, both the
compressive strength and compressive modulus have decreased significantly by
41.6% (4.6±0.8 MPa) and 51.3% (10.8±2.75) MPa respectively. Likewise for the
lipase-treated scaffolds, at 12 hours, both the compressive strength and
compressive modulus have reduced significantly by 44.7% (4.36±1.64 MPa) and
46.4% (11.87±2.58 MPa) respectively. As expected, the reduction in mechanical
properties was accompanied following in increases in the scaffold porosity. Refer to
Appendix A2 – Table A.2 and Table A.3 for the complete data.
Compressive Strength (MPa)
10
8
3M NaOH
0.1% lipase
7.88
7.01
6.85
6.94
7.25
6.90
6.66
7.21
6
4.60
4.36
4
3.19
2.35
2
2.22
1.57
0
0
0
0
10
20
30
40
50
60
70
0
80
0
90
100
0
110
-2
Degradation Time (Hours)
Figure 3.9: Compressive strength of PCL-TCP scaffolds when treated with
NaOH and lipase at pre-determined time intervals.
37
120
30
Compressive Modulus (MPa)
3M NaOH
25
0.1% lipase
22.15
20
19.47
19.00
20.17
18.07
17.32
17.48
15.38
15
11.87
10.78
10
7.98
7.15
7.10
5.65
5
0
0
0
10
20
30
40
50
60
0
70
0
80
0
90
100
0
110
120
-5
Degradation Time (Hours)
Figure 3.10: Compressive modulus of PCL-TCP scaffolds when treated with
NaOH and lipase at pre-determined time intervals.
3.3.4 Surface morphology analysis
Electron microscopy analysis was conducted to investigate the surface morphology
of the PCL-TCP scaffolds. Figure 3.11 shows the SEM micrograph of the original
untreated scaffold (that is, scaffold at 0 hour). In general, the surface was smooth
with numerous small TCP particles protruding out.
Figure 3.11: Electron micrographs of original scaffold (of porosity 75%) at 0 hour:
(L) overall view, and (R) higher-magnification view.
38
Figures 3.12 and 3.13 display SEM micrographs taken at 96 hours immersion in
NaOH and at 12 hours in lipase respectively. Viewed under higher magnification, it
could be observed that the smooth surface of the scaffolds have roughened and
displayed small cracks in both cases. Numerous tiny and shallow pores have
appeared as well. However, the degree of the degradation seemed to be slightly
higher in lipase-treated scaffolds as seen from the more significant reduction in the
diameter of the rods. Refer to Appendix A3 for the complete images.
Figure 3.12: Electron micrographs of scaffold after 96 hours immersion in
3M NaOH: (L) overall view, and (R) higher-magnification view.
Figure 3.13: Electron micrographs of scaffold after 12 hours immersion in
0.1% lipase: (L) overall view, and (R) higher-magnification view.
39
3.3.5 Molecular weight analysis
The values of Mw, Mn, and PDI are shown in Table 3.1. Overall, the NaOH-treated
PCL–TCP scaffolds exhibited no appreciable decrease in molecular weight
throughout the degradation period of up to 108 hours. At 96 hours immersion, the
value of Mw, Mn, and PDI is 58940 Daltons, 41279 Daltons, and 1.43 respectively.
On the other hand, PCL-TCP scaffolds immersed in lipase maintained their
molecular weights for up to 72 hours and showed a sudden drop when it reached 96
hours with nearly 50% reduction in the values. At 12 hours, the lipase-treated PCLTCP scaffolds declined in its Mw, Mn, and PDI to 54491 Daltons, 39763 Daltons, and
1.37 respectively.
Table 3.1: Mw, Mn, and PDI of NaOH-treated and lipase-treated PCL-TCP scaffolds.
Hour 0
Hour 12
NaOH-treated PCL-TCP Scaffolds
54548
54457
Mw (Dalton)
Hour 24
Hour 48
Hour 72
Hour 96
56575
55330
59849
58940
Mn (Dalton)
40415
38010
42283
40090
43041
41279
PDI
1.35
1.43
1.34
1.38
1.39
1.43
54180
50364
54526
29337
Lipase-treated PCL-TCP Scaffolds
53548
54491
Mw (Dalton)
Mn (Dalton)
40415
39763
39842
34859
38529
24667
PDI
1.35
1.37
1.36
1.44
1.42
1.19
3.4 DISCUSSION
In this part of the study, sodium hydroxide solution and lipase enzyme were used to
treat the original 75% porosity scaffolds with the objective of obtaining customized
40
scaffolds of higher porosity value. The desired 85% porosity was targeted for in this
study. The concentration of sodium hydroxide and lipase was selected as 3M and
0.1% respectively, which was considered low-medium in terms of the strength. The
scaffolds were allowed to degrade but in a slower and controlled manner. This was
to ensure that the treated scaffolds still retained most of their essential properties
such as strength, stiffness, pores interconnectivity, molecular weight, and others,
prior immersion in the DMEM growth media or implantation in vivo in rats.
Sodium hydroxide solution is a strong alkaline reagent. Previous studies have shown
that it has the capability to enhance the hydrophilicity and accelerate the degradation
of PCL polymer under laboratory condition [Park, 2005]. However not many have
specifically focused on PCL-TCP scaffolds. The non-enzymatic breakdown of the
scaffolds due to sodium hydroxide involves mainly surface erosion. Reports have
also demonstrated that PCL, as opposed to its related aliphatic polymers, such as
PGA and PLA, have the capability to undergo enzymatic degradation [Gan, 1999].
Among the various types of enzymes, lipase is found to be the most widely
investigated, and that lipase PS is considered the best candidate [Gan, 1997; He,
2003]. And for this reason, Lipase PS was chosen in this experiment. In the case like
this study, whereby the enzyme was water-soluble but the PCL-TCP composite as
substrate was water-insoluble, the degradation mechanism will most likely to follow
the heterogenous kinetics model. The model proposed that the enzyme being
soluble in water will first bind to the polymer substrate, and then catalyze the
hydrolytic scission of polymer chains [Mukai, 1993]. Due to this catalytic property,
thus it is hypothesized that once occurred, enzymatic degradation is a rapid process.
One report showed that the biodegradation of a macroscopic PCL film with a
41
dimension of 10 x 10 x 0.1 mm3 could be completed within 4 days in a buffer solution
containing 5.0 x 10-4 g/ml Lipase PS [Gan, 1997]. As expected, it has been
demonstrated in this experiment that lipase-treated scaffolds are the most rapidly
degraded compared to NaOH-treated scaffolds within the same degradation time
frame. After 7-8 days of immersion in lipase solution, a total breakdown of the PCLTCP scaffolds was actually observed. Findings from the pre-degradation section of
the study also demonstrated that scaffolds with the desired 85% porosity were
obtained after being submerged in 3M NaOH for 96 hours (83.41±3.04 %) and in
0.1% lipase for 12 hours (84.44±2.80 %).
Porosity is one of the key parameters to be considered when designing a scaffold as
it would determine degradation rate, cell seeding efficiency, diffusion and the
mechanical strength of the scaffold [Rai, 2006]. A high porosity and high surface
area to volume ratio are required for uniform cell delivery, cellular attachment and
neo-tissue in growth. However, it was reported that scaffolds with a too high porosity
would possess very low mechanical properties. Based on this, compressive
mechanical testings were conducted and the results showed that the decrease in the
strength and stiffness recorded for the NaOH-treated and lipase-treated scaffolds at
85% porosity were still within the acceptable range to support load bearing tissues
such as bone and cartilage. For the NaOH-treated scaffolds at 96 hours, the
compressive strength and stiffness decreased by 41.6% (4.6±0.8 MPa) and 51.3%
(10.8±2.75) MPa respectively. In the lipase-treated group, both the compressive
strength and compressive modulus of the scaffolds were reduced by 44.7%
(4.36±1.64 MPa) and 46.4% (11.87±2.58 MPa) respectively after 12 hours.
42
Meanwhile, the treated scaffolds still retained their usual framework with proper
interconnected pore networks as desired. This is advantageous as a high pore
interconnectivity would encourage the development of a vascular network to allow
for the infiltration of nutrients, cells and growth factors within the system in vivo. As
for the molecular weights, the results for NaOH-treated scaffolds were relatively
constant and exhibited no particular trend throughout the degradation period of up to
108 hours. This finding then further confirmed that the chemical action of sodium
hydroxide mainly involves surface erosion. In contrast, lipase enzyme causes
scission of the polymer chain backbone and as observed, the molecular weights of
the lipase-treated scaffolds showed a decreasing trend and demonstrated a fall
when it reached a certain time point. In addition, scanning electron microscopy were
conducted and the results displayed that there is an increase in the surface
roughness of the treated scaffolds as clearly shown from the increase in indentations
on the surfaces of the scaffold rods. This is an encouraging morphology as bone
forming cells, like osteoblasts, favour rough surfaces which is optimal for promoting
better cell attachment, proliferation, and differentiation [Park, 2005].
3.5 CONCLUSION
PCL-TCP scaffolds have shown to undergo both chemical and enzymatic
degradation in this part of the study. The objective of the experiment was achieved
with scaffolds of approximately 85% porosity obtained after 96 hours of treatment in
3M NaOH and 12 hours in 0.1% lipase. These pre-treated scaffolds demonstrated
acceptable mechanical strength, structure, and surface morphology.
43
CHAPTER 4: OPTIMIZATION OF
NATIVE AND CUSTOMIZED SCAFFOLDS IN VITRO
AND THEIR EFFECTS IN INITIAL BONE HEALING
4.1 INTRODUCTION
In Chapter 3, the PCL-TCP scaffolds have been subjected to both chemical and
enzymatic degradation. Scaffolds with desirable higher porosity with acceptable
mechanical strength, structure, and surface morphology were successfully obtained.
However the degradation behavior of these customized scaffolds must be further
tested. It was then the interest of this present study to investigate the degradation
kinetics of treated and untreated PCL-TCP scaffolds when immersed in standard
culture medium and when implanted subcutaneously to the back of rats. Both these
in vitro and in vivo studies were described under this chapter for better comparison
purpose.
4.1.1 In vitro degradation study
The in vitro degradation study of the PCL–TCP scaffolds was done closely in
accordance to the ISO 10993 standards. The medium used was Dulbecco’s Modified
Eagle Media (DMEM D1152, Sigma, USA. Refer to Appendix B.5 – Figure B.19)
growth medium. In this study, three group of scaffolds namely native, sodium
hydroxide-treated, and lipase-treated of 85% porosity were fully submerged
separately in 1ml DMEM solution. The samples were placed individually in small
44
cryogenic tubes and they were stored in an oven desiccator at a temperature of 37ºC
inside the cleanroom for the duration of 6, 12, 18, and 24 weeks. 10 samples were
taken and characterized at each time points. In addition, the DMEM solution was
replaced with a fresh batch twice a week (every Monday and Thursday) to avoid any
built-up excess or bacterial contamination.
Figure 4.1: Native (left), NaOH-treated (middle), and lipase-treated (right) scaffolds.
4.1.2 In vivo degradation study
In order to acquire better undertanding of the degradation behaviour of the scaffolds
in the living system and prior conducting larger animal studies, an in vivo study on
smaller animal model is a necessity. In this project, rat models were chosen. A total
of 24 untreated PCL-TCP (native) and 48 treated (24 lipase-treated and 24 NaOHtreated) scaffolds with porosity of 85% were implanted in the subcutaneous backs of
rats for the duration of 3 and 6 months.
45
Figure 4.2: Rat at the start of experiment (left) and at the end after 6 months (right).
4.2 MATERIALS AND METHODS
4.2.1 Scaffold design and fabrication
Scaffold specimens were fabricated with PCL-TCP
(80:20%) filaments by using a fused deposition
modeling (FDM) 3D Modeler RP system from
Stratasys Inc (Eden Prairie, MN). Blocks of 50 x 50
x 3 mm were purchased directly from Osteopore
International Pte Ltd, Singapore. Each composite
Figure 4.3: 50x50x3mm
PCL-TCP scaffold.
manifested a lay-down pattern of 0/60/120º with a
typical
honeycomb
array
of
interconnected
equilateral triangle, and porosity of about 75%. TCP existed as non-uniformly
distributed particles on the rods of PCL. The specimens were then cut into smaller
blocks of 5 x 5 x 3mm dimension by using a cutter under aseptic conditions.
46
4.2.2 Sterilization of scaffolds
The raw pieces of scaffold blocks were pre–treated before used in the degradation
experiment. After being rinsed 3x with phosphate buffered
saline (PBS, 137mM NaCl, 2.7mM KCL, 10mM Na2HPO4,
1.8mM KH2PO4, pH7.4), the PCL-TCP scaffolds were
sterilized in 70% ethanol for 24h. This was followed by
rinsing twice in PBS with centrifugation at 1000 rpm for 10
min. The scaffolds were dried in humidified atmosphere at
37oC and 5% CO2 for 1h and soaked for 3h in PBS before
Figure 4.4:
5x5x3mm PCL-TCP
scaffold.
loading. This latter process of pre–wetting the porous scaffolds was to ensure that
PBS solution had permeated through all the pores of the scaffolds and to let the
scaffolds became more hydrophillic.
