characterization of a biodegradable electrospun - T-Space

CHARACTERIZATION OF A BIODEGRADABLE ELECTROSPUN POLYURETHANE NANOFIBER SCAFFOLD SUITABLE FOR

ANNULUS FIBROSUS TISSUE ENGINEERING

by

Masoud Yeganegi

A thesis submitted in conformity with the requirements for the degree of Masters of Applied Science and Engineering

Department of Materials Science and Engineering and Institute of Biomaterials and Biomedical Engineering

University of Toronto

© Copyright by Masoud Yeganegi 2009

ii

Characterization of a Biodegradable Electrospun Polyurethane Nanofiber Scaffold Suitable for Annulus Fibrosus Tissue Engineering

Masoud Yeganegi

Masters of Applied Science and Engineering

Department of Materials Science and Engineering

and Institute of Biomaterials and Biomedical Engineering University of Toronto

2009

I: Abstract

The current study characterizes the mechanical and biodegradation properties of a

polycarbonate polyurethane (PU) electrospun nanofiber scaffold intended for use in the growth

of a tissue engineered annulus fibrosus (AF) intervertebral disc component. Both the tensile

strength and initial modulus of aligned scaffolds were higher than those of random scaffolds and

remained unaffected during a 4 week biodegradation study, suggesting a surface-mediated

degradation mechanism. The resulting degradation products were non-toxic. Confined

compressive mechanical force of 1kPa, was applied at 1Hz to in vitro bovine AF tissue grown on

the scaffolds to investigate the influence of mechanical force on AF tissue production, which was

found to decrease significantly at 72 hours relative to 24 hours, independent of any effects from

mechanical forces. Overall, the consistent rate of PU degradation, along with mechanical

properties comparable to those of native AF tissue, and the absence of cytotoxic effects, make

this polymer suitable for further investigation for use in tissue-engineering the AF.

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II. Acknowledgements

I would like to thank Dr. Jian Wang, Dr. Liu Yang, Dr. Meilin Yang, Dr. Amritha De Croos,

Douglas Holmyard, and Robert Temkin for their guidance throughout the various stages of this

Masters work. With the members of the Santerre and Kandel lab, I have enjoyed years of

cooperative team effort and will hopefully share many more years of continued friendship.

Most importantly, I’d like to take thank my dedicated mentors, Dr. Rita Kandel and Prof. Paul

Santerre. Their excellence in their respective fields, commitment to research, and attention to

detail has forged a standard in my mind that I will forever be pursuing. This unwavering

commitment has been balanced by their understanding toward several tragedies in my life in the

past few years. Their invaluable academic, professional and personal guidance will be greatly

missed. Thank you.

I’d like to dedicate this work to my dad, my sisters, and my mom, whom I recently came so

close to losing. You have encouraged me to achieve my best, and helped guide me toward that

goal. I will forever be grateful.

Masoud Yeganegi

Spring 2009

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II. Table of Contents

I: Abstract ..................................................................................................................................... ii

II. Acknowledgements ................................................................................................................ iii

II. Table of Contents ................................................................................................................... iv

III. Table of Figures ................................................................................................................... vii

IV. Table of Statistics................................................................................................................. xii

VI: Introduction ............................................................................................................................1

A. Annulus Fibrosus: ...............................................................................................................4

B. Mechanical Properties: .......................................................................................................4

C. Mechanobiology of Annulus Fibrosus: .............................................................................5

D. Degenerative Disc Disease: ...............................................................................................10

E. Tissue Engineering using Biodegradable Polymers: ......................................................12

F. Electrospinning: .................................................................................................................14

G. Proposed Work: ................................................................................................................15

H. References: .........................................................................................................................17

VII: Characterization of a Biodegradable Electrospun Polyurethane Nanofiber Scaffold: Mechanical Properties and Cytotoxicity .............................................................30

A. Introduction .......................................................................................................................32

B. Experimental Section ........................................................................................................34

1. Materials ........................................................................................................................34

2. Biodegradation Study ...................................................................................................36

3. Assessment of Mechanical Properties ..........................................................................36

4. Differential Scanning Calorimetry...............................................................................37

5. Evaluation of Degradation Products............................................................................37

6. Cytotoxicity Study .........................................................................................................38

7. Statistical analysis .........................................................................................................39

C. Results and Discussion ......................................................................................................39

D. Conclusion .........................................................................................................................43

E. Acknowledgements ............................................................................................................44

F. References: .........................................................................................................................45

G. Figures ................................................................................................................................51

v

VIII: Application of Dynamic Compressive Forces on Annulus Fibrosus Cells Grown on a Biodegradable Electrospun Nanofiber Scaffold ............................................61

A. Introduction .......................................................................................................................63

B. Experimental Section ........................................................................................................67

1. Scaffold fabrication ......................................................................................................67

2. Annulus fibrosus cell culture .......................................................................................68

3. Mechanical Stimulation ...............................................................................................68

4. AF cell morphology ......................................................................................................69

5. DNA content ..................................................................................................................70

6. Quantification of Proteoglycan and Collagen Synthesis and Proliferation ...............70

7. Statistical analysis .........................................................................................................71

C. Results ................................................................................................................................71

1. Effect of Mechanical Stimulation on AF Cells ...........................................................71

2. Effect of Mechanical Stimulation on AF Matrix Synthesis ........................................71

D. Discussion...........................................................................................................................72

E. Conclusions ........................................................................................................................73

F. Acknowledgements ............................................................................................................73

G. References: .........................................................................................................................75

H. Figures ................................................................................................................................83

IX: Conclusions and Future Work ............................................................................................90

Appendix A: Scaffold Preparation ............................................................................................95

A. Electrospinning .................................................................................................................95

1. Preparation of nanofibrous polycarbonate urethane scaffolds (random scaffolds) ......95

2. Preparation of nanofibrous polycarbonate urethane scaffolds (aligned scaffolds) ......96

3. Humidity Effects .............................................................................................................97

Appendix B: Biodegradation ...................................................................................................102

A. Reagent Preparation:........................................................................................................102

B. Standard Curve for CE Enzyme Activity ...........................................................................102

C. Calculating CE Activity .....................................................................................................103

D. Scaffold Thickness .............................................................................................................104

Appendix C: Mechanical Testing ............................................................................................105

Appendix D: Annulus Fibrosus Tissue Culture .....................................................................110

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A. Optimization of protocol for cell seeding on scaffolds: ....................................................112

Appendix E: Cytotoxicity Evaluation .....................................................................................116

A. Cytotoxic evaluation of degradation products was performed using the MTT and Live/Dead Assays: ...........................................................................................................116

1. MTT Assay ...................................................................................................................116

2. Live/Dead Assay ..........................................................................................................117

3. Live / Dead Assay Images: ...........................................................................................119

Appendix F: Statistics Tables ..................................................................................................124

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III. Table of Figures

Introduction

Figure 1. The anatomy of the human vertebral column and the intervertebral disc. (DePuy Spine) ......................................................................................................... 2

Figure 2. Mechanical loading of the disc results in a complex set of physical changes that may be transduced as mechanical stimuli to the cells. 4 .................................. 3

Figure 3. Current lumbar disc prostheses. SB Charite´ III (A). Prodisc II (B). Maverick (C). 67 .................................................................................................... 12

Figure 4. Stress induced strain along with material morphology and chemistry, can affect the environmental degradation of the material 84 ........................................ 14

Characterization of a Biodegradable Electrospun Polyurethane Nanofiber Scaffold: Mechanical Properties and Cytotoxicity

Figure 1. Scanning Electron Microscopy Images of aligned (a) and random (b) electrospun Polycarbonate Urethane Nanofiber Scaffolds. (Solution: 16 wt.% PU, injection rate: 0.5 ml/hr, potential difference: 18 kilovolts) ................. 51

Figure 2. Determination of Cholesterol Esterase (CE) Half Life. It was determined that the half life of CE in the presence of the aligned PU scaffolds was approximately 12 hours. Thus CE was added daily to adjust the enzyme activity (n = 3). ...................................................................................................... 52

Figure 3. Cumulative absolute mass loss (a) and cumulative relative mass loss (b) during biodegradation. Scaffolds were incubated in 100 units/ml CE over 4 weeks. Data are reported as mean ± standard error (n=6). (*) Absolute mass loss was found to increase significantly at every week (p<0.05) for all groups, while no statistical differences were observed between the scaffold groups within each week. Relative mass loss was found to increase significantly in the case of aligned scaffolds. ....................................................... 53

Figure 4. (a) Elastic Modulus and (b) Tensile Strength of the electrospun polyurethane nanofiber scaffolds following the pre-wetting (for one week in pH 7.0 PBS at 37ºC) and drying process, comparing non-ADO vs. ADO, as well as aligned vs. random scaffolds. Data are reported as mean ± standard error (n=6). ............................................................................................................ 54

Figure 5. Differential Scanning Calorimetry for as-made and pre-wet/dried samples for (a) aligned PU, (b) aligned PU+0.5%ADO and (c) random PU+0.5%ADO. Ti is the glass transition temperature for the polycarbonate soft segment. Tii indicates the onset of the soft segment melt phase. Tiii and

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Tiv are soft-segment melting transition temperatures. Tv indicates the hard-segment melting transition temperature. ............................................................... 55

Figure 6. a) Initial Modulus and (b) Tensile Strength of the electrospun polyurethane nanofiber scaffolds over four weeks of biodegradation in CE (100 units/ml) at 37ºC, PBS pH=7.0. Data are reported as mean ± standard error (n=6). Aligned scaffolds showed significantly higher modulus than random scaffolds at all time points. Ultimate stress of aligned polymers decreased in the first week of degradation, but remained stable thereafter. .............................. 56

Figure 7. Transmission Electron Microscopy of a non-soluble degradation product .......... 57

Figure 8. Assessing the cytotoxicity of PU degradation products: MTT Assay was used to evaluate potential cytotoxicity of various concentrations of (a) non-soluble and (b) soluble degradation products on bovine annulus fibrosus cells. The experiment was repeated 4 times (n=8 per condition). Data are expressed as mean ± SEM. H2O2 was used as a positive control. ........................ 58

Figure 9. Cell viability of PU degradation products: AF cells were incubated for 24 hours with various concentrations of (a) non-soluble and (b) soluble degradation products. Live/Dead Assay was used to assess cell viability. The number of dead cells were counted and expressed as percent of total number of cells. The experiment was repeated 4 times (n=8 per condition) and data expressed as mean±SEM. H2O2 was used as a positive control. ............ 59

Figure 10. Representatipve images of Live/Dead Assay of AF cells treated with (a) untreated negative control (media with carrier); (b) H2O2-treated positive control; (c) 0.1 wt. % non-soluble degradation products; (d) 100 volume% soluble degradation products ................................................................................ 60

Application of Dynamic Compressive Forces on Annulus Fibrosus Cells Grown on a Biodegradable Electrospun Nanofiber Scaffold

Figure 1. Scanning Electron Microscopy Images of Aligned (a) and Random (b) Electrospun Polycarbonate Urethane Nanofiber Scaffolds ................................... 83

Figure 2. Mechanical Stimulation Apparatus: The polymer is held in place over the porous titanium base by Tygon tubing, which in turn allows seeding media to remain in contact with the scaffold during cell seeding. At the time of mechanical stimulation, an agarose plug is placed above the cells to transmit the force through to the cells. ................................................................................ 84

Figure 3. SEM images of AF cells immediately (b), 6 hr (d); 24 hr (f); 72 hr (h) post-stimulation. The corresponding non-stimulated controls are denoted (a, c, e, g). Stimulated samples are more spread than control samples, particularly at early time points. ................................................................................................... 86

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Figure 4. DNA Content (µg) at various time points following mechanical stimulation (3 day Tissue, stimulated for 1hr at 1Hz and 1kPa). No significant changes were observed between the control and stimulated groups nor between the different time points (n=3, α=0.05). ...................................................................... 87

Figure 5. Evaluation of DNA content (a) and Thymidine Incorporation (b) at 24 hours and 72 hours post-stimulation. Controls were treated similarly but were not stimulated. The results are expressed as ± SEM (n=15, α=0.05).. No significant differences were detected between the two conditions. ...................... 88

Figure 6. Collagen (a) and Proteoglycan (b) synthesis at 24 hours and 72 hours post-stimulation. No significant changes were observed between the control and stimulated groups at either time point. However, there is a significant decrease in matrix accumulation, by 72 hours compared to 24 hours post-stimulation, for the combined group of stimulated and non-stimulated samples, (N=15, α=0.05). ..................................................................................... 89

Appendices

Figure A.1. Various instrumental apparatus (A to C) constructed to attempt to control humidity using a dehumidifier (full and partial air flow into a close/open enclosure) with corresponding SEM images of the resultant scaffolds. The effects of air current and increase in temperature (due to the heat carried from the pump by the dehumidified air), scaffold alignment and fiber diameter was found to be inferior under all conditions. ....................................... 98

Figure A.2. Final apparatus for regulating humidity to 30% R.H. The effects of air current were reduced by introducing a porous wooden base on which the electrospinning apparatus was placed. Further, a cooling reservoir was used to cool the dehumidified air from 50 ºC to 25ºC. ................................................. 99

Figure A.3. The humidity profile using the final equipment set-up. Humidity remains stabilized at the desired level (<30% R.H.) after approximately 10-20 minutes of starting the dehumidification. ........................................................... 100

Figure A.4. Scanning electron microscopy indicating processed fiber dimension, their alignment, and confirming that transverse fibers were not a significant occurrence ........................................................................................................... 101

Figure B.1. Changes in the thickness of scaffolds throughout the biodegradation process ... 104

Figure C.1. Weekly tensile testing of scaffolds: Each polymer sample was clamped on either side and tested using an Instron® model 8501 under a tensile strain of 10 mm/min to the breaking point. ....................................................................... 106

Figure C.2. Stress-strain curves for PU aligned scaffolds under tensile mechanical stress. The various curves are repeats of the same sample group. The initial

x

modulus was found by calculating the slope of the stress-strain curve in the initial elastic portion of each curve. Ultimate stress was also reported on. The ultimate strain is defined by the strain experienced by the sample at the ultimate stress...................................................................................................... 107

Figure C.3. Stress-strain curves for PU + 0.5% ADO aligned scaffolds under tensile mechanical stress: The various curves are repeats of the same sample group. The initial modulus was found by calculating the slope of the stress-strain curve in the initial elastic portion of each curve. Ultimate stress was also reported on. The ultimate strain is defined by the strain experienced by the sample at the ultimate stress. .............................................................................. 108

Figure C.4. Stress-strain curves for PU + 0.5% ADO random scaffolds under tensile mechanical stress: The various curves are repeats of the same sample group. The initial modulus was found by calculating the slope of the stress-strain curve in the initial elastic portion of each curve. Ultimate stress was also reported on. The ultimate strain is defined by the strain experienced by the sample at the ultimate stress. .............................................................................. 109

Figure D.1. Multiple discs were dissected from a single tail and the isolated AF cells were combined to provide sufficient cells for an experiment and improve consistency. Only outer AF cells were used in all experiments. ........................ 111

Figure D.2. Methods evaluated (A to D): (A) Cell suspension on the polymer scaffold alone, (B) Cell suspension on scaffold, supported by an agarose gel base, (C) cell suspension confined by a Teflon insert, (D) cell confinement through the use of Tygon tubing; (E) The seeding method selected for all subsequent experiments which consisted of using a Tygon tubing and a porous titanium disc and (F) the corresponding apparatus for mechanical stimulation of tissue. ........................................................................................... 113

Figure D.3. SEM images at low (A) and higher magnification (B) showing scaffolds seeded at 0.8 million cells / cm2. This density produced cellular layers, where cell-cell contact dominated cell-polymer contact. It was therefore decided to reduce cell seeding density to 0.8 million cells / cm2 ....................... 114

Figure D.4. Percent cell attachment to determine optimal seeding density: DNA content was measured 24 hours after seeding. The attachment level dropped significantly at the highest seeding density. The lower seeding density of 0.8 million cells / cm2 was chosen for subsequent mechanical stimulation studies (N = 6 per condition). ............................................................................. 115

Figure E.1. MTT Optimization: Absorbance vs. Cell Number vs. Incubation Period .......... 118

Figure E.2. Representative images of Live/Dead assay of AF Cells incubated for 24 hours (37 ºC, 5% CO2): in either (A) F12 Ham’s Media containing 5% FBS (negative control); or (B) Ham’s F12 Media containing 0.01 wt% H2O2 (Positive Control) ................................................................................................ 119

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Figure E.3. Live/Dead Assay: Photomicrograph of AF cells subjected to Non-Soluble Degradation Products of PU aligned polymers at various concentrations [(A) 0.001 wt. %, (B) 0.005 wt. %, (C) 0.01 wt. %, (D) 0.025 wt. %, (E) 0.05 wt. %, or (F) 0.1 wt. % (g/100mL)] ............................................................ 120

Figure E.4. Live/Dead Assay: Photomicrograph of AF cells subjected to Non-Soluble Degradation Products of PU + 0.05% ADO aligned polymers at various concentrations [(A) 0.001 wt. %, (B) 0.005 wt. %, (C) 0.01 wt. %, (D) 0.025 wt. %, (E) 0.05 wt. %, or (F) 0.1 wt. % (g/100mL)] ........................................... 121

Figure E.5. Live/Dead Assay: Photomicrograph of AF cells subjected to Buffer Soluble Degradation Products of PU aligned polymers at various concentrations [(A) 20 %, (B) 40 %, (C) 50 %, (D) 60 %, (E) 80 %, (F) 100 % (percent by volume)] .............................................................................................................. 122

Figure E.6. Live/Dead Assay: Photomicrograph of AF cells subjected to Buffer Soluble Degradation Products of PU + 0.05% ADO aligned polymers at various concentrations [(A) 20 %, (B) 40 %, (C) 50 %, (D) 60 %, (E) 80 %, (F) 100 % (percent by volume)] ...................................................................................... 123

xii

IV. Table of Statistics

Introduction

Table 1 - Annulus fibrosus response to a selection of mechanobiological stimuli ................ 8

Appendices

Table F.1 - Cumulative Mass Loss Statistics: Week (Fig. 3, Ch. VII) .................................. 124

Table F.2 - Cumulative Mass Loss Statistics: Material Type (Overall) (Fig. 3, Ch. VII) ..... 124

Table F.3 - Cumulative Mass Loss Statistics: Material Type (in week 1, Fig. 3, Ch. VII) ... 124



Table F.6 - Cumulative Mass Loss Statistics: Material Type (in week 4, Fig. 3, Ch. VII) ...................................................................................................................... 125

Table F.7 - Initial Modulus: Material Type (within as-made, Fig. 4, Ch. VII) ..................... 125

Table F.8 - Initial Modulus: Material Type (within prewet, Fig. 4, Ch. VII) ........................ 125

Table F.9 - Initial Modulus: Material Type (within week 1, Fig. 6, Ch. VII) ....................... 125




Table F.13 - Ultimate Tensile Stress (within Aligned PU + 0.05% ADO, Fig. 6, Ch. VII) ... 126

Table F.14 - Ultimate Tensile Stress (within Aligned PU, Fig. 6, Ch. VII) ............................ 127

Table F.15 - Ultimate Tensile Stress (within Random PU + 0.05% ADO, Fig. 6, Ch. VII) ... 127

Table F.16 - DNA Content: (within groups: Stimulated and Control, Fig. 4, Ch. VIII) ......... 128

Table F.17 - DNA Content (within groups: 0hr, 6hr, 12hr, 24hr, Fig. 4, Ch. VIII) ................ 128

Table F.18 - Relative Thymidine Incorporation Ratio Comparison: 24hr vs 72hr (within groups: Stimulated and Control, Fig. 6, Ch. VIII) .............................................. 129

Table F.19 - Relative Thymidine Incorporation Ratio Comparison: Stimulated vs. Control (within groups: 24 hr and 72 hr, Fig. 6, Ch. VIII) ................................. 129

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Table F.20 - Relative Collagen Content Ratio Comparison: 24hr vs 72hr (within groups: Stimulated and Control, Fig. 6, Ch. VIII) ........................................................... 129

Table F.21 – Relative Collagen Content Ratio Comparison: Stimulated vs. Control (within groups: 24 hr and 72 hr, Fig. 6, Ch. VIII) .............................................. 129

Table F.22 - Relative Proteoglycan Content Ratio Comparison: 24hr vs 72hr (within groups: Stimulated and Control, Fig. 6, Ch. VIII) .............................................. 130

Table F.23 - Relative Proteoglycan Content Ratio Comparison: Stimulated vs. Control (within groups: 24 hr and 72 hr, Fig. 6, Ch. VIII) .............................................. 130

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VI: Introduction

2

II. Introduction

The human vertebral column provides axial support to the body and protects the spinal cord.

