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PJM UNIVERSITY DE
10 SHERBROOKE Faculte de genie
Departement de genie chimique et de genie biotechnologique
DEVELOPPEMENT D'UN SYSTEME TRIDIMENSIONNEL DE CULTURE CELLULAIRE
POUR ORIENTER LA FORMATION DE MICROVAISSEAUX
DEVELOPMENT OF A TRIDIMENSIONAL CELL CULTURE SYSTEM TO ORIENT
MICRO VESSEL FORMATION
These de doctorat Speciality : genie chimique
Irza SUKMANA
Jury : Professeur Patrick VERMETTE (Directeur) Professeur Pierre PROULX (Rapporteur) Professeur Jean-Francois BEAULIEU Professeur Elizabeth JONES
S herbrooke (Quebec) Canada Mai 2010
y^D^l
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Canada
As a milestone of our 13 anniversary I dedicate this work to my wife,
the other half of my soul Ika Nury Wahidah.
To my mother and the memory of my late father (Almarhum) To my beloved son and daughters
To brothers and sisters.
RESUME
Le developpement de micro-vaisseaux fonctionnels dans la culture de substituts tissulaires
constitue un enorme defi du genie tissulaire et de la medecine regeneratrice. Celui-ci est
justifie par la necessite de permettre une meilleure alimentation en nutriments et oxygene des
tissus cultives et une bonne elimination des dechets a l'interieur de ceux-ci. C'est la raison
pour laquelle seulement quelques substituts de tissus sont disponibles pour des applications
medicales. Par consequent, de nombreuses etudes de recherche dans le domaine de l'ingenierie
tissulaire ont mis l'accent sur la culture de micro-vaisseaux a l'interieur de matrices tissulaires
et ce avant leur implantation, tel que presente dans le Chapitre 1 de cette these. Le Chapitre 2
est consacre a l'etude bibliographique presentant revolution recente et les defis futurs lies au
processus de micro-vascularisation de substituts tissulaires. Ce processus est souvent appele
pre-vascularisation.
Le travail experimental presente dans cette these est base sur l'hypothese que des mono
filaments polymeriques separes les uns des autres d'une facon precise peuvent etre utilises
pour guider des cellules endotheliales dispersees dans un gel de fibrine et ainsi induire des
structures tubulaires de facon directionnelle.
Le Chapitre 3 de cette these presente la validation de la formation de micro-vaisseaux dans un
environnement tridimensionnel. La methode de culture tissulaire presentee ici utilise des fibres
de polymere pour guider des cellules endotheliales et ce afin d'orienter la formation des
micro-vaisseaux. La methode d'attachement cellulaire decrite au Chapitre 3 a permis
l'adherence et la migration de cellules endotheliales humaines de la veine ombilicale
(HUVEC) sur des fibres de poly(ethylene terephtalate) (PET) non-modifiees. Les fibres
recouvertes de HUVEC ont ete « sandwichees » entre deux gels de fibrine pour permettre le
developpement des micro-vaisseaux. Le developpement de l'angiogenese a ensuite ete
caracterise a l'aide d'un certain nombre de techniques d'imagerie, y compris la microscopie par
fluorescence et la microscopie confocale. Apres 4 jours de culture, les micro-vaisseaux
formaient de grandes structures en forme de tube (diametre d'environ 100|im) entre les fibres
adjacentes distancees de 0.1mm.
ii
Le Chapitre 4 presente d'un cote l'etude de l'effet des fibroblastes humains et, d'un autre cote,
l'influence de deux facteurs de croissance angiogeniques (i.e., Vascular Endothelial Growth
Factor (VEGF) et le basic Fibroblast Growth Factor (bFGF)) sur les reponses des HUVEC
recouvrant les fibres de PET cultivees en sandwich dans des gels de fibrine. Le developpement
et la maturation des micro-vaisseaux ainsi que la formation des lumens ont ete etudies par
microscopie confocale en utilisant une matrice de fibrine fluorescente et par histologic
L'histologie et les experiences menees avec la fibrine fluorescente ont confirme que ces
pseudo-micro-vaisseaux formes entre les fibres de polymere incorporees dans la fibrine
avaient un lumen. Les effets du VEGF et du bFGF etaient fonction de la concentration de ces
molecules. L'utilisation de fibroblastes a considerablement ameliore la maturation des micro-
vaisseaux comparativement aux echantillons controles et ceux cultives avec le VEGF et le
bFGF.
La conclusion et les propositions pour des travaux fiiturs constituent le Chapitre 5.
Les Annexe A et B presentent, respectivement, les protocoles detailles utilises dans cette etude
pour marquer les cellules et des informations generates sur le cytosquelette.
Mots-cles: polyethylene terephtalate; orientation cellulaire; angiogenese directionnelle;
micro-vaisseaux; lumens multi-cellulaires; cultures en bioreacteur.
HI
ABSTRACT The development of functional and oriented microvessels within tissue constructs is a
tremendous challenge in tissue engineering and regenerative medicine. It is justified by the
need to allow better nutrient and oxygen supply and waste removal within the core of tissue
constructs. It is one of the reasons why only few tissue substitutes are available to clinicians.
Therefore, the focus of many research studies in the field of tissue engineering has been placed
on promoting microvessel ingrowth inside tissue constructs prior to their implantation, as
presented in Chapter 1. This process is often referred to as pre-vascularization. As such,
Chapter 2 of this thesis is devoted to review the recent development and future challenges
related to the micro vascularization process of tissue constructs.
The experimental work in this thesis is based on the hypothesis that polymer monofilaments
precisely distanced from each others can be used to guide endothelial cells when dispersed in a
(fibrin) gel and to induce tube-like structures in a directional fashion.
In Chapter 3 of this thesis, the use of polymer fibres as contact guidance of endothelial cells to
orient microvessel formation and development in a three-dimensional environment was
validated. The novel cell attachment method described in Chapter 3 has resulted in an increase
in Human Umbilical Vein Endothelial Cell (HUVEC) adhesion and spreading on bare
poly(ethylene terephthalate) (PET) fibres. Furthermore, HUVEC-covered fibres were
sandwiched between fibrin gels to allow the development of microvessels. Angiogenesis
development was characterized and evaluated using a number of imaging techniques,
including fluorescence microscopy and confocal microscopy. After 4 days of culture,
microvessels formed large tube-like structures (diameter of approximately 100 um) between
adjacent fibres distanced by 0.1mm. This proof-of-concept opens the door to other
experiments and to possibility of scaling the system to bioreactor cultures.
In Chapter 4, the effect of human fibroblasts and in another set of experiments, the influence
of two angiogenic growth factors (i.e., Vascular Endothelial Growth Factor (VEGF) and basic
Fibroblast Growth Factor (bFGF)) over the responses of the HUVEC-covered fibres
IV
sandwiched in fibrin were investigated. The development and maturation of microvessels as
well as the assessment of lumen formation were evaluated using a fluorescent fibrin matrix,
confocal microscopy and histology. Histology and fluorescent fibrin experiments confirmed
that these microvessel-like structures formed between polymer monofilaments embedded in
fibrin contained a lumen. The effects of VEGF and bFGF were dose dependent. Furthermore,
the use of fibroblasts significantly improved the maturation of the microvessels compared to
control and to samples cultured with VEGF and bFGF.
Conclusions and suggested future work are presented in Chapter 5.
Appendices A and B present the detailed protocols used to label cells in this study and general
information about cell cytoskeleton, respectively.
Keywords: Polyethylene terephthalate; Contact guidance; Cell patterning; Directional
angiogenesis; Microvessel formation; multi-cellular lumen; bioreactor cultures.
v
Acknowledgements
First of all, I would like to express my sincere gratitude to my supervisor Professor Patrick
Vermette, whose enthusiastic support and good advices helped me to go through my
challenging project.
I am very honoured to have Professor Pierre Proulx and Professor Jean-Francois Beaulieu for
their guidance and for being my pre-Doctoral examiners.
A very special thanks to Mr. Yannick Laplante and Dr. Aftab Ahamed, former members of the
Laboratoire de Bioingenierie et de Biophysique de l'Universite de Sherbrooke, for their
precious help when I started my Ph.D. I thank them also for their hints on general technical
and scientific aspects.
I also want to thank all my friends in the group for sharing their ideas and for scientific
discussions on daily basis during this course.
I should thank all the technicians in the Department of Chemical and Biotechnological
Engineering, especially Mr. Mark Couture for his help in designing and producing frames and
culture chambers for my research. Also, I want to thank Mr. Gilles Grondin and Dr. Leonid
Volkov for their contribution in confocal microscopy and Mrs. Johanne Dussault and Dr.
Sameh Geha who helped me with histology.
I am particularly thankful to my sponsor, the Islamic Development Bank (IDB), for all
financial support and living guidance for me and my family.
I am also very grateful to the Muslim community in Sherbrooke who has given me an
invaluable support and help in countless ways. I will never forget the support provided by dear
brothers: Hendra Hermawan, Abdelfettah and Rachid Bannari, Brahim Selma, Mbark El
vi
Morsli, Rachid Kadouche, Taha Abd-El-Rahman, Ahmad Kamal, Hamdy and Habibie, Deyab
El-Gamal, Ahmed Omran, Said and Hoccain and others, "Jazakumullahu khairan katsiran".
Finally, my most acknowledgment goes to my adorable wife, adik Ika Nury Wahidah, and our
lovely kids: Muhammad, Wafa and Sky. The word "Thanks" is never enough for their love,
patience and encouragement. May Allah SWT give you all the best rewards. Amen.
vn
TABLE OF CONTENT
ABSTRACT ii
RESUME iv
Acknowledgements vi
Table of Content viii
List of Figures xi
List of Tables xii
List of Abbreviations xiii
CHAPTER 1 Introduction 1
1.1 Thesis Objectives 1
1.2 Thesis Structure 1
1.3 Contributions 3
CHAPTER 2 Vascularization strategies to engineer tissue substitutes: a challenge to support
blood microvessel development 5
2.1 Abstract 6
2.2 Introduction 6
2.3 Scaffolds in tissue engineering 8
2.3.1 Synthetic polymers 10
2.3.2 Natural polymers 11
2.4 Material properties 21
2.4.1 Scaffold porosity 21
2.4.2 Matrix stiffness 23
2.5 Scaffold vascularisation 24
2.5.1 Vasculogenesis and angiogenesis 28
2.5.1.1 Model 1: Cell death and phagocytosis 29
2.5.1.2 Model 2: Wrapping spaces around ECM 29
2.5.1.3 Model 3: Vacuole formation from the coalescence of intracellular vacuoles 31
2.5.2 Promoting angiogenesis in three-dimensional constructs 31
2.5.2.1 Micro-patterning 32
via
2.5.2.2 Surface modification 33
2.5.2.3 ECM modification 34
2.5.3 Co-culture systems and microvessel maturation 35
2.6 Bioreactor to support vascularization 37
2.7 Concluding remarks 39
CHAPTER 3 Polymer fibres as contact guidance to orient microvascularization in a 3D
Environment 57
3.1 Abstract 58
3.2 Introduction 58
3.3 Materials and methods 59
3.3.1 Materials 59
3.3.2 Cell culture : 60
3.3.3 PET fibres and fibrin gel preparation 60
3.3.4 Cell adhesion assay 61
3.3.5 Cell culture in 3D environment 61
3.3.6 Visualization of microvessels 62
3.3.7 Images and image processing 62
3.3.8 Statistical analysis 63
3.3.9 Confocal microscopy imaging of microvessels 63
3.4 Results 64
3.4.1 Cell adhesion on untreated PET fibres 64
3.4.2 Effect of fibre-to-fibre distance over cell connections and microvessel formation 65
3.4.3 Microvessel formation 66
3.5 Discussion 67
3.6 Conclusions 69
CHAPTER 4 The effects of co-culture with fibroblasts and angiogenic growth factors on
microvascular maturation and multi-cellular lumen formation in HUVEC-oriented
polymer fibre constructs 82
4.1 Abstract.... 83
4.2 Introduction 83
4.3 Materials and methods 85
ix
4.3.1 Materials 85
4.3.2 Cell culture 85
4.3.3 Culture system and fibrin preparation 86
4.3.4 Cell adhesion and angiogenesis assay 86
4.3.5 Visualization of microvessel and lumen formation 87
4.3.6 Histological section 88
4.3.7 Imaging microvessels by confocal microscopy 88
4.3.8 Statistical analysis 89
4.4 Results and discussion 89
4.5 Conclusions 93
CHAPTER 5 Conclusions and suggestions 107
5.1 Conclusions 107
5.2 Conclusion (en francais) 108
5.3 Suggestions 109
5.3.1 Further studies of the angiogenesis assessment 109
5.3.2 The effect of dynamic culture system over microvessel development and proof of
microvessel functionality 111
APPENDIX A Protocol for cell staining 116
APPENDIX B Cell cytoskeleton 119
x
LIST OF FIGURES
Figure 2.1: Fibrinogen structure and fibrin clot process 16
Figure 2.2: Models of the development of tube-like structures during vasculogenesis 30
Figure 2.3: Blood vessel structure and development 36
Figure 3.1: Schematic illustration of the in vitro method designed to guide and orient
microvessel formation 75
Figure 3.2: HUVEC attachment on PET fibres 76
Figure 3.3: Effect of fibre-to-fibre distance over cell connections and microvessel
development 77
Figure 3.4: Confocal and inverted microscopy image 78
Figure 3.5: Schematic of the microvascularization process 79
Supplementary Figure 3.i: Scanning electron microscopy (SEM) image of a PET fibre 80
Supplementary Figure 3.ii: Progression of cell connections and microvessel reorganization
along and in between adjacent fibres sandwiched between two fibrin gels 81
Figure 4.1: Effect of fibroblasts over microvessel development 100
Figure 4.2: Immunofluorescence picture illustrating the effect of fibroblasts on microvessel
development 101
Figure 4.3: Immunofluorescence picture (F-actin and nuclei staining) illustrating the effect of
VEGF and bFGF on microvessel development 102
Figure 4.4: Illustration of multi-cellular lumen formation for cultures carried out for a period
of 4 days with fibroblasts 103
Supplementary Figure 4.i: Comparison between systems using cell-free fibres (left) and
HUVEC-covered fibres (right) over cell-to-cell connections & microvessel
development 104
Supplementary Figure 4.ii: Immunofluorescence confocal pictures of a sample cultured with
no fibroblasts 105
Supplementary Figure 4.iii: Fibroblasts stained with the CFSE CellTracer 106
Figure 5.1: Simple set-up for dynamic cell cultures 113
XI
LIST OF TABLES
Table 2.1: Integrins and non-integrin receptors which can bind to fibrin(ogen) 18
Table 2.2: The use of synthetic and natural polymers to support tissue vascularisation 19
Table 3.1: Effects of fibre-to-fibre distance on cells 74
Table 4.1: Effects of VEGF and bFGF concentrations over the number of cells and cell-cell
connections following 2 days of culture 98
Table 4.2: Comparison of the effects of fibroblasts, VEGF, and bFGF over the number of
cells and cell-cell connections 98
Table 4.3: Review of some studies on the effects of VEGF and bFGF on endothelial cells and angiogenesis 99
xn
LIST OF ABBREVIATIONS
3D
Agp-1 ANOVA
BAEC bFGF
BSA CAM CLS CSTR
DLLA EC ECGS
ECM ePTFE
ERG
ESAM
FBS FDA FGF
GAG HA HBSS HDMEC
HUVEC
HVEC
ICAM-1
MC
three-dimensional or tridimensional angiopoietin-1 analysis of variance
bovine aortic endothelial cell basic fibroblast growth factor
bovine serum albumin chorioallontoic membrane capillary-like structure continuous stirred-tank reactor (D,L-lactic acid) endothelial cell endothelial cell growth supplement extracellular matrix expanded poly(tetrafluoroethylene) early response genes
endothelial cell selective adhesion molecule foetal bovine serum Food and drug administration fibroblast growth factor
glycosaminoglycan hyaluronic acid hanks balances salt saline human dermal endothelial cell human umbilical vein endothelial cell human vascular endothelial cell inter-cellular adhesion molecule-1 messenchymal cell
o-HA
PBS PDGF-BB PDLA PECAM-1 PEG PET PGA PLA
PLG PLGA PLLA
RAOEC RGD
RHAMM
SEM
SF SMC SM-MHC SMocA STLV TMC VE-cadherin VEGF
VSMC
vWF
oligosaccharides of HA
phosphate buffer saline platelet-derived growth factor-BB poly(D,L-lactic acid) platelet endothelial cell adhesion molecule-1 poly(ethylene glycol) poly(ethylene terephthalate) poly(glycolic acid) poly(lactic acid)
poly(lactic glycolic) poly(lactic-co-glycolic acid) poly(L-lactic acid)
rat aortic endothelial cell arginine-glycine-aspartic acid peptide receptor for HA mediated motility scanning electron microscopy skin fibroblast smooth muscle cell smooth muscle myosin heavy chain smooth mucle a-actin slow turning lateral vessel trimethylene carbonate vascular endothelial cadherin
vascular endothelial growth factor vascular smooth muscle cell
von Willebrand factor
X1I1
CHAPTER 1
Introduction The development of functional and oriented microvessels within engineered human tissue
substitutes constitutes a promising hope in tissue engineering and regenerative medicine
[Stock and Vacanti 2001; Eichmann et al. 2005]. To date, the development of thick tissues,
such as pancreas, liver, heart, and kidneys is problematic due to the lack of construct
vascularisation, resulting in cell and tissue death [Atala 2004; Jain et al. 2005; Kannan et al.