4.2.3 Animal husbandry
Twelve, 7-8 week old male Wistar rats (260.83 ±
39.43 g) were employed for this in vivo study. The
animals were housed in the animal holding facility at
the
SingHealth
Experimental
Medicine
Centre
(SEMC), Singapore General Hospital, for the entire
duration of the experiment. Housing and feeding were
according to standard animal-care protocols. The
study has been approved by the Animal Welfare
Figure 4.5: Rat cages.
Committee of the Singapore General Hospital and
47
has been licensed by the National Institute of Health’s Guide for Care and Use of
Laboratory Animals.
4.2.4 Scaffold implantation
The rats were randomly divided into two groups of 6 rats each for the implantation of
scaffolds. They were operated on under general
anaesthesia which consists of an intraperitoneal
injection of ketamine and xylazine mixture (75
mg/kg + 10 mg/kg). Under anesthesia, the neck
and back region of the rat was shaved and
Figure 4.6: Rat shaved and
scrubbed with iodine.
scrubbed with iodine, followed by disinfection with 70 % ethyl alcohol (shown in
Figure 4.6). A midline incision was made in the skin of the back with scissors
paralleling the line connecting the apical most point of the shoulder blades and to the
point most coronal to the line. A cylinder shaped dissection was then made subdermal with a pair of needle holders and the cavities were exposed for the
placement of the samples in the right and left flanks of the dorsal lumbar regions.
For each rat, 6 samples of either treated or
untreated PCL-TCP scaffolds (2 from each
group) with dimensions of 5 x 5 x 3 mm were
placed into the empty spaces in the cavity
Figure 4.7: Scaffolds’ positioning.
and the skin was closed with sutures. The
rats were given buprenorphine (100-500 ug/kg) and cephalexin (15-20 mg/kg)
subcutaneously for 3 and 5 days respectively.
48
Figure 4.8: Incision made (left), implanted scaffold (left, inset), and scaffold
positions (right).
A group of 6 rats were euthanised by an overdose of carbon dioxide inhalation at 3
and 6 months respectively. The tissues surrounding the implanted scaffolds were
removed and fixed in either 10 % neutral buffered formalin or saline solution. After
sufficient fixing, the samples were processed accordingly for their degradation and
physical properties.
Figure 4.9: Sacrifice of rats.
Figure 4.10: Removal of scaffolds.
4.2.5 Scaffold characterizations
Upon retrieval from the respective mediums, the scaffold specimens were subjected
to characterizations for analysis of their porosity, structure, percentage weight loss,
49
compressive mechanical properties, surface morphology, and molecular weight.
Several methods employed to characterize the PCL-TCP scaffold samples were:
4.2.5.1 Micro-computed tomography analysis (n = 3)
Please refer to section 3.2.3.1 for the details.
4.2.5.2 Gravimetric analysis (n = 3)
Please refer to section 3.2.3.2 for the details.
4.2.5.3 Compressive mechanical testing (n = 5)
Please refer to section 3.2.3.3 for the details.
4.2.5.4 Electron microscopy preparation and analysis (n = 2)
Please refer to section 3.2.3.4 for the details.
4.2.5.5 Molecular weight testing (n = 3)
Please refer to section 3.2.3.5 for the details.
4.2.5.5 Histological preparation and analysis (n = 2)
The two specimens removed after 3 months and stored in neutral buffered formalin
were dehydrated in ascending series of alcohol rinses and embedded using a
process that produced ground sections with the glycol metacrylate resin.
Once
polymerized, the block was trimmed to remove excess plastic with an industrial
vertical band saw and cut along its long axis with a diamond band saw (EXAKT
standard saw). Ground polished sections of 10 µm thickness were made using the
50
EXAKT micro grinder system from EXAKT Technologies, Inc., Oklahoma City, OK.
Two slides were created for each scaffold. The slides were stained with Hematoxylin
& Eosin (H & E).
4.2.6 Statistical analysis
All quantitative data (the mechanical strength and molecular weight loss) were
expressed as mean values ± the standard deviation (SD) of the mean. Data
analyses and comparisons were performed using Student’s paired t-test. A value of
p< 0.05 was considered to be statistically significant.
4.3 RESULTS - IN VITRO DEGRADATION STUDY
4.3.1 Porosity measurements and 3D model analysis
Upon retrieval from the growth culture medium at each respective time points, the
porosity of the PCL-TCP scaffolds were measured. The graph as shown in Figure
4.11 revealed an increasing trend in the porosity of all three scaffold groups. The
increasing rate in the porosity of the lipase-treated scaffolds has been noted to be
the highest, followed by NaOH-treated scaffolds and native scaffolds respectively.
After 6, 12, and 18 weeks of immersion, the porosity change of the lipase-treated
scaffolds is 9.82%, 13.49%, and 10.37% respectively. For NaOH-treated scaffolds,
the porosity change after 6, 12, and 18 weeks of immersion is 4.02%, 2.77%, 4.62%
respectively. The porosity change of the native scaffolds after 6, 12, and 18 weeks of
51
immersion was the least among the three, which is 1.03%, 4.18%, and 3.27%
respectively. Refer to Appendix B1 – Table B.1 for the complete data.
140
Native
NaOH-treated
120
Lipase-treated
Porosity (%)
100
92.73
85.24 83.41 84.44
86.12 86.76
95.83
88.80
85.72
88.03 87.26
93.20
80
60
40
20
0
0
6
12
18
Degradation Time (Weeks)
Figure 4.11: Porosity measurements of native, NaOH-treated, and lipase-treated
PCL-TCP scaffolds after immersion in DMEM for 6, 12, and 18 weeks.
Figure 4.12: 3D model of native scaffold (of 85% porosity) at week 0:
(L) top view, and (R) tilted view.
52
Figure 4.13: 3D model of native scaffold after 6 weeks immersion in DMEM:
(L) top view, and (R) tilted view.
Figure 4.14: 3D model of NaOH-treated scaffold after 6 weeks immersion in DMEM:
(L) top view, and (R) tilted view.
Figure 4.15: 3D model of lipase-treated scaffold after 6 weeks immersion in DMEM:
(L) top view, and (R) tilted view.
53
Figure 4.16: 3D model of native scaffold after 12 weeks immersion in DMEM:
(L) top view, and (R) tilted view.
Figure 4.17: 3D model of NaOH-treated scaffold after 12 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
Figure 4.18: 3D model of lipase-treated scaffold after 12 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
54
Figure 4.19: 3D model of native scaffold after 18 weeks immersion in DMEM:
(L) top view, and (R) tilted view.
Figure 4.20: 3D model of NaOH-treated scaffold after 18 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
Figure 4.21: 3D model of lipase-treated scaffold after 18 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
55
Figure 4.22: 3D model of native scaffold after 24 weeks immersion in DMEM:
(L) top view, and (R) tilted view.
Figure 4.23: 3D model of NaOH-treated scaffold after 24 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
Figure 4.24: 3D model of lipase-treated scaffold after 24 weeks immersion in
DMEM: (L) top view, and (R) tilted view.
56
Figure 4.12 displayed the 3D construct of native PCL-TCP scaffold which was
manufactured of 85% porosity at week 0 (before immersion in DMEM). For the 85%
porosity NaOH-treated and lipase-treated scaffolds used at week 0 (prior immersion
in DMEM), please refer to Figure 3.6 and 3.7 respectively.
Figure 4.13 to 4.24 illustrate the 3D models of ±85% porosity native, NaOH-treated,
and lipase-treated PCL-TCP scaffolds after being submerged for 6, 12, 18, and 24
weeks in DMEM. There were obvious changes to the 3D morphology of the scaffolds.
The rods of the scaffolds of all three groups have became thinner and looked fragile
over time. In the case of native scaffolds, up to 18 weeks of immersion, the finding
shows that they still retained the original framework. The same goes to NaOHtreated scaffolds, although slight disconnectivity of the rods started to be seen only
later at week 18. In contrast to these two groups, lipase-treated scaffolds seemed to
have undergone significant structural distortion. Loss of original shape and
honeycomb-like pattern, as well as disconnectivity of the rods, were observed as
early as 6 weeks of immersion. By week 24, all scaffolds were significantly degraded.
4.3.2 Weight loss analysis
Figure 4.25 show the percentage weight loss of scaffolds in DMEM growth medium
over various time intervals. Overall there was an increase in the percentage of
weight loss for the native, NaOH-treated, and lipase-treated scaffolds. Again, the
highest being lipase-treated scaffolds, which have undergone significant degradation
during the immersion. The huge loss of mass was mainly due to the reduction in
volume caused by the disconnection of the rods, which correlate to the results
57
obtained from the Micro-CT analysis as discussed earlier. Apart from lipase-treated
scaffolds, the other two scaffold groups have demonstrated an increase in the weight
loss as well. Although the amount is much lower compared to lipase-treated
scaffolds, the student’s t-test show that the rise in percentage weight loss for native
and NaOH-treated scaffolds was statistically significant. Refer to Appendix B4 –
Table B.7 for the complete data.
100
Native
90
NaOH-treated
76.77
80
Lipase-treated
70.77
66.11
Weight loss (%)
70
60
50
40
30
20
22.38
14.29
13.07
6.23
10
10.89
6.56
0
6
12
18
-10
Degradation Time (Weeks)
Figure 4.25: Weight loss of PCL-TCP Scaffolds In vitro.
4.3.3 Compressive mechanical properties
Compressive properties of degrading porous PCL-TCP scaffolds from three different
groups as a function of degradation time are illustrated in Figure 4.26 and 4.27.
58
Figure 4.26: Relative compressive strength of PCL-TCP Scaffolds In vitro.
Figure 4.27: Relative compressive modulus of PCL-TCP Scaffolds In vitro.
Referring to the graphs, the mechanical properties were shown to decrease over the
course of the study up to 18 weeks. But there are variations in the decreasing rate in
the properties across the three groups. While compressive strength and modulus of
59
the native scaffold decline in a gradual manner (close to exponentially), those of the
NaOH-treated and lipase-treated scaffolds behaved much different within the same
observation period. The latter two demonstrated a drastic fall between week 0 and
week 6, with reduction in compressive strength and modulus by 57.17% and 45.08%
respectively for NaOH-treated scaffolds, and by 72.94% and 84.08% respectively for
lipase-treated scaffolds. This sharp decline have then supported the finding in the
previous section and could be explained that it is likely due to the extreme weight
loss of the scaffolds during the degradation process. However interestingly between
week 6 up to week 18, the compressive strength and modulus of both NaOH-treated
and lipase-treated scaffolds seemed to stall around their value at week 6, with only
slight fluctuations observed. By the end of the study at week 18, the compressive
strength of the native, NaOH-treated, and lipase-treated scaffolds have diminished
by 41.89%, 55%, and 70.64% respectively. As for the stiffness, the values for native,
NaOH-treated, and lipase-treated scaffolds have reduced by 48.45%, 37.94%, and
88.04% respectively. Refer to Appendix B2 – Table B.3 and Table B.4 for the
complete data.
4.3.4 Surface morphology analysis
The PCL-TCP scaffolds immersed in culture medium demonstrated significant
changes to their surface morphology over time as depicted in Figure 4.28 to 4.31. At
week 0, all the three scaffold groups had a consistent interconnected architecture.
For native scaffolds (manufactured 85% porosity), the TCP was evident as particles
protruding out of the rods’ surfaces, contributing to the rough texture (Figure 4.32).
Whereas for the other two groups, their surfaces have some cracks and microporous
60
as resulted from being treated with NaOH and lipase prior immersion in the growth
media (refer to Figure 3.12 and 3.13 respectively).
After soaking in growth medium for 6 weeks, thinning of the rods were observed
across the three groups, and more significantly in NaOH-treated and lipase-treated
scaffolds. In fact for the case of lipase-treated scaffolds, which has undergone the
highest degradation rate, disconnectivity of the rods can actually be seen as early as
6 weeks (Figure 4.28 (e) and (f)). Viewed under higher magnification, it is observed
that the surface of the rods of native scaffolds have roughened to a small extent, but
no significant micropores can be seen yet. The surface of the rods in NaOH-treated
and lipase-treated scaffolds still remained microporous however the size of the pores
have slightly increased. This implies that to a certain extent, degradation of the
scaffolds have occured. Refer to Appendix B3 for the complete images.
At week 12, the surfaces of the scaffold from all groups seemed to be increasingly
eroded, enhancing the surface contact area with the culture medium. The
degradation of the scaffolds continued to accelerate and by the end of week 24,
there was an obvious shrinkage in the size of the three scaffold groups indicating
significant loss of material. Surface line cracks with considerable gaps, and slight
distortion has started to appear on the rods of native scaffolds. In addition, the
lipase-treated scaffolds have severely lost their architexture, although those of the
other two groups still retained their framework.
61
(a)
(b)
(c)
(d)
(e)
(f)
Figure 4.28: Electron micrographs taken after 6 weeks immersion in DMEM for:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated scaffolds.
(L) overall view, and (R) higher-magnification view.