The intervertebral discs lying between the vertebrae provide flexibility and help prevent damage

to the vertebral column by dissipating mechanical loads and shocks.1 The hyaline cartilage

endplates found at each end, represent the anatomical limits of the disc. The nucleus pulposus, a

remnant of the embryonic notochord, contains few cartilage-like cells dispersed in the

proteoglycan-rich matrix and forms the gelatinous central zone of intervertebral discs. The

nucleus pulposus’ fluid nature allows it to deform under pressure transmitting any applied forces.

Surrounding the nucleus pulposus are about 20 sheets of concentric fibrocartilaginous lamellae

called the annulus fibrosus, which are formed from embryonic mesynchymal tissue.2 The

lamellae consist of fibers that are oriented at 60º to the vertical axis of the disc. The alignment of

these fibers alternates between successive lamellae and is of great importance to the stability of

the annulus fibrosus. The lamellae are not fully continuous, with 40-50% of lamellae failing to

completely circumscribe the nucleus pulposus. 2

Figure 1. The anatomy of the human vertebral column and the intervertebral disc. (DePuy

Spine)

Lumbar

Cervical

Thoracic

Vertebral Body

IntervertebralDisc

Annulus Fibrosus

NucleusPulposus

3

Both the annulus fibrosus and nucleus pulposus are responsible for weight-bearing functions.

The annulus fibrosus resists buckling under stress and is mechanically capable of sustaining

compressive stress independently of the nucleus pulposus. 3 However, the annulus is unable to

withstand prolonged compressive stress on its own, and the nucleus pulposus provides additional

mechanical stability. The nucleus pulposus and the inner annulus fibrosus transmit compressive

mechanical forces outwards resulting in tensile deformation of the outer annulus fibrosus and

preventing the buckling of its lamellae. Part of the compressive force is transmitted by the

nucleus from one vertebral body to the next, and lessening the load borne by the annulus. In a

healthy disc, compressive forces are balanced with minimum radial expansion on the part of the

nucleus.1 The nucleus pulposus helps to absorb shock, or rapid changes in stress. Any sudden

increases in compressive stress will take a longer time to propagate through the vertebral column

due to the presence of the disc, as changes in compressive stress are initially diverted to tensile

stain in the annulus. 3 By slowing the rate at which the applied force is transmitted in the

vertebral column, the intervertebral discs protect the vertebra. The cooperative action of nucleus

and annulus, allows the disc to withstand forces that would otherwise result in buckling/failure of

the disc. Because of the intricacies of the intervertebral disc mechanics, any biochemical changes

in the tissue have profound effects on the mechanical stability of the disc.

Figure 2. Mechanical loading of the disc results in a complex set of physical changes that

may be transduced as mechanical stimuli to the cells. 4

4

Annulus Fibrosus:

The outer annulus fibrosus (AF) consists of concentric lamella, mainly made up of collagen

fibers, and oriented at approximately 60º to the vertical. The AF extracellular matrix contains

collagen fibrils, proteoglycans and water. Water makes up 60% of the annulus fibrosus, while

collagen and proteoglycans account for 50-70% and 10-20% of the dry weight respectively. 5

The composition of the annulus fibrosus is radially non-uniform in that collagen type I is

restricted to the annulus fibrosus and not present in the healthy nucleus pulposus. Within the

annulus fibrosus, the concentration of collagen type I is highest in the outer lamellae and lowest

at the inner lamellae nearest the NP.6 Conversely, collagen type II which is the main collagen

type present in nucleus pulposus decreases in concentration radially towards the annulus

fibrosus.5 The relative proportion of collagen type I to collagen type II in the annulus fibrosus

varies from 70:30 in the innermost layers and 85:15 in the outer layers.5 Other types of collagen

also exist in smaller amounts in the annulus fibrosus, with Collagens V, VI, IX, XI, XII and XIV

all contributing to the matrix.7 The interlamellar space also contains proteoglycan aggregates,

with water imbibing properties similar to those found within the nucleus of the disc, while the

lamellar layers are comprised of proteoglycan monomers which interact with, and modulate the

behaviour of the collagen fibrils. 7

Mechanical Properties:

The mechanical properties of intervertebral discs are complex and the literature on this topic

shows significant variation.8-11 In addition, due to its structural arrangement, the mechanical

behavior of AF is relatively complex. Studies indicate that the annulus fibrosus exhibits both

matrix viscoelastic and biphasic viscoelastic behavior.11 Matrix viscoelastic behavior indicates

the intrinsic flow-independent viscoelastic properties of the solid extracellular matrix, whereas

5

the biphasic viscoelastic properties reflect time and rate-dependent effects due to fluid-solid

interactions in the tissue. The elastic modulus of single lamellae of annulus fibrosus has been

found to vary with radial position in the disc with values ranging from 5±4 MPa for posterior

inner AF, 20±12 MPa for posterior outer AF, 10±6 MPa for anterior inner AF, and 49±32 MPa

for anterior outer AF 12. The ultimate stress of single lamellae of annulus fibrosus similarly

varies radially from 0.9±0.3 MPa for posterior inner AF, 1.1±0.3 MPa for posterior outer AF,

0.9±0.7 MPa for anterior inner AF, and 3.3±1.3 MPa for anterior outer AF. 12 The AF

experiences a variety of forces in vivo, and thus the impact of mechanical forces on AF tissue is

of great interest. It has been well established that mechanical loading plays an important role in

regulating behavior in various tissues.13-17 Muscle forces, general loading of the joints and

movement of joints relative to each other act to apply a range of stresses on the disc, including

compressive, shear, tensile, osmotic and hydrostatic forces, which along with other forces,

initiate a response from the AF cells.18-26 A number of studies demonstrating the

mechanosensitivity of the AF are outlined below.

Mechanobiology of Annulus Fibrosus:

The in vivo biological response of annulus fibrosus in the rat tail to short-term dynamic

compression have been investigated by MacLean et al. The expression of anabolic and catabolic

genes was affected by a 2 hour dynamic compression of the intervertebral disc, under an applied

stress of 1 MPa (12.6N) at 0.2 Hz. Anabolic genes for collagen I and collagen II were

downregulated while catabolic genes for collagenase and aggrecanase were upregulated in the

annulus.27 Compressive stresses of 1 MPa and 0.2 MPa were applied in another in vivo study at

three frequencies of 0.01 Hz, 0.2 Hz, and 1 Hz. It was found that the application of 1 MPa at all

frequencies significantly increased catabolic genes for collagenase (21-, 7-, 8-fold respectively)

6

and aggrecanase (5-, 1-, 7-fold respectively) with only slightly increased collagen I expression at

1 Hz (3.5- fold). On the other hand stress of 0.2 MPa at 1 Hz resulted in a slightly elevated levels

of expression for collagen (4-fold) and aggrecan (2-fold), with minor non-significant increases in

collagenase and aggrecanase. The study demonstrated the frequency and magnitude dependence

of the biological response to compressive mechanical stress.18,28 Lotz and Walsh et al. have also

explored the load- and frequency-dependant response of the intervertebral disc to dynamic

stresses.19 Peak compressive stresses of 0.9 MPa and 1.3 MPa were applied at frequencies of 0.1

Hz and 0.01 Hz to in vivo mice tail discs. Under these conditions, little apoptosis (5%) was found

generally at the higher frequency and the lower stress, compared to 30% apoptosis at lower

frequencies. Aggrecan gene expression in the inner annulus increased under lower frequency and

higher stress loading. 19 A static compressive force of 1.0 MPa for 24 hr applied to ex-vivo

cocygeal discs caused apoptosis in the annulus fibrosus.29 Annulus fibrosus cells in an alginate

culture system subjected to 30 hours of static 25% unconfined compressive strain, responded

with increased gene expression for types I and II collagen, and aggrecan.30 Application of

hydrostatic pressure on caudal bovine and human IVD explants, while affecting the nucleus

pulposus and the inner AF, did not produced any significant changes in the outer AF.24,31,32

Osmotic pressure has also been shown to affect mRNA levels of aggrecan, collagen-I, and

collagen-II in AF 3D-cultures.25,33,34 In addition, cellular response to hydrostatic and cyclic

tensile strain was found to be dependent on the osmotic environment.25 Since the AF is

physiologically subjected to tensile forces, it is not surprising that it would respond to this type

of mechanical stimulation. Dynamic tensile strain (5% at 1Hz for 24 hours) of monolayer AF

cells grown on a collagen substrate resulted in decreased proteoglycan synthesis, while

increasing nitrogen oxide production.35 Higher tensile strains at lower frequencies (15% at 0.1Hz

for 24 hours) increased cellular apoptosis in the AF.36 However, in the case of nucleus pulposus,

7

higher tensile forces and lower frequencies (20% at 0.05Hz for 24 hours) produced higher

collagen synthesis and increased cellular proliferation. Axial traction tensile forces applied to

intact IVD explants (at 0.8MPa for 4 hours) resulted in a decrease in proteoglycan synthesis in

the AF.23 Mechano-biological response of AF cells to a collection of stimuli has been

summarized in Table 1 below.

8

Table 1 - Annulus fibrosus response to a selection of mechanobiological stimuli

Ref. Species Experiment

Setup Stress Type

Strain Value

Stress Value

Frequency Length Setup Notes

Effects

37 Porcine Lumbar (4-5 months)

In Vitro Monolayer

Tensile 20% - STATIC 60 s Type I

Collagen Substrate

Lower Cell Death, Increased Proliferation at 12-24hrs ; gene expression: No change MMP-1, TIMP-1,2 at 12-24hrs, increased TGF-b1, decreased

TNF-a at 12-24hrs

8-40 Porcine Lumbar (4-5 months) 38-40

In Vitro Monolayer

Tensile 5-8% - 0.5 Hz 24 h

Gelatin or Type I

Collagen Substrate

No change in cell viability, Col 1, 2, aggrecan decreased at 6hr, but increased at 9hr and no change at 24hr. ; MMP-1,2,3 and TIMP-1,2

unchanged at 24h

36 Rabbit Lumbar (4wks)

In Vitro Monolayer

Tensile 15% - 0.1Hz 24 h Type I

Collagen Substrate

Significantly increased cell death

35 Rabbit Lumbar (4wks)

In Vitro Monolayer

Tensile 5% - 1Hz 24 h Type I

Collagen Substrate

Proteoglycan synthesis decreased at 8-24hrs, no change in aggrecan

23 Porcine Intact IVD (6 months)

In Vitro Intact IVD

Traction Stress

- 0.8

MPa STATIC 4h -

Proteoglycan synthesis decreased in outer anulus, but no change in inner anulus

30,41 Porcine Lumbar (4-5 months)

In Vitro In Alginate Gel

Comp. 25% (1% ctrl.)

<100 kPa

STATIC 30 h - No change (NC) in cell viability ; increased col-1, col-2, aggrecan,

vimentin gene expression

42 Bovine Intact IVD (2 yr)

In Vitro Intact IVD

Comp. -

0.5-15 kg (0.2-

0.6 MPa)

STATIC 8 h - 3H-pro incorporation: NC (no change) at 0.2-0.4 MPa, decreased at

0.6MPa ; 35S-incorporation: increased at 0.2-0.6MPa

43 Rabbit Cells In Vitro

Monolayer Vibratory

Comp. 0.1 g 6 Hz

2, 4, 6, 8 hr

Tissue Culture Plate

Supressed gene expression for Aggrecan, Collagen 3, MMP-3

18 Murine In Vivo Comp. 1 and 0.2

MPa

1 , 0.2, 0.01 Hz

2h -

1 Hz, 1MPa: small increase in aggrecan, large increases in aggrecanase, collagenase, MMP-3 0.01Hz, 1MPa: large increases in aggrecan, col-1, col-2, small increases in aggrecanase and collagenase 1 MPa in general up-regulated aggrecanase (except at 0.2Hz), collagenase and MMP-3 (at

all frequencies), with small changes in anabolic gene expression (3.5 increase in col-1 at 1Hz) Catabolic gene levels lower at 0.2 Hz compared to 1 and 0.01 Hz 0.2 MPa: The only significant changes were at 1Hz: a

small (2x and 4x) increase in aggrecan , col-1

44 Murine In Vivo Comp. 1 MPa 1 Hz 0.5, 2, 4

hr -

Increasing load duration caused increase for Collagen 1, Collagen 2, MMP3, MMP13.

27 Murine In Vivo Comp. 1 MPa 0.2 Hz 2h - Upregulated collagenase and MMP-3 ; Decrease in Collagen 1 and

Collagen 2, No significant changes in aggrecan (slight decrease)

9

Ref. Species Experiment

Setup Stress Type

Strain Value

Stress Value

Len. Setup Notes

Effects

31 Human Tissue Explant

In Vitro Hydrostatic - 1-10 MPa 20 s and 2

h -

10 atm: upregulation of all ECM protein genes. Increase in Collagen-1 (141% of controls), aggrecan (121%). 30 atm: Collagen-2 similar to 10atm, collagen-1 reduced to 42% of controls. MMP-1 and TGFb-1

down-regulated to 71% and 54% of controls. 31

Human Tissue Explant In Vitro Hydrostatic -

2.5, 7.5 MPa

20 s - Proteoglycan synthesis did not change at 2.5 or 7.5 MPa

24,32 Human Tissue Explant In Vitro Hydrostatic - 1-30 atm 2 h -

Collagen and proteoglycan synthesis did not change || noc changes observed for MMP-3 and TIMP-1

45 Canine Lumbar (3-6 yrs) and Rabbit Lumbar

In Vitro Osmotic Pressure

15-25% PEG

loading-swelling pressure

- 5 h - Proteoglycan synthesis decreased in both 15, 25% PEG

34 Porcine Lumbar (4-5 months)

In Vitro in alginate

Osmotic Pressure

255-450 mOsm

- 4 h - 255 mOsm: Increased collagen 2, aggrecan 450 mOsm: increased

biglycan and decorin mRNA

33 Human Lumbar (29-62 yr)

In Vitro in alginate

Osmotic Pressure

255-450 mOsm

- 4 h - 450 mOsm: Increased ADAMTs, decreased IL-6

10

Degenerative Disc Disease:

Back pain is ranked the most prevalent chronic disease for people under 60, slightly above

arthritis and rheumatism.46 Degenerative Disc Disease contributes to the pathogenesis of lower

back pain and involves the progressive degeneration of the intervertebral disc (IVD). The intact

disc is necessary to support compressive and bending stresses while providing flexibility to the

spine.47 Degenerative Disc Disease (DDD) is marked by increased cell proliferation as well as

cell death. Changes in the production and distribution of structural matrix molecules such as

collagen, elastin, fibronectin is also observed. Macroscopic changes in the matrix, including

increased lamellar disorganization and the appearance of fissures along with increased degree of

invasive vascularization and innervation are associated with DDD. 48,49 Further, with increasing

age, the water content and proteoglycan conent of the nucleus and partly the inner annulus,

decrease.

While the reasons behind DDD are not fully understood, a number of contributing factors have

been suggested. Environmental and occupational factors can contribute to DDD but genetics

seem to be a major contributing factor to the predisposition to develop DDD. Heritability has

been shown to be a significant factor in twin studies, even while adjusting for other factors such

as age, weight, height, smoking, occupational manual work and exercise. 50-52 Of course, these

observations of hereditary factors could be the result of hereditary influence on size and shape of

spinal structures, and thus its internal mechanics, or biological and genetic processes that

ultimately affect synthesis and breakdown of matrix components. 52 The vasculature, present at

birth in the intervertebral disc, diminishes over time and the adult disc is left with little blood

supply. This loss of vasculature may contribute to the unusually early degeneration that occurs in

the disc compared to other tissues. 53 The decreased nutrient supply that results, limits the ability

11

of cells to synthesize new matrix and may limit cell division and could account for the decline in

cell density. Apart from the sparse vascular supply in the outer annulus, diffusion across the

cartilage endplates provides much of the essential solutes for nutrition and metabolic exchange.