2005]. The focus of current research efforts in tissue engineering has been mainly on
developing strategies to study and promote microvascularization within tissue constructs
[Kannan et al. 2005; Neuman et al. 2003].
1.1 Thesis objectives
This project was designed to constitute the initial part of a larger research program aiming to
create an oriented functional blood microvessel network within three-dimensional tissue
scaffolds. One of the aims of this research program is to scale-up tissue engineering methods
by the use of dynamic culture conditions using an automated bioreactor system. The principal
objective of this thesis was to design and validate static cell culture processes to guide
endothelial cell responses and the subsequent microvessel development and orientation.
1.2 Thesis Structure
This thesis has been written in a manner that chapters correspond to scientific articles resulting
from this work. At the time of submission, two articles were published and one under
preparation for submission. Below is a brief description of their content.
Chapter 1: Introduction. This section contains thesis objectives and structure, and
contributions.
1
Chapter 2: Vascularization strategies to engineer tissue substitutes: a challenge to
support blood microvessel development. This chapter is a literature review on advancements
in the field of three-dimensional tissue engineering with an emphasis on cellular and
angiogenesis guidance as well as on blood microvessel development. Moreover, the effects of
cell types, co-cultures, and growth factors on microvessel development are discussed. The use
of bioreactors to culture tissue substitutes is also briefly addressed.
Chapter 3: Polymer fibers as contact guidance to orient microvascularization in a 3D
environment. This part of the thesis presents a novel culture system for angiogenesis
guidance using poly(ethylene terephthalate) (PET) fibres. Briefly, in an effort to increase the
bioactivity of PET fibres, cells are first allowed to attach and cover the fibres using a novel
culture method. Then, HUVEC-covered fibres are sandwiched in fibrin and microvessel
structures are examined by confocal microscopy. In addition, the effect of fibre-to-fibre
distance over angiogenesis development is compared. Results from this chapter reveal that this
culture system can be used to orient microvessel structures. The optimum fibre-to-fibre
distance found here is further used in the subsequent part of this thesis.
Chapter 4: The effect of co-culture with fibroblasts and angiogenic growth factors on
microvascular maturation and multi-cellular lumen formation in HUVEC-oriented
polymer fibre constructs. This chapter further validates the use of the culture system
developed in Chapter 3 using a co-culture method with human skin fibroblasts or with
angiogenic growth factors (i.e., Vascular Endothelial Growth Factor (VEGF) and Basic
Fibroblast Growth Factor (bFGF)). The effects of these two parameters over microvessel
development and maturation are investigated. This study is also aiming to determine whether
or not these microvessels contained multi-cellular lumen. VEGF and bFGF are shown to
enhance HUVEC proliferation and migration, thus improving the development of cell-cell
connections and subsequent microvessel development. The use of fibroblasts results in more
mature microvessels containing more cell-cell connections, which also contain lumens.
Chapter 5: Conclusions and Suggestions. This part presents a general conclusion and
suggests future direction.
2
1.3 Contributions
During the course of this thesis, I have had important help from several people, to whom I am
very grateful. Firstly, former members of the Laboratoire de Bioingenierie et de Biophysique
de l'Universite de Sherbrooke, Mr. Yannick Laplante, Dr. Aftab Ahamed, and Dr. Mbark El
Morsli, have helped me to gain technical know-how and scientific knowledge.
Mr. Yannick Laplante has taught me how to prepare protocols and techniques to carry out cell
culture. Furthermore, Mr. Laplante has assisted me in the preparation and evaluation of three-
dimensional culture systems. Dr. Aftab Ahamed has tough me statistics and how to prepare a
scientific article. Dr. Mbark El Morsli has shared his knowledge in advance mathematics and
chemical engineering.
The confocal microscopy was carried out at the Department of Biology (Universite de
Sherbrooke) and at the Centre de recherche clinique Etienne-Le Bel (Centre hospitalier
universitaire de Sherbrooke (CHUS)), respectively, where Mr. Gilles Grondin and Dr. Leonid
Volkov have shared knowledge and helped me to obtain good quality pictures for my
publications. Mrs Johanne Dussault and Dr. Sameh Geha have assisted me with histology.
After fixation, all my histology samples were sent to the Pathology Department (CHUS). The
sectioning and staining have been done by Mrs Johanne Dussault, while further expert
interpretation have been done by Dr. Sameh Geha.
All other laboratory and analytical works were carried out by me.
3
References
Atala, A. (2004) Tissue engineering and regenerative medicine: concepts for clinical
application. Rejuvenation Res. 7, p. 15-31.
Eichmann, A., Noble, F. L., Autiero, M. and Carmeliet, P. (2005) Guidance of vascular and
neural network formation. Current Opinion in Neurobiology 15, p. 108-15.
Jain, R. K., Au, P., Tarn, J., Duda, D. G. and Fukumura, D. (2005) Engineering vascularized
tissue. Nat. Biotechnol. 23, p. 821-23.
Kannan, R. Y., Salacinski, H. J., Sales, K., Butler, P. and Seifalian, A. M. (2005) The roles of
tissue engineering and vascularisation in the development of micro-vascular networks: a
review. Biomaterials 26, p. 1857-75.
Neuman, T., Nicholson, B. S. and Sanders, J. E. (2003) Tissue engineering of perfused
microvessels. Microvasc. Res. 66, p. 59-67.
Stock, U. and Vacanti, J. P. (2001) Tissue engineering: current state and prospects. Annu. Rev.
Med. 52, p. 443-51.
4
CHAPTER 2
Vascularization strategies to engineer tissue substitutes:
a challenge to support blood microvessel development
Irza Sukmana1'2
1. Laboratoire de Bioingenierie et de Biophysique de l'Universite de Sherbrooke, Department
of Chemical and Biotechnological Engineering, Universite de Sherbrooke, 2500, blvd de
l'Universite, Sherbrooke, QC, Canada, J1K 2R1.
2. Research Centre on Aging, Institut universitaire de geriatrie de Sherbrooke, 1036, rue
Belvedere Sud, Sherbrooke, QC, Canada, J1H 4C4.
5
2.1 Abstract
The guidance of endothelial cell organization into a capillary network has been a long standing
challenge in tissue engineering. Some research efforts have been made to develop methods to
promote blood capillary networks inside engineered tissue constructs. Mature microvessels
that would mimic blood microvessel function can be used to subsequently facilitate oxygen
and nutrient transfer as well as waste removal. Microvascularization of engineered tissue-
scaffold appears to be one of the most favorable approaches to overpass nutrient and oxygen
supply limitation, which is often the major hurdle in developing thick and complex tissue or
organ substitutes. This review chapter addresses recent advances and future challenges in
developing and using bioactive scaffolds and 3D culture systems to promote tissue construct
vascularization allowing mimicking blood microvessel development and function encountered
in vivo. Vascularization can be achieved through strategies in which endothelial cells are
cultured in bioactive scaffolds and/or extracellular matrix (ECM)-like environments, with or
without angiogenic growth factors as well as using cell co-cultures and combinations of those.
In the near future, bioreactors will be used to create fully vascularized functional tissue and
organ substitutes.
2.2 Introduction
Tissue engineering is an emerging field. In 1987, the National Science Foundation defined
tissue engineering as "an interdisciplinary field that applies the principles of engineering and
the life sciences towards the development of biological substitutes that restore, maintain or
improve tissue function" [Langer and Vacanti 2003]. Tissue engineering involve the repair
and restoration of various tissue and organ functions, while limiting host rejection and side
effects for the patient by delivering cells and/or biomolecules through the use of 3D scaffolds
[Fuchs et al. 2001; Mooney et al. 1997].
Although there have been some successes to replace and restore some tissues using tissue
engineering approaches, these have been mostly limited to thin or avascular tissues, such as
cartilage, skin or bladder [Mooney and Mikos 1999]. However, for thick and vascular tissues
and for most organs, the lack of a sufficient supply of nutrients and oxygen to growing tissues,
6
as well as the waste removal represent two important factors that limit the successful
development and implantation of engineered tissue constructs [Arthur et al. 1998; Neumann et
al. 2003]. These issues of poor mass transport and mass transfer have often led to the failure of
the culture process or even to that of implants [Arthur et al. 1998; Sipe 2002].
One possible strategy for creating thick engineered tissue substitutes in vitro is to use a
bioactive scaffold that allows the guidance of endothelial cells promoting microvessel
development in a directional fashion in order to vascularized the tissue construct [Godbey and
Atala 2002]. It is quite well accepted among the scientific community that pre-vascularization
of tissue construct appears to be one of the most favorable and efficient approaches to address
the problem of tissue survival due to a lack of oxygen and nutrient supply [Atala 2004; Mikos
et al. 1993; Montano et al. 2009; Nomi et al. 2002]. The concept of pre-vascularization mainly
involves the incorporation of endothelial cells into a bioactive scaffold to form a capillary
network inside the structure prior to its implantation [Atala 2004; Langer et al. 1995]. This
could accelerate the formation of functional microvessels within the core of an implant
[Godbey and Atala 2002; Neumann et al. 2003].
The idea of using pre-vascularized engineered tissue substitutes was first suggested by Mikos
et al. (1993) when comparing the performance of pre-vascularized tissues to non-vascularized
ones. Later, Sakakibara et al. (2002) concluded that pre-vascularization enhanced the benefits
of cardiomyocyte transplantation. More recently, Levenberg et al. (2005) demonstrated that
pre-vascularization improved the performance of skeletal muscle tissue constructs when
implanted in mice. Furthermore, studies aiming to induce and control vascularization may also
advance general knowledge on the development of. therapeutics targeting angiogenesis.
Numerous pathological conditions are associated with insufficient oxygen and blood supply
[Atala 2004; Carmeliet 2005; Neumann et al. 2003]. Also, uncontrolled angiogenesis is
associated with many diseases, including rheumatoid arthritis, macular degeneration, and
tumour growth [Carmeliet and Conway 2001].
Advances in tissue engineering have brought significant knowledge on the mechanisms and
parameters related to vascularization and angiogenesis development and blood microvessel
7
network formation [Levenberg and Langer 2004; Nerem 2006]. This review will report status
of the current research and development related to bioactive scaffolds and culture systems
designed to promote angiogenesis and vascularization. It is necessary to improve our
understanding of the mechanisms behind angiogenesis and to apply that knowledge to guide
microvessel growth in a regulated manner. Other aspects including the challenges in
angiogenesis guidance, assessment of angiogenesis and lumen formation and the use of
bioreactor system to culture vascularized tissue constructs will also be outlined.
2.3 Scaffolds in tissue engineering
The needs for engineered tissue substitutes are important. Currently, the demand for organ
transplants is higher than the supply. In the United States alone, there were 79,512 patients on
the transplantation waiting list in 2002, and only 24,422 received organs while 6,297 died
while waiting [Nerem 2006]. In addition, although organ transplantation is one of the less
expensive therapies in regenerative medicine, tissue engineering offers hope for more
consistent and rapid treatment of those in need [Lanza and Chick 1997].
As an interdisciplinary approach between engineering and life science, tissue engineering
seeks an opportunity to develop suitable biomaterial-cell hybrid constructs to support the
regeneration and restoration of tissue structure and function. The critical challenge facing
tissue engineering today relies on our knowledge and ability to fabricate tissue and organ
replacements that would fulfill physiological function [Koh et al. 2009; Nerem 2006]. Also,
the success of tissue engineering methods relies on the ability of the construct to integrate at
the implantation site with the native tissue. Tissue engineering is facing important clinical and
practical problems, such as cell sourcing, rejection, healing, and cell/tissue death [Conway and
Carmeliet 2004; Gimble and Guilak 2003].
Various key concepts in tissue engineering and regenerative medicine have been pursued to
overcome those problems and these concepts include injection of tissue-specific viable cells
directly into a damaged tissue (for example brain cells in the case of Parkinson or Alzheimer),
encapsulation of specific cell types within synthetic permeable matrices that allow release of
therapeutics (e.g., the release of insulin or dopamine from pancreatic islets in the treatment of
8
diabetes), and to seed scaffolds with living cells in vitro, allowing their maturation before
being implanted [Lanza et al. 2000; Yang et al. 2001a]. In the present thesis, we are mainly
interested with the last concept.
If an isolated cell population can be expanded in vitro using cell culture and bioreactor
techniques, in theory, only a very small number of cells from donors would be necessary to
prepare such biological implants. Since the isolated cells cannot form new tissue by
themselves, a (temporary) template is needed, which we refer here to as scaffold. Scaffolds are
expected to provide a control over tissue architecture and mechanical properties. They can
allow cells to adhere, proliferate, and migrate [Howe et al. 1998; Huttenlocher et al. 1995] in
order to form a required structure and to synthesize their own extracellular matrix (ECM)
molecules [Chang and Werb 2001], thus hopefully allowing tissue regeneration or repair
[Nerem 2007; Yang et al. 2001b].
It is believed that the success to develop tissue constructs depends on many aspects, such as
cell sourcing, type of biomaterials used to make scaffolds, tissue culture methods, only to
name a few. For example, using cells from other species, such as pig, remains a debate since
there is a risk of transferring diseases from animals to humans [Bach 1998]. Using cells from
the same genotype or close relatives of a patient could avoid problems associated with
immune rejection, which can result in tissue death [Bach 1998; Lee et al. 2005; Ye et al.
2000b].
The behaviour of individual cells and the dynamic state of multi-cellular tissues are regulated
by the interaction between cells and their surrounding matrix. Therefore, the design of
scaffolds from the macroscopic scale (e.g., pore structure) to the nanoscopic level (e.g.,
surface properties) is very important. Firstly, decision of using either natural or synthetic
scaffolds should be based on their capability to provide a specific microenvironment that
mimics the natural environment of the targeted anatomical site [Garcia et al. 2005]. Secondly,
the three-dimensional scaffold should fulfill some requirements with respect to:
biocompatibility, degradation rate, porosity, mechanical properties (e.g., stiffness), and
chemistry (e.g., surface chemical/protein composition) [Chaikof et al. 2002].
9
In the in vivo environment, cells interact with their ECM in a dynamic manner. The concept of
dynamic "communication" between cells and their matrix has opened a wide exploration of
the ECM molecules and scaffold materials that can be used in tissue engineering. Scaffolds
can be made from synthetic polymers, naturally occurring materials, or a combination of both.
2.3.1 Synthetic polymers
Synthetic polymers have been investigated and used to make scaffolds in tissue engineering
for a variety of possible applications. The principal advantage of using synthetic polymers is
that their properties (e.g., biodegradation, physicochemistry and mechanical stiffness) can be
controlled by manipulating their molecular weight and compositions, for example [Chaikof et
al. 2002]. Among them, synthetic degradable polymers from the poly(a-esters) family, such as
poly(lactic acid) (PLA), poly(glycolic acid) (PGA) and their co-polymers, have been
extensively investigated in biomaterials and tissue engineering.
Synthetic polymers from the poly(a-esters) group are degraded mainly through chemical
hydrolysis and are mostly insensitive to enzymatic attack [Larson and Chu 2006], and often,
the degradation profile does not vary between patients [Mooney et al. 1997]. For example, it
was recognized that the constituting monomers of PLA and PGA are nontoxic and are
metabolized in the body [Tabata 2006; Tabata 2009]. Therefore, PLA, PGA and their
copolymers (e.g., poly(lactic-co-glycolic acid) (PLGA)) are FDA-approved and they can be
produced to form a variety of implants ranging from screws, meshes, and sutures to porous
scaffolds [Larson and Chu 2006; Lee et al, 2008; Tabata 2006].