62
(a)
(b)
(c)
(d)
(e)
(f)
Figure 4.29: Electron micrographs taken after 12 weeks immersion in DMEM for:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated scaffolds.
(L) overall view, and (R) higher-magnification view.
63
(a)
(b)
(c)
(d)
(e)
(f)
Figure 4.30: Electron micrographs taken after 18 weeks immersion in DMEM for:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated scaffolds.
(L) overall view, and (R) higher-magnification view.
64
(a)
(b)
(c)
(d)
(e)
(f)
Figure 4.31: Electron micrographs taken after 24 weeks immersion in DMEM for:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated scaffolds.
(L) overall view, and (R) higher-magnification view.
65
Figure 4.32: Electron micrographs of native scaffold (of 85% porosity) at week 0:
(L) overall view, and (R) higher-magnification view.
4.3.5 Molecular weight analysis
Table 4.1: Mw, Mn, and PDI of native, NaOH-treated, and lipase-treated PCL-TCP
Scaffolds in vitro.
Week
0
6
12
55489
40023
1.39
55905
40880
1.37
NaOH-treated PCL-TCP scaffolds in DMEM
58940
50213
Mw (Dalton)
41279
36788
Mn (Dalton)
1.43
1.36
PDI
53315
37225
1.43
Lipase-treated PCL-TCP scaffolds in DMEM
54491
51717
Mw (Dalton)
39763
35215
Mn (Dalton)
1.37
1.47
PDI
52769
36598
1.44
Native PCL-TCP scaffolds in DMEM
56437
Mw (Dalton)
42101
Mn (Dalton)
1.34
PDI
The molecular weights of the PCL-TCP scaffolds immersed in culture medium were
measured with gel permeation chromatography for the degradation of up to 12
weeks and the results were tabulated as given in Table 4.1. The molecular weights
remained relatively constant for all the three scaffold groups, with only slight
decreasing trend observed for NaOH-treated scaffolds and lipase-treated scaffolds.
66
4.4 RESULTS - IN VIVO DEGRADATION STUDY
4.4.1 Porosity measurements and 3D models analysis
After 12 and 24 weeks of implantation in rats, the PCL-TCP scaffolds were retrieved
and their porosity was measured. The graph as shown in Figure 4.33 revealed that
the porosity values demonstrated an increasing trend for all three scaffold groups.
The increasing rate in the porosity of the lipase-treated scaffolds has been found to
be the highest, followed by NaOH-treated scaffolds and native scaffolds respectively.
At 12 and 24 weeks, the change in the porosity percentage recorded were 0.11%
and 2.28% for native scaffolds, 2.42% and 5.56% for NaOH-treated scaffolds, and
5.97% and 6.87% for lipase-treated scaffolds respectively. Refer to Appendix B1 –
Table B.2 for the complete data.
150
Native
NaOH-treated
120
Porosity (%)
Lipase-treated
90
85.24
83.41
84.44
85.33
85.43
89.48
87.18
88.05
60
30
0
0
12
Degradation Time (Weeks)
Figure 4.33: Porosity of PCL-TCP Scaffolds In vivo.
67
24
90.24
Figure 4.34: 3D model of native scaffold after 3 months implantation:
(L) top view, and (R) tilted view.
Figure 4.35: 3D model of NaOH-treated scaffold after 3 months implantation:
(L) top view, and (R) tilted view.
Figure 4.36: 3D model of lipase-treated scaffold after 3 months implantation:
(L) top view, and (R) tilted view.
68
Figure 4.37: 3D model of native scaffold after 6 months implantation:
(L) top view, and (R) tilted view.
Figure 4.38: 3D model of NaOH-treated scaffold after 6 months implantation:
(L) top view, and (R) tilted view.
Figure 4.39: 3D model of lipase-treated scaffold after 6 months implantation:
(L) top view, and (R) tilted view.
69
4.4.2 Weight loss analysis
The percentage weight loss of the PCL-TCP scaffolds harvested from rats after
implantation for 12 and 24 weeks were shown in Figure 4.40. As expected, the graph
revealed an increasing trend for all the three scaffold groups. After 12 weeks, the
percentage loss in weight for native, NaOH-treated, and lipase-treated scaffolds
were 2.54%, 7.59%, and 6.63% respectively. As the degradation period increased,
the weight loss percentage also increased. A sudden leap in the weight loss
indicating a significant loss of mass were observed at 24 weeks for lipase-treated
scaffolds. By the end of the in vivo study, the weight loss recorded were 4.12%,
6.14%, and 17.98% for native, NaOH-treated, and lipase-treated scaffolds
respectively. Refer to Appendix B4 – Table B.8 for the complete data.
35
30
Weight loss (%)
25
17.98
20
15
7.59
10
5
6.63
4.12
2.54
6.14
0
Native
-5
NaOH-treated
-10
Lipase-treated
-15
12
24
Degradation Time (Weeks)
Figure 4.40: Weight loss of PCL-TCP Scaffolds In vivo.
70
4.4.3 Compressive mechanical properties
The compressive properties of the scaffolds as a function of degradation time are
depicted in Figure 4.41 and 4.42. Both the compressive strength and modulus have
decreased significantly during the first 12 weeks of implantation and then continued
declining at a much slower rate as the degradation period increased. After 3 months
of implantation, the compressive strength of native, NaOH-treated, and lipasetreated scaffolds have reduced by 13.41%, 49.78%, and 57.34% respectively (Figure
4.41). Likewise, the compressive modulus have decreased by 25.15%, 37.01%, and
62.93% for native, NaOH-treated, and lipase-treated scaffolds respectively (Figure
4.42). By the end of the study at 24 weeks, the decrease in the compressive strength
and modulus recorded were 60.89% and 82.61% for native scaffolds, 52.61% and
46.75% for NaOH-treated scaffolds, and 58.03% and 69.25% for lipase-treated
scaffolds respectively. Refer to Appendix B2 – Table B.5 and Table B.6 for the
complete data.
Figure 4.41: Compressive strength of PCL-TCP Scaffolds In vivo.
71
Figure 4.42: Compressive modulus of PCL-TCP Scaffolds In vivo.
4.4.4 Surface morphology analysis
Figures 4.43 and 4.44 displayed SEM micrographs of the three scaffold groups after
implantation for 12 and 24 weeks in vivo, under low and high magnifications
respectively. The honeycomb-like pattern of triangular pores present in scaffolds at
week 0 was lost at week 12 and subsequently at week 24, with the diagonal rod-like
structures appeared to be melted and fused together. It is observed as well that the
rods’ diameter have decreased in size as the degradation occured. This is
particularly obvious for the lipase-treated scaffolds whose surface has became rough
and highly distorted (Figure 4.43 and 4.44 (e) and (f)). This correlates with the
findings from micro-CT analysis. Refer to Appendix B3 for the complete images.
72
(a)
(b)
(c)
(d)
(e)
(f)
Figure 4.43: Electron micrographs taken after 3 months implantation:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated scaffolds.
(L) overall view, and (R) higher-magnification view.
73
(a)
(b)
(c)
(d)
(e)
(f)
Figure 4.44: Electron micrographs taken after 6 months implantation:
(a,b) native, (c,d) NaOH-treated, and (e,f) lipase-treated scaffolds.
(L) overall view, and (R) higher-magnification view.
74
4.4.5 Molecular weight analysis
The tabulated molecular weight values for the PCL-TCP scaffolds tested in vivo (up
to 12 weeks) were shown in Table 4.2. In general, the values obtained did not reveal
any trend. The molecular weights remained relatively constant for all the three
scaffold groups.
Table 4.2: Mw, Mn, and PDI of native, NaOH-treated, and lipase-treated PCL-TCP
Scaffolds in vivo.
Month
0
3
Native PCL-TCP scaffolds in rat
56437
Mw (Dalton)
61054
Mn (Dalton)
42101
48094
PDI
1.34
1.27
NaOH-treated PCL-TCP scaffolds in rat
58940
Mw (Dalton)
55875
Mn (Dalton)
41279
41002
PDI
1.43
1.36
Lipase-treated PCL-TCP scaffolds in rat
54491
Mw (Dalton)
56240
Mn (Dalton)
39763
46196
PDI
1.37
1.22
75
4.4.6 Histology analysis
Slides of horizontal cross-sections of the
scaffold blocks were prepared and
stained with Hematoxylin and Eosin (H &
E) upon retrieval of the three scaffold
groups after 3 months implantation in rat
models. Figure 4.45 up to 4.50 displays
(a) 5x
representative slides of the native,
N a O H - t r e a t e d , a n d l i p a s e - t re a t e d
scaffolds after H & E staining. The pink
and blue colour seen is a positive stain
of cytoplasm due to Eosin and of nuclei
due to Hematoxylin respectively. As
observed, soft tissues, specifically
(b) 20x
adipose tissues as identified by their
Figure 4.45: H&E stain of native
scaffolds after 3 months implantation.
distinct anatomical features, were
(a) 10x
(b) 20x
Figure 4.46: H&E stain of native scaffolds after 6 months implantation.
76
evident throughout the scaffold. They
were found covering the surface of the
rods as well as infiltrating the pores of
the scaffolds. Moreover, several vascular
vessels were also detected. Some of
them were highlighted in orange circle in
Figure 4.45 (b), 4.47(b), and 4.49(b). No
(a) 5x
overt microscopic signs of inflammatory
cells
or
fibrous
encapsulation
were
recorded. Multinucleated giant cells or
macro-phages were also absent. The
lower magnification images captured
were also able to provide a rough
demonstration of the density of the rods
(b) 20x
of the three scaffold groups. The blacker
Figure 4.47: H&E stain of NaOHtreated scaffolds after 3 months
implantation.
the region is observed (as pointed by the
(a) 10x
(b) 20x
Figure 4.48: H&E stain of NaOH-treated scaffolds after 6 months implantation.
77
green arrows), the higher the density of the rod is, as white region constitute empty
spaces. Out of the three groups, NaOH-treated scaffolds appear to have the highest
density (Figure 4.47(a)), followed by native scaffolds (Figure 4.45(a)) and lipasetreated scaffolds (Figure 4.49(a)).
(a) 5x
(b) 40x
Figure 4.49: H&E stain of lipase-treated scaffolds after 3 months implantation.
(a) 10x
(b) 20x
Figure 4.50: H&E stain of lipase-treated scaffolds after 6 months implantation.
78
4.5 DISCUSSION
The first objective for this chapter of study was to monitor the in vitro degradation
profile of PCL-TCP scaffolds when immersed in standard culture medium that was
used as it contains vitamins, amino acid and glucose and is closer to the human
blood composition [Rai, 2006; Cheong, 2005]. The findings then would be more
clinically relevant. Results demonstrated that when immersed in DMEM culture
medium, PCL-TCP scaffolds in all three experimental groups displayed distinct
degradation behaviours. Micro-CT analysis revealed that although there is an
increasing trend in the porosity for all three scaffold groups, the rate has been noted
to be the highest for lipase-treated scaffolds, and then followed by NaOH-treated
scaffolds and native scaffolds respectively. By the end of week 18, the porosity has
reached to considerably high values of 93.20%, 87.26%, and 88.03% respectively.
This is in accordance to the gravimetric analysis, which showed a relatively huge
jump in the weight loss percentage of the lipase-treated as compared to the other
two groups. Further increase in surface roughness and thinning of the rods were
observed from the scanning electron micrographs for all the three scaffold groups as
the degradation time increased. In general, no significant nanopores can be detected
yet on the surface of the native scaffolds at week 6 and 12. Surface line cracks with
considerable gaps, and slight distortion has started to appear only after
approximately 18 weeks. In contrast, for lipase-treated scaffolds which has
undergone the highest degradation rate, disconnection of the rods can actually be
seen as early as 6 weeks and by 24 weeks, severe degradation and reduction in the
dimensions of the scaffolds were reported. In addition, compressive tests have also
corresponded well with the degradation results. There was a drastic fall in the
79
strength and stiffness of the lipase-treated scaffolds during the first 6 weeks of
immersion to an unfavourably low value (< 2MPa). For the native scaffolds, both
their strength and stiffness have been low, with the values decreasing thereafter. As
for the NaOH-treated scaffolds, a sudden decrease was noted between week 0 and
week 6, after which the values of the strength and stiffness fluctuated around 2-3
MPa and 5-6 MPa. The data suggest that the NaOH-treated scaffolds were able to
withstand considerable stress. Results here suggest that NaOH-treated scaffolds
demonstrate better physical properties compared to the lipase-treated and native
untreated ones. Lipase-treated scaffolds were shown to degrade in a much faster
but uncontrollable manner, whereas the untreated native scaffolds displayed the lack
of favorable surface properties.
The next objective of the study was to investigate the in vivo degradation profile of
untreated and treated PCL-TCP scaffolds implanted in the subcutaneous back of
rats for 12 and 24 weeks (3 and 6 months respectively). During the period of
implantation of up to 24 weeks, the PCL-TCP scaffolds exhibited major degradation
and changes in the surface morphology.