Proteoglycan content of the cartilage endplates are very important to transport and control of

water content in the disc, and especially in the nucleus pulposus. Calcification of the cartilage

endplate would affect not only diffusion of nutrients into the disc, but extrusion of metabolic

degradation products that could be toxic to cells. Notochordal cells, which are believed to be

involved in the formation and preservation of nucleus pulposus, gradually disappear with age.54

Changes in the cartilage endplate such as calcification correlate with degeneration of the disc and

particularly that of nucleus pulposus. 55-57 Overall, the emergence of degenerative disc disease

has been linked to lack of vascularity, mechanical trauma to the vertebral body or the disc tissue,

loss of notochordal cells and influence by genetic predispositions, age, gender and other

environmental factors.50-53,55,56,58,59

Currently, existing treatments include discectomy, the use of a prosthetic substitute, or the

fusion of adjacent vertebrae, none of which is optimal. Spinal fusion of degenerated disc may be

effective in some cases, but a number of patients can develop degeneration, due to reduced

flexibility, loss of disc height, and increased stress, in adjacent segments. 60-66 Complications can

also occur in patients undergoing disc replacement with synthetic substrates. Dislocations and

mechanical failure, although rare, have been reported.67 The formation of wear debris can induce

an inflammatory response mediated by various cytokines, leading to pain, osteolysis, fibrous

tissue formation, prosthetic loosening and pain.67-69

12

Figure 3. Current lumbar disc prostheses. SB Charite´ III (A). Prodisc II (B). Maverick (C).

67

Tissue Engineering using Biodegradable Polymers:

One alternative strategy in response to intervertebral disc degeneration is the replacement of

the diseased tissue by a tissue-engineered substitute.70-72 Tissue engineering of annulus fibrosus

is particularly challenging due to the complex structure of the tissue. A significant portion of

biodegradable polymers considered for tissue engineering, belong to the polyester family.

Among these poly(α-hydroxy acids), such as poly(glycolic acid) (PGA), poly(lactic acid) (PLA),

and their copolymers have been closely studied and have been used as synthetic biodegradable

materials.73 Efforts in producing AF tissue in vitro has involved various polymeric scaffolds

including PDLLA/45S5 Bioglass® films, polyglycolic acid, collagen/hyaluronan,

collagen/glycosaminoglycans (GAGs), atelocollagen, and alginate scaffolds.74-79 In addition to

inadequate tissue formation, some biomaterials, such as polyglycolic-based polymers, generate

acidic byproducts throughout their biodegradation, which can significantly alter cell behavior,

tissue production and possibly cause cell death.80,81 Studies have shown that porous PLA-PGA

scaffolds produce toxic solutions as a result of acidic degradation, which may illicit adverse

responses during the tissue repair process. 73 Further, the release of small particles can also

trigger an undesirable inflammatory response. 82

13

In this study, polycarbonate-urethane polymers were used because of their expected

biocompatible and biodegradable nature. Polyurethanes (PUs) have been used in biomedical

devices since the 1960s. Traditionally, research by investigators in the 80’s and 90’s had been

directed at producing biostable polyurethanes in an effort to shield them from biodegradation

processes. However in the past decade, the focus has shifted to utilizing the flexible chemistry of

PUs in developing bioactive/biocompatible and biodegradable polyurethanes for the purpose of

tissue engineering or regeneration.83,84 As an elastomer, the mechanical properties of PU can be

carefully controlled. Polyurethanes can have a broad range of mechanical properties depending

on the chemistry of the specific copolymer. Tensile strengths of PUs have been found to be in the

range of 6-40 MPa.85 The shift to biodegradable PU-based materials has been accompanied with

a change in the diisocyanates used in their synthesis. Aromatic diisocyanates, which are prone to

carcinogenic effects on tissue, have been replaced with diisocyanates such as hexane

diisocyanate, whose ultimate degradation products are more likely to be non-toxic.86-90 Early

studies on the biodegradation of polyurethanes cited environmental stress cracking, driven by

factors including surface oxidation, residual stress, polyether soft-segment chemistry, molecular

morphology, the presence of MDM and foreign body giant cells (FBGC), as well as an unknown

biological element.84 However, a more inclusive approach to polyurethane biodegradation,

termed environmental biodegradation, has been proposed which accounts for biodegradation due

to hydrolytic enzymes. Santerre and Labow were first to study PU degradation using

physiologically relevant enzymes.91 Enzymes such as cholesterol esterase (CE) were shown to

preferably degrade ester linkages immediately adjacent to the hard segment. CE is present in

monocytes as they differentiate into macrophages and has been reported to degrade PUs.92,93

Further, CE has been shown to exceed the degradation potential of many other enzymes by more

14

than 100-fold. 94 The results of these studies have also demonstrated that PU biodegradation is

affected by a variety of factors such as hard-segment chemistry and stress induced strain. 84

Figure 4. Stress induced strain along with material morphology and chemistry, can affect the

environmental degradation of the material 84

Short-term studies on in vitro and in vivo biocompatibility of biodegradable polyurethane

polymers have shown no abnormal growth behaviour, nor morphological changes or inhibition in

metabolic activity.95 Considering the collection of previous work in this area, it was

hypothesized that the byproducts resulting from the degradation of polycarbonate urethanes in

this study would likely be non-toxic to AF cells as well.

Electrospinning:

Polycarbonate urethanes can be fabricated in many different forms and most importantly, they

may be suitable for use in the process of electrospinning.96 In this process, the polymer is

dissolved in a volatile solvent and subjected to a high voltage compared to a rotating deposition

surface.96. This electrical field overcomes surface tension of the solution and causes the solution

15

to separate into fine fibers. The produced fibers mimic the aligned nature of the annulus

fibrosus, and thus provide a more appropriate surface for growth of such a tissue. The high

surface to volume ratio of these scaffolds is expected to favor cell attachment and retention of

cell phenotype. 97,98 Studies have shown that by depositing electrospun nanofibers onto a rotating

mandrel, one can dictate the mechanical anisotropy of scaffolds, addressing the importance of

mechanical strength of fibrous scaffolds for AF tissue regeneration.99

Proposed Work:

To achieve biological repair of a degenerate disc using a tissue-engineered construct, it is

necessary to develop methods that encourage production of tissue that closely mimics its

physiological counterpart. The overall strategy behind the use of an electrospun polyurethane

nanofiber scaffold for tissue engineering the AF, involves the growth of an AF tissue layer on the

surface of an aligned scaffold. Multiple layers of the resulting aligned AF tissue can then be

combined to produce a tissue engineered AF construct. We hypothesize that PU is an appropriate

scaffold to use for tissue engineering the annulus fibrosus. This will be determined by

characterizing the mechanical, biodegradation and cytotoxic characteristics of an elastomeric

polycarbonate-urethane. The four objectives of this work are 1) to determine the mechanical

properties of the aligned and random electrospun polycarbonate-urethane nanofiber scaffold. It

was anticipated that aligned scaffolds would have superior mechanical properties to random

scaffolds. The mechanical properties of these polymers in relation to those of native AF tissue

were of particular interest; 2) to study the effects of biodegradation on PU’s mechanical

properties to determine the level to which the scaffold can provide mechanical support

throughout the biodegradation process; 3) to determine the cytotoxic effects of the PU

degradation products on AF cells. It was anticipated that PU should produce non-toxic

16

degradation byproducts given previous studies of related biomaterials 84; 4) to investigate the

response by AF cells grown on polyurethane electrospun scaffolds, to cyclic compressive

mechanical forces. Previous studies have shown some mechanical forces to be detrimental and

others beneficial to AF tissue development, making it difficult to anticipate the response of AF

tissue to the proposed mechanical forces. Compression was chosen as a starting point for

analysis of mechanical forces on the AF tissue as they have been shown to initiate a response

from AF cells.

17

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nucleus pulposus and annulus fibrosus cells in alginate gel. Biochem.Soc.Trans.,

2002, Pt 6, 874-878.

(76) Wilda, H. and Gough, J. E. In vitro studies of annulus fibrosus disc cell attachment,

differentiation and matrix production on PDLLA/45S5 Bioglass composite films.

Biomaterials, 2006, 30, 5220-5229.

(77) Rong, Y., Sugumaran, G., Silbert, J. E., and Spector, M. Proteoglycans synthesized by

canine intervertebral disc cells grown in a type I collagen-glycosaminoglycan matrix.

Tissue Eng, 2002, 6, 1037-1047.

(78) Alini, M., Li, W., Markovic, P. et al. The potential and limitations of a cell-seeded

collagen/hyaluronan scaffold to engineer an intervertebral disc-like matrix. Spine, 1-

3-2003, 5, 446-454.

27

(79) Mizuno, H., Roy, A. K., Vacanti, C. A. et al. Tissue-engineered composites of anulus

fibrosus and nucleus pulposus for intervertebral disc replacement. Spine, 15-6-2004,

12, 1290-1297.

(80) Ishihara, H. and Urban, J. P. Effects of low oxygen concentrations and metabolic

inhibitors on proteoglycan and protein synthesis rates in the intervertebral disc.

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degradation of PLGA. Composites Science and Technology, 2005, 14, 2226-2232.

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in vitro acute toxicity of accumulated degradation products. J.Appl.Biomater., 1994,

2, 151-157.

(83) Guelcher, S. A. Biodegradable polyurethanes: synthesis and applications in regenerative

medicine. Tissue Eng Part B Rev., 2008, 1, 3-17.

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biodegradation of polyurethanes: from classical implants to tissue engineering

materials. Biomaterials, 2005, 35, 7457-7470.

(85) P.Bruin, G.J.Veenstra, A.J.Nijenhuis, and A.J.Pennings Design and synthesis of

biodegradable poly(ester-urethane) elastomer networks composed of non-toxic

building blocks. Die Makromolekulare Chemie, Rapid Communications, 1988, 8,

589-594.

28

(86) Zhang, J. Y., Beckman, E. J., Piesco, N. P., and Agarwal, S. A new peptide-based

urethane polymer: synthesis, biodegradation, and potential to support cell growth in

vitro. Biomaterials, 2000, 12, 1247-1258.

(87) Skarja, G. A. and Woodhouse, K. A. Synthesis and characterization of degradable

polyurethane elastomers containing and amino acid-based chain extender.

J.Biomater.Sci.Polym.Ed, 1998, 3, 271-295.

(88) Saad, B., Ciardelli, G., Matter, S. et al. Degradable and highly porous polyesterurethane

foam as biomaterial: effects and phagocytosis of degradation products in osteoblasts.

J.Biomed.Mater.Res., 15-3-1998, 4, 594-602.

(89) Cohn, D., Stern, T., Gonzalez, M. F., and Epstein, J. Biodegradable poly(ethylene

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2002, 2, 273-281.

(90) Woo, G. L., Mittelman, M. W., and Santerre, J. P. Synthesis and characterization of a

novel biodegradable antimicrobial polymer. Biomaterials, 2000, 12, 1235-1246.

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urethane by cholesterol esterase: isolation and identification of principal

biodegradation products. J.Biomed.Mater.Res., 5-9-1997, 3, 407-417.

(92) Labow, R. S., Meek, E., and Santerre, J. P. Synthesis of cholesterol esterase by

monocyte-derived macrophages: a potential role in the biodegradation of

poly(urethane)s. J.Biomater.Appl., 1999, 3, 187-205.

29

(93) Labow, R. S., Sa, D., Matheson, L. A., and Santerre, J. P. Polycarbonate-urethane hard

segment type influences esterase substrate specificity for human-macrophage-

mediated biodegradation. J.Biomater.Sci.Polym.Ed, 2005, 9, 1167-1177.

(94) Tang, Y. W., Labow, R. S., and Santerre, J. P. Enzyme-induced biodegradation of

polycarbonate-polyurethanes: dependence on hard-segment chemistry.


(95) van, Minnen B., van Leeuwen, M. B., Stegenga, B. et al. Short-term in vitro and in vivo

biocompatibility of a biodegradable polyurethane foam based on 1,4-

butanediisocyanate. J.Mater.Sci.Mater.Med., 2005, 3, 221-227.

(96) Stankus, J. J., Guan, J., and Wagner, W. R. Fabrication of biodegradable elastomeric

scaffolds with sub-micron morphologies. J.Biomed.Mater.Res.A, 15-9-2004, 4, 603-

614.

(97) Thapa, A., Miller, D. C., Webster, T. J., and Haberstroh, K. M. Nano-structured polymers

enhance bladder smooth muscle cell function. Biomaterials, 2003, 17, 2915-2926.

(98) Yang, L., Kandel, R. A., Chang, G., and Santerre, J. P. Polar Surface Chemistry of

Nanofibrous Polyurethane Scaffold Affects Annulus Fibrosus Cell Attachment and

Early Matrix Accumulation. J.Biomed.Mater.Res.A, 2008,

http://www3.interscience.wiley.com/journal/121582889/.

(99) Nerurkar, N. L., Elliott, D. M., and Mauck, R. L. Mechanics of oriented electrospun

nanofibrous scaffolds for annulus fibrosus tissue engineering. J.Orthop Res, 2007, 8,

1018-1028.

30

VII: Characterization of a Biodegradable Electrospun

Polyurethane Nanofiber Scaffold: Mechanical Properties

and Cytotoxicity

31

Characterization of a Biodegradable Electrospun

Polyurethane Nanofiber Scaffold: Mechanical

Properties and Cytotoxicity

Masoud Yeganegi1, 2, Rita A Kandel 1, 2, 3, and J Paul Santerre2, 4

1CIHR- Bioengineering of Skeletal Tissues Team, Mount Sinai Hospital, Toronto, M5G 1X5

Canada

2Institute of Biomaterials and Biomedical Engineering and Department of Materials Science and

Engineering, University of Toronto, Toronto, M5S 3G9 Canada

3 Department of Pathobiology and Laboratory Medicine, Mt. Sinai Hospital, University of

Toronto, Toronto, M5G 1X5 Canada

4Faculty of Dentistry, University of Toronto, Toronto, M5G 1G6 Canada

To whom correspondence should be sent:

Dr. Paul Santerre

Department of Biological and Diagnostic Sciences

Faculty of Dentistry


124 Edward St., Toronto, Ontario, Canada

M5G 1G6

Phone: (416) 979 4903 x4341

Email: [email protected]

32

Introduction

The human vertebral column is made up of 26 vertebral bodies that provide support to the

body and protect the spinal cord. The intervertebral discs lying between the vertebrae provide

flexibility and help dissipate mechanical loads and shocks that would otherwise damage the

vertebral column.1 The intervertebral discs are composed of the annulus fibrosus, a

fibrocartilaginous tissue, which surrounds the gelatinous inner nucleus pulposus. The hyaline

cartilage endplates found at each end represent the anatomical limits of the disc and contribute to

the interface of the disc and bone.

The annulus fibrosus (AF) is responsible for withstanding circumferential tensile forces and to

a lesser extent compressive forces.2 The outer AF consists of concentric lamella, made up of

collagen fibers, and oriented at approximately 60º to the vertical. The alignment of these fibers

alternates between successive lamellae and is of great importance to the functional nature of the

annulus fibrosus.3 The AF extracellular matrix contains collagen fibrils, proteoglycans and water.

Water makes up 60% of the annulus fibrosus, while collagen and proteoglycans account for 50-

70% and 10-20% of the dry weight respectively. 4 The composition of the annulus fibrosus is

radially non-uniform in that collagen type I decreases in concentration radially towards the

center of the disc.5 Conversely, collagen type II which is present in small quantities in the AF,

increases in concentration toward its innermost lamellae. The relative proportion of collagen type

I to collagen type II in the annulus fibrosus varies from 70:30 in the innermost layers and 85:15

in the outer layers.4 Other types of collagen also exist in smaller amounts in the annulus

fibrosus.6

33

The mechanical properties of intervertebral discs are complex and the literature on this topic

shows significant variability, 7-10 perhaps in part due to the diversity of experimental conditions

and mechanical models used to measure material properties. Studies indicate that the annulus

fibrosus exhibits both matrix viscoelastic (flow-independent) and biphasic viscoelastic (flow-

dependent) behaviour. 10 The elastic modulus of a single lamella of annulus fibrosus has been

found to vary with radial position in the disc with values ranging from 5±4 MPa for posterior

inner AF, 20±12 MPa for posterior outer AF, 10±6 MPa for anterior inner AF, and 49±32 MPa

for anterior outer AF. 2 The ultimate stress of a single lamella of annulus fibrosus also varies

radially from 0.9±0.3 MPa for posterior inner AF, 1.1±0.3 MPa for posterior outer AF, 0.9±0.7

MPa for anterior inner AF, and 3.3±1.3 MPa for anterior outer AF. 2 Thus the forces experienced

by the AF tissue can be quite high and therefore an intact AF is critical to proper disc function.

Back pain is ranked the most prevalent chronic disease for people under 60.11 Degenerative

disc disease may contribute to the pathogenesis of lower back pain and involves the progressive

degeneration of the intervertebral disc (IVD). The intact disc is necessary to support compressive

and bending stresses while providing flexibility to the spine.12 Currently, existing treatments

include, discectomy, the use of a prosthetic substitute, or the fusion of adjacent vertebrae, none

of which are optimal. Spinal fusion may be effective in some cases, but a number of patients can

develop degeneration, due to reduced flexibility and increased stress in adjacent segments. 13-16

Complications, can also occur in patients undergoing disc replacement with synthetic substrates.