PGA is an inelastic polyester with a high crystallinity (46-50%) and is degraded by water
(through hydrolysis) to form glycolic acid [Kikuchi et al. 1997; Vert et al. 1994]. PLA is less
crystalline, more hydrophobic, and less susceptible to hydrolysis than PGA [Larson III and
Chu 2006; Vert et al. 1994]. For tissue engineering applications, their copolymers, such as
PLLA (poly(L-lactic acid)), PLGA (poly(lactic-co-glycolic acid)) and PDLLA (poly(D,L-lactic
acid) are more popular, since their properties can be tailored [Vert et al. 1994]. For example,
the degradation time of PLLA has been reported to be very slow, while PDLLA hydrolysed in
a matter of weeks [Yang et al. 2001a]. Even a small amount of D,L-LA in the polymer chain of
10
PLA can accelerate the degradation time dramatically [Bramfeld et al. 2007; Grizzi et al.
1995; Tabata 2006].
Several investigators have explored the potential use of polymers from the PLA and PGA
family to fabricate scaffolds for the promotion of microvascularization and angiogenesis. For
example, Grizzi et al. (1995) have produced poly(D,L-lactic-co-glicolic acid) scaffolds to
engineer tubular tissues. Furthermore, when endothelial cells were seeded in these constructs,
they were able to generate a capillary network inside the scaffold [Mooney et al. 1994;
Mooney et al. 1996]. Also, in a more recent study, Levenberg et al. (2005) have successfully
pre-vascularized PLLA/PLGA sponges with pore size ranging from 225 to 500um. Then, they
implanted the pre-vascularized scaffold in skeletal muscles of mice and found that the method
was able to promote angiogenesis in the implant.
In another application, D,L-lactic acid (DLLA) was combined with 1,3-trimethylene carbonate
(TMC) at a specific molecular weight ratio of 81:19 (DLLA:TMC) in order to produce a
TMC-DLLA copolymer. This copolymer was processed to make a scaffold with lOOum
average pore size having a high interconnectivity. This scaffold was tested in vitro and shown
to support cardiomyocyte survival [Pego et al. 2003b]. When this scaffold seeded with cells
was subcutaneously implanted in rats, it elicited an acute inflammatory reaction [Pego et al.
2003a].
2.3.2 Natural polymers
Comparing to synthetic polymers, natural polymers have longer history and have been broadly
used in many applications in the biomedical, pharmaceutical, and tissue engineering fields.
While synthetic scaffolds offer good mechanical properties and less product variability with a
high level of control, natural scaffolds provide a better environment for cell attachment and
signalling, resulting in more efficient regulation of cell structures and functions [Chiu et al.
2009; Grizzi et al. 1995]. Natural polymers can be made from proteins (e.g., collagens, gelatin,
albumin, and fibrinogen), polysaccharides (e.g., chitosan, hyaluronic acid, alginate, cellulose
and dextran), and their chemical derivatives.
11
ECM-derived polymers are attractive materials to make bioactive scaffolds, since they can
provide cells with an environment more similar to the cell native ECM [Howe et al. 1998;
Larson III and Chu 2006]. The ECM can be defined as a complex protein structure outside the
cells, which mainly consists of collagens and proteoglycans. The primary function of the ECM
is to support the cellular structure. Some ECM components regulate cellular processes, such as
cell proliferation, motility, differentiation, migration and adhesion [Chiu et al. 2009].
Each tissue has a unique ECM composition and environment. Therefore, the design of ECM-
derived scaffold should mimic certain features and functions of the ECM for the targeted end
use. For example, in the case of scaffold vascularization, the matrix should provide an
environment such as in the connective tissue for the endothelial cells to adhere and proliferate
as well as to form and remodel vascular structures [Becker et al. 2009]. Furthermore, as
endothelial cells are lining the innermost layer of blood vessels and capillary microvessels,
their interaction with the underlying ECM is essential to maintain cellular integrity and
functional activity for the development of functional and mature blood vessels [Hutchings
2003; Jain 2003]. To date, natural polymers such as hyaluronic acid, chitosan, alginate,
collagens, and fibrin are the most important biodegradable materials to fabricate scaffolds.
Hyaluronic acid (HA), also known as hyaluronan, is a glycosaminoglycan (GAG) that has a
linear polysaccharide branch (glucuronic acid N-acetyl D-glucosamine). Hyaluronic acid is the
embryo's first ECM material and is present in nearly all adult mammalian tissues [Brekke et
al. 2006; Toole 2004]. Hyaluronic acid, with high molecular mass (ranging between 10 to
1,000 kDa), has a unique characteristic [Brekke et al. 2006; Slevin et al. 2002]. Indeed, HA
shows poor cell adherence and inhibits endothelial cell proliferation, while its degradation
products (e.g., oligosaccharides of HA, o-HA) are pro-angiogenic and, through chemotaxis,
can stimulate cell migration, differentiation and the overall angiogenesis process [Rao et al.
1997; Slevin et al. 2002]. For example, West et al. (1985) have demonstrated that o-HA
(molecular mass < 10 kDa) induced angiogenesis with human umbilical endothelial cells
(HUVEC) in the chorioallontoic membrane (CAM) assay. Furthermore, the CD44 receptors in
endothelial cells was found to bind o-HA and to initiate the expression of early response genes
(ERG), resulting in cell proliferation and migration [Lesley et al. 2000; Miletti-Gonzalez et al.
12
2005]. More recently, o-HA was reported to stimulate angiogenesis, either in vitro or in vivo,
with vascular endothelial cell through both CD44 and RHAMM (receptor for HA mediated
motility) [Lokeshwar and Selzer 2000; Miletti-Gonzalez et al. 2005; Slevin et al. 2002]. In
addition, added Fibroblast Growth Factor (FGF) in the HA matrix improved the
neovascularization of the construct [Rauh et al. 2006].
Other polysaccharides, such as chitosan and alginate, have also been investigated for various
tissue engineering and biomedical applications. Chitosan is a linear polysaccharide, composed
of iV-acetyl and D-glucosamine, and has been investigated for some applications, such as to
make contact lens [Park et al. 2003], matrix to encapsulate cells [Zielinski and Aebischer
1994], drug release device, and to engineer cartilage and bone substitutes [Rauh et al. 2006].
Chitosan is an attractive biomaterial to fabricate scaffolds [Park et al. 2003]. For example,
Black et al. (1999) and Slevin and Kumar (2002) have produced a scaffold made of chitosan
cross-linked with HA. This scaffold improved endothelial cell proliferation and induced
capillary network and angiogenesis development inside the construct [Black et al. 1999;
Slevin et al. 2002]. Alginate gel is a hydrogel that is not affected by temperature changes
[Draget et al. 1997]. It has been tested for drug delivery and cell transplantation [Draget et al.
1997; Drury and Mooney 2003; Smidsrod and Skjak-Braek 1990]. Furthermore, incorporating
vascular endothelial growth factor (VEGF) in alginate gels promoted neovascularization in the
matrix [Chang et al. 2010; Drury and Mooney 2003]. This system has been suggested as a
promising approach for clinical applications [Chang et al. 2010].
Collagens are the most abundant proteins, being found in nearly all tissues in mammalians
[Becker et al. 2009]. Type I, II, III, and IV are the most abundant forms and make up
approximately 90% of the collagens in the human body [Becker et al. 2009; Haarer and Dee
2006]. To date, over 25 types of collagens have been identified and have been processed into
various forms, including films, sponges, fibres, and gels [Haarer and Dee 2006]. Collagen type
I, II, III, V and XI can self-assemble into fibrils. Other collagens (e.g., type IV, VIII, and X)
form network and are found in the basement membrane [Becker et al. 2009; Chen et al. 1995].
13
In tissue engineering, physical and chemical cross-linking is often used to increase the
mechanical strength and to avoid rapid degradation of collagens [Chen et al. 1995]. Photo-
oxidation, de-hydrothermal treatment, and ultraviolet irradiation are examples of physical
cross-linking methods, while chemical methods include treatment with carbodiimides,
glutaraldehydes, and poly(glycidyl ether) [Ma et al. 2004]. Often, chemical methods result in
higher cross-linking degree; they are therefore more common than physical methods. Also,
chemical treatment parameters (e.g., time and temperature) as well as catalyst concentration
can be adapted to vary mechanical and degradation properties of collagen scaffolds [Angele et
al. 2004; Ma et al. 2004]. On the other hand, chemical methods could leave some potentially
toxic chemical residues [Ma et al. 2004].
Collagens can be purified from animal and human sources, but the concern of the
immunologic and disease transmissions, especially for animal collagens, still remains. To
avoid those risks, Toman et al. (1999) have suggested a method to produce recombinant
collagens. Recombinant human collagen types I and II are commercially available (e.g.,
FibroGen Inc, San Francisco, CA) [Becker et al. 2009; Toman et al. 1999]. Engineered tissue
substitutes for skin replacement called Apligraft™ (Organogenesis Inc, Canton,
Massachusetts, USA) is made from collagen type I and was the first commercialized man-
made tissue substitute [Zisch et al. 2003]. Collagen can also be extracted from tilapia
(Oreochromis niloticas). Indeed, Sugiura et al. (2009) have produced collagen sponge from
tilapia. In vivo implantation of the scaffold into rabbit muscle revealed that tilapia collagen
caused less inflammatory responses, when compared to porcine collagen [Sugiura et al. 2009].
Other studies have reported the use of type I and IV collagens to carry out angiogenesis and
vasculogenesis assays [Soker et al. 2000]. In vitro culture of endothelial cells in 3D matrix
made of type I collagen resulted in an increased number of tube-like structures and supported
angiogenesis development [Francis et al. 2008]. Also, with FGF, collagen type IV scaffold
supported endothelial cell growth and differentiation, thus regulating capillary development
[Ingber and Folkman 1989]. Xu et al. (2001) concluded that denaturation of collagen type IV
can promote a specific angiogenic cryptic epitope (i.e., HUIV26), which can bind to the
cellular integrin ctvP3. To date, at least four different collagen binding integrins on endothelial
14
cells are known and these include ocipi, oc2pl, al0(3l, and a l i p i [Davis and Senger 2005;
Gulberg and Lundgren-Akerlund 2002; Senger et al. 1997; Xu et al. 2001].
Human fibrinogen is a large, complex and fibrous glycoprotein with a molecular weight of
340 kDa. It is 45nm in length and composed of two symmetric "D" domain molecules and a
central "E" domain. Each domain consists of one set of three different poly-peptide chains
termed Aa, Bp\ and y chain (see Figure 2.1.a) [Herrick et al. 1999; Tawil 2006]. In the body,
fibrinogen is present in human blood plasma at a concentration of approximately 2.5 g/L. The
protein is essential for haemostasis, wound healing, inflammation, angiogenesis, and other
biological events. Fibrinogen is a soluble macromolecule, which can be converted to an
insoluble gel (i.e., fibrin) to stabilize the haemostatic plug and to provide a temporary matrix
for subsequent cellular responses involved in wound healing [Janmey et al. 2009; Tawil 2006].
The role of fibrin in this process is not passive, but the protein rather actively directs cellular
responses through specific receptor-mediated interactions with blood cells (e.g., leucocytes) as
well as endothelial cells of the vessel wall [Herrick et al. 1999]. Therefore, the use of fibrin as
a bioactive scaffold to support tissue vascularization is of interest.
The formation of fibrin clot during wound healing is initiated by the release of thrombin, a
serine protease enzyme, which subsequently activates the coagulation cascade [Herrick et al.
1999; Rowe et al. 2007; Tawil 2006]. After the release, thrombin cleaves peptide fragments
from fibrinogen to generate the fibrin monomer (Fig. 2.1.b) by the clotting cascade into
protofibrils (Fig. 2.1.c). Afterward, in the presence of chloride ion and transglutaminase factor
XIII or factor XHIa, protofibrils undergo intermolecular cross-linking to form a stable fibrin
gel [Janmey et al. 2009; Litvinov et al, 2005; Rowe et al. 2007]. See Figure 2.1.d for details on
the fibrin formation as well as the degradation processes.
Changing fibrinogen or thrombin concentration can change the resulting fibrin material,
affecting both biochemical and mechanical properties [Ferrenq et al. 1997]. For example,
Vailhe et al. (1998) have shown that capillary-like structures made from HUVEC seeded on
fibrin depended on the mechanical factor of the gel. Harder gel, made using higher
concentration of fibrinogen, led to a decreased number of capillary-like structures. No
15
capillary-like structures was found in softer matrix (< 0.5mg/mL of fibrinogen) as well as in
too rigid one (> 4mg/mL of fibrinogen) [Ferrenq et al. 1997; Vailhe et al. 1998].
a
/».*feY"^-'."% v chain
D domain E domain O domain « * * ** *
two A audi two & "kiwlis* IK ., }B T-nunluk • msH|Mk. ^ \ Ii ^ 0-m,vlnk jr-«i"lufc
"ibrm unworncr
|— [)-ii£i>iii—~-| —~— J -ivgj n —— j—~l)-rfjjttin
i>'«f»« D*ejih>fl protofibril
i"**,\. Am<mi €»**
c»-'
»»&*
P»'i*?IS»ril»
r-*Jt.» fasm *rt
Uteri*
Figure 2.1: Fibrinogen structure and fibrin clot process. Adapted from Herrick et al. (1999)
and Litvinov et al. (2005).
16
Increasing fibrinogen concentration can reduce the matrix pore size, thus hindering endothelial
cell migration and capillary formation [Nehls and Herrmann 1996]. In addition, Rowe et al.
(2007) found that decreasing thrombin concentration resulted in both an increased gel
compaction and micro-fibres size, thus causing different cellular morphology and alignment of
vascular smooth muscle cells. These examples illustrate that fibrin gel properties and
subsequent cell responses can be modulated, to some extent, opening the door to more
applications [Cox et al. 2004; Rowe et al. 2007].
To date, fibrin is commercially available as fibrin sealant or fibrin glue (e.g., Tisseel™, Baxter
AG, Vienna, Austria) for surgical applications [Tawil 2006]. Also, fibrin scaffolds have been
used in many tissue engineering applications, including as matrix to treat bone and skin
defects [Catelas et al. 2006], for drug delivery in neurological and cardiovascular disorders
[Ferguson et al. 2003; Tassipoulos et al. 2000], and for three-dimensional angiogenesis assays
[Vailhe et al. 1998; Nakatsu and Hughes 2008]. For example, Montesano et al. (1986) and
Dejana et al. (1987) found that fibrinogen induced adhesion, spreading, and microfilament
organization of human endothelial cells either in 2D or 3D in vitro culture system.
Also, culturing endothelial cells on microcarrier beads and then embedding these beads in
fibrin have been proposed by Nehls and Drenckhahn (1995). This system resulted in formation
of capillary structures and sprouting [Nehls and Drenckhahn 1995]. However, the system
failed to model sprouting angiogenesis containing multi-cellular lumen surrounded by
polarized endothelial cells, which is of importance during blood microvessel development
[Montano et al 2009; Carmeliet and Conway 2001; Yancopoulos et al. 2000]. More recent
vascularization studies using fibrin gels have been presented by Chen et al. (2009), Nakatsu et
al. (2003a), and Sukmana and Vermette (2010a). In our study (see Chapter 3 of this thesis),
endothelial cells proliferate and migrate along patterned polymer fibres and then fibrin is
degraded along with the formation of cell-cell interactions, leading to the formation of tube
like structures, and eventually to sprouting and lumen formation with adjacent vessels
[Sukmana and Vermette 2010b].
17
Unlike synthetic hydrogels, fibrin is an active matrix for cells. It can bind many growth factors
and bioactive cloth components including fibronectin, hyaluronic acid, and von Willebrand
factor [Herrick et al. 1999; Weisel 2005]. Human fibrinogen can bind to endothelial cells
through either integrins or non-integrin binding sites (Table 2.1). For example, fibrin has two
pairs of RGD binding sites and a non-RGD site at the y chain, which can interact with
endothelial cell integrins (i.e., cc5pi, av(33, and anb(33) as well as with leucocyte integrins
(i.e., aMp3 and ccxp2) [Cheresh et al. 1989; Lishko et al. 2002; Tawil, 200; Weisel 2005]. The
location of integrins binding sites in fibrinogen is presented in Figure 2. La. Other non-integrin
receptors that can bind to endothelial cells include ICAM-1, CD-44 surface receptor, and
platelet endothelial cell adhesion molecule-1 (PECAM-1, also known as CD-31) [Cheresh et
al. 1989; Herrick et al. 1999; Monchaux and Vermette 2010].