Scanning electron micrographs
demonstrated obvious distortion and increased surface roughness of the scaffolds,
especially for the lipase-treated scaffolds. Also, the distinct honeycomb-like
architecture of the pores present in original PCL –TCP scaffolds was lost. Micro-CT
scans revealed similar changes to the overall 3D framework and considerable
reduction in size. The percentage of porosity reported an increasing trend for all
three scaffold groups, with the highest rate seen for the lipase-treated scaffolds,
followed by NaOH-treated scaffolds and native scaffolds respectively. By the end of
week 24, the porosity has reached to considerably high values of 90.24%, 88.05%,
80
and 87.18% respectively. The weight loss recorded were 17.98%, 6.14%, and 4.12%
for lipase-treated, NaOH-treated, and native scaffolds respectively. This increase
helps to explain the porosity changes mentioned previously as weight-loss changes
are related to porosity i.e the higher the porosity, the greater the weight loss [Wan,
2005]. In line with the increase in porosity and weight loss, the mechanical properties
decreased significantly after 12 weeks of implantation. At the end of the in vivo study
of 24 weeks, the strength of the lipase-treated scaffolds dropped to < 2MPa. As for
the NaOH-treated scaffolds, the strength and stiffness were found to be above 2
MPa and 5 MPa respectively, indicating a more favourable tolerance in load bearing
sites. Interestingly, the molecular weight of the three groups remained relatively
constant during the degradation up to 12 weeks.
Histological sections of the scaffolds after implantation in vivo for 3 and 6 months
were stained with Hematoxylin and Eosin and analyzed. Throughout all scaffolds,
the adipose tissues were found covering the surface of the rods as well as infiltrating
the pores of the scaffolds. These cells were observed in direct apposition to the rods
of the scaffolds and are generally presented with a healthy appearance, suggesting
their biocompatibility nature. In addition, several vascular vessels were also
detected; thus indicating that vascularization has occured. At the implantation sites,
vasculature provides the main mode of transport. Molecular transport would include
the exchange of oxygen, nutrient, metabolic wastes and molecular signaling. And
these biochemical exchanges are essential for cell migration and proliferation.
Moreover, the presence of overt inflammatory cells or fibrous encapsulation was not
detected. One of the main characteristics of the pathological response (foreign body
reaction) to a biomaterial is the presence of multinucleated giant cells. Macrophages
81
or multinucleated giant cells were not detected suggesting the absence of an
inflammatory response by the scaffolds in vivo in a rat model. Based on this
histology findings as well as the physical properties results for the PCL-TCP
scaffolds during the in vivo study, NaOH-treated scaffolds appeared to provide a
better performance compared to the other two groups (native and lipase-treated
scaffolds) as they demonstrated a more favorable surface morphology and
maintained sufficient mechanical properties while degraded predictably to a higher
porosity value.
4.5.1 Comparison between in vitro and in vivo studies
The studies in this chapter showed that in vitro and in vivo degradation of PCL-TCP
scaffolds behaved quite similarly over the course of the degradation period. The
extent of degradation of PCL-TCP scaffolds in vitro was found to be comparable to
that in vivo. Data at a similar timepoint of 3 and 6 months were used in making the
comparison to provide a fair analysis. When immersed in the DMEM growth medium,
the scaffolds were found to degrade in a similar rate as when they were implanted
subcutaneously into the back of rats. This was reflected through the relatively similar
porosity increase, mechanical properties, and weight loss seen between in vitro and
in vivo experiments. Results from SEM also showed that both groups similarly
demonstrated significant changes to the surface morphology of the PCL-TCP
scaffolds.
The in vivo environment is much more complex and difficult to predict than the in
vitro setting, particularly compared with acellular in vitro experiments [Hedberg,
82
2005]. Implant size and location, health of the animal, and enzymatic and local
cellular activity are all factors that can influence the rate of degradation of a given
polymer scaffold [Perrin, 1997]. Accordingly, in general, correlations between in vitro
and in vivo results can be difficult to make. Nevertheless, in this study the in vivo
degradation of the PCL-TCP composite scaffolds appears similar to that seen in the
in vitro setting in the absence of any cells. Hence, the findings may suggest the
possibility of conducting in vitro experiment by using DMEM solution to observe the
degradation behaviour of PCL-TCP scaffolds in lieu of in vivo experiment in the
future.
In a study conducted previously, PCL-TCP scaffolds were implanted in the abdomen
of rat model and results showed that the degradation rate was higher in vivo than in
vitro [Yeo, 2007]. However, this is most likely due to the fact that rat’s abdomen was
an aggressive environment and hence caused the scaffolds to degrade within a
shorter time frame compared when they were immersed in the culture medium or
when implanted subcutaneously to the back of rats.
4.6 CONCLUSION
Some exciting observations were realized from this chapter of the study. The
outcome demonstrated that the untreated and treated PCL-TCP scaffolds, which
were separately immersed in culture medium for up to 24 weeks or implanted for 24
weeks inside rat model, had undergone a significant degradation. From the 12 and
24 weeks data, the extent of degradation in vivo was comparable to that observed in
83
vitro. However, regardless of in vitro or in vivo experiments, the degradation rate of
the lipase-treated scaffolds was noted to be the highest, followed by NaOH-treated
scaffolds and native scaffolds. Results from histology also supported that all the
three group of scaffolds promote healthy cellular attachment as well as
vascularizations. The absence of overt inflammatory response by the scaffolds in
vivo in the rat model was also reported.
In conclusion, our findings demonstrated that the NaOH-treated scaffolds performed
most favourably as compared to the rest. In contrast, lipase-treated scaffolds
degraded in a much faster and uncontrollable manner, and native scaffolds
displayed the lack of favourable surface properties. For oral and maxillofacial
applications, a scaffold that degrades around 6 months with controlled degradation
rate and favorable mechanical properties is ideal for bone regeneration.
84
CHAPTER 5: EVALUATION OF PCL-TCP SCAFFOLDS
IN A CLINICALLY RELEVANT DEFECT MODEL
5.1 INTRODUCTION
In the previous chapter, the degradation behavior of PCL-TCP scaffolds in smaller
animal model has been evaluated. We discovered that the native and customized
scaffolds were biocompatible and that scaffolds with faster degradation rate were
needed when used for oral and maxillofacial applications. However the rat study
alone was still considered insufficient if we planned to present the technique for
clinical applications; that is to treat dentoalveolar defects in human. The main
concern was that in the previous animal study, the implantation of the scaffolds
(subcutaneous back of rats) was not exactly at clinically relevant sites. Ideally prior
to human clinical study, a more demanding and clinically representative larger
animal model was required. Hence with this in mind, another in vivo study was
conducted whereby PCL-TCP scaffolds were implanted in the mandibles of
micropigs. The possibility of the PCL-TCP scaffold for use as a bone substitute in
bone regeneration would then be compared to the current gold standard of using
autogenous bone [Betz, 2002; Horch, 2006; Schuckert, 2009].
In addition to scaffolds, the latest guided bone regeneration (GBR) technique for
localized ridge augmentation also involves the usage of barrier membranes.
Extensive experimental studies have been conducted and results from these studies
85
have demonstrated that the placement of a membrane promotes the osseous
healing of bone defects, since competing non-osteogenic soft tissue cells are
excluded
from
defect
healing
by
the
presence
of
the
physical
barrier.
Simultaneously, membranes allow the ingrowth of angiogenic and osteogenic cells
to populate and regenerate these defects with bone [Schenk, 1994; Buser, 1996;
Dahlin, 1988; Dahlin, 1990; Seibert, 1990]. On the other hand, noncovered control
sites demonstrated incomplete bone regeneration and the presence of scar tissue
within the defects [Buser, 2002]. These observations were confirmed in a recently
published study by Schenk et al. that gave detailed information about the sequence
and pattern of bone regeneration underneath barrier membranes. This study in the
canine mandible demonstrated that bone regeneration in membrane-protected
defects closely followed the pattern of normal bone growth and development, and
that tissue which had formed beneath membranes was normal bone [Schenk, 1994;
von Arx, 2001]. Recent experimental results also indicate that the use of barrier
membranes can direct bone fill not only to contour defects of the alveolar ridge, but
also to grow beyond the level of the surrounding bone, thus forming excess bone to
a considerable extent [Schliephake, 1994; Kostopoulos, 1994; Schliephake, 1998].
For this study, we have decided to test for the first time, the feasibility of PCL-TCP
sheets when used as barrier membranes. They were designed to be 3 layers thick
with varying porosity, and would be compared to the widely used resorbable
collagen membranes, which are currently the gold standard.
The specific aim of this study was then to evaluate PCL-TCP scaffolds and sheets
as defect fillers and barrier membranes respectively for novel guided bone
regeneration technique in the reconstruction of localized dentoalveolar defects in a
86
micropig model. A total of 10 micropigs were employed for the study. All premolars
(P1-P4) and the 1st molars (M1) were initially extracted from the posterior mandible
and
removed
bilaterally
to
create
partially
edentulous
alveolar
ridges.
Simultaneously, 2 bilateral bone defects were created on each side (15 x 10 x 8 mm)
of the mandible by removing the buccal cortex. The defects were left to heal for a
period of 2 months. Upon re-entry, the defects were planned for lateral ridge
augmentation using either autogenous graft or PCL-TCP scaffolds. A collagen
membrane or a prototype PCL-TCP sheet was also used. The membranes or sheets
were trimmed, immersed in a hot water bath to mold it into shape and placed over
the augmented sites and the defect margins by about 2mm. Each micropig then had
the 4 defects randomly treated with one of the following GBR techniques:
Site 1: Collagen membrane + PCL-TCP scaffold
Site 2: Collagen membrane + autograft
Site 3: PCL-TCP sheet + autograft
Site 4: PCL-TCP sheet + PCL-TCP scaffold
After a healing period of 6 months following GBR (Figure 5.1), all micropigs were
sacrificed and analyses of the bone regenerated were performed.
Surgery 1 (Extraction and defect creation)
2 months
Surgery 2 (Ridge augmentation)
6 months
Sacrifice + Analysis
Figure 5.1: Timeline for the complete micropig study.
87
5.2 MATERIALS AND METHODS
5.2.1 Implant design and fabrication
Bioresorbable scaffold and sheet specimens were fabricated with PCL-TCP
(80:20%) filaments by using a fused deposition modeling (FDM) 3D Modeler RP
system from Stratasys Inc (Eden Prairie, MN). They were purchased directly from
Osteopore International Pte Ltd, Singapore. Each composite manifested a lay-down
pattern of 0/60/120º with a typical honeycomb array of interconnected equilateral
triangle. TCP existed as non-uniformly distributed particles on the rods of PCL. The
scaffolds came in a block of 15 x 10 x 8 mm with 70% porosity for the outer 2 mm
and 85% porosity for the inner 3 mm. On the other hand, the sheets were 25 x 25 x 1
mm consisting of 3 layers: 1 outer layer of 30% porosity and 2 inner layers of 70%
porosity. The sheets were light, resilient and malleable. All specimens (Figure 5.2)
were produced in a class 10K clean room environment and sterility was achieved via
ethidium oxide treatment.
Figure 5.2: 15x10x8mm PCL-TCP scaffold (left) and 25x25x1mm
PCL-TCP sheet (right).
88
The bioresorbable collagen membranes were purchased from BioGide, Geistlich
Pharma AG, Wolhusen, Switzerland (Figure 5.3, left). Bio-Gide’s collagen membrane
was developed particularly for periodontal, peri-implant applications or to improve
the ossification of bone defects of any origin. It is a bilayer membrane; one compact
and smooth layer is covered by a particularly dense film, designed to prevent the
invasion of soft tissue in a membrane-protected bone defect. The other rough side is
placed towards the bone defect in order to make bone ingrowth possible [Zhao,
2000].
Figure 5.3: Bioresorbable collagen membrane from BioGide (left) and temperaturecontrolled hot water bath (right).
5.2.2 Animal husbandry
A total of 10 male micropigs were employed for the study. At the beginning, these
animals were about 1-2 years old and weighed approximately 40-50 kg. The animals
were housed in the animal holding facility at the SingHealth Experimental Medicine
Centre (SEMC), Singapore General Hospital (SGH), for the entire duration of the
experiment. Housing and feeding were according to standard animal-care protocols.
The study was conducted according to the guidelines of the SingHealth
Experimental Medicine Centre, Singapore General Hospital.
89
Figure 5.4: Micropig housing facility at SEMC, SGH (left) and weighing of micropig
prior to the experiment (right).