Dislocations, slippage, wear and mechanical failure, although rare, have been reported.17 The

formation of wear debris can induce an inflammatory response mediated by various cytokines,

leading to pain, osteolysis, fibrous tissue formation, prosthetic loosening and pain.17,18

34

One alternative strategy for the treatment of intervertebral disc degeneration is the replacement

of the diseased tissue by a tissue-engineered substitute.19-21 Tissue engineering of the annulus

fibrosus is particularly difficult due to the complex structure of the tissue. Efforts in producing

AF tissue in vitro have involved various polymeric scaffolds including PDLLA/45S5 Bioglass®

films, polyglycolic acid, collagen/hyaluronan, collagen/gag, atelocollagen, and alginate

scaffolds.22-27 In addition to inadequate tissue formation, some biomaterials, such as

polyglycolic-based polymers, create acidic byproducts throughout their biodegradation, which

can significantly alter cell behavior, tissue production and possibly cause cell death.28,29

In this study, a potentially more suitable alternative, polycarbonate-urethane polymers, were

used owing to their biocompatible, biodegradable and reproducible nature. Further, this polymer

can be synthesized with the addition of molecules that may enhance cell attachment. It is

possible to use electrospin this polymer and fabricate aligned fibers, which mimic the aligned

nature of the annulus fibrosus, and thus providing a more appropriate scaffold to support the

growth of such a tissue, especially as topographical properties have been shown to influence cell

alignment.30 The high surface to volume ratio of these scaffolds is expected to favor cell

attachment and retention of cell phenotype. 31,32 The purpose of this study was to characterize the

electrospun PU nanofiber biomaterial, and determine its suitability for use in repairing the AF.

Experimental Section

Materials

Polyurethanes have been used in biomedical devices since the 1960s. Efforts in the 80’s and

90’s had previously been directed at producing biostable polyurethanes, however in the past

decade, the focus has shifted to developing bioactive/biocompatible and biodegradable

35

polyurethanes for the purpose of tissue engineering or regeneration.33,34 In this study,

polycarbonate urethane (PU) was synthesized, as has been described, with hexane

diisocyanate:polycarbonate diol:butane diol molar ratio of 3:2:1 35. In addition, an anionic

dihydroxyl oligomer (ADO) additive was synthesized through the reaction of lysine

diisocyanate, polytetramethylene oxide, and hydroxyethyl methacrylate (HEMA). 32 This anionic

additive has been previously shown to increase AF cellular adhesion mediated by protein

adsorption to the surface. 32

Electospinning was employed to fabricate either random or aligned nanofiber scaffolds using

the base polymer (PU) with or without ADO (0.5 wt.%). The polymer was dissolved in

1,1,1,3,3,3-hexafluora-2-propanol at a concentration of 16 wt.%. The scaffold was then

electrospun, by injecting the polymer solution from a metallic syringe at a rate of 0.5 ml/hr onto

the collecting surface. An 18 kilovolts difference was applied across the syringe and the

collecting surface. In the case of aligned scaffolds, the collecting surface consisted of a

cylindrical aluminum mandrel rotating at 1250 rpm, such that the surface of the mandrel moved

at 10 meters/s.32,36 For the random scaffold, a stationary aluminum plate was used. Relative

humidity (<30%) and temperature (approximately 25ºC) were controlled to minimize adverse

effects on the scaffold quality and reproducibility. The resulting electrospun films were

approximately 106 ± 5µm and 550 ± 50µm thick for aligned and random scaffolds respectively.

The fibers ranged from 200-400 nm in diameter as measured by scanning electron microscopy

(Figure 1).

36

Biodegradation Study

Cholesterol Esterase (CE) has been shown to be present in monocytes as they differentiate into

macrophages and has been reported to degrade PUs.37,38 Thus this enzyme was selected to

evaluate the biodegradation of the PU scaffolds. CE (C3766, Sigma Aldrich, St. Louis, MO) was

adjusted to 10 units/ml, with 1 unit defined as generating 1 nmol/min of p-nitrophenol from p-

nitrophenylbutyrate (as measured colorimetrically by the DU 800 Beckman Coulter

Spectrophotometer).37 This enzyme concentration was selected based on previous CE dose

response studies, and was chosen to ensure that adequate degradation would occur throughout

the four week study. 39 The half-life of CE in the presence of the polymer was approximately 12

hours (Figure 2), thus, throughout the 4-week biodegradation study, appropriate volumes of a

concentrated enzyme solution (100 units/ml) were added daily to adjust the CE activity to a final

concentration of 10 units/ml in a phosphate buffer solution (PBS, pH 7.0, 37ºC) containing the

samples.

Assessment of Mechanical Properties

To evaluate their biodegradation properties, dried scaffolds were cut into 6mm x 30mm pieces

(thickness of 106 ± 5µm and 550 ± 50µm for aligned and random scaffolds respectively) and

weighed (Ohaus Explorer analytical balance) prior to placing in the enzymatic solution. The

differences in thickness between the aligned and random scaffolds stem from the fact that

following the fabrication process, thinner random polymers are difficult to remove from the

deposition surface due to their inferior mechanical properties, and thus thicker polymers were

required to prevent polymer deformation prior to mechanical testing. The scaffolds were

collected weekly, washed in dH2O, lyophilized and weighed to determine average mass loss. The

dried scaffolds underwent tensile testing prior to and during the 4-week biodegradation period to

37

assess mechanical properties. Each polymer sample was carefully immobilized on either end

using metallic clamps, and the tensile strength was evaluated using a mechanical testing device

(Instron® model 8501) under a tensile strain of 10 mm/min to breaking point. The initial

modulus and the ultimate stress were used as measures of intrinsic mechanical properties

throughout the degradation period.

Differential Scanning Calorimetry

Differential scanning calorimetry (DSC) was performed to assess the crystallinity of scaffolds

and changes in their microstructure due to the pre-wetting and drying process. The samples were

pre-wet in phosphate buffer in the absence of CE for one week, washed in distilled water and

lyophilized. The dried samples were then compared to as-made controls to understand the effects

of the wetting and drying processes on the scaffolds. A small section of each sample, weighing

approximately 2–3 mg, was cooled using liquid nitrogen and the thermograms were recorded

between −100 to 200°C at a heating rate of 15°C/min (DSC was performed using the TA

Instruments differential scanning calorimeter (model 2910) at the Brockhouse Institute for

Material Research, McMaster University, Hamilton, Ontario, Canada). Data recorded for the first

of two consecutive heating cycles was used to analyze possible crystalline state changes.

Evaluation of Degradation Products

The soluble and non-soluble degradation products were collected. Phosphate buffered saline

(PBS) containing the degradation products, was spun down at 3,000 RCF to isolate the non-

soluble particulate. Aliquots of the non-soluble degradation products were placed directly in

formvar-coated grids and imaged using transmission electron microscopy (TEM, FEI Tecnai 20,

38

Hillsboro, Oregon) to visualize particulate size and morphology. Following the separation of

soluble and non-soluble degradation products, cytotoxicity studies were performed.

Cytotoxicity Study

To evaluate if the degradation products were cytotoxic, bovine annulus fibrous cells were

chosen as a valid model for human lumbar AF 40-42. Briefly, to obtain AF cells, intervertebral

discs from bovine caudal spines (6–9 months old) were dissected out aseptically. Five discs from

one spine were combined to obtain sufficient cells for an experiment. The outer annulus was

separated from the disc and minced into small pieces. To isolate the cells, the tissues underwent

serial digestion with 0.5% protease (Sigma, St. Louis, MO) for 1 hour at 37° C, followed by

0.25% collagenase A (Roche, Laval, Quebec, Canada) overnight at 37° C. The cell suspension

was filtered through a sterile mesh (pore size: 70µm), and re-suspended in Ham’s F12 media

supplemented with 5% fetal bovine serum (FBS). AF cells were placed in monolayer culture

(seeding density of 1x105/cm2) and grown for 48 hours in Ham’s F12 media containing 5% fetal

bovine serum. The non-soluble degradation products were added to the tissue culture media at

concentrations ranging from 0.001 to 0.1 wt %. Soluble degradation products (in PBS) were

added to the media at various concentrations ranging from 20% to 100%. Cytotoxicity of the

degradation products was assessed using the MTT assay and Live/Dead fluorescent imaging. In

the MTT assay, mitochondrial dehydrogenase of viable cells cleave the tetrazolium ring of 3-

(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) yielding purple formazan

crystals which are insoluble in aqueous solutions. The resulting crystals were dissolved in

methoxyethanol (acidified with HCL to pH: 3). The resulting purple solution was measured

spectrophotometrically at a wavelength of 570nm (Thermo Scientific model Multiskan Ex

photometer). Hydrogen peroxide (0.01%) served as a positive control.43 In the live/dead assay

39

(L-3224, Invitrogen, Burlington, ON) Calcein AM is enzymatically converted to its fluorescent

variant, which in cells with intact membranes, is concentrated within the cytoplasm. Ethidium

homodimer-1 enters cells with damaged membranes and binds to nucleic acids and becomes

fluorescent. ImageJ (ver. 1.4g) was used to find regions of interest (ROI) by isolating local

maxima, which targeted the intensified Calcein AM observed around the nucleus of viable cells.

These ROIs were then manually examined to ensure that only viable cells were detected and

counted. This procedure was similarly used to detect dead cells stained by Ethidium homodimer-

1. On average, nearly 600 cells were counted for each data point.

Statistical analysis

All test groups contained at least four samples and each experiment was repeated at least 3

times. The results from all experiments were combined and expressed as the mean ± standard

error of the mean. The data were analyzed using a one-way analysis of variance and all pair-wise

comparisons between groups were conducted using the Scheffe post hoc test. Significance was

assigned at p-values < 0.05.

Results and Discussion

Comparing the mass of polymers before and after degradation, it was observed that

degradation by CE reached an average of 2.0 mg per sample for all groups by week four. This

enzyme concentration had specifically been adjusted to produce such an extent of degradation as

appropriate for the purposes of evaluating any changes in mechanical properties and subsequent

assessment of degradation product cytotoxicity. The rate of biodegradation was 0.56 ± 0.05

mg/week and was relatively linear over the four weeks (r2=0.86) (Figure 3a), reaching a

maximum cumulative mass loss of 30±2% for aligned scaffolds and 7±2% for random scaffolds

40

at four weeks. The absolute mass loss was similar for both random and aligned scaffolds despite

differences in relative mass loss (Figure 3b). This, combined with the fact that all samples

possessed the same surface area available for enzymatic degradation, suggests that surface-

mediated degradation took place rather than bulk material breakdown. These findings, likely

explained by diffusion limitations imposed by the scaffold on cholesterol esterase, are in

agreement with previous studies on polyurethane biomaterials in which CE mediated surface

degradation was primarily observed.44,45

The aligned samples experienced a significant decrease (45.6 MPa to 8.9 MPa, p<0.05) in the

initial modulus following a wetting and drying process and prior to degradation (Figure 4a). In

addition, a significant drop in tensile strength (13.8 MPa to 6.6 MPa, p<0.05) was also observed

in aligned scaffolds (Figure 4b). There was no observed difference between PU and

PU+0.5%ADO samples, suggesting that ADO did not appear to have any plasticizing or

hardening effects on the mechanical behaviour of the scaffolds. In order to further study the

structural transformations introduced in the materials during the pre-wetting and drying process,

samples were analyzed using differential scanning calorimetry (DSC). The thermal transition

temperatures were relatively similar for both PU and PU+0.5%ADO scaffolds (Figure 5 a and b).

The Tv hard segment melting transition temperature remained relatively unchanged near 93ºC

across all samples and was unaffected by the pre-wetting process. Likewise, the Tg values for

polycarbonate near -36°C remained relatively unchanged. This temperature corresponds to the

crystalline segments containing polar urethane groups, which are anticipated to be the most

stable regions of the polymer due to extensive hydrogen bonding within this domain. Perhaps the

most striking difference between the as-made controls and the pre-wet samples was the shift in

the soft-segment melting transition temperature (Tiii and Tiv in Figure 5) from approximately

41

51ºC to 68ºC respectively. The shift of this phase towards higher temperatures for the PCN phase

of the polymer suggests a loss of PCN phase purity (pure polycarbonate has a melt transition

temperature of 45ºC 35) and an increase in phase mixing of hard segment with the soft segment

content. More evidence that the pre-wetting process disrupted the crystalline state of the soft

segment polycarbonate (PCN) phase was observed near 33ºC (Tii, Figure 5). The appearance of

the onset of the soft segment melt phase for the prewet sample is distant from the endotherm at

67ºC. Such a broad transition is characteristic of a heterogeneous phase. These transformations

may be disrupting the organization of the crystalline matrix for the soft segments and are

believed to be contributing in part to the softening of the material post exposure to aqueous

medium and the resulting deterioration in its mechanical properties. In the case of random

scaffolds, no significant drop in mechanical properties were observed with exposure to water.

This may be explained in part by the slower buffer uptake by random scaffolds due to their

substantially greater thickness (550 ± 50µm for random vs 106 ± 5µm for oriented scaffolds).

Further, DSC results for random scaffolds show a slightly lower shift in the soft-segment melt

temperature (a shift of 12°C as compared to 17°C), which may partially account for the

differences in the effects of buffer uptake. Aligned scaffolds are subjected to stress induced

strain due to the forces experienced by polymer fibers as they deposit onto the rotating mandrel.

Such stress induced strain in polyurethanes, which has been recognized as affecting material

morphology and chemistry, may contribute to the varying responses of aligned and random

scaffolds to buffer uptake.46

After the initial exposure to the aqueous medium, the mechanical properties of the scaffolds

showed no further significant changes during the course of the four week biodegradation study,

despite showing a degradation rate of 0.56 ± 0.05 mg/week (Figures 3 and 6). The surface-

42

mediated biodegradation mechanism helps explain the lack of a significant drop in mechanical

properties, since the observed mass losses are accompanied by similar drops in thickness of the

scaffolds. Throughout the four week biodegradation process, corresponding to a maximum 30 %

mass loss, the aligned scaffolds retained values of approximately 8.0 MPa for the modulus and

2.3 MPa for the tensile strength, thereby remaining within the same range as those of native

tissue (Figure 6). Random scaffolds were similarly unaffected by the four week biodegradation

study, although the relative mass loss was significantly lower which could partly explain the

absence of any changes in mechanical properties. The reported literature values of elastic

modulus for a single lamella of native human AF are 5±4 MPa for posterior inner AF, 20±12

MPa for posterior outer AF, 10±6 MPa for anterior inner AF, and 49±32 MPa for anterior outer

AF. 2 The ultimate stress of single lamellae of annulus fibrosus similarly varies radially from

0.9±0.3 MPa for posterior inner AF, 1.1±0.3 MPa for posterior outer AF, 0.9±0.7 MPa for

anterior inner AF, and 3.3±1.3 MPa for anterior outer AF. 2 Overall, the initial moduli and

ultimate stress of the different scaffold groups were comparable and in some cases superior to

those of native AF tissue.

In this study, cholesterol esterase was used to degrade the polymer samples. Although this

gives an indication of what may occur in vivo as CE is known to be secreted by macrophages

37,38, the latter also secrete other proteases as well as oxidative agents, which would introduce

oxidative degradation in addition to the hydrolytic degradation cause by CE. Hence, further In

vivo studies are required to fully assess the biodegradation behavior. The degradation products

were spun down and separated into soluble and non-soluble products. The non-soluble products,

imaged by transition electron microscopy, varied in shape and size (Figure 7). The thickness of

the fiber fragment was found to be approximately on average 400 nm, which is comparable to

43

that of the electrospun nanofibers (Figure a). Different concentration gradients of the soluble and

non-soluble degradation products were applied to bovine annulus fibrosus cells grown in

monolayer culture. The MTT colorimetric assay indicated no significant cytotoxic effects from

either non-soluble or soluble degradation products, from the scaffolds made from PU or

PU+ADO (Figure 8a and b). Although the MTT assay is used to measure cytotoxicity, it is an

indicator of metabolic activity. Thus, a live/dead assay was performed concurrently with the

MTT assay to confirm the findings. The live/dead assay confirmed that the material’s

degradation products did not induce cytotoxicity (Figures 9 and 10). These results are in

agreement with our hypothesis that degradation of PU would likely produce non-toxic

byproducts given previous studies of related biomaterials. 34 This work has concentrated on

investigating possible effects of degradation products on cell viability and morphology. Future

studies are required to explore additional indicators of toxicity such as the effect degradation

fragments on cell proliferation and extracellular matrix synthesis.

Conclusion

The findings in this study have shown that electrospun aligned scaffolds produced superior

mechanical properties in relation to random scaffolds and suggest this formulation as a more

appropriate scaffold for engineering annulus fibrosus tissue. An important consideration in the

design of such scaffolds is the issue of structural changes due to water uptake into the material,

particularly as it relates to the rate of degradation. It was established that exposure to aqueous

media disrupted the material’s chemical structure and resulted in a reduction of its mechanical

properties. DSC showed the appearance of disrupted soft segment crystal phase with changes in

the degree of phase mixing of hard and soft segments as well as the PCN crystal state within the

polymer. The degradation of the polymer by cholesterol esterase provided a useful model for

44

biodegradation, yielding a controlled and consistent mass loss rate. Of particular importance, the

degradation of the materials, which resulted in mass losses as high as 30% over four weeks, did

not result in a significant deterioration of mechanical properties, indicating a surface degradation

process rather than bulk material breakdown. This finding is supported by the fact that absolute

mass loss appears to be independent of the thickness of the scaffold and appears to be in

agreement with previous studies in which CE mediated surface degradation has been observed.44

The degradation products did not cause significant acute cytotoxicity in-vitro. Additional

mechanical studies to explore the changes in mechanical properties of the aligned polymer in the

presence of AF tissue grown on the PU substrate will provide further understanding of the role of

this polymer for AF tissue engineering. In summary, the results of this report, namely the

relatively constant rate of material degradation, the observed mechanical behavior resembling

that of AF tissue, and the absence of cytotoxic effects make this polymer a suitable biomaterial

candidate for use in the formation of tissue-engineered annulus fibrosus.

Acknowledgements

The funding for this work was provided by a University of Toronto Fellowship Award, an

Ontario Graduate Scholarship in Science and Technology (OGSST), in addition to an NSERC-

CIHR Collaborative Health Research Program (CHRP) grant (312882) and a CIHR operating

grant (MOP86723).

45

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874-878.



Biomaterials, 2006, 30, 5220-5229.



Tissue Eng, 2002, 6, 1037-1047.

48


collagen/hyaluronan scaffold to engineer an intervertebral disc-like matrix. Spine, 3-1-

2003, 5, 446-454.


fibrosus and nucleus pulposus for intervertebral disc replacement. Spine, 6-15-2004, 12,

1290-1297.

(28) Ishihara, H. and Urban, J. P. Effects of low oxygen concentrations and metabolic inhibitors

on proteoglycan and protein synthesis rates in the intervertebral disc. J.Orthop.Res.,

1999, 6, 829-835.