Table 2.1: Integrins and non-integrin receptors which can bind to fibrin(ogen).
Integrins
ccvp3
oc5pl
ociibp3
ocMp3
«xP2
Non-integrin receptor
PECAM-1 (CD31)
ICAM-1
CD-44
Cell types
endothelial cells and fibroblasts
endothelial cells
platelets and endothelial cells
leucocytes and monocytes
lymphocytes and leucocytes
Cell types
endothelial cells
fibroblasts, endothelial cells
endothelial cells, fibroblasts and tumorous cells
Cellular function
adhesion and spreading
adhesion
adhesion and cloth retraction
adhesion and phagocytosis
adhesion
Cellular function
adhesion and von Willebrand factor release proliferation and adhesion
adhesion, migration, and proliferation
References
Cheresh et al. 1989; Cox et al. 2004; Monchaux and Vermette 2010. Cheresh et al. 1989; Herrick et al. 1999; Weisel 2005. Janmey et al. 2009; Monchaux and Vermette 2010. Herrick et al. 1999; Lishko et al 2002; Tawil 2006. Herrick et al. 1999; Janmey et al. 2009.
References
Monchaux and Vermette 2010; Nakatsu et al. 2003a; Cheresh et al. 1989; Herrick et al. 1999; Rowe et al. 2007. Herrick et al. 1999; Weisel 2005.
18
In other tissue engineering applications, fibrin was combined with collagen for the
development of blood vessel substitutes [Cummings et al. 2004; Stegemann et al. 2006].
Collagen type I is the predominant structural component of the media as well as the adventia
layers of blood vessels, while the inner layer of natural blood vessels, called the intima layer,
is lined by endothelial cells [Monchaux and Vermette 2010]. Therefore, the combination of
collagen-fibrin can be used to make scaffolds with good mechanical and biochemical
properties [Yao et al. 2005; Cummings et al. 2004]. For example, Isenberg et al. (2006) have
investigated a tubular scaffold made of type I collagen and fibrinogen to engineer small-
diameter artificial arteries.
A summary of the use of synthetic and natural polymers that have been applied to support
tissue vascularization is presented in Table 2.2.
Table 2.2: The use of synthetic and natural polymers to support tissue vascularization.
Scaffold Materials
Poly(D,L-lactic-co-glicolic acid)
PLLA/PLGA sponges
DLLA/TMC porous scaffold
Hyaluronic acid (HA)
HA cross-linked with chitosan
Purpose and Methods
• Scaffold designed for tubular tissues.
• EC were seeded in the construct.
• Scaffolds designed with pores from 225 to 500um for skeletal muscle tissue.
• Pre-seeded with MC, then scaffolds were implanted in skeletal muscles of mice.
• For heart tissue engineering with pore size oflOOum.
• Cardiomyocytes were seeded in the scaffold. Then scaffolds were subcutaneously implanted in rats.
Fibroblast growth factor (FGF) was added in the HA matrix.
Scaffold designed to produce a human skin
Results
EC were able to generate capillary network inside the scaffolds.
Scaffold pre-vascularization promoted angiogenesis in the implant.
This method elicited an acute inflammatory reaction.
FGF improved neovascularization.
The hybrid scaffold improved EC
References
Mooney et al. 1994; Mooney et al. 1996.
Levenberg et al. 2005.
Pego et al. 2003a; Pego et al. 2003b.
Rauh et al. 2006.
Black et al. 1999; Slevin et
19
Alginate gel
Marine collagen from tilapia (Oreochromis niloticas)
Collagen type I
Collagen type IV
Fibrin gel
equivalent. • Endothelial cells were
seeded in the construct.
• The construct was made for several applications including cell encapsulation.
• Vascular endothelial growth factor (VEGF) was incorporated in the scaffold.
Sponges were tested in vivo.
• In vitro angiogenesis assay.
• EC were seeded in the matrix.
Fibroblast growth factor (FGF) was incorporated in the scaffold for in vitro study of angiogenesis.
Fibrinogen and thrombin concentrations were changed to study angiogenesis, in vitro, using HUVEC and VSMC.
proliferation and induced al. 2002. capillary network formation inside the scaffold. VEGF promoted neovascularization.
Tilapia collagen caused less inflammatory responses when compared to porcine collagen. Type I of collagen increased the number of tube-like structures and supported angiogenesis development. • Addition of FGF in
type IV collagen scaffold supported EC growth and capillary formation.
• Degradation product of collagen type IV promoted a specific angiogenic epitope.
• Capillary-like structures (CLS) made of HUVEC depended on the matrix rigidity.
• Higher rigidity decreased CLS number.
• No CLS were found in very soft or very rigid matrices.
• Decreasing thrombin concentration caused different morphology and alignment of VSMC.
Chang et al. 2010; Drury and Mooney 2003.
Sugiura et al. 2009.
Francis et al. 2008; Soker et al. 2000.
Ingber and Folkman 1989; Xuetal. 2001.
Ferrenq et al. 1997; Nehls and Herrmann 1996; Rowe et al. 2007; Vailhe et al. 1998.
20
Fibrin gel in which microcarrier beads were embedded
Polymer monofilaments (i.e., PET) embedded in fibrin.
Fibrin gel combined with collagen type I
Aim: to generate angiogenesis development. Microcarrier beads were pre-coated with HUVEC and subsequently embedded in fibrin. In vitro study of angiogenesis guidance. PET fibres were pre-coated with HUVEC, then sandwiched in fibrin.
SMC were grown on a sheet-like scaffold, and then wrapped around tubular vessel. The vessel allowed to transfer nutrient and oxygen.
This in vitro culture system provided a step-by-step process of the capillary development containing multi-cellular lumen.
• Pre-coating PET with cells enabled increasing the fibres bioactivity.
• PET fibres were able to guide EC to orient microvessels.
This in vitro culture system enabled producing a better environment for the vascularization of small-caliber arterial substitutes.
Chen et al. 2009; Nakatsu et al. 2003a; Nehls and Drenckhahn 1995.
Sukmana and Vermette 2010a; Sukmana and Vermette 2010b.
Cummings et al. 2004; Isenberg et al. 2006; Yao et al. 2005.
EC: endothelial cell smooth muscle cell;
MC: mesenchymal cell; VSMC: vascular smooth muscle cell; SMC: PET: poly(ethylene terephthalate).
2.4 Material properties
Scaffold and ECM materials are selected based on bulk and surface properties, which can be
tuned with the aim to modulate cell adhesion and proliferation as well as phenotypic cell
expression [Francis et al. 2008]. Among the properties of importance, scaffold porosity and
matrix stiffness play significant roles in cell and tissue responses and these will be briefly
discussed below.
2.4.1 Scaffold porosity
Porosity is defined as the fraction of the void space over the total volume of a scaffold. Pore
size corresponds to the distance between two solid sections of the porous matrix [Lawrence et
al. 2008]. In tissue engineering, a highly porous scaffold (about 90% porosity) is more
desirable, since it should increase mass and nutrient transport [Kannan et al. 2005]. Higher
porosity can increase cell adhesion and provide a sufficient area for cell-matrix interactions
21
and new ECM production by cells [Agrawal and Ray 2001; Kannan et al. 2005]. However,
bulk and mechanical properties should also be considered. At higher porosity, the total solid
volume of the solid part of the scaffold is lower, when compared to scaffolds with lower
porosity, thus resulting in weaker mechanical support [Kannan et al. 2005].
The effect of pore size over cell behaviour has been investigated in culturing bone tissue
substitutes. Pores in the range of 300-400um have been found to be optimal for osteoblast
attachment, growth and proliferation [Linnes et al. 2007]. The important role of such porous
structures in endothelial cell organization and angiogenesis development was pioneered by
Clowes et al. (1986) more than 20 years ago. In a more recent study, pore size was found to
have a significant effect over cell binding, morphology, and phenotype, thus inducing
endothelial cell migration and capillary formation inside the scaffold [O'Brien et al. 2005].
In the scientific literature, there are some suggestions concerning the optimum pore size to
support vascularization. For example, many mature cell types, including fibroblasts and
endothelial cells, have been found to be unable to spread and completely colonize the bulk of
scaffolds with pore size higher than 300um, due to the difficulty on bridging the distance
[Yanmas et al. 1989; Zeltinger et al. 2001]. In an in vitro angiogenesis study, it was shown
that a cell's ability to bridge the distance in 3D scaffolds is important to support the
vascularization process [Sukmana and Vermette 2010a]. Using HUVEC-covered
poly(ethylene terephthalate) (PET) monofilaments, as contact guidance in HUVEC-seeded
fibrin, it was suggested that the optimum fibre-fibre distance to support microvessel
development was lOOum [Sukmana and Vermette 2010a].
Furthermore, hepatocytes were also reported to spread well on gelatin-chitosan scaffold (3:1)
with pore size of 20um, while fibroblasts and endothelial cells spread better on the same
matrix but with pores ranging between 100-150um, compared to pores from 20-80um [Huang
et al. 2005]. The role of pore size to promote endothelial cell lining has been investigated in
vascular grafts. For example, Zhang et al. (2004) found that an external pore size of 30um was
preferable over 20um or smaller pores in terms of promoting rapid tissue ingrowth and
endothelial cell growth in the expanded poly(tetrafluoroethylene) (ePTFE) graft.
22
In addition, Marshall et al. (2004) found that fibrin with 35um pores significantly supported
angiogenesis development, when compared to fibrin with either 20um or 70um pores. Fibrin
with pore sizes of approximately 30um promotes the ingrowth of vascularized fibrous tissue in
engineered blood vessels [Harding et al. 2002]. However, the optimum porosity and pore size
of the scaffold is still open to question and is based on the application as well as the cell type
[Kannan et al. 2005; Zeltinger et al. 2001; Zhang et al. 2004].
2.4.2 Matrix stiffness
Substrate mechanical properties, such as stiffness, are known to be important parameters
affecting cell responses. In cell biology, matrix stiffness is sensed by cell receptors and
integrins transmit mechanical stress across the cell surface to the cell cytoskeleton (see
•Appendix B for a brief description), converting mechanical signals into (bio)chemical ones
[Becker et al. 2009; Geiger et al. 2001; Ingber et al. 1995].
Therefore, the ECM stiffness will influence cellular functions, including cell adhesion,
proliferation, migration as well as phenotype differentiation [Geiger et al. 2001; Ingber et al.
1995]. For example, Pelham and Wang (1997) have examined the effect of collagen-coated
poly(acrylamide) scaffold on the behaviour of rat epithelial and fibroblast cells. They found
that on more rigid (higher stiffness) surfaces, cells were more spread and showed increased
motility and focal adhesion contacts [Pelham and Wang 1997]. Furthermore, increasing
surface stiffness was found to result in increased cell contractility [Li et al. 2005], more
organized cells cytoskeleton and actin stress fibres, and higher adhesion strength [Discher et
al. 2005]. The phenomenon related to the effect of environment stiffness over cell behaviour is
known to as durotaxis [Lo et al. 2000].
Substrate stiffness and cell contractility also have a significant role over microvascular
development. For example, while endothelial cells proliferate more on rigid surfaces, they
form tube-like structures on softer substrates [Deroanne et al. 2001; Discher et al. 2005].
Vasculogenesis decreased with an increase in matrix stiffness, which was a result of an
increase in collagen [Angele et al. 2004; Haarer and Dee 2006; Yeung et al. 2005] or
fibrinogen [Duong et al. 2009; Tawil 2006; Vailhe et al. 1998] concentration.
23
2.5 Scaffold vascularization
Cells can be isolated from a patient or donor and seeded into a scaffold to allow cell
proliferation and support pre-vascularization process [Lanza et al. 2000; Mikos et al. 1993].
Subsequently, these constructs can be implanted in the same patient. After implantation,
hopefully endothelial cells would develop connections with the existing blood microvessels of
the surrounding tissue, forming a microvascular network and allowing adequate tissue
perfusion [Lanza et al. 2000; Ye et al. 2000a].
When tissues become thicker, cells located more than a few hundred microns (about 200 to
300um) from capillaries suffer from a lack of oxygen and nutrients, resulting into necrotic
conditions [Hutmatcher 2001; Ko et al. 2007]. Currently, the development of thick and
complex tissues and organs such as the heart, muscles, kidneys, liver and lung relies, in part,
on the knowledge and ability to stimulate microvascular network formation within tissue
constructs [Ko et al. 2007; Mooney and Mikos 1999]. Therefore, knowing how to carry out
cell seeding within a scaffold and to perform successful in vitro cultures to prevascularize
tissue constructs prior to their implantation is highly important. As such, researchers rely on
the increasing knowledge related to neovascularization (i.e., vasculogenesis and angiogenesis)
process to stimulate vascular network formation within three-dimensional tissue constructs
[Bramfeld et al. submitted-2010]. Before discussing vasculogenesis and angiogenesis
processes, I will highlight the cell types available and/or used to carry out these experiments.
The existence of capillaries in blood vessels and lung of frogs was first reported by Marcello
Malpighi in 1661 [Hillen et al. 2006]. After the invention of biological microscopes by
Antonie van Leeuwenhoek (1632-1727) and his colleagues and further development of
fluorescence techniques, the anatomy of blood vessels was revealed [Hillen et al. 2006]. Blood
vessels are composed of several cell types and the main ones are smooth muscle cells (SMC),
pericytes, fibroblasts and endothelial cells [Jain 2003; Ko et al. 2007].
Smooth muscle cells are found in blood vessels, such as in middle layer (i.e., tunica media) of
large and small blood vessels, in lymphatic vessels, uterus, and in the gastrointestinal and
respiratory systems [Hillen et al. 2006; Jain 2003]. Behind the basic function of vascular SMC
24
in blood vessels i.e., to maintain vessels integrity and support the endothelium [Auerbach et al.
2003; Hillen et al. 2006], they are highly specialized cells for the regulation of blood vessel
diameter, vessel contraction, blood pressure and flow distribution [Auerbach et al. 2003;
Owens et al. 2004]. Furthermore, SMC synthesize the connective tissue matrix of the vessel
wall, which is composed of elastin, collagen, and proteoglycans [Auerbach et al. 2003; Hillen
et al. 2006]. SMC show a very low level of proliferation during normal development but
become more proliferative in response to vessel injury [Jain 2003].
There are some markers to identify smooth muscle cells, including smooth muscle cc-actin
(SMaA) [Owens et al. 2004], smooth muscle myosin heavy chain (SM-MHC) [Bramfeld et al.
submitted-2010; Hungerford and Little 1999], SM22cc, and calponin [Duband et al. 1993]. To
date, SMaA is the most commonly used marker of SMC, which represents up to 70% of the
actin population in vascular SMC [Bramfeld et al. submitted-2010; Hungerford and Little
1999; Jung etal. 2002].
Pericytes are perivascular-specific cells that are associated with capillaries and blood
microvessel development [Hillen 2006; Jain 2003]. Pericytes have the capacity to differentiate
into other cell types, including SMC, fibroblasts, and osteoblasts [Jung et al. 2002; Owens et
al. 2004]. During blood microvessel development, the coverage of capillary by pericytes is
important for the maturation, remodeling and maintenance of the vascular system via the
secretion of growth factors and/or modulation of the ECM [Allt and Lawrenson 2001; Jung et
al. 2002]. The important role of pericytes in capillary development in vitro has been studied by
some researchers. For example, endothelial cells co-cultured with pericytes, separated by a
cellulose membrane, resulted in inhibition of capillary growth, while when pericytes were in
contact or close to endothelial cells; endothelial cell growth and capillary development were
observed [Orlidge et al. 1987].
Fibroblasts are a cell type mostly found in connective tissues, including blood vessels,
cartilage, soft tissues (e.g., dermis) and bone. A connective tissue can be defined as a tissue
that wraps, connects, nourishes and supports all other tissues and organs [Becker et al. 2009].