5.2.3 Pre- and postoperative medication
All surgical procedures were performed under general anesthesia in an operating
room. For premedication, the following agents were used: Ketamine (11-15 mg/kg,
Ketapex®, Apex Laboratories, Australia) and Atropine (0.4-0.5 mg/kg, Pharmacia
Pte Ltd, Australia) IM. Subsequently, the micropigs were intubated and were
administered an inhalation of 5% Isoflurane initially and thereafter maintained with 13% Isoflurane (Abbott Laboratories Ltd, England) in 02-maintenance. After
disinfection of the surgical site with 10% povidone-iodine solution (Clinidine®,
Clinipad Co., Guilford, CT, USA), local anesthetic (Lidocaine HCL 2% with
epinephrine 1:100,000, Henry Schein Inc., Port Washington, NY, USA) was
administered by infiltration at the respective buccal and lingual sites. Postoperatively,
90
the micropigs received Caprofen 2-4 mg/kg every 12-24 hours for 3 days (Vericore
Ltd, Scotland, UK) as an analgesic IM. For antibiotic cover, 2.5 mg/ 50kg of
Benzathine-Penicillin 150,000 + Procaine-Penicillin G 150,000 was delivered every
48 hours for 7-10 days IM (Pen-B®, Pfizer Inc., Lee’s summit, MO, USA). In addition,
1-2 mg/kg of the antibiotic Gentamicin was administered IM every 12 hours for 7-10
days (Il Dong, Pharmaceutical Co Ltd, Korea). For suture removal, Ketamine (11-15
mg/kg Ketapex®, Apex Laboratories, Australia) and Atropine (0.4-0.5 mg/kg,
Pharmacia Pte Ltd, Australia) were administered IM and thereafter maintained with
1-3% Isoflurane (Abbott Laboratories Ltd, England) in 02-maintenance. Oral hygiene
procedures were carried out two times a week using 0.2% chlorhexidine gel (PlakOut® Gel, Hawe Neos Dental, Biaggio, Switzerland). A soft diet was maintained
throughout the study.
5.2.4 Surgery 1 (Extraction and defect creation)
Sulcular incisions were made with subsequent reflection of full mucoperiosteal flaps.
In the mandible, all premolars (P1-P4) and the first molars (M1) were removed,
whereas in the maxilla the 2nd and 3rd premolars (P2 and P3) were also extracted.
Prior to removal, all two-rooted teeth were sectioned and separated individually,
employing a separating disk, prior to root extraction. Subsequently, a large “chronictype” bone defect (Length 45 mm, Height 12 mm, Depth 5 mm) were created in the
mandible by removing the buccal bone plate. The large defect encompassed
approximately the extraction sites of P2, P3, and P4. A small round bur was used to
outline the defect margins on the buccal bone plate. Subsequently, the bur holes
were connected employing a fissure bur and the buccal bone plate was removed
91
with a chisel placed in the cut groove. In order to accentuate the defect, a pearshaped bur was utilized. Caution was exercised to retain the lingual cortex and the
height of the crest. All drilling was done with sterile saline irrigation. Finally, the flaps
were re-approximated with single interrupted resorbable 4.0 Vicryl sutures (Ethicon,
Norderstedt, Germany). These were removed two weeks postoperatively. Oral
hygiene procedures were carried out two times a week using 0.2% chlorhexidine gel
(Plak-Out® Gel, Hawe Neos Dental, Biaggio, Switzerland). A soft diet was
maintained throughout the study. During the first week of post-operative healing, the
animals were checked daily for signs of infection.
Figure 5.5: Removal of all premolars and first molar (left), and
the extraction sites (right).
Figure 5.6: The flaps were re-approximated with Vicryl sutures (left), and the defect
sites were closed (right).
92
5.2.5 Surgery 2 (Ridge augmentation)
After a healing period of 2 months, the defect sites in the mandible were reopened
using a mid-crestal incision from P1 to M1. Vertical releasing incisions enabled full
access to the area. All granulation tissue was carefully removed from the formerly
created ridge defects. To open up the bone marrow space around the chronic-type
bone defect, small holes were drilled into the surrounding cancellous compartment.
The bone defects were then augmented in four different ways with random
assignment of each grafting treatment.
Site 1: Collagen membrane + PCL-TCP scaffold
Site 2: Collagen membrane + autograft
Site 3: PCL-TCP sheet + autograft
Site 4: PCL-TCP sheet + PCL-TCP scaffold
Site 1
Site 2
PCL-TCP scaffold + collagen membrane
Autograft + collagen membrane
Site 3
Site 4
Autograft + PCL-TCP sheet
PCL-TCP scaffold + PCL-TCP sheet
Figure 5.7: Schematic illustrations of the four tested grafting procedures.
93
The autografts were harvested from the site of the formerly extracted M1 (12 x 8 x 5
mm), using the same method as before when the mandibular defects were created.
Cortico-cancellous block grafts were procured from the buccal aspect using a small
round bur to outline the grafts with a series of perforations. These were then
connected with a side-cutting fissure bur and the fragments relieved with a chisel
placed in the cut groove. Before the placement of the grafts and scaffolds, multiple
small perforations were made into the recipient wall of the defect to encourage
bleeding and the release of growth factors and cells into the defect sites.
Figure 5.8: Placement of PCL-TCP scaffolds and autografts (left), followed by
PCL-TCP sheets and collagen membranes (right).
The block grafts were then immediately transplanted to their assigned defect sites. A
porous PCL-TCP scaffold (12 x 8 x 5 mm) was used as an alternative bone
substitute and placed in the assigned defects. The grafts and scaffolds were secured
using a centrally located miniscrew. A prototype PCL-TCP sheet and a bioresorbable
collagen membrane (BioGide®, Geistlich Pharma AG, Wolhusen, Switzerland) were
again randomly selected and individually trimmed to overlap the defect margins by
about 2-3 mm. To facilitate a fluid-tight and tension-free wound closure, the
periosteum was released at its base. Wound margins were then re-approximated
94
and closed with horizontal mattress and interrupted resorbable 4.0 Vicryl sutures
(Ethicon, Norderstedt, Germany). Sutures were removed two weeks postoperatively. Oral hygiene procedures were carried out two times a week using 0.2%
chlorhexidine gel (Plak-Out® Gel, Hawe Neos Dental, Biaggio, Switzerland). A soft
diet was maintained throughout the study. During the first week of post-operative
healing, the animals were checked daily for signs of infection.
5.2.6 Sacrifice
Figure 5.9: Micropig under euthanasia (left), and the mandible was block resected
using an oscillating autopsy saw (right).
Figure 5.10: The recovered segment of mandible (left), the site after removal (right).
95
All 10 micropigs were sacrificed 6 months after lateral ridge augmentation.
Euthanasia was performed with an overdose of pentobarbital sodium 0.2 ml i.v. (=65
mg/kg, Euthanasia-5®, Henry Schein Inc). Subsequently, the mandibles were block
resected using an oscillating autopsy saw and the recovered segments were
immediately immersed in a solution of formaldehyde 4% combined with CaCl2 1%.
5.2.7 Micro-computed tomography analysis
Upon retrieval from the animals, the PCL-TCP scaffold specimens were
characterized using Micro-computed tomography (Micro-CT Skyscan 1076, Belgium).
The specimens were placed in a sample holder and scanned through 180° at a
spatial resolution of 30µm. The image data from the scanned planes were
subsequently thresholded and reconstructed to create 3-D images for quantitative
analysis. The 3D volumes were evaluated by direct transformation methods and
subsequently the total scaffold volume was calculated within a volume defined by the
boundaries of the constructs. All parameters were measured on the buccal and
lingual aspects of the specimens. The bone volume fraction (BVF) obtained from
BV/TV, represents the percentage of a volume of interest that was mineralized.
5.3 RESULTS
5.3.1 Gross examinations
All 10 animals used in the study remained healthy throughout the experiment and
survived the surgical procedure with minimal adverse effects from the implantation of
96
the autografts and the PCL-TCP constructs. Opened sutures, or soft tissue
dehiscences, were noted for the majority of grafts covered with PCL-TCP sheets
(70% occurrence for autograft + PCL-TCP sheet combination, and 90% occurrence
for PCL-TCP scaffold + PCL-TCP sheet combination). However, only some were
observed for those covered with collagen membranes (10% occurrence for autograft
+ collagen membrane combination, and 20% occurrence for PCL-TCP scaffold +
collagen membrane combination).
Figure 5.11: The recovered segment of the mandible of a micropig.
Figure 5.12: Soft tissue dehiscence observed for the majority of grafts
covered with PCL-TCP sheets.
97
Table 5.1: Number of sites with soft tissue dehiscence for the implanted autograft,
collagen membranes, PCL-TCP scaffolds, and PCL-TCP sheets.
Group
Combinations
Number of sites with soft
tissue dehiscence (out of 10)
1
Autograft + Collagen membrane
1
2
Autograft + PCL-TCP sheet
7
3
PCL-TCP scaffold + Collagen membrane
2
4
PCL-TCP scaffold + PCL-TCP sheet
9
Due to the high occurrence of the soft tissue dehiscence for PCL-TCP sheets groups,
it was decided that samples from this group would be omitted for analysis. Only
those covered with collagen membranes that would be discussed from this point
onwards (i.e. autograft means autograft covered with collagen membrane).
5.3.2 New bone formation
Micro-CT was utilized to determine the value of early bone volume ingrowth detected
at the 6 months after implantation of the autografts and the scaffolds. Results of new
bone formation at 6 months showed a higher trend in the volume of bone ingrowth
by the autografts than the scaffolds (as illustrated in Figure 5.13). Due to the missing
screw in the autograft site covered with collagen membrane in P7, values from P7
were omitted from the computation for better comparison purpose.
98
Figure 5.13: Bone volume fraction detected after 6 months of implantation of
autografts and PCL-TCP scaffolds for individual micropigs.
The bone volume fraction represents the amount of mineral present at the defect site.
The volume fraction of newly formed bone within the defect was calculated to be
22.67% for PCL-TCP scaffolds under collagen membrane, and therefore the pores
of the PCL-TCP scaffolds were not completely filled with mineralized tissue after 6
months. Bone formation in the autograft site covered with collagen membrane was
higher at 36.22% volume fraction. These values (shown in Figure 5.14) were based
on the average of 9 micropigs, as data from P7 was incomplete and hence omitted.
The considerably high standard deviation was likely due to the different healing
pattern and capacity of each micropig tested.
99
Figure 5.14: The average values of bone volume fraction detected after 6 months of
implantation of autografts and PCL-TCP scaffolds.
5.3.3 Ratio of bone volume fraction for PCL-TCP scaffolds with respect
to autografts
In order to quantify the efficiency of PCL-TCP scaffolds as compared to autograft in
the bone reconstruction application, the ratio of bone volume fraction for PCL-TCP
scaffolds with respect to autografts were computed. This was obtained by dividing
the BV/TV value of a PCL-TCP scaffold with the BV/TV value of an autograft. Figure
5.14 displayed the calculated ratio for each individual micropigs tested. As shown
from the graph, the efficiency for all the micropigs was found to be comparable. The
average, based on 9 micropigs, was 0.64 ± 0.16. This means that when compared to
the use of autografts in the reconstruction of bone, the PCL-TCP scaffolds were
about 64% efficient.
100
Figure 5.15: The ratio of bone volume fraction for PCL-TCP scaffolds with respect
to autografts for individual micropigs.
5.3.4 3D model analysis
Representative 3D micro-CT images of autograft-treated and PCL-TCP scaffoldtreated defects under collagen membrane were constructed and examined to
observe the formation of new bone at the defect area. It must be noted that the
images were captured at a density that only picked up bone and miniscrews, and not
scaffolds. The entire height of the defect was not fully occupied with bone. Regularsized gaps were evident in the new bone that corresponded to the scaffold porous
architecture.
101
Figure 5.16: PCL-TCP scaffold treated site: overview (left) and cross-section (right).
Figure 5.17: Autograft-treated site: overview (left) and cross-section (right).
5.3.5 Two-dimensional x-ray radiographs evaluation
Radiographs of the defect sites after 6 months implantation of both autograft and
PCL-TCP scaffold were also taken and findings similar to those from the Micro-CT
images were revealed. The volume of mineralized tissue at the defect sites treated
with autografts were observed to be higher than those treated with PCL-TCP
scaffolds as shown from Figure 5.18 to 5.20 below. The images below show the side
102
of the mandible that has been treated with both autograft and PCL-TCP scaffold,
and covered with collagen membrane. “P” and “A” refers to the posterior and anterior
section of the mandible respectively.
Figure 5.18: X-ray image of a micropig’s left mandible treated with autograft
(posterior) and PCL-TCP scaffold (anterior), and covered with collagen membrane.
Figure 5.19: X-ray image of a micropig’s right mandible treated with PCL-TCP
scaffold (posterior) and autograft (anterior), and covered with collagen membrane.
103
Figure 5.20: X-ray image of a micropig’s left mandible treated with autograft
(posterior) and PCL-TCP scaffold (anterior), and covered with collagen membrane.
5.4 DISCUSSION
The present study has evaluated the use of PCL-TCP scaffolds and sheets for ridge
augmentation in an experimental micropig model. A pig model has been chosen, as
it possesses similar healing properties to that of the human. The anatomy of the
pig’s jaws and their dentition, bone metabolism, clotting parameters closely
resembles to that of a human. Being omnivorous animals, the dietary pattern would
also be similar. Initially the domestic pig model was chosen due to their availability
and low cost. However, potential logistic and surgical challenges at sacrifice when
the final weights of these pigs measure approximately 130-150 kg must be taken into
consideration. A smaller micropig model was then finally decided.
A clinically frequent situation with mandibular bone atrophy was simulated in this
study by creating chronic bone defects on the buccal aspect of the alveolar ridge.