49









(36) Stankus, J. J., Guan, J., and Wagner, W. R. Fabrication of biodegradable elastomeric

scaffolds with sub-micron morphologies. J.Biomed.Mater.Res.A, 9-15-2004, 4, 603-

614.

(37) Labow, R. S., Meek, E., and Santerre, J. P. Synthesis of cholesterol esterase by monocyte-

derived macrophages: a potential role in the biodegradation of poly(urethane)s.

J.Biomater.Appl., 1999, 3, 187-205.

(38) Labow, R. S., Sa, D., Matheson, L. A., and Santerre, J. P. Polycarbonate-urethane hard

segment type influences esterase substrate specificity for human-macrophage-mediated

biodegradation. J.Biomater.Sci.Polym.Ed, 2005, 9, 1167-1177.

(39) Tang, Y. W., Labow, R. S., and Santerre, J. P. Enzyme induced biodegradation of

polycarbonate-polyurethanes: dose dependence effect of cholesterol esterase.

Biomaterials, 2003, 12, 2003-2011.

50

(40) Demers, C. N., Antoniou, J., and Mwale, F. Value and limitations of using the bovine tail

as a model for the human lumbar spine. Spine, 12-15-2004, 24, 2793-2799.

(41) Cotterill, P. C., Kostuik, J. P., D'Angelo, G., Fernie, G. R., and Maki, B. E. An anatomical

comparison of the human and bovine thoracolumbar spine. J.Orthop.Res., 1986, 3, 298-

303.

(42) Wilke, H. J., Krischak, S., and Claes, L. Biomechanical comparison of calf and human

spines. J.Orthop.Res., 1996, 3, 500-503.

(43) Hoffmann, M. E. and Meneghini, R. Action of hydrogen peroxide on human fibroblast in

culture. Photochem.Photobiol., 1979, 1, 151-155.

(44) Christenson, E. M., Patel, S., Anderson, J. M., and Hiltner, A. Enzymatic degradation of

poly(ether urethane) and poly(carbonate urethane) by cholesterol esterase.

Biomaterials, 2006, 21, 3920-3926.

(45) Skarja, G. A. and Woodhouse, K. A. In vitro degradation and erosion of degradable,

segmented polyurethanes containing an amino acid-based chain extender.

J.Biomater.Sci.Polym.Ed, 2001, 8, 851-873.




51

Figures

Figure 1. Scanning Electron Microscopy Images of aligned (a) and random (b) electrospun

Polycarbonate Urethane Nanofiber Scaffolds. (Solution: 16 wt.% PU, injection rate:

0.5 ml/hr, potential difference: 18 kilovolts)

52

Figure 2. Determination of Cholesterol Esterase (CE) Half Life. It was determined that the half

life of CE in the presence of the aligned PU scaffolds was approximately 12 hours.

Thus CE was added daily to adjust the enzyme activity (n = 3).

0

20

40

60

80

100

120

140

0 5 10 15

CE Activity (units/mL)

Hours

CE

CE+PU Scaffold

53

Figure 3. Cumulative absolute mass loss (a) and cumulative relative mass loss (b) during

biodegradation. Scaffolds were incubated in 100 units/ml CE over 4 weeks. Data are

reported as mean ± standard error (n=6). (*) Absolute mass loss was found to increase

significantly at every week (p<0.05) for all groups, while no statistical differences

were observed between the scaffold groups within each week. Relative mass loss was

found to increase significantly in the case of aligned scaffolds.

0

0.5

1

1.5

2

2.5

3

Week 1 Week 2 Week 3 Week 4

Weight Loss (mg)

Aligned PU+0.5%ADO

Aligned PU Random PU+0.5%ADO

*

*

*

(a)

0%

5%

10%

15%

20%

25%

30%

35%

Week 1 Week 2 Week 3 Week 4

Weight Loss (% of Original)

Aligned PU+0.5%ADO

Aligned PU Random PU+0.5%ADO

*

*

*

(b)

54

Figure 4. (a) Elastic Modulus and (b) Tensile Strength of the electrospun polyurethane

nanofiber scaffolds following the pre-wetting (for one week in pH 7.0 PBS at 37ºC)

and drying process, comparing non-ADO vs. ADO, as well as aligned vs. random

scaffolds. Data are reported as mean ± standard error (n=6).

0

10

20

30

40

50

60

Aligned PU Aligned PU

+ ADO

Random ADO

Elastic Modulus (M

Pa) As Made

Pre‐wet

(a) *

****

0

2

4

6

8

10

12

14

16

18

Aligned PU Aligned PU

+ ADO

Random ADO

Tensile Strength (M

Pa) As Made

Pre‐wet

(b) *

****

55

Figure 5. Differential Scanning Calorimetry for as-made and pre-wet/dried samples for (a)

aligned PU, (b) aligned PU+0.5%ADO and (c) random PU+0.5%ADO. Ti is the glass

transition temperature for the polycarbonate soft segment. Tii indicates the onset of

the soft segment melt phase. Tiii and Tiv are soft-segment melting transition

temperatures. Tv indicates the hard-segment melting transition temperature.

‐35 C32 C

67 C93 C

51 C

‐4

‐3

‐2

‐1

0

‐100 ‐50 0 50 100 150 200

Heat Flow (mW)

Temperature (°C)

DSC ‐ Aligned PU

prewetas‐made

ii

iii iv v

i

(a)

‐34 C 33 C

68 C

94 C

51 C

‐4

‐3

‐2

‐1

0

‐100 ‐50 0 50 100 150 200

Heat Flow (mW)

Temperature (°C)

DSC ‐ Aligned PU + 0.5% ADO

prewet

as‐madeii

iii iv v

i

(b)

‐36 C 33 C

67 C

55 C

93 C

‐4

‐3

‐2

‐1

0

‐100 ‐50 0 50 100 150 200

Heat Flow (mW)

Temperature (°C)

DSC ‐ Random PU + 0.5% ADO

prewet

as‐madeii

iii iv v

i

(c)

56

Figure 6. a) Initial Modulus and (b) Tensile Strength of the electrospun polyurethane nanofiber

scaffolds over four weeks of biodegradation in CE (100 units/ml) at 37ºC, PBS

pH=7.0. Data are reported as mean ± standard error (n=6). Aligned scaffolds showed

significantly higher modulus than random scaffolds at all time points. Ultimate stress

of aligned polymers decreased in the first week of degradation, but remained stable

thereafter.

* ** * *

0

2

4

6

8

10

12

14

Pre‐wet Week 1 Week 2 Week 3 Week 4

Initial M

odulus (M

Pa)

Aligned PU

Aligned PU+ADO

Random PU+ADO

(a)

0

2

4

6

8

10

Pre‐wet Week 1 Week 2 Week 3 Week 4

Ultim

ate Stress (M

Pa)

Aligned PU

Aligned PU+ADO

Random PU+ADO

**

**

(b)

57

Figure 7. Transmission Electron Microscopy of a non-soluble degradation product

58

Figure 8. Assessing the cytotoxicity of PU degradation products: MTT Assay was used to

evaluate potential cytotoxicity of various concentrations of (a) non-soluble and (b)

soluble degradation products on bovine annulus fibrosus cells. The experiment was

repeated 4 times (n=8 per condition). Data are expressed as mean ± SEM. H2O2 was

used as a positive control.

0%

20%

40%

60%

80%

100%

120%

Relative

Metabolic Activity

% Weight ( g / 100 ml) Non‐Soluble

Degradation Products in Feeding Media

Control PU ADO

a)

0%

20%

40%

60%

80%

100%

120%

Relative

Metabolic Activity

% Volume (Soluble Degradation

Products in PBS : Feeding Media)Control PU ADO

b)

59

Figure 9. Cell viability of PU degradation products: AF cells were incubated for 24 hours with

various concentrations of (a) non-soluble and (b) soluble degradation products.

Live/Dead Assay was used to assess cell viability. The number of dead cells were

counted and expressed as percent of total number of cells. The experiment was

repeated 4 times (n=8 per condition) and data expressed as mean±SEM. H2O2 was

used as a positive control.

0%

20%

40%

60%

80%

100%

% Live Cells (Live

/ Total)

% Weight ( g / 100 ml) Non‐Soluble

Degradation Products in Feeding Media

a)

Control PU ADO

0%

20%

40%

60%

80%

100%

% Live Cells (Live

/ Total)

% Volume (Soluble DegradationProducts in PBS: Feeding Media)

b)

Control PU ADO

60

Figure 10. Representatipve images of Live/Dead Assay of AF cells treated with (a) untreated

negative control (media with carrier); (b) H2O2-treated positive control; (c) 0.1 wt. %

non-soluble degradation products; (d) 100 volume% soluble degradation products

61

VIII: Application of Dynamic Compressive Forces on

Annulus Fibrosus Cells Grown on a Biodegradable

Electrospun Nanofiber Scaffold

62

Application of Dynamic Compressive Forces on

Annulus Fibrosus Cells Grown on a Biodegradable

Electrospun Nanofiber Scaffold

Masoud Yeganegi1, 2, J Paul Santerre2, 4, and Rita A Kandel 1, 2, 3

1CIHR- Bioengineering of Skeletal Tissues Team, Mount Sinai Hospital, Toronto, M5G 1X5

Canada

2Institute of Biomaterials and Biomedical Engineering and Department of Materials Science and

Engineering, University of Toronto, Toronto, M5S 3G9 Canada

3Department of Pathobiology and Laboratory Medicine, Mt. Sinai Hospital, University of

Toronto, Toronto, M5G 1X5 Canada

4Faculty of Dentistry, University of Toronto, Toronto, M5G 1G6 Canada

To whom correspondence should be sent:

Rita Kandel, M.D.

Professor, Dept. of Laboratory Medicine and Pathobiology


600 University Ave. Toronto, Ontario, Canada

M5G 1X5

Phone: 416-586-8516

Fax: 416-586-8628

E-mail: [email protected]

63

Introduction

The human vertebral column provides axial support to the body and protects the spinal cord.

The intervertebral discs lying between the vertebrae provide flexibility and help to prevent

damage to the vertebral column by dissipating mechanical loads and shocks.1 The hyaline

cartilage endplates found at each end represent the anatomical limits of the disc. The nucleus

pulposus forms the gelatinous central zone of intervertebral disc. Surrounding the nucleus

pulposus is a series of concentric fibrocartilaginous lamellae called the annulus fibrosus. Each

lamella consists of collagen fibers that are oriented at 60º to the vertical axis of the disc. The

alignment of these fibers alternates between successive lamellae and is of great importance to the

functionality of the annulus fibrosus.2 The composition of the AF is radially non-uniform.

Within the annulus fibrosus, the concentration of collagen type I is highest in the outer lamellae

and lowest toward in the centre.3 Conversely, collagen type II, which is also present in nucleus

pulposus increases in concentration toward the innermost lamellae in the annulus fibrosus. The

relative proportion of collagen type I to collagen type II in the annulus fibrosus varies from 70:30

in the innermost layers and 85:15 in the outer layers.4

Due to its structural arrangement, the mechanical behaviour of AF is relatively complex.

Studies indicate that the annulus fibrosus exhibits both matrix viscoelastic (flow-independent)

and biphasic viscoelastic (flow-dependent) behaviour. 5-8 The elastic modulus of a single lamella

of annulus fibrosus has been found to vary with the radial position in the disc displaying values

ranging from 5±4 MPa for posterior inner AF, 20±12 MPa for posterior outer AF, 10±6 MPa for

anterior inner AF, and 49±32 MPa for anterior outer AF 9. The ultimate stress of a single lamella

of annulus fibrosus similarly varies radially from 0.9±0.3 MPa for posterior inner AF, 1.1±0.3

MPa for posterior outer AF, 0.9±0.7 MPa for anterior inner AF, and 3.3±1.3 MPa for anterior

64

outer AF (n=15). 9 The forces experienced by AF can be quite high, and thus the impact of

mechanical forces on AF tissue is of great interest. It has been well established that mechanical

loading plays an important role in regulating the behavior of various tissues, including the AF.

Muscle forces, general loading of the joints, and movement of joints relative to each other act to

apply a range of stresses on the disc, including compressive, shear, tensile, osmotic and

hydrostatic. The AF cells sense these forces and respond through changes in gene expression.9

A number of studies have demonstrated the mechanosensitivity of the AF. The in vivo

biological response of annulus fibrosus in the rat tail, to short-term dynamic compression, has

been investigated by MacLean et al. 10-12 The expression of anabolic and catabolic genes was

affected by a 2 hour dynamic compression of the intervertebral disc, under an applied stress of 1

MPa (12.6N) at 0.2 Hz. Matrix genes, collagen I and collagen II, were slightly downregulated

while catabolic genes, collagenase and aggrecanase, were upregulated in the annulus.10 Stresses

of 1 MPa and 0.2 MPa were applied in another disc study at three frequencies of 0.01 Hz, 0.2 Hz,

and 1 Hz. It was found that the application of 1 MPa at all frequencies significantly increased the

catabolic genes, collagenase (21-, 7-, 8-fold respectively) and aggrecanase (5-, 1-, 7-fold

respectively) with only slightly increased collagen I expression at 1 Hz (3.5- fold) in the AF. On

the other hand, stress of 0.2 MPa at 1 Hz resulted in a slightly elevated level of expression for

collagen (4-fold) and aggrecan (2-fold), with minor non-significant increases in collagenase and

aggrecanase. This study has demonstrated the frequency and magnitude dependence of the

biological response to compressive mechanical stress. 11,12 Lotz and Walsh et. al. have also

explored the load- and frequency-dependant response of the intervertebral disc to dynamic

stresses. 13 Peak stresses of 0.8 MPa and 1.3 MPa were applied at frequencies of 0.1 Hz and 0.01

Hz to in vivo mice tail discs. Under these conditions, little apoptosis of AF cells was found

65

generally at the higher frequency and the lower stress. Aggrecan gene expression in the inner

annulus increased under lower frequency and higher stress loading. Similar results were

observed under almost identical experimental conditions, with peak compressive stresses of 0.9

MPa and 1.3 MPa applied at 0.1 Hz and 0.01 Hz. 13 Static compressive force of 1.0 MPa for 24

hr applied to ex-vivo cocygeal discs caused apoptosis in the annulus fibrosus.14 Annulus fibrosus

cells in an alginate culture system subjected to 30 hours of static 25% unconfined compressive

strain, responded with increased gene expression for types I and II collagen, and aggrecan.15

Application of hydrostatic pressure on caudal bovine and human IVD explants, while affecting

the nucleus pulposus and the inner AF, did not produce any significant changes in the outer

AF.16-18 Osmotic pressure has also been shown to affect mRNA levels of aggrecan, collagen-I,

and collagen-II in AF 3D-cultures.19-21 Interestingly, cellular response to hydrostatic and cyclic

tensile strain was found to be dependent on the osmotic environment.19 The AF is

physiologically subjected to tensile forces 9, and thus a response to tensile mechanical

stimulation is expected. Dynamic tensile strain (5% at 1Hz for 24 hours) of monolayer AF cells

grown on a collagen substrate resulted in decreased proteoglycan synthesis, while increasing

nitrogen oxide production.22 Higher tensile strains at lower frequencies (15% at 0.1Hz for 24

hours) increased cellular apoptosis in the AF.23 Axial traction tensile forces applied to intact IVD

explants (at 0.8MPa for 4 hours) resulted in a decrease in proteoglycan synthesis in the AF.24

Thus the effects of mechanical forces on AF cells can be variable depending on the conditions.

Back pain is ranked as the most prevalent chronic disease for people under 60.25 Degenerative

disc disease is associated with lower back pain and involves the progressive degeneration of the

intervertebral disc (IVD). An intact disc is necessary to support compressive and bending

stresses while providing flexibility to the spine.26 Currently, existing treatments include the use

66

of a prosthetic substitute, or the fusion of neighbouring vertebrae, neither of which is optimal.

Spinal fusion of degenerated disc may be effective in some cases, but a number of patients can

develop degeneration, due to reduced flexibility and increased stress, in adjacent segments. 27-30

Complications, such as slippage, wear and mechanical failure of the device materials can also

occur in patients undergoing disc replacement with synthetic substrates.31,32

One strategy towards addressing intervertebral disc degeneration is through the replacement of

the diseased tissue by a tissue-engineered substitute.33-35 Tissue engineering of annulus fibrosus

is of particularly challenging due to the complex structure of the tissue.2 Efforts in producing AF

tissue in vitro has involved various polymefric scaffolds including PDLLA/45S5 Bioglass®

films, polyglycolic acid, collagen/hyaluronan, collagen/gag, atelocollagen, and alginate

scaffolds.36-41 In addition to inadequate tissue formation, some biomaterials, such as

polyglycolic-based polymers, create acidic byproducts throughout their biodegradation, which

can significantly alter cell behavior, tissue production and possibly cause cell death.42,43 In this

study, degradable polycarbonate-urethane (PU) polymers were used due to their biocompatible,

biodegradable and reproducible nature. Electrospinning was employed to fabricate aligned fibers,

mimicking the aligned nature of the annulus fibrosus, and thus providing a more appropriate

surface for growth of such a tissue.

In a previous study, we demonstrated that AF cells can be grown on electrospun PU fibers and

they could synthesize and accumulate extracellular matrix.44 The high surface to volume ratio of

these scaffolds favors cell attachment and retention of cell phenotype. 44,45 Given the mechano-

sensitivity of AF cells, the purpose of this study was to assess the effect of dynamic compressive

67

mechanical forces on AF cells grown on an aligned biodegradable electrospun nanofiber

scaffold.