Fibroblasts originate from mesenchymal cells - mesenchymal cells are progenitor cells
25
capable of forming connective tissues and the lymphatic system [Auerbach et al. 2003; Jain
2003]. During normal development and wound healing process of connective tissues,
fibroblasts produce fibres and secrete factors to maintain the ECM and provide structural
support for the tissues [Auerbach et al. 2003]. To date, fibroblasts from human skin or skin
fibroblasts have been co-cultured with endothelial cells in many angiogenesis studies [Allt and
Lawrenson 2001; Chen et al. 2009; Nakatsu et al. 2003a; Sukmana and Vermette 2010b].
Also, 3T3 cells that were originally obtained from Swiss mouse embryo tissues have become a
standard fibroblast cell line [Allt and Lawrenson 2001]. In an in vitro study, skin fibroblasts
have been reported to have a significant effect over microvascular formation and angiogenesis
development from HUVEC [Nakatsu et al. 2003a; Sukmana and Vermette 2010b].
In the entire circulatory system, endothelial cells line the inner layer of the vessels (i.e., tunica
intima) and consist of more than 1013 cells for approximately 1kg of vessel [Jain 2003;
Sumpio et al. 2002]. The first culture of endothelial cells was reported by Shibuya around
1930, and then various techniques and sources of endothelial cells were investigated [Hillen et
al. 2006]. For in vitro studies, endothelial cells from animal or human sources can be used.
Endothelial cells from animals include canine jugular endothelial cells and aortic endothelial
cells either from bovine (BAEG), porcine (PAOEC) or rat (RAOEC) [Ko et al. 2007] sources.
Human endothelial cells include human umbilical vein (HUVEC), human dermal
microvascular (HDMEC) and human vascular (HVEC) [Tonello et al. 2003]. Since endothelial
cells from animals can result in immune responses if used in humans, and due to the available
sources of endothelial cells, HUVEC have been utilized by many research groups [Ko et al.
2007].
Heterogeneity between endothelial cells has been reported not only between large vessel-
derived cells and those of microvascular origin, but also between organs [Hillen et al. 2006;
Sumpio et al. 2002]. For example, aortic endothelial cells are thicker and cover a small area
compared to those from human pulmonary artery [Fuchs et al. 2001]. Also, endothelial cells
from microvascular (e.g., human dermal microvascular endothelial cells (HDMEC)) are
elongated, while those from human umbilical vein (HUVEC) are polygonal [Fuchs et al. 2001;
Lang et al. 2003].
26
Furthermore, endothelial cells derived from arteries are different than those isolated from
veins (Figure 2a) [Aird 2007; Lang et al. 2003]. In arteries, endothelial cells are long and
narrow, aligned in the direction of blood flow and form a continuous endothelium with many
tight junctions. Endothelial cells from veins are shorter and wider, and not aligned in the
direction of blood flow [Aird 2007]. These heterogeneities reflect their difference in
functionality in term of the release of vasoactive substances and their interaction with
leucocytes during normal vessel development and wound healing [Aird 2007; Lang et al.
2003; Otto etal. 2001].
Some unique molecular markers as well as genes, preferentially expressed by endothelial cells,
are listed in Table 2.1. For example, the expression of CD31 (also known as platelet
endothelial cell adhesion molecule-1 or PECAM-1), CD34, and von Willebrand factor (vWF)
as well as Dil-acetylated LDL uptake has been used in many studies to distinguish endothelial
cells from other cell types [Bramfeld et al. Submitted-2010; Hillen et al. 2006; Aird 2007].
Also, cell adhesion molecules (CAM), such as ICAM-1 (intercellular adhesion molecule-1),
VCAM-1 (vascular cell adhesion molecule-1), E- and P-selectins can be used to identify
endothelial cells during wound healing and angiogenesis [Monchaux and Vermette 2010; Otto
et al. 2001; Peters et al. 2008; Kirkpatrick et al. 2003].
Other cell-cell adhesive proteins, such as ESAM (endothelial cell selective adhesion
molecule), VE-cadherin (vascular endothelial cadherin) and N-cadherin, are known to have a
significant in angiogenesis regulation [Dejana 2004; Fuchs et al. 2001; Hillen et al. 2006].
ESAM is a tight junction molecule that is responsible for the regulation of cellular
permeability and for maintaining the polarity of endothelial cells [Dejana 2004; Fuchs et al.
2001]. VE-cadherin is an adherens junction molecule that plays an important role in regulating
cell growth and in the organization of new vessels during angiogenesis [Bazzoni and Dejana
2004; Hillen et al. 2006]. Another member of the cadherin family, N-cadherin is localized on
the basal side of endothelial cells [Hillen et al 2006]. During vessel maturation, N-cadherin is
in contact with pericytes or smooth muscle cells [Bazzoni and Dejana 2004; Dejana 2004].
27
Furthermore, in other angiogenesis studies, the expression of some genes to assess if
endothelial cells are undergoing angiogenesis has been suggested. Some genes that are
preferentially expressed during endothelial cell sprouting, lumen formation and capillary
network establishment are HESR-1, notch 1, notch 4 and delta 4 [Nakatsu et al. 2003a;
Sainson et al. 2005; Henderson et al. 2001; Taylor et al. 2002].
2.5.1 Vasculogenesis and angiogenesis
Recent studies on blood microvessels have provided essential information to develop
strategies allowing neovascularization. Microvessels can develop through two processes:
vasculogenesis and angiogenesis. Both can be potentially applied to vascularize tissue
constructs. Although in some papers both terms are often used to mean the same thing and are
simply referred to as angiogenesis, the specific role of the micro-environmental factors and the
overall mechanisms are different [Francis et al. 2008].
Vasculogenesis refers to the process of differentiation of endothelial progenitor cells (i.e.,
mesodermal, mesenchymal or bone marrow cells) to form new blood vessels. For example, in
dermal tissues, vasculogenesis occurs in three main steps: 1) differentiation of mesodermal
cells into angioblasts or hemangioblasts; 2) differentiation of angioblasts or hemangioblasts
into endothelial cells; and 3) the organization of new endothelial cells into a primary capillary
plexus [Moon and West 2008].
Angiogenesis can occur in two different ways: 1) intussusceptive, the longitudinal splitting of
existing vessels, and 2) sprouting angiogenesis, the outgrowth of a new branch from a pre
existing vessel [Cockerill et al. 1995; Levin et al. 2007; Risau 1997]. Compared to the
intussusceptive angiogenesis that mostly occurs during new organ formation as well as during
tumour development, sprouting angiogenesis is relatively well characterized [Levin et al.
2007]. Here, I will focus on sprouting angiogenesis. Sprouting angiogenesis refers to the
formation of new capillaries from pre-existing blood vessels, which is mainly initiated by a
sprouting process [Cockerill et al. 1995; Risau 1997].
28
Angiogenesis involves four different stages: 1) endothelial cells interact with their ECM and
the underlying basement membrane; 2) they proliferate, migrate and communicate with each
others; 3) they form cell-cell connections and tube-like structures; and 4) tube sprouting and
remodelling occur to form microvessels containing multi-cellular lumen [Sukmana and
Vermette 2010a; Risau 1997].
Our knowledge of angiogenesis as well as the availability of numerous models was pioneered
about 30 years ago by Folkman and Haudenschild (1980) when they demonstrated that new
capillary blood vessels form in tumor progression. They observed that capillary tube formation
was initiated from a vacuole structure within the endothelial cells that subsequently develop
multi-cellular capillary lumen [Folkman and Haudenschild 1980]. More recently, Kucera et al.
(2007) have proposed three models explaining how tube-like structures developed during the
vascularization process: 1) Cell death and phagocytosis. 2) Wrapping of the spaces around the
extracellular matrix, and 3) vacuole formation from the coalescence of intracellular vacuoles.
Below, I will briefly discuss these three models [Kucera et al. 2007]. The models are presented
on the Figure 2.2.
2.5.1.1 Model 1: cell death and phagocytosis
Some in vitro as well as in vivo studies of angiogenesis report that lumen formation was
associated with apoptotic and cell death events [Fierlbeck et al. 2003; Meyer et al. 1997]. This
model proposed the following steps in vascular lumen formation: endothelial cells organize
themselves into a multi-cellular cord and cells in the middle of the cord become apoptotic and
die. Finally, endothelial cells at the edge phagocytise the apoptotic cells, subsequently forming
a multi-cellular vessel containing lumen (see Figure la). According to this model, dead cells
define the lumen of the tube-like structures. However, this model fails to explain how the
tube-like structures continue their formation into a continuous vessel [Kucera et al. 2007].
2.5.1.2 Model 2: wrapping spaces around ECM
This model mainly suggests that most capillary are initiated by the elongation of endothelial
cells to open multi-cellular structures. Subsequently, vascular tube can form when the
elongated cell sheets are closed, even without vacuole formation (see Figure lb) [Kucera et al.
29
2007]. This concept was initiated by Hirakow and Hiruma (1983), when they studied the
development of vascular lumen made of endothelial cells without the evidence of vacuole-like
structures. In a recent study, Parker et al. (2004) have shown that the vascular development
was initiated by the proliferation and migration of endothelial cells to form a cord-like
structure, and vascular cords then undergo tubulogenesis to form vessels containing lumen.
However, this model does not explain the polarity of endothelial cells during the development
of capillary structures [Kucera et al. 2007].
' Lumen %'
Phagocytosis" and exocytosis
Lumen levelopmen
b
Elongation * Closure and IS lumen formation
m Membrane *
invagination
fff Vacuole
Vacuole formation
Lumen
Vacuole fusion & lumen development
"f^ Extracellular matrix
^,r Endothelial cell
Apoptoticcell
Figure 2.2: Models of the development of tube-like structures during vasculogenesis. Adapted
from Kucera et al. (2007).
30
2.5.1.3 Model 3: vacuole formation from the coalescence of intracellular vacuoles
The idea behind this model was proposed by Folkman and Haudenschild (1980) when
explaining angiogenesis and lumen formation within 3D constructs. This model involves
several steps: formation either of single or multiple vacuole(s) inside the endothelial cells and
vacuoles subsequently fuse with each others. Finally, a continuous multi-cellular lumen will
form when two or more cells with their vacuoles adhere to each others, therefore fusing (see
Figure lc) [Folkman and Haudenschild 1980; Davis et al. 2000].
Davis et al. (2000) have reported that endothelial cell lumens were forming in three-
dimensional collagen and fibrin matrices. The authors reported that this process was controlled
by the formation and coalescence of intracellular vacuoles and in the absence of cellular
junctions [Davis et al. 2000]. Furthermore, the expression of cytoskeletal regulators
controlling various cellular functions of endothelial cells, such as Cdc42 and Racl GTPases,
' were involved in morphogenesis and vascularization processes [Connolly et al. 2002; Bayless
and Davis 2002].
Even though this model can explain the role of vacuoles in capillary tube structure and lumen
formation, some questions remain, such as how endothelial cells compensate their basal side
during the vacuole coalescence and vacuole fusion, and how the apical-basal polarity of
endothelial cells can be established during tube stabilization [Kucera et al. 2007; Yancopoulos
2000]. Since none of these three models seem to include all the cellular events observed
during vascular tube development, it can thus be hypothesized that all these three models can
be combined, in some way, to explain observations made in angiogenesis and vasculogenesis
assays [Darland and D'Amore 2001; Hubbel 2003; Lutolf and Hubbel 2005].
2.5.2 Promoting angiogenesis in three-dimensional constructs
During vasculogenesis and angiogenesis development, cell-cell as well as cell-ECM
interactions are complex [Darland and D'Amore 2001; Lutolf and Hubbel 2005]. While the
roles of some individual factors during neovascularization have been investigated, the
optimum combination between cell and their ECM can significantly vary from one material to
31
another [Hubbel 2003]. Therefore, some strategies have been proposed to promote and direct
angiogenesis in 3D environments.
Endothelial cells have been cultured in various matrices that have been pre-coated or not with
adhesive matrix proteins or cell binding peptides. Surface chemistry and the immobilization
mode can affect protein conformation, orientation and anchorage strength, thus influencing
cell behaviour [Hubbel 2003]. Formation of capillary-like structures made of endothelial cells
depends on the particular biological environment that will direct cell-cell as well as cell-
biomaterial adhesion strength, which relies on biochemical and mechanical signals [Kannan et
al. 2005]. To date, there are three major strategies that have been used to study how
biomaterials properties affect angiogenesis formation: 1) Micro-patterning, 2) Surface
modification, and 3) extracellular matrix modifications [Kannan et al. 2005; Moon and West
2008].
2.5.2.1 Micro-patterning
Photolithography, micro-moulding, and micro-printing are some examples of techniques used
in micro-patterning. Among them, laser micro-contact printing is the most frequently studied
[Hahn et al. 2006; Kannan et al. 2005]. It is a process in which biological molecules are
printed directly onto a scaffold surface, and it has been broadly utilized by tissue engineering
scientists. For example, a poly(ethylene glycol)-diacrylate (PEGDA) hydrogel, known to resist
protein adsorption, has been micro-printed with the cell adhesive ligand, Arg-Gly-Asp-Ser
(RGDS) in different concentrations and using different patterns. Endothelial cells were
cultured on these RGDS patterns and enhanced tube-like structure formation was observed on
50u,m-wide stripes [Moon and West 2008].
In another study, micro-contact printing was used to form various patterns of fibronectin
molecules on gold [Moon and West 2008]. Endothelial cells cultured on these fibronectin
patterns formed tubular structures on lOum-wide lines of fibronectin [Moldovan and Ferrari
2002]. In a different study, when coated with gelatin, 20u.m lines promoted endothelial cells to
undergo capillary morphogenesis [Medalie et al. 1996]. However, applying this technique to
3D scaffolds is problematic [Kannan et al. 2005].
32
2.5.2.2 Surface modification
Proteins such as fibronectin, gelatin, collagens, and fibrinogen, only to name a few, have been
used to coat synthetic polymers to increase their surface bioactivity [Ye et al. 2000a]. For
example, collagen coating on the surface of electrospun poly(L-lactic acid)-co-poly(e-
caprolactone) scaffold increased endothelial cell adhesion and spreading [He et al. 2005; Pham
et al. 2006]. Collagen type I has been known to increase the number of tube-like structures
made of endothelial cells, thus supporting angiogenesis and vasculogenesis in in vitro
experiments [Davis and Sanger 2005]. Furthermore, as previously described, the proteolytic
cleavage of collagen type IV provides an important binding site for endothelial cells
undergoing angiogenesis in vitro [Xu et al. 2001].
Gelatin is used by many research groups to pre-coat tissue culture dishes. In tissue
engineering, gelatin was used to coat poly(glycolic acid) (PGA) scaffold for the controlled
release of some angiogenic growth factors [Dcada and Tabata 1998; Tabata et al. 1994; Tabata
and Ikada 1999]. When PGA scaffolds covered by bFGF- and VEGF-containing gelatin
hydrogel were implanted in mice, significant angiogenic effect was observed at the implanted
site [Dcada and Tabata 1998; Tabata 2009]. Further clinical test to treat diabetic foot ulcer
using gelatin matrix to release bFGF confirmed that the matrix allowed better wound healing
[Tabata 2009].
Fibronectin has been investigated in many studies and found to enhance endothelial cell
adhesion and spreading [Kannan et al. 2005; Monchaux and Vermette 2010]. Fibronectin also
supports the formation of focal adhesion and induces the organization of actin filaments into
stress fibre via the interaction with its main receptor i.e., a5|31 and avP3 [Burmeister et al.
1996; van Wachem et al. 1997]. Furthermore, the lack of fibronectin during vascular
development caused an abnormality in the vascular formation in mice [Burmeister et al. 1996;
George et al. 1997].
Another protein, fibrinogen, has been incorporated in poly(ethylene glycol) (PEG) hydrogels
and it found to increase the bioactivity of the scaffold by enhancing endothelial cell and SMC
adhesion [Almany and Seliktar 2005] as well as mesenchymal stem cell adhesion [Zhang et al.
33
2006]. Furthermore, fibrinogen has many binding sites for endothelial cells. Also, fibrin gel
binds many growth factors and bioactive cloth components [Herrick et al. 1999; Weisel 2005].
Therefore, the use of fibrin(ogen) to modify biomaterials surfaces and to encapsulate cells in
tissue engineering applications is of importance [Janmey et al. 2009; Tawil 2006].