The defects were treated using either autogenous grafts or PCL-TCP scaffolds in
104
combination with collagen membranes or prototype PCL-TCP sheets. Attention was
then given to achieving a close adaptation of the membrane to the surrounding bone
and good membrane stabilization. Throughout the experiment, all the autograft bone
blocks and PCL-TCP scaffolds that were implanted within the assigned defects
remained in situ with no signs of migration. The survival rate of the micropigs was
excellent and they healed and recovered well after the surgeries. There were no
reports of any complications detected, such as overt edema and infection during the
healing period, suggesting the biocompatibility of PCL-TCP scaffolds in promoting a
conducive in vivo environment for healing to take place. However, mid-way through
the 6 months healing period, almost all the 10 micropigs presented with soft tissue
dehiscence and biomaterial exposure in majority of sites that utilized PCL-TCP
sheets as barrier membranes. This occurred despite the stringent post-operative
care that was prescribed to the animals following GBR surgeries. It appears that the
nature of the PCL-TCP sheets when moulded around the augmented sites had some
form of ‘elastic memory’ which resulted in a partial relapsed of the PCL-TCP sheets
and caused unwanted tension and perforation of the overlying soft tissue whilst
healing. When a soft tissue dehiscence occurs, the exposure of the membrane could
lead to its contamination with bacteria from the oral cavity and frequently to an
infection in the membrane site [Buser, 1999; Buser, 2002]. For this reason, samples
from the PCL-TCP sheets groups were omitted. Interestingly, GBR procedures
utilizing collagen membranes with autografts or PCL blocks showed few soft tissue
complications. The reason could be because collagen membrane is pliable when
moist and conforms well to the surgical area. It provides a thrombogenic surface that
is sealed coronally to the root surface by a fibrin clot, and does not elicit any allergic
responses [Blumenthal, 1993]. Collagen itself has several advantages because it is
105
absorbable, does not require a second surgical procedure for removal, and has
some unique biologic properties. It is the major extracellular macromolecule of the
periodontal connective tissue and bone and is physiologically metabolized by these
tissues; it is chemotactic for fibroblasts; it has been reported to act as a barrier for
migrating epithelial cells in vitro; and it has been used experimentally in animals and
humans [Pitaru, 1988; Owens, 2001].
The results from this study demonstrated that the combined placement of collagen
membranes with autografts at the defect sites promotes the best osseous healing of
bone defects as shown from the higher volume of mineralized tissue detected. This
observation is consistent with the previous study conducted by Buser et al,
comparing autografts with four alternative bone fillers in such defects in the mandible
of micropigs. Utilizing autografts, bone regeneration is optimized during the initial
bone healing period due to the presence of particulate grafts with excellent
osteoconductivity [Buser, 1999; Lyford, 2003]. In addition, autografts also have
osteoinductive properties in the sense of osteogenic transfer. With graft application,
osteoblasts and osteoblast precursor cells as well as growth factors (transforming
growth factor β) and bone-inducing factors (bone morphogenetic protein) entrapped
in the grafted bone matrix are transferred to the augmentation site resulting in an
activation of bone formation [Burchardt, 1983; Buser 1999, Buser, 1996]. This is
crucial as bone formation is mainly activated by the release of growth factors and
bone-inducing substances. This activation is manifested as a stimulation of
neoangiogenesis, the recruitment of osteoblasts, and in the onset of bone matrix
deposition, provided that the activators act upon committed responding cells. An
influencing mechanism of collagen membrane placement may also be that
106
stimulating growth factors are locally concentrated in the osseous wound at inductive
doses, leading to osseous repair of such defects that normally would not heal
spontaneously [Schenk, 1994; Dahlin, 1993].
It is also essential to understand the biological behavior of autografts with respect to
graft incorporation and repair and the differences between cortical and cancellous
autografts. These details have been intensively studied in numerous experimental
studies in orthopedic surgery [Burchardt, 1983; Burchardt, 1987]. Cancellous
autografts are rapidly revascularized, and they are completely repaired by creeping
substitution. In contrast, revascularization of cortical autografts is slow and occurs
through existing haversian canals. Remodeling of cortical autografts is also slow and
results in a mixture of necrotic and new viable bone [Lyford, 2003; Buser, 1999].
Based on this biological knowledge of graft incorporation and graft repair,
corticocancellous block grafts placed in the center of the augmentation area were
subsequently used in this study. They were appropriately applied to the recipient site
with rigid fixation of the graft. A bone-graft fixation screw should be used because it
allows precise positioning of the graft and prevents micromovements of the graft
underneath the membrane during healing. In addition, the block graft must be placed
with its cortical layer facing buccally and the cancellous portion of the graft in direct
contact of the host bone. This surgical approach is based on two assumptions. First,
the cortical portion of the graft facing to the buccal aspect of the crest is used to
reestablish the missing buccal cortex. Although this new cortex will be a mixture of
necrotic and new viable bone, it offers good mechanical stability and is less
susceptible to resorption than cancellous bone. Second, the cancellous portion of
the graft is placed in direct contact to the host bone in the area where the implant will
107
be placed during next surgery. The host bone surface is perforated during the
surgical procedure to activate bone formation and to open the marrow space,
allowing fast ingrowth of blood vessels. It can be expected that this portion of the
graft will undergo rapid revascularization and graft remodeling. These assumptions,
however, are based on orthopedic literature, and histologic details of graft
incorporation and repair underneath barrier membranes are not yet documented
[Buser, 2002; Buser, 1999].
Both autografts and PCL-TCP scaffolds are found to be able to support an applied
membrane, thus preventing a membrane collapse and maintaining the created
space. There is an agreement in the literature that the maintenance of the
membrane-protected space is one of the essential prerequisites for a successful
treatment outcome with guided bone regeneration procedures [Buser, 1990;
Rominger, 1994; Buser, 1999]. When compared to the use of autografts in the
reconstruction of bone, the PCL-TCP scaffolds were only about 64% efficient.
However the application of PCL-TCP scaffolds, instead of autografts, would be
highly preferable as this option would avoid the harvesting of autogenous bone from
the patient. Hence, more studies and modifications were needed to further enhance
the efficiency of PCL-TCP scaffolds as a bone substitute in bone regeneration for
localized ridge augmentation. The development of a longer-lasting bioabsorbable
membrane with the same qualities as the current collagen membrane concerning
tissue compatibility, mechanical attributes, and intrasurgical handling would be highly
desirable too.
108
5.5 CONCLUSION
This part of the study was to investigate the in vivo behavior of early matrix
deposition and early bone formation of PCL-TCP scaffolds as compared to the use
of autografts, in conjunction with membrane coverage in the mandible of micropigs
after 6 months of implantation. A clinically frequent situation with mandibular bone
atrophy was simulated by creating chronic bone defects on the buccal aspect of the
alveolar ridge. Healing was uneventful in all micropigs showed that the PCL-TCP
scaffolds exhibited good biocompatibility. Across the tested treatment options in this
study of 10 animals, defect sites augmented with autografts and collagen
membranes showed the most promising results. The collagen membranes were
found to offer the advantage of a reduced frequency of soft tissue dehiscence. More
improvements are needed to increase the efficiency of the PCL-TCP scaffolds in
bone healing as they could ruled out the need for harvesting grafts.
109
CHAPTER 6: FINAL CONCLUSIONS AND
RECOMMENDATIONS
6.1 FINAL CONCLUSIONS
The general aims of the present study were to investigate the degradation and loadbearing profile of 3D bioresorable polycaprolactone-20% tricalcium phosphate (PCLTCP) scaffolds under enzymatic and hydrolytic conditions and subsequently to
evaluate the efficacy of the scaffolds in both small and large animal models. The
purpose was to develop scaffolds with desirable customized properties and
increased degradation rates suitable for application in dentoalveolar defects
treatment.
PCL-TCP scaffolds have shown to undergo both chemical and enzymatic
degradation in the first part of the study when degraded in sodium hydroxide and
lipase solution respectively for up to 108 hours. The objective of the experiment was
achieved with scaffolds of approximately 85% porosity obtained after 96 hours of
treatment in 3M NaOH and 12 hours in 0.1% lipase. These pre-treated scaffolds
demonstrated acceptable mechanical strength, structure, and surface morphology.
Some exciting observations were also realized from the second part of the study.
The outcome demonstrated that the untreated and treated PCL-TCP scaffolds,
which were separately immersed in culture medium for up to 24 weeks or implanted
110
for 24 weeks inside rat model, had undergone a significant degradation. From the 12
weeks data, the extent of degradation in vivo was comparable to that observed in
vitro. However, regardless of in vitro or in vivo experiments, the degradation rate of
the lipase-treated scaffolds was noted to be the highest, followed by NaOH-treated
scaffolds and native scaffolds. Results from histology also supported that all the
three group of scaffolds promote healthy cellular attachment as well as
vascularizations. The absence of overt inflammatory response by the scaffolds in
vivo in the rat model was also reported. Our findings demonstrated that the NaOHtreated scaffolds performed most favourably as compared to the rest. In contrast,
lipase-treated scaffolds degraded in a much faster and uncontrollable manner, and
native scaffolds displayed the lack of favourable surface properties.
Lastly, another in vivo study was conducted whereby PCL-TCP scaffolds and sheets
were evaluated as defect fillers and barrier membranes respectively for novel guided
bone regeneration technique in the reconstruction of localized dentoalveolar defects
in a micropig model for up to 6 months. This was essential as prior to human clinical
study, a more demanding and clinically representative larger animal model was
required. The possibility of the PCL-TCP scaffold for use as a bone substitute was
compared to the current gold standard of using autogenous bone. Healing was found
to be uneventful in all micropigs and this showed that the PCL-TCP scaffolds
exhibited good biocompatibility. Across the tested treatment options, defect sites
augmented with autografts and collagen membranes showed the most promising
results with greater bone formation detected as compared to PCL-TCP scaffolds and
collagen membranes which were about 64% efficient. The collagen membranes
were found to offer the advantage of a reduced frequency of soft tissue dehiscence
111
in comparison to PCL-TCP sheets. For oral and maxillofacial applications, a scaffold
that degrades around 6 months with controlled degradation rate and favorable
mechanical properties is ideal for bone regeneration. More improvements are
needed to increase the efficiency of the PCL-TCP scaffolds in bone healing as they
could ruled out the need for harvesting grafts.
6.2 RECOMMENDATIONS FOR FUTURE WORK
Future work may also entail the investigation of in vivo degradation behaviour of the
bioactive factors-loaded PCL-TCP scaffolds in bone mandibular defects of large
animal model such as micropig. The growth factor of choice could be bone
morphogenetic proteins (BMPs), one of the most potent factors that regulate
osteoblasts to form mineralized tissue [Yamaguchi, 2000]. The key research
objective here is to investigate the capacity of BMP-loaded PCL-TCP composite
scaffolds in improving the stimulation of bone repair and regeneration upon
implantation into large segmental bone defects for a long-term period and ultimately
to rule out the need for harvesting autografts. A large animal model is vital for
advancement into clinical trials and a long time period is required to acquire useful
insights of the in vivo degradation properties of the scaffolds and the time required
for mature bone deposition and remodeling to occur. BMPs have the unique
functions of inducing the differentiation of cells of the osteoblastic lineage, thus
increasing the pool of mature cells, and of enhancing the differentiated function of
the osteoblasts [Canalis, 2003; Yamaguchi, 2000]. They were originally identified as
growth factors that regulate growth and differentiation of chondroblast and osteoblast
112
lineage cells in vitro. In accordance with their in vitro effects, BMPs induce bone and
cartilage formation when implanted at ectopic sites in rats [Reddi, 1997; Urist, 1965].
PCL-TCP scaffolds seeded with recombinant human bone morphogenetic protein-2
(rhBMP-2) have previously been investigated both in vitro and in small animal
models, and results have shown that the incorporation of rhBMP-2 seems to
accelerate mineralization. When implanted for guided bone regeneration technique
in the reconstruction of localized dentoalveolar defects in a micropig model, specific
objectives could include investigating the amount and quality of bone formation in
terms of the bone volume fraction, bone union formation and mechanical properties;
the amount and type of cellular and vascular infiltration into the defect site; the longterm effects of the material on host tissue; and the long-term effects of the host
environment on the material biodegradation. It is also interesting to compare the
concentration of growth factor released at the local site of implantation against the
amount that is transported in the bloodstream and excreted in urine. It is hoped that
the local, controlled release of rhBMP-2 from PCL-TCP scaffolds in vivo will enhance
the osteoinductivity of the scaffolds and that this strategy will provide a different and
more sophisticated way of enhancing the efficacy of the PCL-TCP scaffolds for bone
tissue engineering in dentoalveolar applications. In addition, more characterization
methods and extensive analyses of the PCL-TCP scaffolds can be performed to
provide better understanding of the changes in the scaffolds’ properties.
113
REFERENCES
Adell R, Eriksson B, Lekholm U, Branemark P I, Jemt T. Long term follow up study of
osseointegrated implants in the treatment of totally edentulous jaws. Int J Oral
Maxillofac Implants 1990; 5: 347-359.
Alsberg E, Kong H.J, Hirano Y, Smith M.K, Albeiruti A, Mooney D.J. Regulating bone
formation via controlled scaffold degradation. J Dent Res 2003; 82 (11): 903-908.