Experimental Section

In recent years, efforts have focused on developing bioactive/biocompatible biodegradable

polyurethanes for the purpose of tissue engineering or regeneration.46,47 In this study, a

degradable polycarbonate urethane (PU) was synthesized, as has been previously described, with

hexane diisocyanate:polycarbonate diol:butane diol ratio of 3:2:1 48. In addition, an anionic

dihydroxyl oligomer (ADO) additive was synthesized through the reaction of lysine

diisocyanate, polytetramethylene oxide, and hydroxyethyl methacrylate (HEMA).44 This anionic

additive has been previously shown to increase AF cellular adhesion mediated by protein

adsorption to the surface.44

Scaffold fabrication

Electospinning was employed to fabricate either random or aligned nanofiber scaffolds using

the base polymer (PU) with or without ADO (0.5 wt.%). The polymer was dissolved in

1,1,1,3,3,3-hexafluora-2-propanol at a concentration of 16 wt.%. The scaffold was then formed

by electrospinning, through the injection of the polymer solution from a metallic syringe at a rate

of 0.5 ml/hr onto the collecting surface. An 18 kilovolts difference was applied across the

syringe and the collecting surface. In the case of aligned scaffolds, the collecting surface

consisted of a cylindrical aluminum mandrel rotating at 1250 rpm, such that the surface of the

mandrel moved at 10 m/s. Relative humidity and temperature were maintained at <30% relative

humidity and approximately 25ºC. The resulting electrospun fibers ranged from 200-400 nm in

diameter as measured by scanning electron microscopy (Figure 1). Polymers were punched into

68

circular sections (D=6mm, and thickness of 80 ± 10µm) and fixated over a porous titanium disc

(D=4mm, h=2mm). The construct was held in place by the Tygon tubing (D=4mm) surrounding

the components. The tubing created a well-like structure to prevent cell spillage and provided

cells with a confined area to attach within a specific region of the scaffold. The porous titanium

base allowed for the application of compression to the scaffold while ensuring media diffusion

from below (Figure 2).

Annulus fibrosus cell culture

Bovine annulus fibrous cells were used as a model for human cells.49-51 Annulus fibrosus from

bovine caudal spines (6–9 months old) were dissected out aseptically. Five discs were combined

from one tail for each experiment. To isolate the cells, the tissues were chopped into 1mm pieces

and underwent serial digestion with 0.5% protease (Sigma, St. Louis, MO) for 1 hour at 37° C,

followed by 0.25% collagenase A (Roche, Laval, Quebec, Canada) overnight at 37° C. The cell

suspension was washed, filtered through a sterile mesh, and resuspended in Ham’s F12

supplemented with 5% fetal bovine serum (FBS). A 40µL aliquot of AF cell suspension was

seeded onto the surface of the scaffold at a density of 8x105/cm2 and allowed to attach for 3

hours. Ham’s F12 media containing 5% fetal bovine serum was then used to submerge the entire

construct. Ascorbic acid was added to media 72 hours post-seeding (final concentration of 100

µg/mL).

Mechanical Stimulation

Cells were mechanically stimulated using a Mach-1 mechanical tester (Biosyntech, Laval, PQ,

Canada) under confined compression. Titanium alloy plates with a porous surface layer (35 vol%

layer of sintered Ti·6Al·4V powders) 52,53 were used as supports for the PU membranes while

69

applying 1kPa of compressive force to cylindrical agarose gels (D=3.5mm, h=4mm, 2% agarose

in Ham’s F12 media), which had been placed over the cells within the tubing. The use of the

porous agarose inserts protected the cells and allowed the culture medium to diffuse to the cells.

The mechanical properties of the agarose gel likely differ from those of the AF tissue layer, and

thus the relative strain experienced is also likely to be different. However, because the layer of

cells lies between the agarose plug and the underlying polymer, the compressive force

experienced throughout the agarose gel and the AF tissue layer is expected to be the same, as

required by a state of mechanical equilibrium. Mechanical loading of the disc results in a

complex set of mechanical stimuli experience by the cells, including tensile, compressive and

shear load.54 Compression was chosen as a starting point for analysis of mechanical forces on the

AF tissue. In all cases, mechanical stimulation was conducted at a frequency of 1 Hz, 1kPa, for 1

hour. The aforementioned parameters for this mechanical stimulation study were chosen upon

examining previous studies in the area. The 1Hz frequency of dynamic compressive stimulation

was chosen since it represents the natural step frequency. Further, a number of studies have

demonstrated the frequency and magnitude dependence of the AF response to compressive

mechanical stress. Lower loads and appropriately higher frequencies have been found to increase

anabolic gene expression, while abnormally higher loads, lower frequency or static compression

have proved detrimental to tissue production and AF cell viability.11-14 Control cells were treated

in an identical manner but did not receive mechanical stimulation.

AF cell morphology

Cell morphology was evaluated at various time points following the application of mechanical

stimulation. The cell-scaffold constructs were washed in Ca2+ and Mg2+ free PBS three times and

fixed in 2.5% gluteraldehyde for 1 hour and stored at 4ºC. They were later dehydrated in

70

increasing concentrations of ethanol (i.e. 50%, 70%, 90%, 95%, 100%) before critical point

drying. All samples were sputter coated with gold and evaluated using scanning electron

microscopy (SEM) (FEI XL30 ESEM, Hillsbro, OR, USA).

DNA content

To determine cellularity at various time points, samples were papain digested (Sigma; 40

μg/mL, 1 mM EDTA, 20 mM ammonium acetate, and 2 mM DL-dithiothreitol) for up to 48

hours at 65°C. The DNA content of the cells was determined using the Hoechst 33258 dye assay

(Polysciences, Warrington, PA) and fluorometry (Thermoscientific model Fluoroskan at an

excitation wavelength of 365 nm, and emission wavelength of 458 nm). Calf thymus DNA

(Sigma, Oakville, ON) was used to generate a standard curve.

Quantification of Proteoglycan and Collagen Synthesis and Proliferation

Collagen and proteoglycan synthesis was determined by incubating cells with [3H]-proline and

[35S]-sulfate (4µCi/well) for 24 hours. The samples were harvested, and washed in Ca2+ and

Mg2+ free PBS three times, papain digested and incorporated radioactivity determined using a β-

liquid scintillation counter (Beckman model LS 6500, Mississauga, Ontario).

To measure proliferation, cells were incubated with [3H]-thymidine (2µCi/well) for 24 hours.

The samples were harvested, and washed in Ca2+ and Mg2+ free PBS three times, papain digested

and counted using a β-liquid scintillation counter. CPM (counts-per-minute) measurements were

normalized to DNA content.

71

Statistical analysis

All conditions were done in at least quadruplet and each experiment repeated at least 3 times.

The results from all experiments were combined and expressed as the mean ± standard error of

the mean. The data were analyzed using a one-way analysis of variance and all pair-wise

comparisons between groups were conducted using the Scheffe post hoc test. Significance was

assigned at p-values < 0.05.

Results

Effect of Mechanical Stimulation on AF Cells

The application of 1kPa of confined cyclic compressive force on 3-day old AF tissue at 1Hz

resulted in changes in cell morphology. Scanning electron microscopy showed that stimulated

cells spread noticeably more immediately post-stimulation and at 6 hours post-stimulation.

Further, AF cell density appeared to be greater in stimulated samples than their un-stimulated

counterparts (Figure 3). These differences were less apparent at later time points and mostly

absent by 72 hours. To verify the observation of varying cell density from the SEM data, DNA

content was measured post-stimulation (Figure 3). Cell density as measure by DNA content was

found to remain unaffected by the mechanical stimulation (Figure 5a). Further, proliferation was

also found to be comparable between stimulated and control samples for both 24 and 72 hour

time points.

Effect of Mechanical Stimulation on AF Matrix Synthesis

Dynamic compressive mechanical stimulation (1kPa, 1Hz, 1hr) similarly did not significantly

affect collagen synthesis (Figure 4a) or proteoglycan synthesis (Figure 4b) in the 24 hours post-

stimulation. Synthesis at 72 hours post-stimulation produced similar results, however, there was

72

a significant decrease in synthesis of collagen and proteoglycans at 72 hours compared to 24

hours post-stimulation. Despite the changes in morphology (Figure 3) which persisted for at least

72 hours, matrix synthesis (normalized to DNA) remained unaffected by the compressive

mechanical stimulation.

Discussion

With regards to cell spreading, previous studies have shown that when compressive

deformation is applied to AF cells in a three-dimensional culture system, an early increase in

vimentin mRNA expression and subunit polymerization occurs, characteristic of changes in the

cytoskeleton of AF cells.55 The observed changes in cell spreading in the current study are also

consistent with the findings by Handa et al. 55 In terms of AF cell density, since DNA content

was not significantly different in stimulated and non-stimulated samples, it was concluded that

the SEM images only provided a qualitative measure of the cellular density and did not reflect a

complete picture of the cell state or tissue surfaces. It is difficult to reconcile the SEM findings

alongside the cell analysis data. It is possible that the dehydration process required for SEM

imaging resulted in dislodging AF cells in un-stimulated samples. This would suggest that AF

cells in stimulated samples may have been more strongly attached than those of the control group

as a result of applied mechanical forces. Future studies would have to evaluate cell retention in

order to validate this hypothesis. Compressive mechanical forces did not result in significant

changes in matrix synthesis. It is possible that mRNA instability prevents changes in gene

expression to remain present long enough to result in detectable changes in matrix synthesis,

without further application of compressive force.56 Further, dynamic compressive stimulation of

AF cells has been shown to alter gene expression in AF in a frequency and magnitude dependant

manner. 10,12,13,57 In one such study, no significant changes were observed at 0.5hr of stimulation

73

(0.2 MPa), while significant changes were observed following 2hr and 4hr of stimulation.

Therefore, it is possible that AF cells simply did not respond strongly to the particular set of

parameters (1 kPa confined compression, 1Hz, 1hr) applied in this study. Future studies may

attempt to vary the applied force and duration of mechanical stimulation. Other parameters that

can be investigated in the future, to determine AF response to mechanical stimulation include

apoptosis, changes in cell-specific matrix genes (types I, and II collagen and aggrecan) and

catabolic genes (MMP-3, MMP-13, and ADAMTs-4). Different modes of mechanical force such

as tensile, shear or hydrostatic forces may also be influential on AF cells and should be explored.

Conclusions

The findings in this study have shown that confined dynamic compressive mechanical

stimulation of 1kPa at 1Hz caused an increase in extent of cell spreading early on as per SEM

images. These effects were observed immediately following stimulation and persisted for at least

72 hours post-stimulation. However, the morphological effects did not result in significant

changes in DNA content, cell proliferation, or matrix synthesis at 24 or 72 hours post-

stimulation. Matrix synthesis decreased for both stimulated and control samples at 72 hours

when compared to 24 hours post-stimulation. The results of this study, suggest that compressive

forces on the AF cells under the aforementioned conditions, have little influence on AF tissue

growth. Additional studies should be aimed at alternative mechanical stimulation parameters to

yield a more significant response from AF cells.

Acknowledgements

The funding for this work was provided by the Ontario Graduate Scholarship in Science and

Technology (OGSST), the University of Toronto Fellowship Award, an NSERC-CIHR

74

Collaborative Health Research Program (CHRP) Grant (312882), and a CIHR Operating Grant

(MOP86723).

75

References:







159-171.





1271.




environment of intervertebral disc cells determined by a finite deformation, anisotropic,

and biphasic finite element model. J.Biomech.Eng, 2003, 1, 1-11.

(8) Wu, H. C. and Yao, R. F. Mechanical behavior of the human annulus fibrosus. J.Biomech.,

1976, 1, 1-7.

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(10) MacLean, J. J., Lee, C. R., Grad, S. et al. Effects of immobilization and dynamic

compression on intervertebral disc cell gene expression in vivo. Spine, 15-5-2003, 10,

973-981.

(11) Iatridis, J. C., MacLean, J. J., Roughley, P. J., and Alini, M. Effects of mechanical loading

on intervertebral disc metabolism in vivo. J.Bone Joint Surg.Am., 2006, 41-46.

(12) MacLean, J. J., Lee, C. R., Alini, M., and Iatridis, J. C. Anabolic and catabolic mRNA

levels of the intervertebral disc vary with the magnitude and frequency of in vivo

dynamic compression. J.Orthop.Res., 2004, 6, 1193-1200.

(13) Walsh, A. J. and Lotz, J. C. Biological response of the intervertebral disc to dynamic

loading. J.Biomech., 2004, 3, 329-337.

(14) Ariga, K., Yonenobu, K., Nakase, T. et al. Mechanical stress-induced apoptosis of endplate

chondrocytes in organ-cultured mouse intervertebral discs: an ex vivo study. Spine, 15-

7-2003, 14, 1528-1533.

(15) Chen, J., Yan, W., and Setton, L. A. Static compression induces zonal-specific changes in

gene expression for extracellular matrix and cytoskeletal proteins in intervertebral disc

cells in vitro. Matrix Biol., 2004, 7, 573-583.



disc. Spine, 15-5-1997, 10, 1085-1091.

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(17) Ishihara, H., McNally, D. S., Urban, J. P., and Hall, A. C. Effects of hydrostatic pressure on

matrix synthesis in different regions of the intervertebral disk. J.Appl.Physiol, 1996, 3,

839-846.

(18) Liu, G. Z., Ishihara, H., Osada, R., Kimura, T., and Tsuji, H. Nitric oxide mediates the

change of proteoglycan synthesis in the human lumbar intervertebral disc in response to

hydrostatic pressure. Spine, 15-1-2001, 2, 134-141.

(19) Wuertz, K., Urban, J. P., Klasen, J. et al. Influence of extracellular osmolarity and

mechanical stimulation on gene expression of intervertebral disc cells. J.Orthop.Res.,

2007, 11, 1513-1522.

(20) Boyd, L. M., Richardson, W. J., Chen, J. et al. Osmolarity regulates gene expression in

intervertebral disc cells determined by gene array and real-time quantitative RT-PCR.

Ann.Biomed.Eng, 2005, 8, 1071-1077.

(21) Chen, J., Baer, A. E., Paik, P. Y., Yan, W., and Setton, L. A. Matrix protein gene

expression in intervertebral disc cells subjected to altered osmolarity.


(22) Rannou, F., Richette, P., Benallaoua, M. et al. Cyclic tensile stretch modulates

proteoglycan production by intervertebral disc annulus fibrosus cells through

production of nitrite oxide. J.Cell Biochem., 1-9-2003, 1, 148-157.



Am.J.Pathol., 2004, 3, 915-924.

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stress on solute transport and proteoglycan synthesis in the porcine intervertebral disc

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1290-1297.

(42) Ishihara, H. and Urban, J. P. Effects of low oxygen concentrations and metabolic inhibitors

on proteoglycan and protein synthesis rates in the intervertebral disc. J.Orthop.Res.,

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comparison of the human and bovine thoracolumbar spine. J.Orthop.Res., 1986, 3, 298-

303.

(51) Wilke, H. J., Krischak, S., and Claes, L. Biomechanical comparison of calf and human

spines. J.Orthop.Res., 1996, 3, 500-503.

(52) Spiteri, C. G., Young, E. W., Simmons, C. A., Kandel, R. A., and Pilliar, R. M. Substrate

architecture and fluid-induced shear stress during chondrocyte seeding: role of

alpha5beta1 integrin. Biomaterials, 2008, 16, 2477-2489.

(53) Bhardwaj, T., Pilliar, R. M., Grynpas, M. D., and Kandel, R. A. Effect of material

geometry on cartilagenous tissue formation in vitro. J.Biomed.Mater.Res., 2001, 2, 190-

199.

(54) Setton, L. A. and Chen, J. Mechanobiology of the intervertebral disc and relevance to disc

degeneration. J.Bone Joint Surg.Am., 2006, 52-57.

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disc. Spine, 15-5-1997, 10, 1085-1091.

(56) MacLean, J. J., Lee, C. R., Alini, M., and Iatridis, J. C. The effects of short-term load

duration on anabolic and catabolic gene expression in the rat tail intervertebral disc.

J.Orthop.Res., 2005, 5, 1120-1127.

(57) Kim, P. K. and Branch, C. L., Jr. The lumbar degenerative disc: confusion, mechanics,

management. Clin.Neurosurg., 2006, 53, 18-25.

83

Figures

Figure 1. Scanning Electron Microscopy Images of Aligned (a) and Random (b) Electrospun

Polycarbonate Urethane Nanofiber Scaffolds

84

Figure 2. Mechanical Stimulation Apparatus: The polymer is held in place over the porous

titanium base by Tygon tubing, which in turn allows seeding media to remain in

contact with the scaffold during cell seeding. At the time of mechanical stimulation,

an agarose plug is placed above the cells to transmit the force through to the cells.

Titanium plate

Agaroseplug

Tygon Tubing

Cells on the Scaffold

Porous Titanium Base

85

a ) b )

c ) d )

e ) f )

86

Figure 3. SEM images of AF cells immediately (b), 6 hr (d); 24 hr (f); 72 hr (h) post-

stimulation. The corresponding non-stimulated controls are denoted (a, c, e, g).

Stimulated samples are more spread than control samples, particularly at early time

points.

a) b)

c) d)

g ) h )

87

Figure 4. DNA Content (µg) at various time points following mechanical stimulation (3 day

Tissue, stimulated for 1hr at 1Hz and 1kPa). No significant changes were observed

between the control and stimulated groups nor between the different time points (n=3,

α=0.05).

0

0.1

0.2

0.3

0.4

0.5

0 hr 6 hr 12 hr 24 hrDNA (ug DNA)

Time (hours) post‐stimulation

Control

Stimlulated

88

Figure 5. Evaluation of DNA content (a) and Thymidine Incorporation (b) at 24 hours and 72

hours post-stimulation. Controls were treated similarly but were not stimulated. The

results are expressed as ± SEM (n=15, α=0.05).. No significant differences were

detected between the two conditions.

0.0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

24 hr

[Stim.]

24 hr

[Ctrl.]

72 hr

[Stim.]

72 hr

[Ctrl.]

DNA Content (ug)

a)

0

500

1000

1500

2000

2500

3000

3500

4000

4500

5000

24 hr [Stim.]

24 hr [Ctrl.]

72 hr [Stim.]

72 hr [Ctrl.]

3H‐Thym

idine‐Incorporation

(CPM / ug DNA)

b)

a) b)

c) d)

e) f)

89

Figure 6. Collagen (a) and Proteoglycan (b) synthesis at 24 hours and 72 hours post-

stimulation. No significant changes were observed between the control and stimulated

groups at either time point. However, there is a significant decrease in matrix

accumulation, by 72 hours compared to 24 hours post-stimulation, for the combined

group of stimulated and non-stimulated samples, (N=15, α=0.05).

0

1000

2000

3000

4000

5000

6000

7000

8000

24 hr [Stim.]

24 hr [Ctrl.]