In other strategies, polymer surfaces can be pre-coated with cells to increase the bioactivity of
the scaffold. For example, expanded poly(tetrafluoroethylene) (ePTFE) was pre-coated with
bladder carcinoma cells before being implanted in mice. This strategy stimulated angiogenesis
and neovascularization in ePTFE vascular grafts [Kidd et al. 2002]. In a more recent study,
pre-coated PET fibres with HUVEC allowed the guidance of the angiogenesis process and
subsequent micro-vascularization in fibrin [Sukmana and Vermette 2010b]. No microvessel
structures were observed when uncovered fibres were utilized (see Chapter 4 of this thesis).
2.5.2.3 ECM modification
The interaction between cells and their matrix is very complex, since there are many proteins
and soluble molecules (including growth factors) present in the ECM [Davis et al. 2002; Davis
and Sanger 2005; Ingber and Folkman 1989]. These molecules interact with cells through
different cell binding domains. In blood vessel development, there are approximately 20
angiogenic growth factors, and among them vascular endothelial growth factor (VEGF), basic
fibroblast growth factor (bFGF), platelet-derived growth factor (PDGF) and angiopoietin-1
(Agp-1) are the most frequently studied [Nakatsu et al. 2003b].
In in vitro angiogenesis assays, growth factors can be added to the culture media and/or
immobilized in the ECM. These strategies have been found to have a significant effect on the
development of tube-like structures and on microvessel maturation [Fourier and Doillon 2007;
Zisch et al. 2003]. For example, endothelial cells cultured in type I collagen gels with culture
media supplemented with fibroblast growth factor-2 (FGF-2), vascular endothelial growth
factor (VEGF), and phorbol ester formed tubes containing lumens, which have similar
structure to microvessels found in vivo [Davis and Camarillo 1996]. In the absence of growth
factors and phorbol ester, the vascular tube structures did not form [Davis and Camarillo 1996;
Kannan 2005]. Other in vitro angiogenesis studies showed that growth factors added to culture
34
media can promote sprouting, lumen formation and better vessel stability [Chen et al. 2009;
Nakatsu et al. 2003b; Pepper 1997; Uemura et al. 2002].
Growth factors are often bound to the ECM and can be released upon interaction with cells
[Moon et al. 2009; Moon and West 2008]. In some studies, growth factors were immobilized
to the ECM that was used to carry out the culture [Hubbel 2007; Lutolf and Hubbel 2005]. In
this approach, growth factors can be released, as encountered in vivo. For example, VEGF was
covalently immobilized into type I collagen gel [Koch et al. 2006]. Using the chicken
chorioallantoic membrane (CAM) assay, this collagen matrix was found to induce capillary
formation and tissue ingrowth [Koch et al. 2006]. Also, basic fibroblast growth factor (bFGF)
was immobilized into poly(ethylene glycol) (PEG) hydrogels to guide cell alignment and
migration [DeLong et al. 2005; Koch et al. 2006; Moon et al. 2009].
As many cellular processes observed during vasculogenesis require numerous signalling
pathways and growth factors, recent research efforts have been focusing on the sequential
delivery of multiple growth factors [Mooney et al. 1997]. For example, the sequential delivery
of VEGF and PDGF-BB (platelet-derived growth factor-BB) in a controlled-release polymer
device made of poly(lactic glycolic) (PLG) (85:15 lactide:glycolide) induced a mature
vascular network covered by smooth muscle cells [Richardson et al. 2001; Sumpio et al.
1998]. Other results also concluded that one growth factor alone is not sufficient to create
mature and stable vasculature [Blau et al. 2001; Mooney et al. 1997].
2.5.3 Co-culture systems and microvessel maturation
Achieving microvessel maturation and stabilization during blood microvessel development
represents another challenge following vascular tube formation by endothelial cells. This is
often referred to as angiogenesis remodelling [Hubbel 2003; Yancopoulos 2000].
Angiogenesis remodelling is the process by which primary vascular tubes or immature vessels
are modified to form an interconnecting branching network, yielding to microvessel structure
stabilization [Ennett and Mooney 2002; Hubbel 2003]. During this phase, tube-like structures
made of endothelial cells and containing lumens recruit other supporting cells, such as
35
pericytes and smooth muscle cells (SMC), to form a close wall and mature blood microvessels
[Hubbel 2003; Jain 2003].
Furthermore, in the vascular tree, endothelial cells are lining on the inner layer of the vessel
wall - this is the tunica intima - and are supported by other cell types [Jain 2003; Korff et al.
2001]. For example, immature vessels consist of tube structures made of endothelial cells.
Capillaries consist of tubes made of endothelial cells covered by pericytes and a basement
membrane. In arterioles and venules, vascular SMC cover large area and are closely associated
with the endothelium basement membrane. Finally, as discussed in a previous section, the
structure of large veins and arteries consists of three distinct layers: tunica intima, tunica
media and tunica adventia [Carmeliet and Conway 2001; Jain 2003]. The reader is referred to
Figure 3 for schematic illustrations of the different vessel structures.
Figure 2.3: Blood vessel structure and development. Adapted from Jain (2003).
36
Therefore, co-culture systems seem to be appealing to recreate the real microvessel
environment and to achieve more mature microvessels. For example, smaller number of
vessel-like structures containing lumens was found when endothelial cells were cultured alone
in PLLA-PLGA (50:50) scaffolds, compared to samples co-cultured with fibroblasts
[Levenberg et al. 2005]. Vascularized engineered skeletal muscle tissue constructs have been
successfully produced by Levenberg et al. using a multiculture system of myoblasts
(progenitor cells of the muscle cells), fibroblasts and endothelial cells into this PLLA-PLGA
scaffold [Levenberg et al. 2005].
2.6 Bioreactors to support vascularization
Many tissue engineering scientists believe that the next generation of functional tissue
replacements truly needs the use of bioreactors, in which the culture conditions (e.g., pH,
temperature, O2 concentration, pressure, pulsation, nutrient transfer and waste removal, etc.)
can be adjusted and studied to find the optimal mechanical, chemical, and biological stimuli
for a given application [Martin et al. 2004]. In addition, bioreactor operations will provide a
rational basis for the structural and functional design of engineered tissues for the use as
model systems to reduce time of innovation, discovery and production in biological and
clinical research [Martin et al. 2004; Martin and Vermette 2005]. Also, the use of bioreactors
should accelerate the development, evaluation, and delivery of engineered tissue products to
patients [Freed et al. 2006].
To date, there are various types of bioreactors that have been designed and tested and these
differ from the applications involved. For example, bioreactors have been used to support the
growth of cartilage [Obradovic et al. 1999; Pazzano et al. 2000; Seidel et al. 2004], bone
[Ignatius et al. 2005; Yu et al. 2004], cardiac [Carrier et al. 2002; Radisic et al. 2004],
ligament [Alltman et al. 2002; Lee et al. 2005], and heart valve [Hoerstrup et al. 2000;
Schenke-Layland et al. 2003] tissues.
Furthermore, there are some types of bioreactors that can be used in large tissue cultures, and
these include the continuous stirred-tank reactor (CSTR), hollow fiber reactors and the slow
turning lateral vessel (STLV) reactor [Martin and Vermette 2005], only to name a few. CSTR
37
reactors are designed to mix oxygen and nutrients within the culture medium and to reduce the
boundary layer at the scaffold surface [Belfort 1989]. Hollow fibre reactors consist of a
chamber perfused with semi-permeable fibres filled with culture medium. Often, cells are
located in the extracapillary space of the chamber. In the past years, this type of reactor was
used to produce proteins made by mammalian cells, and to expand mammalian cells in vitro
[Jauregui et al. 1996; Schwarz et al. 1992]. STLV reactor was proposed by NASA (National
Aeronautics and Space Administration) at the Johnson Space Center (USA) for earth-based as
well as space experiments [Schwarz et al. 1992].
In the context of vascularization and blood microvessel development, ideally the bioreactor
should enable to control basic environmental parameters such as the dissolved oxygen
concentration, pH, and temperature, as well as tissue factors including cellular structures and
function [Bilodeau and Mantovani 2006]. For example, the system should allow the transport
of nutrients and oxygen from the reservoir to the cell location within the scaffold. As it has
been long known, the maximum tissue thickness achievable by diffusive transport alone is
approximately 1mm [Martin and Vermette 2005]. The optimization of scaffold material
properties as well as the design of bioreactor chamber is therefore of upmost importance
[Chouinard et al. 2009].
Furthermore, bioreactors can also enable control over the mechanical stimulation for cellular
guidance inside the scaffold in order to improve tissue vascularization. As endothelial cells are
lining the inner surface of blood vessels of the entire capillary and circulatory system, they
experience fluid shear stress and dynamic flow conditions [Aird 2007; Xu et al. 2001].
Therefore, the use of a dynamic culture system can allow some cell mechanical stimulation
prior to the use of the grown tissue construct [Levenberg et al. 2005; Lawrence and Madihally
2008].
Dynamic culture parameters such as pulsation, pressure, and flow rate that mimic in vivo
conditions often result in better cellular organization as well as mechanical properties of the
tissue constructs when compared to constructs cultured in static environment [Sodian et al.
2002; Yao et al. 2005]. To date, several studies of endothelial cell monolayers have revealed
38
that cell orientation reflects the direction of the flow [Lesman et al. 2010; Li et al. 2005;
Stegemann and Nerem 2003]. Also, in endothelial cells, shear stress increased VEGF [Conklin
et al. 2002] and PDGF-BB expression [Sumpio et al. 1998] compared to those cultured in
static conditions. In addition, pulsatile flow can improve structural organization of SMC and
this is an essential step in the vascularization process [Akhyari et al. 2002; Kofidis et al. 2002;
Niklason et al. 1999].
2.7 Concluding remarks
Over the past years, significant advances in tissue engineering research have provided hope for
the commercial availability of human bioartificial materials. More simple engineered tissue
substitutes (e.g., skin, bone, and cartilage) have been successfully applied and implanted,
while for more complex and vascularized tissues, many problems still remain before tissue
substitutes can be available. Consequently, the development of biomaterials to engineer tissue
constructs as well as strategies to enhance vascularization inside tissue constructs becomes
important to provide scaffolds with improved oxygen and nutrient transfer as well as waste
removal. This could allow production of thicker tissue substitutes as well as artificial organs.
Tissue engineering research relies on the increasing knowledge of angiogenesis and
vasculogenesis mechanisms occurring during capillary tube formation and blood microvessel
development. Furthermore, to overcome the bottleneck of the complex interplay between
various factors influencing tissue vascularization, the use of bioreactor system is necessary.
39
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56
CHAPTER 3
Polymer Fibres as Contact Guidance to Orient
Microvascularization in a 3D Environment
Irza Sukmana1'2, Patrick Vermette1'2
1. Laboratoire de Bioingenierie et de Biophysique de l'Universite de Sherbrooke, Department
of Chemical Engineering, Universite de Sherbrooke, 2500, blvd de l'Universite, Sherbrooke,
QC, Canada, J1K2R1.
2. Research Centre on Aging, Institut universitaire de geriatrie de Sherbrooke, 1036, rue
Belvedere Sud, Sherbrooke, QC, Canada, J1H 4C4.
Article published in J Biomed Mater Res A 92 (2010), p. 1587-1597.
DOI: 10.1002/jbm.a.32479
http://www3.interscience.wiley.com/iournal/122381455/abstract
57
3.1 Abstract
We describe an in vitro culture process that uses 100-um diameter poly (ethylene
terephthalate) monofilaments as contact guidance of human umbilical vein endothelial cells
(HUVECs) to orient the development of microvessels in a 3D environment. Untreated fibres,
distanced either by 0.05, 0.1, 0.15, or 0.2 mm, were first covered with HUVECs then,
sandwiched between two layers of fibrin gel containing HUVECs. After 2 and 4 days of
culture, cell connections and microvessels were evaluated. Cell connections formed massively
along and in between adjacent fibres that were distanced by 0.05 and 0.1 mm, whereas with
fibres separated by larger distances, connections were rare. After 4 days of culture, the
optimum fibre-to-fibre distance to form microvessels was 0.1 mm. This study reveals that
polymer fibres embedded in gel can be used as guides to direct the microvascularization
process, with potential applications in cancer and cardiovascular research and tissue
engineering.
3.2 Introduction
One of the recent developments in the field of biotechnology is tissue engineering, which is
the application of engineering principles and medical sciences to develop tissue substitutes
that can be used to replace, repair or regenerate damaged, injured, or missing body parts [1-3].
Although massive progress has been made, some major problems are still left to culture
human tissues. The lack of a sufficient supply of nutrients and oxygen to growing tissues in
vitro, as well as waste removal are two important factors that often lead to the failure of the
culture process or even to that of implants [3,4]. One possible strategy for creating three-
dimensional (3D) engineered tissue substitutes in vitro is to develop a scaffold allowing the
guidance of cells in order to promote microvascularization in directional fashion.
Pre-vascularization of tissue-scaffold appears to be the most favorable and efficient approach
to address the problem of tissue survival due to a lack of oxygen supply. Pre-vascularization
mainly involves the incorporation of endothelial cells or vascular-like structures into a scaffold
and then to implant this scaffold at the desired anatomical site. This approach could favor a
link between the existing vasculature from the host and the endothelialized tissue construct.
58
Furthermore, it could accelerate the formation of functional microvessels in the interior of an
implant [5,6].
Angiogenesis is a key mechanism involved in the development of new blood vessels as well as
during wound healing [7,8]. Several assays have been described to grow networks of
capillaries and to promote angiogenesis in 3D environment, often composed of gel [9-12].
Although some steps of the angiogenesis process have been identified, the origin and
mechanisms involved in lumen formation has been the subject of some debates [13]. The term
lumen is sometimes used to describe features composed of cells forming circular structures in
a nearly 2D plane rather than 3D tube-like structures made of multiple and interconnected
cells, as also suggested by others [14].
Other approaches have been developed to modulate angiogenesis with the use of pre-
vascularized scaffolds made of either natural fibres [15,16] or polymer fibres [17]. Some cells
can orient and often migrate along fibres axis. Cells can also bridge from one fibre to another,
if fibres are separated by distances small enough [18-21]. The fibre-to-fibre distance and the
pattern by which fibres are presented to cells can be adjusted to modulate cell guidance [19-
21]. Furthermore, as shown in a previous study, fibres pre-coated with cell-adhesive
molecules, such as the RGD peptide, were used to guide cell responses [21].
The objective of this study was to develop and validate a culture process that can be
subsequently scaled to bioreactor cultures and that uses polymer fibres to guide the
development of microvessels. We hypothesize that the orientation of microvessels would
subsequently facilitate the flow induction within microvessels lumen for microvascularization
and angiogenesis studies.
3.3 Materials and Methods
3.3.1 Materials
100-um diameter monofilaments made of poly(ethylene terephthalate) (PET, ES305910, Good
Fellow, Devon, USA) were used and fixed onto polycarbonate (Boedeker Plastics Inc., Texas,
USA) frames that fitted into wells of 6-well plates traditionally used in cell culture. The
59
frames were 32.50 mm in diameter and 3.00 mm thick (Fig. 3.1b). A scanning electron
microscopy (SEM) picture of a PET fibre is presented as Supplementary Figure 3.i.
The phosphate buffered saline solution (PBS IX, pH 7.4) used in these experiments was
prepared from Milli-Q water and NaCl (0.137 M), KC1 (0.003 M), Na2HP04 (0.008 M), and
KH2P04 (0.002 M). This 150 mM solution was diluted in Milli-Q water to make a 10 mM
solution. Hank's balances salt solution (HBSS, H1387) and albumin from bovine serum (BSA,
CAS 9048-46-8) were purchased from Sigma-Aldrich Inc. (St Louis, USA).
3.3.2 Cell culture
Human umbilical vein endothelial cells (HUVECs) were purchased from Cambrex (cc-2519,
Walkersville, MD USA). HUVECs were cultured at 37°C and 5% C0 2 in Ml99 culture
medium (M5017, Sigma-Aldrich) containing 2.2 mg mL"1 sodium bicarbonate (Fisher, Fair
Lawn, USA), 90 \ig mL"1 sodium heparin (HI027, Sigma-Aldrich), 100 U/100 ug mL"1
penicillin/streptomycin (15140-122, Invitrogen Corporation, NY, Grand Island, USA), 10%
fetal bovine serum (FBS, F1051, Sigma-Aldrich), 2 mM L-glutamine (25030149, Invitrogen
Corporation), and 15 (Xg mL"1 endothelial growth factor supplement (ECGS, 356006, BD
Biosciences, San Jose, CA, USA). HUVECs between passages 2 and 6 were used in all
experiments.