Barckhaus R.H. and Hohling H.J. Electron microprobe analysis of freeze dried and
unstained mineralized epiphyseal cartilage. Cell Tissue Res 1978; 18693: 541-549.
Baron R. Anatomy and ultrastructure of bone. In: Favus MJ, editor. Primer on
Metabolic and Bone Diseases and Disorders of Mineral Metabolism. Philadelphia:
Lippincott-Raven. 1996. p3-9.
Barrère F, Ni M, Habibovic P, Ducheyne P, and de Groot K. Scaffolds: Degradation
of bioceramics, in Tissue Engineering, CA van Blitterswijk et al, Editors, 2008,
Academic Press, p.223-254.
Barrère F, Stigter M, et al. In vitro dissolution of various calcium-phosphate coatings
on Ti6Al4V. Bioceramics Key Engineering Materials 2000.
Betz R.R. Limitations of autograft and allograft: new synthetic solutions. Orthopedics
2002; 25: 561.
Blumenthal N.M. A clinical comparison of collagen membranes with e-PTFE
membranes in the treatment of human mandibular Class II furcations defects. J
Periodontol 1993; 64: 925-33.
Boskey A.L. Biomineralization: an overview. Connect Tissue Res 2003; 44: 5.
Brickley M. and Ives, R. The bioarchaeology of metabolic bone disease. San Diego:
Academic Press. 2008.
Bronner F, Worrell R.V (eds.), Orthopaedics: Principles of basic and clinical science.
Boca Raton, FL; 1999, CRC.
Burchardt H. Biology of bone transplantation. Orthodont Clin North Am. 1987; 18:
187-196.
Burchardt H. The biology of bone graft repair. Clin Orthop Res. 1983; 174: 28-36.
Buser D, Bragger U, Lang N.B, Nyman S. Regeneration and enlargement of jaw
bone using guided tissue regeneration. Clin Oral Implants Res. 1990; 1: 22-30.
114
Buser D, Dula K, Hess D, Hirt H.P, Belser U.C. Localized ridge augmentation with
autografts and barrier membranes. In: Wikesjoe UME, Selvig KA (ed.): Periodontal
wound healing and regeneration. Periodontology 2000, 1999; 19: 151.
Buser D, Dula K, Hirt H.P, Schenk R.K. Lateral ridge augmentation using autografts
and barrier membranes. A clinical study in 40 partially edentulous patients. J Oral
Maxillofac Surg 1996; 54: 420.
Buser D, Weingart D, Weber H.P. Localized ridge augmentation using guided bone
regeneration in deficient implant sites. In: Greenberg AM, Prein J (eds.):
Craniomaxillofacial Reconstructive and Corrective Bone Surgery: Principles of
internal fixation using the AO/ASIF technique. Springer Verlag, New York. 2002;
chapt 16: 155-163.
Canalis E, Economides A.N, Gazzerro E. Bone morphogenetic proteins, their
antagonists and the skeleton. Endocr Rev 2003; 24: 218-35.
Celil A.B, Guelcher S, Hollinger J.O, Miller M. Tissue engineering applications –
Bone. In Tissue engineering and artificial organs, edited by Joseph D. Bronzino.
Boca Raton: CRC/Taylor & Francis, 2006
Celil A.B, Guelcher S, Hollinger J.O, Miller M.J. Tissue engineering applications Bone. In CRC Biomedical Engineering Handbook. New York, NY: CRC. 2006.
p5001-5022.
Chemical Land 21. Retrieved 10 March 2007 from:
http://www.chemicalland21.com/industrialchem/inorganic/TRICALCIUM%20PHOSP
HATE.htm
Chenu C, Delmas P.D. Physiology of bone remodeling. Advances in organ
biology1998; 5 (1): 45-64.
Cheong J.J. Mechanical properties of polycaprolactone-based scaffolds and its
deterioration studies. B.Eng Thesis. National University of Singapore. 2006.
Chim H, Hutmacher D.W, Chou A.M, Oliveira A.L, Reis R.L, Lim T.C, Schantz J.T. A
comparative analysis of scaffold material modifications for load-bearing applications
in bone tissue engineering. Int. J. Oral Maxillofac. Surg. 2006; 35: 928–934.
Coombes A.G.A, Rizzi S.C, Williamson M, Barralet J.E, Downes S, Wallace W.A.
Precipitation casting of polycaprolactone for applications in tissue engineering and
drug delivery. Biomaterials 2004; 25: 315-325.
Dahlin C, Gottlow J, Linde A, et al. Healing of maxillary and mandibular bone defects
using a membrane technique. Stand J Plast Reconstr Hand Surg 1990; 24: 13.
115
Dahlin C, Hansson H.A, Linde A. Expression of growth factors during healing of rat
mandibular trephine lesions treated by the osteopromotive membrane technique. In:
Dahlin C. Osteopromotion. Thesis. University of Göteborg, Sweden. 1993. p51-63.
Dahlin C, Linde A, Gottlow J, et al. Healing of bone defects by guided tissue
regeneration. Plast Reconstr Surg 1988; 81: 672.
de Groot K. Ceramics of calcium phosphates: preparation and properties. In
Bioceramics of calcium phosphates (de Groot K, ed.). CRC Press Inc 1983; pp. 100111.
Den Hollander, W, Patka, P, et al. Macroporous calcium phosphate ceramics for
bone substitution: a tracer study on biodegradation with 45Ca tracer. Biomaterials
1991; 12(6): 569-573.
Elliot J.C. Structure and Chemistry of the
Orthophosphates. Amsterdam : Elsevier. 1994.
Apatites
and
other
Calcium
Endres M, Hutmacher D.W, Salgado A.J, Kaps C, Ringe J, Reis R.L, Sittinger M,
Brandwood A, Schantz J.T. Osteogenic induction of human bone marrow derived
mesenchymal progenitor cells in novel synthetic polymer hydrogel matrices. TE
2003; 9: 689-701.
Ferrer M.C. Development and characterisation of completely degradable composite
tissue engineering scaffolds. Ph.D Thesis. Barcelona. 2007.
Fleisch H. Bisphosphonates in Bone Disease. From the Laboratory to the Patient.
New York: Academic Press. 2000.
Gan Z, Fung J.T, Jing X, Wu C, Kuliche W.K. A novel laser light-scattering study of
enzymatic biodegradation of poly (ε-caprolactone) nanoparticles. Polymer 1999; 40:
1961-1967.
Gan Z, Liang Q, Zhang J, Jing X. Enzymatic degradation of poly(ε-caprolactone) film
in phosphate buffer solution containing lipases. Polymer Degradation and Stability
1997; 56 (2): 209-213.
Gan Z, Yu D, Zhong Z, Liang Q, Jing X. Enzymatic degradation of poly(εcaprolactone)/poly(DL-lactide) blends in phosphate buffer solution. Polymer 1999;
40: 2859-2862.
Glimcher M.J. Mechanism of calcification: role of collagen fibrils and collagen
phosphoprotein complexes in vitro and in vivo. Anat Rec 1989; 224(2): 139-153.
Glimcher M.J. The nature of the mineral phase in bone: Biological and clinical
implications. In: Avioli LV, Krane SM, editors. Metabolic Bone Disease and Clinically
Related Disorders. St. Louis: Academic Press. 1998. p23-50.
116
Ha J.H, Kim S.H, and Han S.Y. Albumin release from bioerodible hydrogels based
on semi-interpenetrating polymer networks composed of poly(ε-caprolactone) and
poly(ethylene glycol) macromer. J. Control Release 1997; 49: 253–262.
He F, Li S, Vert M, Zhuo R. Enzyme-catalyzed polymerization and degradation of
copolymers prepared from e-caprolactone and poly(ethylene glycol). Polymer 2003;
44: 5145-5151.
Hedberg E.L, Kroese-Deutman H.C, Shih C.K, Crowther R.S, Carney D.H, Mikos
A.G, Jansen J.A. In vivo degradation of porous poly(propylene fumarate)/poly(DLlactic-co-glycolic acid) composite scaffolds. Biomaterials 2005; 26: 4616-4623.
Hedberg E.L, Shih C.K, Lemoine J.J, Timmer M.D, Liebschner M.A, Jansen J.A, and
Mikos A.G, In Vitro Degradation of Porous Poly(propylene fumarate)/Poly(DL-lacticco-glycolic acid) Composite Scaffolds, Biomaterials 2005; 26: 3215-3225.
Horch H-H, Pautke C. Regeneration instead of reparation. Mund Kiefer Gesichtschir
2006;10: 213.
Hu J, Liu X, Ma P.X. Biomineralization and bone regeneration. In: Anthony Atala,
Robert Lanza, James A. Thomson, and Robert M. Nerem (Editors) Principles of
Regenerative Medicine. Elsevier. 2007. p744-755.
Hutmacher D, Woodfield T, Dalton P.D, Lewis J.A. Scaffold design and fabrication in
tissue engineering. C Van Blitterswijk, J de Bruijn, R Cancedda, P Thomsen, J
Hubbell, D Williams, A Lindahl, and J Sohier (Ed.) Elsevier. 2008. p403-450.
Hutmacher D.W, Garcia A.J. Scaffold-based bone engineering by using genetically
modified cells. Gene 2005; 347: 1-10.
Hutmacher D.W, Schantz T, Zein I, Ng K.W, Teoh S.H, Tan K.C. Mechanical
properties and cell culture response of polycaprolactone scaffolds designed and
fabricated via fused deposition modeling. J. Biomed Mater Res 2001; 55(2): 203–16.
Hutmacher D.W, Sittinger M. and Risbud M.V. Scaffold-based tissue engineering:
rationale for computer-aided design and solid free-form fabrication systems. Trends
Biotechnol 2004; 22: 354 – 362.
Hutmacher D.W. Scaffolds in tissue engineering bone and cartilage. Biomaterials
2000; 21(24): 2529–43.
Jemt T, Patterson P. A 3-year follow up study on single implant treatment. J Dent.
1993; 21: 203-208.
Jeong J.C, Lee J, Cho K. Effects of crystalline microstructure on drug release
behavior of poly (ε-caprolactone) microspheres. J Control Release 2003; 92: 249258.
117
Jeong S.I, Kim B.S, Kang S.W, Kwon J.H, Lee Y.M, Kim S.H, Kim Y.H. In vivo
biocompatibility and degradation behavior of elastic poly(L-lactide-co-ε-caprolactone)
scaffolds. Biomaterials 2004; 25: 5939-5946.
Kartsogiannis V. and Ng K.W. Cell lines and primary cell cultures in the study of
bone cell biology. Mol Cell Endocrinol 2004; 228 (1-2): 79-102.
Kostopoulos L, Karring T. Augmentation of the rat mandible using guided tissue
regeneration. Clin Oral Implants Res 1994; 5: 75-84.
Lai W, Garino J, et al. Excretion of resorption products from bioactive glass
implanted in rabbit muscle. J Biomed Mater Res Part A 2005; 75A(2): 398-407.
LeGeros R. Calcium Phosphates in oral biology and medicine. San Francisco : Calif,
Karger. 1991.
LeGeros R.Z. Properties of osteoconductive biomaterials: calcium phosphates. Clin
Orthop Relat Res 2002; (395): 81-98.
Lei Y, Rai B, Ho K.H, Teoh S.H. In vitro degradation of novel bioactive
polycaprolactone-20% tricalcium phosphate composite scaffolds for bone
engineering. Materials Science and Engineering C 2006. Article in press.
Lowry K.J, Hamson K.R, Peng L.B, Calaluce R, Evans M.L, Anglen J.O, Allen W.C.
Polycaprolactone/glass bioabsorbable implant in a rabbit humerus fracture model. J
Biomed Mater Res 1997; 36: 536-541.
Lyford R.H, Mills M.P, Knapp C.I, Scheyer E.T, Mellonig J.T. Clinical evaluation of
freeze-dried block allografts for alveolar ridge augmentation: A case series. The
International Journal of Periodontics & Restorative Dentistry 2003; 23(5): 417-425.
Mano J.F, Sousa R.A, Boesel L.F, Neves N.M, Reis R.L. Bioinert, biodegradable and
injectable polymeric matrix composites for hard tissue replacement: state of the art
and recent developments. Composites Science and Technology 2004; 64: 789-817.
Mirtchi A.A, Lemaitre J, et al. Calcium phosphate cements: study of the betatricalcium phosphate-monocalcium phosphate system. Biomaterials 1989; 10 (7):
475 – 480.
Mukai K, Yamada K, Doi Y. Kinetics and mechanism of heterogeneous hydrolysis of
poly[(R)-3-hydroxybutyrate] film by PHA depolymerases. Int. J. Biol. Macromol.
1993; 15: 361-366.
Owens K.W, Yukna R.A. Collagen membrane resorption in dogs: a comparative
study. Implant Dent. 2001; 10: 49-50.
Park G.E, Pattison M.A, Park K, Webster T.J. Accelerated chondrocyte functions on
NaOH-treated PLGA scaffolds. Biomaterials 2005; 26: 3075-3082.