72 hr [Stim.]

72 hr [Ctrl.]

3H‐OH‐Proline accumulation

(CPM / ug DNA)

a)*

0

500

1000

1500

2000

2500

3000

24 hr

[Stim.]

24 hr

[Ctrl.]

72 hr

[Stim.]

72 hr

[Ctrl.]

35SO

4Incorporation

(CPM / ug DNA)

b)*

90

IX: Conclusions and Future Work

91

Conclusions and Future Work

The findings in this thesis have demonstrated the basic characteristics that make the

biodegradable electrospun polycarbonate urethane nanofiber scaffold suitable for producing a

tissue-engineered Annulus Fibrosus. Aligned scaffolds were found to possess superior

mechanical properties in relation to random scaffolds, suggesting that this formulation is a more

appropriate scaffold for engineering annulus fibrosus tissue. While it was found that exposure to

aqueous media disrupted the material structure and resulted in a reduction of the mechanical

properties, the mechanical properties remained comparable to those of native tissue itself. The

observed drop in mechanical strength immediately following the start of incubation for the

aligned scaffolds was explained by differential scanning calorimetry, which showed the

appearance of a disrupted soft segment crystal phase with changes in the degree of phase mixing

of hard and soft segments as well as the PCN crystal state within the polymer.

Of particular importance, it was found that the degradation of the materials, which resulted in

mass losses as high as 30% over four weeks, did not result in a significant deterioration of

mechanical properties, indicating a surface degradation process rather than bulk material

breakdown. The presence of surface degradation may result in a more predictable degradation

rate and rate of release for the degradation products, since diffusion through the bulk is not a

contributing factor.

The degradation of the polymer by cholesterol esterase provided a useful model for

biodegradation, yielding a controlled and consistent mass loss rate, at which CE mediated

surface degradation was primarily observed. The degradation products did not cause significant

acute cytotoxicity in-vitro, indicating that this biodegradable polymer may be appropriate as a

92

substrate for AF cells. Confined compressive mechanical forces were shown to instigate slight

changes to cell morphology, while leaving matrix production unaffected and the dynamic

compressive mechanical stimulation of 1kPa at 1Hz appeared to cause an increase in extent of

cell spreading early on, and an unconfirmed increase in cell density at later time points. These

effects were observed immediately following stimulation and persisted for at least 72 hours post-

stimulation. However, these morphological effects did not result in significant changes in DNA

content, cell proliferation, or matrix synthesis at 24 or 72 hours post-stimulation. It is therefore

recommended that the morphological differences observed under SEM be confirmed by looking

at more specific indicators of AF cellular adhesion.

Future studies may attempt to vary the applied force and duration of mechanical stimulation.

Other parameters that can be investigated to determine the AF response to mechanical

stimulation include apoptosis, changes in cell-specific matrix genes (types I, and II collagen and

aggrecan) and catabolic genes (MMP-3, MMP-13, and ADAMTs-4). Further, different modes of

mechanical force such as tensile, shear or hydrostatic forces may be more influential on AF cell

functionality and should be explored. Additional mechanical studies to explore the changes in

mechanical properties of the aligned polymer in the presence of AF tissue grown on the PU

substrate will provide further understanding of the role of this polymer for AF tissue engineering.

The results of this report, including the relatively constant rate of material degradation, the

observed mechanical behavior resembling that of AF tissue, and the absence of cytotoxic effects

make this polymer a suitable biomaterial candidate for use in the formation of tissue-engineered

annulus fibrosus. Studies have shown annulus fibrosus cells to be responsive to tensile

mechanical stimulation.1-4 While compressive forces on the AF cells under the aforementioned

93

conditions were found to have little influence on tissue growth, future work involving varied

conditions for compressive stimulation or the use of tensile mechanical stimulation, may be more

helpful to promote tissue production that more closely mimics that of the native annulus fibrosus.

94

References:

(1) Benallaoua, M., Richette, P., Francois, M. et al. Modulation of proteoglycan production by

cyclic tensile stretch in intervertebral disc cells through a post-translational mechanism.

Biorheology, 2006, 3-4, 303-310.


stress on solute transport and proteoglycan synthesis in the porcine intervertebral disc in

vitro. Eur.Spine J., 1994, 6, 325-330.

(3) Rannou, F., Richette, P., Benallaoua, M. et al. Cyclic tensile stretch modulates proteoglycan

production by intervertebral disc annulus fibrosus cells through production of nitrite

oxide. J.Cell Biochem., 9-1-2003, 1, 148-157.



Am.J.Pathol., 2004, 3, 915-924.

95

95

Appendix A: Scaffold Preparation

Electrospinning

Electrospinning is a process that produces polymer fibers with diameters ranging from a few

nanometers to a few microns. The high voltage power supply produces a voltage in the range of

0–30 kV or higher. For safety, the current should be limited to a few hundred microamperes.

When a strong electrostatic field is applied to the syringe needle or a capillary, a droplet of the

polymer solution held at the tip by surface tension is deformed into a conical shape, which is

called the Taylor cone. When electrostatic force overcomes the surface tension of the solution, a

liquid jet is ejected from the tip of Taylor cone. A polymer nanofiber can be formed after

solvent evaporates while the jet moves from the tip to the grounded collector. There are usually

four parameters affecting the formation of nano-scale polymer fibers, i.e. concentration of

polymer solution, flow rate of polymer solution, distance between needle and collector, and

voltage.

Preparation of nanofibrous polycarbonate urethane scaffolds (random scaffolds)

16% polymer solution (can vary between 15-20%) was prepared by dissolving

polycarbonate in 1,1,1,3,3,3-hexafluora-2-propanol (i.e. 0.4 g in 2.5 ml of HFP).

The solution viscosity was compared relatively to previously prepared solutions using a

simple gravity-based capillary viscometer consisting of a hypodermic needle and 1mL

syringe to ensure consistency between different scaffold fabrications

The polymer solution was transferred to a BD 10 ml syringe, to which an 18G stainless

steel needle was attached. All of air bubbles were removed.

The syringe was fixed to the syringe pump and the wire supplying the voltage

difference was connected to the metallic needle far from its tip.

A foil paper covered collector was placed 18 cm below the needle and connected to the

ground terminal of the voltage generator.

96

96

The syringe pump was turned on at 0.5 ml/hr and the desired volume (1.5-2.5mL) was

delivered.

The power generator was powered up to 18 kilovolts.

Preparation of nanofibrous polycarbonate urethane scaffolds (aligned scaffolds)

16% polymer solution (can vary between 15-20%) was prepared by dissolving

polycarbonate in 1,1,1,3,3,3-hexafluora-2-propanol (i.e. 0.4 g in 2.5 ml of HFP).

The solution viscosity was compared relatively to previously prepared solutions using a

simple capillary viscometer consisting of a hypodermic needle and 1mL syringe to

ensure consistency between different scaffold fabrications

The polymer solution was transferred to a BD 10 ml syringe, to which an 18G stainless

steel needle was attached. Traces of air bubbles were removed.

The syringe was fixed to the syringe pump and the wire supplying the voltage

difference was connected to the metallic needle far from its tip.

The mandrel was placed directly 18cm below the needle and attach the ground terminal

to the mandrel axle.

The surface of the mandrel edge was covered with a strip of foil paper (~48cm x 2cm).

The mandrel speed was adjusted to 1250 rpm (may be varied to optimize conditions)

The syringe pump was set to 0.5 ml/hr and the desired volume (1.5-2.5mL) was

delivered.

Turn on the power generator and slowly increase the voltage to 18 kilovolts.

The power generator was powered up to 18 kilovolts.

Note: 0.05% ADO was added to the solution where required. Following the electrospinning

procedures, all scaffolds were γ-irradiated at 4 MRad (Gammacell 220 Research Irradiator, MDS

Nordion, Canada).

97

97

Humidity Effects

It was found that increased humidity resulted in detrimental polymer quality, by affecting the

degree of solvent evaporation. A number of instrument set ups were devised in an attempt to

lower the humidity of the enclosure containing the electrospinning system. A dehumidifier was

used to pump dehumidified air into the enclosure. However these did not succeed due to

disruption of fiber deposition by the air current, and the increased temperature within the

enclosure (Figure A.1). The final design (Figure A.2) involved the delivery of dehumidified air

in a uniform and controlled manner (Figure A.3), such that the fiber deposition remained

unaffected (Figure A.4). In addition, the dehumidified air was cooled from 50ºC to 25ºC via a

cooling reservoir.

98

98

Figure A.1. Various instrumental apparatus (A to C) constructed to attempt to control

humidity using a dehumidifier (full and partial air flow into a close/open enclosure) with

corresponding SEM images of the resultant scaffolds. The effects of air current and increase in

temperature (due to the heat carried from the pump by the dehumidified air), scaffold alignment

and fiber diameter was found to be inferior under all conditions.

A'

B'

C'

A

B

C

Sealed Enclosure Dehumidifier

Partially Open Enclosure

Sealed Enclosure Dehumidifier

Dehumidifier

99

99

Figure A.2. Final apparatus for regulating humidity to 30% R.H. The effects of air current

were reduced by introducing a porous wooden base on which the electrospinning apparatus was

placed. Further, a cooling reservoir was used to cool the dehumidified air from 50 ºC to 25ºC.

Dehumidifier

48°C

Flexible Plastic Duct

Cooling Reservoir Containing Ice-Water (0-4° C)

25°C

Electrospinning Enclosure

100

100

Figure A.3. The humidity profile using the final equipment set-up. Humidity remains

stabilized at the desired level (<30% R.H.) after approximately 10-20 minutes of starting the

dehumidification.

0%

10%

20%

30%

40%

50%

60%

70%

0 10 20 30

Relative

Humidty

Time (Minutes)

Electrospinning Enclosure

Fume Hood (Containing the Enclosure)

101

101

Figure A.4. Scanning electron microscopy indicating processed fiber dimension, their

alignment, and confirming that transverse fibers were not a significant occurrence

102

102

Appendix B: Biodegradation

The activity of the cholesterol esterase enzyme was measured by analyzing its degradation of

p-nitrophenylbutyrate (p-NPB) into a non-soluble precipitate. 1 CE unit was defined as

generation of 1nmol/min of p-nitrophenol from p-nitrophenylbutyrate.

Reagent Preparation:

1. 1 liter 50mM sodium phosphate buffer was prepared by dissolving 2.6908 gram of

NaH2PO4.H2O; 4.3298 gram Na2HPO4 in 1L of Millipore filtered water

2. 4mM p-nitrophenylbutyrate (p-NPB) substrate solution was prepared by dissolving

17.75l of p-NPB (F.W. 209.2, density: 1.2g/ml in 25 C) to 5.5 ml of acetonitrile in a

25ml glass tube, and adding 19.5ml 50mM Sodium phosphate buffer. The solution was

stored at -70C.

3. The initial standard enzyme solution was prepared by dissolving 15mg of Cholesterol

Esterase (CE) (Sigma; 683U/mg, C-3766) in 50 ml 50mM phosphate buffer, pH 7.0.

Standard Curve for CE Enzyme Activity

1) The Tungsten lamp of DU800 device was warmed up for 20 min before use and set to

401nm.

2) In a 1.5 mL cuvette, the following was added:

50 L of Enzyme Solution

950 L of 50mM Sodium phosphate buffer

500 L of 4mM p-NPB

CE p-nitrophenylbutyrate (p-NPB) p-nitrophenol (yellow) + butyrate

103

103

3) Optical density of the solutions at 401nm was measured every 30 seconds for 300

seconds

4) The average OD/minute can be determined using the plot.

Calculating CE Activity

The absorbance of samples at 401nm is related to the concentration of the generated p-

nitrophenol by the Beer–Lambert law: A= LC, where C is the molar concentration

(mol/L) of p-nitrophenol in sample, L is the path-length of light (1cm), is the molar

extinction coefficient (16,000 L/mol/cm at 7.0 pH and 401nm for p-nitrophenol)

Let T2 and T1 represent different time points with corresponding absorbance of A2 and A1.

The following equations describe the relationship between the absorbance plot and changes

in p-NPB concentration).

/ /·

10 · /16000

Activity of CE working solution (units/ml) = 0.0015 · 10 · /16000 /0.05

The activity of CE was measured in the presence and absence of the polymer. It was

determined that the half-life of CE in the presence of the polymer was approximately 12 hours.

Thus, small volumes of concentrated CE solution were added daily to the polymer solutions to

adjust the enzyme activity to 10 units/ml.

104

104

Scaffold Thickness

The thickness of the various scaffold groups are shown in Figure B.1. The differences in

thickness between the aligned and random scaffolds stem from the fact that following the

fabrication process, thinner random polymers are difficult to remove from the deposition surface

without being subjected to deformation, due to their inferior mechanical properties. Thus thicker

random scaffolds were fabricated to prevent polymer plastic deformation prior to mechanical

testing. On the other hand, thicker aligned scaffolds could not be fabricated, since additional

deposited fibers would begin to lose their aligned nature. It was therefore not possible to

overcome this disparity in polymer thickness, as would be ideal for such an experiment. A

consistent decline in thickness was observed throughout the four weeks of biodegradation, as

would be expected given the presented mass loss. Further, the suspected surface-mediated

degradation seems to be supported by this data as well. The cumulative decline in thickness

appears to be independent of the original thickness of the scaffolds.

Figure B.1. Changes in the thickness of scaffolds throughout the biodegradation process

0

0.1

0.2

0.3

0.4

0.5

0.6

As‐made Pre‐wet Week 1 Week 2 Week 3 Week 4

Thickn

ess (m

m)

Aligned PU+ADO Aligned PU Random PU+ADO

105

105

Appendix C: Mechanical Testing

The nanofiber scaffolds, which measured about 3cm in length, were held at both ends in an

Instron® model 8501 mechanical testing device. A tensile force was applied such that the strain

rate remained at 10 mm/min. Displacement and force data was collected, the dimensions of the

polymers noted and used to convert these raw measurements to stress and strain data. Initial

modulus was determined through analyzing the initial linear behavior of the stress strain curves.

It is important to note that aligned scaffolds were prone to the presence of folds upon

immobilization within the clamps. This folding phenomenon was not observed in the case of

random fibers due to their higher thickness and easier handling. Thus during the mechanical

testing procedure for it was difficult to begin applying the tensile stress in such a manner that all

points along the cross-section of an aligned sample begin to experience strain at the same

moment. Soon after the test begins, as the entire sample begins to experience strain at a uniform

rate with the disappearance of the folds within the scaffold surface. This phenomenon explains

the very small initial toe region visible in some stress-strain curves (such as in Figure C.2A). Due

to the expected fluctuations in the raw data, the linearity of the initial portion of the curve was

extracted by reducing as much as possible, the random variations from the raw data. It was found

that the initial modulus (the longest linear portion found at the outset of the stress strain curves)

was a consistent and relevant measure of material behavior. The ultimate stress, while not as

consistent, was nonetheless analyzed due to its importance.

106

106

Figure C.1. Weekly tensile testing of scaffolds: Each polymer sample was clamped on either

side and tested using an Instron® model 8501 under a tensile strain of 10 mm/min to the

breaking point.

clamp

scaffold

Site of failure for a valid test

sample

107

107

Figure C.2. Stress-strain curves for PU aligned scaffolds under tensile mechanical stress. The

various curves are repeats of the same sample group. The initial modulus was found by

calculating the slope of the stress-strain curve in the initial elastic portion of each curve. Ultimate

stress was also reported on. The ultimate strain is defined by the strain experienced by the

sample at the ultimate stress.

0

5

10

15

20

0 0.5 1 1.5

Stress (M

Pa)

Strain

0

5

10

15

20

0 0.5 1 1.5 2

Stress (M

Pa)

Strain

0

1

2

3

4

5

0 0.2 0.4 0.6 0.8 1

Stress (M

Pa)

Strain

0

0.5

1

1.5

2

2.5

3

3.5

0 0.2 0.4 0.6 0.8 1Stress (M

Pa)

Strain

0

0.5

1

1.5

2

2.5

0 0.2 0.4 0.6 0.8

Stress (M

Pa)

Strain

0

0.5

1

1.5

2

2.5

0 0.2 0.4 0.6 0.8 1

Stress (M

Pa)

Strain

A: as-made

C: Week 1

E: Week 3

B: prewetted

D: Week 2

F: Week 4

108

108

Figure C.3. Stress-strain curves for PU + 0.5% ADO aligned scaffolds under tensile

mechanical stress: The various curves are repeats of the same sample group. The initial modulus

was found by calculating the slope of the stress-strain curve in the initial elastic portion of each

curve. Ultimate stress was also reported on. The ultimate strain is defined by the strain

experienced by the sample at the ultimate stress.

0

5

10

15

20

0 0.5 1 1.5

Stress (M

Pa)

Strain

0

5

10

15

20

0 0.5 1 1.5 2

Stress (M

Pa)

Strain

0

1

2

3

4

5

0 0.2 0.4 0.6 0.8 1

Stress (M

Pa)

Strain

0

1

2

3

4

5

0 0.2 0.4 0.6 0.8 1 1.2 1.4Stress (M

Pa)

Strain

0

1

2

3

4

5

0 0.2 0.4 0.6 0.8 1

Stress (M

Pa)

Strain

0

1

2

3

4

5

0 0.2 0.4 0.6 0.8 1

Stress (M

Pa)

Strain

A: as-made

C: Week 1

E: Week 3

B: prewetted

D: Week 2

F: Week 4

109

109

Figure C.4. Stress-strain curves for PU + 0.5% ADO random scaffolds under tensile

mechanical stress: The various curves are repeats of the same sample group. The initial modulus

was found by calculating the slope of the stress-strain curve in the initial elastic portion of each

curve. Ultimate stress was also reported on. The ultimate strain is defined by the strain

experienced by the sample at the ultimate stress.

0

1

2

3

4

0 5 10

Stress (M

Pa)

Strain

0

1

2

3

4

0 5 10 15

Stress (M

Pa)

Strain

0

0.5

1

1.5

2

0 1 2 3 4 5

Stress (M

Pa)

Strain

0

0.5

1

1.5

2

2.5

3

0 2 4 6

Stress (M

Pa)

Strain

0

0.5

1

1.5

2

2.5

0 2 4 6

Stress (M

Pa)

Strain

0

0.5

1

1.5

2

2.5

0 1 2 3 4 5

Stress (M

Pa)

Strain

A: as-made

C: Week 1

E: Week 3

B: prewetted

D: Week 2

F: Week 4

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Appendix D: Annulus Fibrosus Tissue Culture

In vitro culture was used in the cytotoxic evaluation and mechanical stimulation studies. The

AF cell seeding procedure is outlined below.