3.3.3 PET fibres and fibrin gel preparation
PET fibres were fixed on frames (Fig. 3.1a and 3.1b) to be precisely distanced from each
others (Fig. 3.1c). Frames bearing the fibres were cleaned in RBS (RBS Detergen 35, Pierce
Biotechnology, Rockfdrd, IL, cat 27952), sonicated for 15 min, rinsed with a flow of Milli-Q
gradient water (Millipore Canada, Nepean, Canada) with a resistivity of 18.2 Mflcm, and
finally blow-dried with 0.2-um filter-sterilized air. Frames bearing the fibres were then
immersed in 2 mL of sterile PBS containing 10% antibiotics (100 U/100 ug mL"'
penicillin/streptomycin) and exposed under a UV light for 30 min, as the final sterilization
step.
60
In 6-well plates, fibrin gels were prepared to be used as the attachment bench, using 1 mL/well
of fibrinogen solution (2.0 mg mU1) in HBSS and supplemented with 350 KIU mL1 of
aprotinin. This solution was mixed directly with 1 mL of a thrombin solution (1 U mL"1 in
HBSS) for the polymerization process of fibrinogen into fibrin (5 min at room temperature
followed by 10-20 min at 37°C and 5 % C02).
3.3.4 Cell adhesion assay
After the polymerization process, sterile frames bearing the fibres were transferred to the top
of this fibrin gel. Then, 100,000 cells were seeded directly over the frames bearing the fibres
for cell adhesion. Two (2) mL of Ml99 medium was finally added to each well to cover the
frames (see Fig. 3.Id).
Cell adhesion was investigated after 4 hrs of incubation, as described elsewhere [21-23]. To
count the number of cells attached on fibres surfaces, culture media were removed from the
wells and the frames bearing fibres were incubated 1 hr with Hoechst 33258 (cat. 23491-45-4,
Sigma-Aldrich) at a dilution of 1:10,000 in PBS containing 20% (wt/v) BSA.
Samples were observed under an inverted microscope (Eclipse TE 2000-S, Nikon Corp.,
Chiyoda-ku, Tokyo, Japan) using fluorescence and phase contrast imaging. Images were taken
with a 10X lens in conjunction with a digital CCD camera (Regita 1300R, QIMAGING,
Burnaby, Canada) and an imaging software (SimplePCI, Compix Inc., Minneapolis, MN). The
numbers of nuclei per fibre, corresponding to the number of attached cells, were counted
manually after 4 hrs and 48 hrs following cell adhesion.
3.3.5 Cell culture in 3D environment
After the adhesion phase, frames bearing fibres were transferred to a new gel with the same
composition as that described above, and then covered with a second layer of fibrin that
contained 50,000 HUVECs per milliliter suspended in heparin-free Ml99 medium. The
fibrinogen solution was allowed to clot for 5 min at room temperature and then, at 37°C and
5% CO2 for 10-20 min. After the polymerization process, 2 mL of Ml99 culture medium were
61
poured over it. Culture media were changed daily and the culture process was evaluated after 2
and 4 days (Figs. 3.1e and 3.If).
3.3.6 Visualization of microvessels
To observe endothelial cell behaviour, samples were observed daily under phase contrast
microscopy and images were recorded.
For cell staining, cell-seeded fibres were gently washed with PBS (3 times) and fixed in a
formaldehyde solution (3.75%, wt/v) in PBS for 20 min. Following three washes with PBS,
cells adhered on fibres were permeabilized with a Triton X-100 solution (0.5% v/v in PBS) for
5 min. Samples were finally rinsed three times in PBS, incubated 1 hr in a PBS solution
containing 20% (wt/v) BSA, and rinsed three times in PBS. Samples were incubated in a
solution containing a mixture of TRITC-phalloidin (1:300 dilution, cat. P1951, Sigma-
Aldrich) and Hoechst 33258 (1:10,000 dilution) made in a blocking buffer solution containing
BSA (20% (wt/v) in PBS) for 1 hr at room temperature, in the dark, and rinsed three times
with PBS and conserved in PBS.
In addition, cells were stained with Dil-Ac-LDL (BT-902, Biomedical Technologies Inc.,
Stoughton, MA, USA) after 2 days of culture to analyze the formation of tube-like structures
and to verify their phenotype.
The number of cell connections was manually quantified after 2 and 4 days of culture by
counting the numbers of sprouts and branch points. Sprout is an elongated structure where a
connection starts, whereas branch points were defined as areas where a single trunk structure
gave rise to two divergent outgrowths.
3.3.7 Images and image processing
Fibres were imaged with an objective of 10X and recorded as high-resolution files (*.tif) to
identify the effects of fibre-to-fibre distance over microvessels formation. An image processing
software (Scion Image; Scion, Frederick, MD) and Picasa2 were used to improve image
quality. These images were used to quantify the number of connections, which originated from
62
the newly formed tube-like connections along fibres axis. The numbers of connections per
fibre surface area as well as those per fibre length were reported.
3.3.8 Statistical analysis
The results were expressed as mean values + standard deviations for each group of samples (n
= 12), and after assessment of significant differences by one-way variance analysis
(ANOVA), the level of significance was considered at ap-value < 0.05 or < 0.01.
3.3.9 Confocal microscopy imaging of micro vessels
To analyze microvessels between adjacent fibres, frames bearing fibres embedded in fibrin
were gently washed with PBS (3 times) and fixed 20 min in a formaldehyde solution (3.75%,
wt/v, PBS). Following three washes with PBS, cells were permeabilized with a Triton X-100
solution (0.5% v/v in PBS) for 5 min. Samples were finally rinsed three times in PBS,
incubated 1 hr in a PBS solution containing 20% (wt/v) BSA, and rinsed three times in PBS.
Samples were incubated in a solution containing a mixture of TRITC-phalloidin (1:300
dilution) made in a blocking buffer solution containing BSA (20% (wt/v) in PBS) for 1 hr at
room temperature, in the dark, and rinsed three times with Milli-Q water. Finally, samples
were incubated with SYTOX Green Nucleic Acid Stain (1:20,000 dilution in Milli-Q water,
cat. S7020, Invitrogen) for 20 min and washed three times with Milli-Q water.
Images of microvessels were taken with a Confocal Laser Scanning Biological Microscope
(Olympus Fluoview FV300, Olympus Optical Co. Ltd. Tokyo, Japan) equipped with an
Olympus 1X70 camera and recorded as high-resolution and as layer-by-layer files. A complete
image reconstruction was made to illustrate microvessels in a 3D fashion. Images were edited
with the Image-Pro Plus Software to identify microvessels and lumen formation along fibres
axis.
63
3.4 Results
3.4.1 Cell adhesion on untreated PET fibres
To better promote cell adhesion on untreated PET fibres, frames bearing fibres were placed on
the top of a fibrin gel and HUVECs were then seeded directly over it (Fig. 3.Id). To evaluate
cell adhesion and cell spreading, fibre frames were transferred to a new tissue culture
poly(styrene) (TCPS) plate (Fig. 3.1e) in order to label cell nuclei. To induce the development
of microvessels, HUVEC-covered PET fibres were then sandwiched in fibrin gel; the lower
gel (i.e., that onto which fibre frames were deposited) was prepared without HUVECs, while
the upper fibrin gel (i.e., that covering fibre frames) was seeded with HUVECs (Fig. 3.If).
HUVECs began to attach to untreated PET fibres surfaces after 4 hrs, as indicated by arrows
in Figure 3.2a. The culture was continued for a period of 2 days in order to allow cells to
proliferate, migrate, and spread along the fibres curvature and mostly at the interface between
the fibre and the fibrin gel, as indicated by arrow heads in Figure 3.2b.
Cell density was compared 4 hrs (Fig. 3.2c) and 2 days (Fig. 3.2d) following the initial
incubation. To observe cell orientation and cell spreading along fibres surfaces following 2
days of culture, HUVECs were double-stained with TRITC-phalloidin for actin filaments and
with Hoechst 33258 for nuclei. The epifluorescence microscopy picture (Fig. 3.2e) revealed
that actin filaments were well elongated along fibres axis.
Cell adhesion on fibres is presented as the mean number of attached cells per fibre surface area
(number of cells/mm2) (Fig. 3.2f). The number of anchored cells per fibre surface area
increased as the fibre-to-fibre distance decreased as well as it increased with time. After 4 hrs
of incubation, the number of cells per fibre surface area was significantly (p<0.01) higher with
a fibre-to-fibre distance of 0.05 mm (179 ± 25 cells/mm ) when compared to fibre-to-fibre
distances of 0.1 mm (148 + 14 cells/mm2), 0.15 mm (99 ±11 cells/mm2), and 0.2 mm (68 ± 13
cells/mm2). However, following 2 days of culture, the mean numbers of anchored cells per
fibre surface area were on par with fibres distanced by 0.05 mm (473 ± 48 cells/mm2) and 0.1
mm (455 ± 6 1 cells/mm ), whereas the mean numbers of cells per surface area for fibres
64
distanced either by 0.15 mm or 0.2 mm were significantly (p<0.01) lowered with values of
257 ± 70 cells/mm2 and 132 + 24 cells/mm2, respectively.
3.4.2 Effect of fibre-to-fibre distance over cell connections and microvessels
formation
When HUVEC-covered fibres separated from each others by 0.05 mm were transferred to new
TCPS plates to be sandwiched in fibrin gel, within 4 hrs some HUVECs formed connection
bridges between adjacent fibres, while for the larger fibre-to-fibre distances tested here, cells
were more elongated along the fibres (Supplementary Figure 3.ii). HUVECs digested fibrin
along fibres. With fibres distanced by 0.1 mm, cell-to-cell connections attached to either one
or two fibres adjacent from each others were formed (Supplementary Figure 3.ii and Fig. 3.3).
When fibres were distanced by 0.15 or 0.2 mm, HUVECs needed more time to develop
connections; even if some connections were found after 2 days, the density was much lower
(Figs. 3.3c and 3.3d).
Phase contrast microscopy pictures of HUVECs labeled with Dil-acetylated-LDL showed a
good cell growth over the fibre curvature and the stability of the cells phenotype over time
(Supplementary Figure 3.ii). Also, testing with Factor VIII at day 2 gives a positive labeling
for endothelial cells (Supplementary Figure 3.ii).
The mean numbers of cell connections between fibres distanced either by 0.05, 0.1, 0.15, or
0.2 mm were compared for periods of 2 and 4 days (Fig. 3.3 and Table 3.1). The mean number
of cell connections found between fibres distanced by 0.1 mm over 2 days (Fig. 3.3c) was
larger than those of fibres distanced either by 0.05, 0.15, or 0.2 mm. A similar trend was
observed after 4 days (Fig. 3.3d).
Cell connections along and in between fibres as well as cell numbers per fibre surface area are
presented in Table 3.1 as mean values + standard deviations. The numbers of connections per
fibre length (number of connections/mm of fibre) after 2 days of culture were on par for fibres
distanced by 0.05 mm (6.4 connections/mm) and 0.1 mm (7.0 connections/mm), whereas it
significantly (p<0.01) decreased with fibres distanced by 0.15 mm (2.6 connections/mm) and
65
0.2 mm (0.9 connections/mm). Following 4 days, the number of connections per fibre length
for fibres distanced by 0.1 mm (12.1 connections/mm) was significantly (p<0.01) larger than
those of fibres distanced either by 0.05 mm (8.1 connections/mm), 0.15 mm (5.6
connections/mm), or 0.2 mm (2.1 connections/mm) (Table 3.1).
The number of connections per fibre surface area (number of connections/mm2) after 2 and 4
days of culture was significantly (p<0.01) larger for the 0.1 mm fibre-to-fibre distance than
those of fibre-to-fibre distances of 0.05, 0.15 and 0.2 mm (Table 3.1).
The number of cells per fibre surface area increased when fibre-to-fibre distance decreased as
well as when the culture period increased. After 2 days of culture, the number of cells per fibre
surface area for fibres separated by 0.05 mm (338 cells/mm ) was significantly (p<0.01) larger
than those of fibres distanced by 0.1 mm (261 cells/mmz), 0.15 mm (167 cells/mm ), and 0.2
mm (108 cells/mm2). After 4 days, the number of cells per fibre length was also significantly
(p<0.05) larger for a fibre-to-fibre distance of 0.05 mm (460 cells/mm2), when compared to
those of 0.2 mm (133 cells/mm2) and 0.1 mm (332 cells/mm2), which the latter was on par
with that of 0.15 mm (321 cells/mm2) (Table 3.1).
3.4.3 Microvessels formation
The confirmation of fully formed microvessels was verified by confocal microscopy for fibres
distanced by 0.1 mm, from layer-by-layer confocal pictures (Figs. 3.4a-3.4e) and from the
complete image reconstruction. The complete image reconstruction (Fig. 3.4f) reveals the
existence of single lumen in between adjacent fibres. The higher magnification image (Fig.
3.4g) more clearly shows the presence of a lumen inside the network of cell connections
(between upper and lower connections). The picture of microvessels taken using an inverted
microscope is also shown, as comparison (Fig. 3.4h).
In this study, we have successfully imaged microvessels with confocal microscopy without the
need to section the gel containing the cells and the fibres. In fact, histological analyses
following embedding and sectioning were not possible because the fibres delaminated during
sectioning, resulting in microvessels damage (data not shown).
66
3.5 Discussion
Although cell adhesion on PET surface is limited or even negligible [21,24] since the surface
is moderately wettable (contact angle of approximately 75 degrees) [25], some procedures
have been successfully developed to improve PET fibres biocompatibility [21]. The use of a
fibrin gel to promote the ability of HUVECs to attach, proliferate, migrate, and spread along
PET fibres surface has been successfully demonstrated in the present study. Our findings are
in good agreement with those of Khang et al.[18] that tested fibroblasts adhesion on PET
fibres surfaces. Also, Hadjizadeh et al.[21] and Mirenghi et al.[24] show that it was possible to
enhance human endothelial cell adhesion on plasma-modified and RGD-covered PET
surfaces.
Comparing to a previous study by our group in which PET fibres were surface modified by
plasma polymerization, the number of attached cells per fibre surface area obtained in the
present study was larger. The better cell adhesion on untreated PET fibres observed with the
method described here, as compared to other studies, can be explained in the following ways:
• The fibrin gel can stimulate cell responses because it contains several binding motifs
such as the RGD binding peptides [21-23], and fibrin plasma proteins that interact with
fibronectin to serve as a provisional extracellular matrix (ECM) for cell adhesion
[26,27].
• Fibres offer rigid interfaces to the cells, and as cells generally prefer rigid surfaces than
soft ones, they will migrate to that interface [28-30].
• The surface free energy provided by the bottom fibrin gel favors cell adhesion [31].
Angiogenesis is referred to as the growth of new microvessels from pre-existing blood vessels.
During angiogenesis, endothelial cells orient and organize themselves into tube-like structures
with the presence of lumens, allowing connections of neighboring cells to subsequently form a
network of tubes and initiate new blood vessels [7,14,32]. The angiogenesis process can be
divided into four stages, which conduct to the formation of microvessels [31]: stage 1, cells
digest their surrounding matrix; stage 2, committed cells proliferate, migrate, and
communicate with each others in order to form tube-like connections; stage 3, sprouting and
lumen formation; stage 4, development of a microvessel.
67
In our system, it can be hypothesized that microvessels formation occurred in the following
steps. Early after HUVEC-covered fibres were embedded in the fibrin gel, spread cells
detached from some areas of the fibres and migrate to the fibrin gel layer, while other
detached cells died. Within 4 hrs after the cell-covered fibres were sandwiched between fibrin
gels, it is hypothesized that cells started to digest fibrin along fibres axis. Afterwards, cells
were able to proliferate and migrate along fibres. Carrying the culture up to 24 hrs, cells on
fibres and those inside fibrin gel interacted and built cell connections. During this phase, cell
cytoskeleton shape became more flatten and elongated to form tube-like structures. The
development of tube-like cell connections was followed by the sprouting and branching of cell
connections until 2 days of culture to form a lumen. The development of tube-like structures
and lumen formation may be associated with a balance between cell migration activity and cell
death, as also reported by other researchers [33,34].