118
Perrin D.E and English J.P. Handbook of Biodegradable Polymers. In Florence T
Alexander, Gregoriadis Gregory (eds) Drug targeting and Delivery, UK University of
London. 1997. Pp 63-67.
Pilliar R.M, Filiaggi M.J, et al. Porous calcium polyphosphate scaffolds for bone
substitute applications – in vitro characterization. Biomaterials 2001; 22 (9): 963 –
972.
Pitaru S, Tal H, Soldinger M, Grosskopf A, Noff M. Partial regeneration of
periodontal tissue using collagen barriers. Initial observation in the canine. J
Periodontol 1988; 59: 380-6.
Pitt C.G, Chasalow F.I, Hibionada Y.M, Klimas D.M, Schindler A. Aliphatic polyesters
1. The degradation of poly (ε-caprolactone) in vivo. J Appl Polym Sci 1981; 26: 3779.
Pollok J.M, Vacanti J.P. Tissue Engineering. Seminars in pediatric surgery 1996; 5
(3): 191-6.
Putnam A.J, Mooney D.J. Tissue engineering using synthetic extracellular matrices.
Nature Medicine 1996; 2 (7): 824-826.
Radin S.R. and Ducheyne P. Effect of bioactive ceramic composition and structure
on in vitro behavior. III. Porous versus dense ceramics. J Biomed Mater Res 1994;
28(11): 1303-1309.
Rai B, Teoh S.H, Ho K.H, Hutmacher D.W, Cao T, Chen F, Yacob K. The effect of
rhBMP-2 onto 3D bioactive polycaprolactone scaffolds. Biomaterials 2004; 25: 54995506.
Rai B, Teoh S.H, Hutmacher D.W, Cao T, Ho K.H. Novel PCL-based honeycomb
scaffolds as drug delivery systems for rhBMP-2. Biomaterials 2005; 26: 3739-3748.
Rai B. The evaluation of bioactive polycaprolactone scaffolds as protein delivery
systems for bone engineering applications. Ph.D Thesis 2006, National University of
Singapore.
Raynaud S, Champion E, et al. Dynamic fatigue and degradation in solution of
hydroxyapatite ceramics. J Mater Sci Mater Med 1998; 9 (4): 221 – 227.
Reddi A.H. Bone morphogenetic proteins: an unconventional approach to isolation of
first mammalian morphogens. Cytokine growth Factor Rev 1997; 8: 11-20.
Rezwan K, Chen Q.Z, Blaker J.J, Boccaccini A.R. Biodegradable and bioactive
porous polymer/inorganic composite scaffolds for bone tissue engineering.
Biomaterials 2006; 27: 3413–31.
Rho J.Y, Kuhn-Spearing L, Zioupos P. Mechanical properties and the hierarchical
structure of bone. Med Eng Phys 1998; 20(2): 92-102.
119
Rominger J.W, Triplett R.G. The use of guided tissue regeneration to improve
implant osseointegration. J Oral Maxillofac Surg 1994; 52: 106-112.
Schantz J.T, Hutmacher D.W, Ng K.W, Teoh S.H, Chim H, Lim T.C. Induction of
ectopic bone formation by using human periosteal cells in combination with a novel
scaffold technology. Cell Transplant 2002; 11: 125-138.
Schantz J.T, Teoh S.H, Lim T.C, Endres M, Lam X.F.C, Hutmacher D.W. Repair of
calvarial defects with customized tissue-engineered bone grafts I. Evaluation of
osteogenesis in a three-dimensional culture system. Tissue Engineering 2003; 9:
113-126 (Suppl 1).
Schenk R.K, Buser D, Hardwick W.R, Dahlin C. Healing pattern of bone
regeneration in membrane-protected defects: A histologic study in the canine
mandible. Int J Oral Maxillofac Implants 1994; 9: 13-29.
Schliephake H, Aleyt J. Mandibular onlay grafting using prefabricated bone grafts
with primary implant placement: An experimental study in minipigs. Int J Oral
Maxillofac Implants 1998; 13: 384–393.
Schliephake H, Neukam F.W, Hutmacher D, Becker J. Enhancement of bone
ingrowth into a porous HA-matrix using a resorbable polylactic membrane. J Oral
Maxillofaci Surg 1994; 52: 57-63.
Schuckert K.H, Jopp S, Teoh S.H. Mandibular defect reconstruction using threedimensional polycaprolactone scaffold in combination with platelet-rich plasma and
recombinant human bone morphogenetic protein-2: De novo synthesis of bone in a
single case. Tissue Eng Part A. Mar 2009; 15(3): 493-499.
Seibert J, Nyman S. Localized ridge augmentation in dogs: A pilot study using
membranes and hydroxylapatite. J Periodonto 1990; 161: 157.
Teoh S.H. Engineering materials for biomedical applications. World Scientific
Publishing. Co. Pte. Ltd. 2004.
Timmins M.R, Lenz R.W, Fuller R.C. Heterogeneous kinetics of the enzymatic
degradation of poly(β-hydroxyalkanoates). Polymer 1997; 38 (3): 551-562.
Urist M.R. Bone: formation by autoinduction. Science 1965; 150: 893-9.
U.S National Cancer Institute's Surveillance, Epidemiology and End Results (SEER)
Program. Retrieved November 2009 from:
http://training.seer.cancer.gov/anatomy/skeletal/tissue.html
Van Gaalen S, Kruyt M, Meijer G, Mistry A.S, Mikos A.G, Van den Beucken J,
Jansen J.A, de Groot K, Cancedda R, Olivo C, Yaszemski M.J, and Dhert W. Tissue
engineering of bone, in Tissue Engineering, C. van Blitterswijk, Ed., Elsevier
Academic Press, San Diego. 2008. p559-610.
120
von Arx T, Cochran D.L, Hermann J.S, Schenk R.K, Buser D. Lateral ridge
augmentation using different bone fillers and barrier membrane application. A
histologic and morphometric pilot study in the canine mandible. Clin Oral Impl Res
2001; 12: 260-269.
Wan Y, Yu A, Wu H, Wang Z, Wen D. Porous-conductive chitosan scaffolds for
tissue engineering II. In vitro and in vivo degradation, J. of Mat. Sci: Mat in Med
2005; Vol 16: 1017–1028.
Wang X, Bank R.A, TeKoppele J.M, Agrawal C.M. The role of collagen in
determining bone mechanical properties. J Orthop Res 2001; 19(6):1021-1026.
Wikimedia. Retrieved 8 March 2007 from:
http://upload.wikimedia.org/wikipedia/en/f/fc/Pcl_synthesis.png
Woodard S.C, Brewer P.S, Moatmed F, Schindler A, Pitt C.G. The intracellular
degradation of poly (ε-caprolactone). J Biomed Mater Res 1985; 19: 1437-1444.
Wu L, Ding J. In vitro degradation of three-dimensional porous poly(D,L-lactide-coglycolide) scaffolds for tissue engineering. Biomaterials 2004; Vol 25: 5821 – 5830.
Wu W.J. and Nancollas G.H. The dissolution and growth of sparingly soluble
inorganic salts: A kinetics and surface energy approach. Pure and Applied Chemistry
1998; 70(10): 1867-1872.
Yamaguchi A, Komochi T, Suda T. Regulation of osteoblast differentiation mediated
by bone morphogenetic proteins, hedgehogs and Cbfa1. Endocr Rev 2000; 21: 393411.
Yeo A, Rai B, Sju E, Cheong J.J, Teoh S.H. The degradation profile of novel,
bioresorbable PCL-TCP scaffolds: An in vitro and in vivo study. J. Biomedical
Materials Research: Part A. Accepted 2007.
Zein I, Hutmacher D.W, Tan K.C, Teoh S.H. Fused deposition modeling of novel
scaffold architectures for tissue engineering applications. Biomaterials 2002; 23(4):
1169–85.
Zerbo I.R, Bronckers A.L.J.J, et al. Localisation of osteogenic and osteoclastic cells
in porous β-tricalcium phosphate particles used for human maxillary sinus floor
elevation. Biomaterials 2005; 26(12): 1445-1451.
Zhao S, Pinholt E.M, Madsen J.E, Donath K. Histological evaluation of different
biodegradable and non-biodegradable membranes implanted subcutaneously in
rats. J Craniomaxillofac Surg 2000; 28: 116-122.
Zhou Y, Chen F, Ho S.T, Woodruff M.A, Lim T.M, Hutmacher D.W. Combined
marrow stromal cell-sheet techniques and high-strength biodegradable composite
scaffolds for engineered functional bone grafts. Biomaterials 2007; 28: 814-824.
121
[...]... an initial stage of a larger project, in order to develop a scaffold of a higher porosity that allows for a more rapid degradation whilst maintaining favourable mechanical properties A final porosity of about 85% was targeted for 1.3.2 Part 2: Optimization of native and customized scaffolds in vitro and their effects in initial bone healing (in Chapter 4) In the second part of the study, PCL-TCP scaffolds. .. channels Forming the outer wall of bones, it bears most of the supportive and protective function of the skeleton Cancellous bone, on the other hand, makes up the remaining 20% of bone mass in the body It consists of trabeculae which form an interconnected lattice Cancellous bone can be found in vertebrae, fracture joints, ends of long bones and in foetuses The whole structure, an outer cortical sheath and. .. of 85% by treating them with 3M NaOH or 0.1% lipase-PBS medium under physiological conditions for up to 108 hours 2 To compare the degradation profile of treated and untreated PCL-TCP scaffolds in vitro when immersed in standard culture medium for up to 24 weeks, and in vivo when implanted in the subcutaneous back of rats for 24 weeks (6 months) 3 To evaluate the rate and extent of bone formation of. .. in bone healing as they could ruled out the need for harvesting grafts x LIST OF TABLES Table 3.1 Mw, Mn, and PDI of NaOH-treated and lipase-treated PCL-TCP Scaffolds 40 Table 4.1 Mw, Mn, and PDI of native, NaOH-treated, and lipase-treated PCL-TCP Scaffolds in vitro 66 Table 4.2 Mw, Mn, and PDI of native, NaOH-treated, and lipase-treated PCL-TCP Scaffolds in vivo 75 Table 5.1 Number of sites with soft... alive, and contains cells which work continuously to regenerate and repair it 9 [Bronner, 1999; Ferrer, 2007] Bone tissue contains five basic types of bone cells: osteogenic cells, osteoblasts, osteocytes, osteoclasts, and bone- lining cells Osteogenic cells respond to traumas, such as fractures, and begin the healing process immediately by giving rise to bone- forming cells (osteoblasts) and bonedestroying... polymers in the field of tissue engineering has been widely investigated in recent years, with advances in the scaffold technology extending their usage to clinical applications such as bone regeneration In particular of such interest is poly(ε-caprolactone) -tricalcium phosphate (PCL-TCP) composite scaffold, a synthetic biodegradable polymer frequently investigated for bone tissue engineering applications... engineering 2.2.1 Degradation of PCL polymer Degradation behaviours of scaffolds play an essential role in the engineering of new tissue, as the rate of degradation is intrinsically linked to many cellular processes including cell viability, tissue growth, as well as the host response [Lei, 2006] Once implanted in the body, a porous scaffold should maintain its mechanical properties and structural integrity... assembly of collagen fibrils and fibers and bone mineral crystals [Rho, 1998] Bone s function is both biomechanical and metabolic Biomechanically, bone acts to: (1) maintain the shape of the skeleton, (2) protect soft tissues in the cranial, thoracic and pelvic cavities, (3) transmit the forces of muscular contraction during movement, and (4) supply a framework for bone marrow Metabolically, bone (1)... regarding bone tissue engineering strategy and the application in implant dentistry, as well as the current drawback of PCL-TCP scaffolds in dentoalveolar defects treatment that lead the author to pursue this research Detailed research objectives and research scope are discussed in the next and last sections respectively 1.1.1 Bone tissue engineering Loss of human tissues or organs is a devastating problem... role of scaffolds come into the picture as they may eliminate the need for an extensive bone harvesting procedure from a donor site However in facing a complex biological system as the human body, the requirements of scaffold materials for bone tissue engineering in dentoalveolar application can be extremely challenging 3 1.1.3 PCL-TCP scaffolds: Current drawback The use of synthetic polymers in the ... submitted for the degree of Master of Engineering in the Department of Mechanical Engineering at the National University of Singapore under the supervision of Professor Teoh Swee Hin and Dr Alvin Yeo... secrete bone tissue and form the tissue around itself like a protective wall of bone tissue They are responsible for the maintenance of healthy bone by secreting enzymes and directing the bone mineral... CHAPTER 4: OPTIMIZATION OF NATIVE AND CUSTOMIZED SCAFFOLDS IN VITRO AND THEIR EFFECTS IN INITIAL BONE HEALING 4.1 INTRODUCTION 44 4.1.1 In vitro degradation study 44 4.1.2 In vivo degradation study
Ngày đăng: 09/10/2015, 11:18
Xem thêm: In vitro and in vivo evaluation of customized polycaprolactone tricalcium phosphate scaffolds for bone tissue engineering, In vitro and in vivo evaluation of customized polycaprolactone tricalcium phosphate scaffolds for bone tissue engineering