1. Biopsy punches were used to cut fibrous PU membranes into pieces 6 mm in diameter

2. Circular polymer pieces were placed on top of cylindrical Tygon tubing (4mm in

diameter and 6mm in height). A porous titanium disc was used to push and fit the

polymer within the Tygon tubing (see Figure D.2E).

3. The constructs were irradiated at 4 MRad

4. The Tygon/membrane/titanium disc constructs were incubated overnight in 24 well

plates containing 1.5 mL of Ham’s F12 media.

5. Bovine tails were dissected (Figure D.); outer AF tissue was removed and chopped into

1mm pieces. The tissue underwent serial digestion with 0.5% protease (Sigma, St.

Louis, MO) for 1 hr at 37° C, followed by 0.25% collagenase A (Roche, Quebec,

Canada) overnight at 37° C.

6. The cell suspension was washed, filtered through a sterile mesh, and resuspended in

Ham’s F12 supplemented with 5% fetal bovine serum (FBS).

7. 20-30 µL Ham’s F12 supplemented with 5% FBS was placed into each well of a 96-

well plate; constructs were then added.

8. 40µL of cell suspension containing the desired AF cell numbers (8x105/cm2) was added

to the top of the scaffolds, ensuring all air bubbles were removed and the cells were

allowed to adhere for 2-4 hours in the incubator.

9. Additional Ham’s F12 supplemented with 5% FBS was then added to submerge

constructs entirely.

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111

10. After 24 hours, the constructs were transferred to a 24-well plate containing 2mL of

media

11. The media was replenished every 2 days with 1.5mL of F12 supplemented with

5%FBS in addition to ascorbic acid (final concentration of 100 µg/mL) starting on day

3.

12. Constructs were washed 3 times in serum-free Ham’s F12 media containing ascorbic

acid and further incubated in serum free F12 media overnight prior to mechanical

stimulation experiments.

Figure D.1. Multiple discs were dissected from a single tail and the isolated AF cells were

combined to provide sufficient cells for an experiment and improve consistency. Only outer AF

cells were used in all experiments.

IVD

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Optimization of protocol for cell seeding on scaffolds:

Several methods were attempted for cell seeding on polymer scaffolds, each with a number of

problems, such as scaffold folding/wrinkling, cell suspension spillage, and the inability to

immobilize the seeded scaffold for mechanical stimulation. These issues were resolved in the

finalized setup where polymers were biopsy-punched into circular sections (D=6mm, and

thickness of 80 ± 10µm) and fixed over a porous titanium disc (D=4mm, h=2mm), by the Tygon

tubing (Figure D.2E). The tubing created a well-like structure to prevent cell spillage and

provided cells with a confined area to attach within a specific region of the scaffold. The porous

titanium base allowed the application of compression to the scaffold while ensuring media

diffusion from below. A cell seeding density optimization study was performed, where cell-

seeded scaffolds were papain digested and cell attachment was measured through analysis of

DNA content. The extent and uniformity of cell attachment was evaluated by SEM analysis.

Lower seeding densities were found to produce higher percent attachment (of original seeded

cell suspension). Further, a seeding density of 8 million cells / cm2 produced thick cellular layers,

where cell-cell contact dominated cell-polymer contact (Figure D.3). A seeding density of 0.8

million cells / cm2 was chosen to ensure that cell-polymer contact was sufficient (Figure D.4).

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113

Figure D.2. Methods evaluated (A to D): (A) Cell suspension on the polymer scaffold alone,

(B) Cell suspension on scaffold, supported by an agarose gel base, (C) cell suspension confined

by a Teflon insert, (D) cell confinement through the use of Tygon tubing; (E) The seeding

method selected for all subsequent experiments which consisted of using a Tygon tubing and a

porous titanium disc and (F) the corresponding apparatus for mechanical stimulation of tissue.

Titanium plate

Agarose plug

Tygon Tubing

Cells on the Scaffold

Porous Titanium Base

A B C

E F

D

Cell suspension

Agarose Gel

Teflon Insert

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Figure D.3. SEM images at low (A) and higher magnification (B) showing scaffolds seeded at

0.8 million cells / cm2. This density produced cellular layers, where cell-cell contact dominated

cell-polymer contact. It was therefore decided to reduce cell seeding density to 0.8 million cells /

cm2

A

B

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115

Figure D.4. Percent cell attachment to determine optimal seeding density: DNA content was

measured 24 hours after seeding. The attachment level dropped significantly at the highest

seeding density. The lower seeding density of 0.8 million cells / cm2 was chosen for subsequent

mechanical stimulation studies (N = 6 per condition).

0%

10%

20%

30%

40%

50%

60%

0.8 2.0 4.0 8.0

Seeding Density: million AF cells / cm2

*% AF Cell Attachmen

t

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Appendix E: Cytotoxicity Evaluation

Cytotoxic evaluation of degradation products was performed using the MTT and Live/Dead

Assays:

Frozen degradation products (previously maintained in PBS), were thawed.

Degradation solutions were mixed and spun down at 3000 RCF to form a non-soluble

pellet

The supernatant was filtered using a syringe filter of size 0.20 um

The non-soluble degradation products were resuspended in F12 containing 5% FBS.

Soluble and non-soluble degradation products were added to 96-well plates containing

200k/cm2 AF cells in monolayer cultures and incubated for 24 hours (160µL per well).

Degradation products were combined from 4 samples for each specific condition.

Following the 24 hour incubation period, the MTT and Live/Dead Assays was

performed

Media was aspirated and 1 mL of fresh medium containing MTT or Live/Dead reagents

were added to the cells

MTT Assay

It was found that the appropriate cell density (near confluent) of 100k/cm2 monolayer

cultures produced satisfactory differences between negative controls and positive controls

(Figure E.). Two hours was found to be sufficient to detect such a difference, without

reaching a saturation point

The solutions were allowed to incubate (at 37ºC and 5% CO2) for 2 hours before

the MTT solution was aspirated

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117

200µl of methylcellusolve (pH3.5) was used to dissolve the formazan precipitate

(plates were shaken for one to two minutes to allow the precipitate to dissolve)

200µl aliquots of the formazan solutions were pipetted into a 96 well plate and read

at 570nm

Live/Dead Assay

The degradation product solution was gently aspirated, and the cells were washed

lightly with Ham’s F12 media. This process was performed for both the positive

and negative controls as well.

160µL of Ham’s F12 media containing 4 μM calcein AM and 4 μM ethidium

homodimer EthD-1 (Invitrogen L-3224, Burlington, Ontario) were added directly

to cells.

Each sample was allowed to incubate (at 37 ºC and 5% CO2) for 15 minutes prior

to confocal imaging (Figure E.2) (Calcein: 494/517 nm, Ethidium homodimer-1:

528/617 nm)

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Figure E.1. MTT Optimization: Absorbance vs. Cell Number vs. Incubation Period

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

0.4

0.45

0 0.1 0.2 0.3 0.4 0.5

Absorbance at 570 nm

Cell Number (Millions)

1 hr

2 hr

3 hr4 hr

5 hr

5 hr (H2O2)

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Live / Dead Assay Images:

Figure E.2. Representative images of Live/Dead assay of AF Cells incubated for 24 hours (37

ºC, 5% CO2): in either (A) F12 Ham’s Media containing 5% FBS (negative control); or (B)

Ham’s F12 Media containing 0.01 wt% H2O2 (Positive Control)

A

B

*

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Figure E.3. Live/Dead Assay: Photomicrograph of AF cells subjected to Non-Soluble

Degradation Products of PU aligned polymers at various concentrations [(A) 0.001 wt. %, (B)

0.005 wt. %, (C) 0.01 wt. %, (D) 0.025 wt. %, (E) 0.05 wt. %, or (F) 0.1 wt. % (g/100mL)]

A B

C D

E F

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Figure E.4. Live/Dead Assay: Photomicrograph of AF cells subjected to Non-Soluble

Degradation Products of PU + 0.05% ADO aligned polymers at various concentrations [(A)

0.001 wt. %, (B) 0.005 wt. %, (C) 0.01 wt. %, (D) 0.025 wt. %, (E) 0.05 wt. %, or (F) 0.1 wt. %

(g/100mL)]

A B

C D

E F

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Figure E.5. Live/Dead Assay: Photomicrograph of AF cells subjected to Buffer Soluble

Degradation Products of PU aligned polymers at various concentrations [(A) 20 %, (B) 40 %,

(C) 50 %, (D) 60 %, (E) 80 %, (F) 100 % (percent by volume)]

A B

C D

E F

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Figure E.6. Live/Dead Assay: Photomicrograph of AF cells subjected to Buffer Soluble

Degradation Products of PU + 0.05% ADO aligned polymers at various concentrations [(A) 20

%, (B) 40 %, (C) 50 %, (D) 60 %, (E) 80 %, (F) 100 % (percent by volume)]

A B

C D

E F

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Appendix F: Statistics Tables

(Significance was established at p < 0.05)

Table F.1 - Cumulative Mass Loss Statistics: Week (Fig. 3, Ch. VII)

Comparison Diff of Means t Unadjusted P Significant?

4 vs. 1 1.624 8.111 1.38E-11 Yes

4 vs. 2 1.218 6.085 6.03E-08 Yes

3 vs. 1 0.944 4.716 0.0000124 Yes

4 vs. 3 0.68 3.396 0.00115 Yes

3 vs. 2 0.538 2.689 9.00E-03 Yes

2 vs. 1 0.406 2.026 0.0467 Yes

Table F.2 - Cumulative Mass Loss Statistics: Material Type (Overall) (Fig. 3, Ch. VII)

Note: A_PU, A_PUADO, and R correspond to Aligned PU, Aligned PU+0.5%ADO and Random PU+0.5%ADO scaffolds


R vs. A_PUADO 0.322 1.819 0.074 No

R vs. A_PU 0.257 1.45 0.152 No

A_PU vs. A_PUADO 0.0654 0.369 0.713 No

Table F.3 - Cumulative Mass Loss Statistics: Material Type (in week 1, Fig. 3, Ch. VII)


R vs. A_PUADO 0.367 1.035 0.305 No

R vs. A_PU 0.317 0.893 0.375 No

A_PU vs. A_PUADO 0.05 0.141 0.888 No



R vs. A_PU 0.133 0.376 0.708 No

A_PUADO vs. A_PU 0.1 0.282 0.779 No

R vs. A_PUADO 0.0333 0.094 0.925 No

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125



R vs. A_PUADO 0.315 0.889 0.378 No

R vs. A_PU 0.307 0.865 0.39 No

A_PU vs. A_PUADO 0.00833 0.0235 0.981 No



R vs. A_PUADO 0.574 1.62 0.11 No

A_PU vs. A_PUADO 0.303 0.856 0.395 No

R vs. A_PU 0.271 0.764 0.448 No

Table F.7 - Initial Modulus: Material Type (within as-made, Fig. 4, Ch. VII)


A_PU vs. R 44646200.38 10.357 2.98E-09 Yes

A_PUADO vs. R 42608198.01 10.208 3.77E-09 Yes

A_PU vs. A_PUADO 2038002.376 0.488 0.631 No

Table F.8 - Initial Modulus: Material Type (within prewet, Fig. 4, Ch. VII)


A_PUADO vs. R 11331747.28 7.289 0.000000899 Yes

A_PU vs. R 10845488.33 7.28 0.000000915 Yes

A_PUADO vs. A_PU 486258.958 0.304 0.765 No

Table F.9 - Initial Modulus: Material Type (within week 1, Fig. 6, Ch. VII)


A_PUADO vs. R 4916816.546 10.072 8.54E-08 Yes

A_PU vs. R 4557873.192 9.793 0.000000121 Yes

A_PUADO vs. A_PU 358943.354 0.735 0.474 No

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A_PU vs. R 2911228.117 5.173 0.000232 Yes

A_PUADO vs. R 2993563.469 4.801 0.000433 Yes

A_PUADO vs. A_PU 82335.353 0.137 0.893 No



A_PUADO vs. R 5550167.556 6.195 0.0000324 Yes

A_PU vs. R 5083444.056 5.674 0.0000761 Yes

A_PUADO vs. A_PU 466723.499 0.499 0.626 No



A_PU vs. R 5288278.147 6.651 0.000036 Yes

A_PUADO vs. R 5054492.706 5.994 0.0000901 Yes

A_PU vs. A_PUADO 233785.441 0.277 0.787 No

Table F.13 - Ultimate Tensile Stress (within Aligned PU + 0.05% ADO, Fig. 6, Ch. VII)


As made vs. Week 2 9970298 9.736 0 Yes


As made vs. Week 1 8842397 8.275 0.00E+00 Yes


Prewet vs. Week 2 7998304 7.266 0 Yes




Week 1 vs. Week 3 1495748 1.248 0.215 No

Week 1 vs. Week 4 1538116 1.099 0.274 No

Week 1 vs. Week 2 1127901 1.025 0.308 No

Week 2 vs. Week 3 367847.5 0.317 0.752 No

Week 2 vs. Week 4 410215.2 0.3 0.764 No

Week 3 vs. Week 4 42367.71 0.0293 0.977 No

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Table F.14 - Ultimate Tensile Stress (within Aligned PU, Fig. 6, Ch. VII)


As made vs. Week 2 12193899.96 12.325 0 Yes




Prewet vs. Week 2 6383812.952 6.453 0 Yes



Prewet vs. As made 5810087.012 5.873 0 Yes


Week 1 vs. Week 4 1310297.163 1.094 0.277 No

Week 1 vs. Week 3 1305240.957 1.089 0.279 No

Week 1 vs. Week 2 766087.413 0.717 0.475 No

Week 2 vs. Week 4 544209.75 0.482 0.631 No

Week 2 vs. Week 3 539153.544 0.478 0.634 No

Week 3 vs. Week 4 5056.205 0.00404 0.997 No

Table F.15 - Ultimate Tensile Stress (within Random PU + 0.05% ADO, Fig. 6, Ch. VII)









As made vs. Prewet 5810087.012 5.873 0 Yes


Week 1 vs. Week 4 1310297.163 1.094 0.277 No

Week 1 vs. Week 3 1305240.957 1.089 0.279 No

Week 1 vs. Week 2 766087.413 0.717 0.475 No

Week 2 vs. Week 4 544209.75 0.482 0.631 No

Week 2 vs. Week 3 539153.544 0.478 0.634 No

Week 3 vs. Week 4 5056.205 0.00404 0.997 No

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Table F.16 - DNA Content: (within groups: Stimulated and Control, Fig. 4, Ch. VIII)

Within Group: Stimulated

Comparison Diff of Means t Unadjusted P Critical Level Significant?

0 vs. 12 1.567 4 2.203 0.410 No

0 vs. 24 0.472 3 0.664 0.886 No

0 vs. 6 0.351 2 0.494 0.728 No

6 vs. 12 1.215 3 1.709 0.453 No

6 vs. 24 0.121 2 0.170 0.905 No

24 vs. 12 1.094 2 1.538 0.281 No

Within Group: Control


12 vs. 24 1.689 4 2.375 0.343 No

12 vs. 6 0.809 3 1.137 0.702 No

12 vs. 0 0.584 2 0.821 0.564 No

0 vs. 24 1.106 3 1.555 0.518 No

0 vs. 6 0.225 2 0.317 0.824 No

6 vs. 24 0.880 2 1.238 0.385 No

Table F.17 - DNA Content (within groups: 0hr, 6hr, 12hr, 24hr, Fig. 4, Ch. VIII)

Within Group: 0hr


Ctrl vs. Stim. 0.199 2 0.280 0.844 No

Within Group: 6hr

Ctrl vs. Stim. 0.0734 2 0.103 0.942 No

Within Group: 12hr

Ctrl vs. Stim. 1.951 2 2.743 0.057 No

Within Group: 24hr

Ctrl vs. Stim. 0.833 2 1.171 0.411 No

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Table F.18 - Relative Thymidine Incorporation Ratio Comparison: 24hr vs 72hr (within groups: Stimulated and Control, Fig. 6, Ch. VIII)



24hr vs. 72hr 0.157 0.482 0.631 0.05 No



24hr vs. 72hr 0.542 1.659 0.1 0.05 No

Table F.19 - Relative Thymidine Incorporation Ratio Comparison: Stimulated vs. Control (within groups: 24 hr and 72 hr, Fig. 6, Ch. VIII)

Within Group: 24 hr


Stim vs. Ctrl 0.185 0.565 0.573 0.05 No

Within Group: 72 hr


Stim vs. Ctrl 0.2 0.612 0.542 0.05 No

Table F.20 - Relative Collagen Content Ratio Comparison: 24hr vs 72hr (within groups: Stimulated and Control, Fig. 6, Ch. VIII)



24hr vs. 72hr 0.392 3.323 0.001 0.050 Yes



24hr vs. 72hr 0.261 2.157 0.034 0.050 Yes

Table F.21 – Relative Collagen Content Ratio Comparison: Stimulated vs. Control (within groups: 24 hr and 72 hr, Fig. 6, Ch. VIII)

Within Group: 24 hr


Stim vs. Ctrl 0.121 1.009 0.316 0.050 No

Within Group: 72 hr


Ctrl vs. Stim 0.0106 0.0890 0.929 0.050 No

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Table F.22 - Relative Proteoglycan Content Ratio Comparison: 24hr vs 72hr (within groups: Stimulated and Control, Fig. 6, Ch. VIII)



24hr vs. 72hr 0.263 2.080 0.041 0.050 Yes



24hr vs. 72hr 0.341 2.633 0.010 0.050 Yes

Table F.23 - Relative Proteoglycan Content Ratio Comparison: Stimulated vs. Control (within groups: 24 hr and 72 hr, Fig. 6, Ch. VIII)

Within Group: 24 hr


Stim vs. Ctrl 0.0293 0.229 0.819 0.050 No

Within Group: 72 hr


Stim vs. Ctrl 0.107 0.840 0.404 0.050 No

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