When the culture process was carried out up to 4 days, cells proliferated and migrated even
more, as indicated by the increasing numbers of cell connections and cells per fibre surface
area. Finally, cells formed one tube-like structure composed of multiple cells bridged between
two adjacent fibres, as shown by confocal microscopy pictures (Figs. 3.4f-3.4g). This finding
suggests that the direction of microvessels formation can be guided by the physical
interactions involved between cells and fibres to form a multi-cellular wall, allowing lumen
formation. These microvessels that form between adjacent fibres distanced by 0.1 mm were
associated by morphological events of the differentiation of endothelial cells into tube-like
structures, to subsequently create mature lumen, as also reported by others [14,34,35]. A
complete schematic illustration of the proposed microvascularization development in the
sandwich fibrin gel system containing PET fibres is shown in Figure 3.5 [36].
The fibre-to-fibre distance is an important parameter affecting the stimulation of cell adhesion
as well as microvessels formation. In the adhesion phase, as compared to the larger distances
tested here, smaller fibre-to-fibre distances give more possibility for HUVECs to sail over the
gel to better attach to the rigid fibres surfaces. In this study, endothelial cells were placed at
the interface between the rigid PET fibres and the soft fibrin gel. HUVECs tend to migrate and
68
attach to the fibres, in particular, integrin cytoplasmic regions of the cells will convert
mechanical forces into biochemical signaling in a process called durotaxis [28,29].
When HUVEC-covered fibres separated by smaller distances (i.e., 0.05 and 0.1 mm) were
sandwiched between two fibrin gels, within 4 hrs cells migrated to form bridges between the
fibres, stabilizing their cytoskeleton, and subsequently building connection strands. In the
larger fibre-to-fibre distances tested in this study (i.e., 0.15 and 0.2 mm), cells required more
time to form and stabilize their connection strands. The numbers of attached cells on the fibres
as well as that of cell connections play an important role in the development of lumen and
micro vessels, as also reported by others [35].
3.6 Conclusions
This study reported the development and validation of a cell adhesion method that uses fibrin
gel to increase HUVECs adhesion and spreading on untreated PET fibres. When fibres pre-
covered with HUVECs were sandwiched between two fibrin gels, microvessels formation
occurred along the fibres. The fibre-to-fibre distance was found to have a significant role in
promoting cell adhesion as well as cell connections between adjacent fibres, thus guiding
microvessels formation along fibres in a 3-dimensional environment. Microvessels formed
large tube-like structures (diameter of approximately 100 urn) between adjacent fibres
distanced by 0.1 mm, after 4 days of culture. When fibres were separated by 0.05 mm, the
distance was too small, so that connections lacked orientation and failed to form a large tube
like microvessel. In the larger fibre-to-fibre distances (i.e., 0.15 and 0.2 mm), cell connection
strands were rare, so microvessels did not form between adjacent fibres. The requirements of
this culture process were dictated by the possibility to scale the system to bioreactor cultures,
which is the next step in our research program. This proof-of-concept study opens the door to
other experiments in which we plan to study by using this culture process the effects of cell
number, fibre .diameter, ECM density and type, cell types, and the presence of angiogenic
growth factors on microvessels formation. Also, the functionality of these microvessels will be
investigated.
69
This system can be applied to culture vascularized tissues such as in the development of a
bioartificial pancreas device. These well oriented microvessels can serve to supply nutrients to
growing cells. If the fibres need to be removed from the constructed tissue before use or
implantation, PET fibres can be replaced by biodegradable fibres.
Acknowledgements
This work was supported by the Canadian Foundation for Innovation through an On-going
New Opportunities Fund (project no.7918), the National Sciences and Engineering Research
Council of Canada (NSERC), and by the Universite de Sherbrooke. Mr. Sukmana was
supported by the IDB Merit Scholarship Program. The authors are also grateful to Yannick
Laplante, Mark G. Couture, Gilles Grondin, and Aftab Ahamed for their technical support.
70
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73
Table 3.1: Effects of fibre-to-fibre distance on cells.
Parameters
Number of cell connections per fibre length (connections/mm) Number of cell connections per fibre surface area (connections/mm2) Number of cells per fibre surface area (cells/mm2)
Culture time
(days)
2
4
2
4
2
4
Fibre-to-fibre distances (mm)
0.05
6.4a
±0.6 8.1b
±2.3 23.8" ±2.1 25.8b
±3.1 338a
±26 460a
±157
0.1
7.0a
±1.8 12.1a
±1.5 28.2a
±7.3 42.8a
±11.3 261" ±18 332b
±52
0.15
2.6b
±0.7 5.6C
±1.7 9.1c
±3.7 14.5C
±2.6 167c
±31 321b
±60
0.2
0.9C
±0.3 2.1d
±0.2 2.4d
±1.1 4.1d
±1.3 108d
±32 133c
±58
CD value
p<0.05
1.41
1.91
5.84
6.78
40.08
125.67
p<0.01
1.99
2.69
8.25
9.57
56.56
177.34
*Means in a row followed by different letters are significantly different.
74
/ P5T
W
Seeding HUVECs
Fibrin gel
Tissue culture poly(styrene) (TCPS) plate
PET fibre HUVEC-covered PET fibre
Fibrin gel + HUVECs
Frame
HUVEC-covered PET fibre
Figure 3.1: Schematic illustration of the in vitro method designed to guide and orient
microvessel formation.
Polycarbonate frames were produced by micro-machining (a and b) to maintain fibres parallel from each others by well defined distances (c). A layer of fibrin gel was prepared to serve as a support for endothelial cells (i.e., HUVECs) adhesion. Polycarbonate frames bearing PET fibres were placed over the gel and HUVECs were seeded directly over fibres and M199 culture medium was added to cover the fibres (d). The culture period was allowed to proceed until HUVECs attached and covered fibres surfaces. When cells reached confluence, frames bearing fibres were transferred to a new plate for further culture (e). PET fibres covered with HUVECs were sandwiched between two fibrin gels and culture medium was added (f); only the upper gel contained HUVECs.
75
N ^
J
0.05 0.1 0.15 0.2 Fibre-to-fibre distances (mm)
Figure 3.2: HUVECs attachment on PET fibres.
HUVECs attached to fibres after 4 hrs (Step Id in Figure 3.1), as shown by arrows (a). PET fibres were used to guide HUVECs proliferation along fibres axis until 48 hrs, as shown by arrowheads (b). To count the number of attached cells, nuclei were stained with Hoechst after 4 hrs (c) and 48 hrs (d). Actin filaments and nuclei were stained with TRITC-phalloidin and Hoechst, respectively (e). The number of attached cells on fibres surfaces increased as the fibre-to-fibre distance decreased and as the time progressed (f). Scale bars correspond to 100 urn
76
Fibre-to-fibre distances
0.05mm
2 days 4 days
" :* fibre
0.1mm
0.15mm
0.2mm
Figure 3.3: Effect of fibre-to-fibre distance over cell connections and micro vessel
development.
HUVECs built cell connections, as observed by phase contrast microscopy by day 2 (left-hand column) as well as by day 4 (right-hand column). Actin filaments (red) and nuclei (blue) were stained with TRITC-phalloidin and Hoechst, respectively. Fibres were distanced by 0.05 mm (a and b), 0.1 mm (c and d), 0.15 mm (e and f), and 0.2 mm (g and h). Scale bars correspond to 100 |am.
77
•• Microvessel formation between fibres. •*• (A) Upper connection. •*• (B) Lower connection. •*• (C) Lumen formation between
upper and lower connections.
Figure 3.4: Confocal and inverted microscopy pictures.
After 4 days of culture, cells were stained for actin filaments, using TRITC-phalloidin, and counterstained with SYTOX Green to highlight the position of the nuclei during confocal analyses. Confocal pictures were recorded layer-by-layer from the top to the bottom of samples (a-e). Complete image reconstructions are presented in (f) and (g) for a higher magnification of the lumen. A picture of microvessels taken using an inverted microscope is also shown, as comparison (h). Scale bars correspond to 100 urn.
78
\5
(
2nd gel S£\ \ +HUVECs ^ \ ~"
/ Bottom gel
HUVECs
HUVECs attachment, migration and elongation on fibres. y view.
HUVEC-eovered PET fibres are sandwiched between fibrin gel. During polymerization process of the fibrin gel, new HUVECs positioned themselves in all levels of the gel down to the first bottom gel.
Fibre \
Fibre 1 ja—r-JJSSsffi" I "*i2
PET fibres
100 um
HUVECs start to make connections with each others between adjacent fibres. Cells become more flatten and elongated and form tube-like structures. y view.
Cell connections and lumen formation between 2 fibres. x view.
Branching, sprouting. y view.
Microvessels formed between adjacent fibres. y view.
Figure 3.5: Schematic illustration of the microvascularization process.
Adapted from Lutolf et al. [36].
79
SSI' 4 E ''••
3 TJHdB*
Supplementary Figure 3.i: Scanning electron microscopy (SEM) image of a PET fibre.
80
Supplementary Figure 3.ii: Progression of cell connections and microvessel reorganization
along and in between adjacent fibres sandwiched between two fibrin gels.
When fibres were distanced by 0.5mm, HUVECs started to communicate with each others and were able to develop a connection bridge between two adjacent fibres after 4 hrs, as shown by arrows (a). At 0.1 mm fibre-to-fibre distance, cell connections were formed between fibres axis after 24hrs, as shown by discontinued arrow (b). Whereas in larger fibre-to-fibre distances (0.15 and 0.2 mm), connections were rare and discontinued (c and d). Phase contrast microscopy revealed a positive phenotype for endothelial cells (Dil-acetylated LDL uptake), which were distributed along fibres (e). Immuno-labelling with Factor VIII antibody also underlined the positive phenotype of endothelial cells (f). Scale bars correspond to lOOjtm.
81
CHAPTER 4
The Effects of Co-culture with Fibroblasts and
Angiogenic Growth Factors on Microvascular
Maturation and Multi-cellular Lumen Formation
in HUVEC-oriented Polymer Fibre Constructs
Irza Sukmana1'2, Patrick Vermette1'2
1. Laboratoire de Bioingenierie et de Biophysique de l'Universite de Sherbrooke,
Department of Chemical and Biotechnological Engineering, Universite de Sherbrooke,
2500, blvd de l'Universite, Sherbrooke, QC, Canada, J1K 2R1.
2. Research Centre on Aging, Institut universitaire de geriatrie de Sherbrooke, 1036, rue
Belvedere Sud, Sherbrooke, QC, Canada, J1H 4C4.
Article published in Biomaterials 31 (2010), p. 5100-5109.
DOI: 10.1016/j.biomaterials.2010.02.076
http://www.sciencedirect.com/science/iournal/01429612
82
4.1 Abstract
In the present study, polymer monofilaments were embedded in fibrin seeded with human
umbilical vein endothelial cells (HUVEC) to guide HUVEC attachment and migration in order
to form oriented vessel-like structures between adjacent monofilaments. Histology and
fluorescent fibrin experiments confirmed that microvessel-like structures, which were
developing between polymer monofilaments embedded in fibrin, contained a lumen. The
effect of human fibroblasts and growth factors (VEGF and bFGF) over the microvessel
formation process was tested. The effects of VEGF and bFGF were dose dependent. The effect
of VEGF was optimum at the lower concentration tested (2ng/mL), while that of bFGF was
optimum at the higher tested concentration (20ng/mL). Furthermore, the use of fibroblasts
significantly improved the maturation of the microvessels compared to control and to samples
cultured with VEGF and bFGF.
4.2 Introduction
Microvascularization is a process in which endothelial cells form microvessels. It is involved
in the progression of cancerous cell masses [1-3]. Also, functional microvessels are necessary
to supply nutrients and remove wastes in cell and/or tissue aggregates in order to improve
tissue substitute survival and function [3,4], therefore justifying the development and
application of methods to support microvessel formation.
Angiogenesis is a multistage process involving the sprouting of blood vessels from pre
existing vasculature [2,4]. Briefly, endothelial cells degrade the extracellular matrix followed
by cell proliferation, migration, and cell-cell interactions leading to the formation of tube-like
structures, and eventually to sprouting and lumen formation with adjacent vessels [4,5].
Several in vitro and in vivo angiogenesis assays have been developed to study and follow the
development of sprouts and lumens and to validate the efficacy of both pro- and anti-
angiogenic therapeutics [5,6]. In vitro assays include endothelial cells grown in different gels
[6,7], microcarrier bead assays [8,9], and aortic rings embedded in gels [10-12], only to name
a few. In vivo assays include the chick chorioallantoic membrane (CAM) [13], the Matrigel
implant [14], and the retinal angiogenesis assay [15].
83
A number of reports have shown that pericytes [16] and smooth muscle cells [3] can maintain
the patent tubular structure of blood vessels, and at the same time provide important factors to
improve the maturation of microvessels [16,17]. Fibroblasts have been found to improve
capillary sprout growth and stabilization in collagen and fibrin gels [16-18]. At some stage of
the angiogenesis process, smooth muscle cells inhibit endothelial cell proliferation and new
vessel formation, implying that mural cells interact with endothelial cells to stabilize newly
formed capillaries [19].
On a different note, other studies reported that angiogenic growth factors, such as vascular
endothelial growth factor (VEGF) [17,20] and basic fibroblast growth factor (bFGF) are
essential for the induction of angiogenesis [21,22]. However, there are some contradictory
data on how VEGF and bFGF concentrations influence cellular responses [12,17].
Although some aspects of angiogenesis, including endothelial cell migration, proliferation and
phenotype differentiation seem to be established among the scientific community, many
studies fail to model the maturation of microvessels up to the stage of lumen formation and
even just to prove the existence of such multi-cellular lumen [17]. In addition, often
microvessel networks are randomly distributed in a dispersing gel environment, and lack any
patterning and specific orientation [22,23]. In tissue engineering applications and cancer
research, the modeling and study of directional sprouting and microvessel formation is
important [23,24]. Firstly, we believe that by orienting the development of microvessels,
induction of fluid flow within the lumen of these microvessels will be facilitated [24].
Secondly, having a system in which microvessels can be patterned opens the door to new in
vitro models of metastasis progression in cancer research [23-26] to better understand how
biosignalling localization affects microvessel development in a 3D environment.
For these reasons, here we have further optimized and validated our in vitro model of
microvessel patterning using polymer monofilaments embedded in fibrin. In a previous study,
we have reported the development of an angiogenesis assay in which microvascularization
process was modulated in a directional fashion [24]. It was demonstrated that polymer fibres
deposited on top of a fibrin gel allowed cell attachment on these monofilaments. Furthermore,
84
when HUVEC-covered fibres were subsequently sandwiched in fibrin, cells established
connections with each others, thus leading to the formation of one microvessel between two
adjacent fibres. Using this culture process, the aims of the present study were to investigate the
effect of human fibroblasts and two angiogenic growth factors (i.e., vascular endothelial
growth factor (VEGF) and basic fibroblast growth factor (bFGF)) on microvessel maturation
and to assess lumen formation.
4.3 Materials and Methods
4.3.1 Materials
100-um diameter poly(ethylene terephthalate) monofilaments (PET, ES 305910, Good Fellow,
Devon, USA) were fixed onto polycarbonate (Boedeker Plastics Inc., Texas, USA) frames that
fitted into traditional 6-well plates used in cell culture. The frames were 22.5 mm in diameter
and 2.0 mm thick [24].
The phosphate buffered saline solution (PBS IX, pH 7.4) used in these experiments was
prepared from Milli-Q water and NaCl (0.137 M), KC1 (0.003 M), Na2HP04 (0.008 M), and
KH2P04 (0.002 M). This 150 mM solution was diluted in Milli-Q water to make a 10 mM
solution. Hank's balances salt solution (HBSS, H1387) and albumin from bovine serum (BSA,
CAS 9048-46-8) were purchased from Sigma-Aldrich Inc. (ON, Canada).
Vascular endothelial growth factor (VEGF) used here is a human recombinant VEGFi65
with a 38.2-kDa disulfide-linked homodimeric protein consisting of two 165 amino acid
polypeptide chains. Basic fibroblast growth factor (bFGF) is a human recombinant with a
17.2-kDa protein consisting of 154 amino acid residues. VEGF165 (cat.# 100-20) as well as
bFGF (cat.# 100-18B) were purchased from PeproTech (PeproTech inc., NJ, USA).
4.3.2 Cell culture
Human umbilical vein endothelial cells (HUVEC) were purchased from PromoCell (C-12200,
Heidelberg, Germany). HUVEC were cultured at 37°C and 5% C02 in Ml99 culture medium
(M5017, Sigma-Aldrich) supplemented with 2.2 mg mL"1 sodium bicarbonate (Fisher, Fair
85
Lawn, USA), 90 ug ml/1 sodium heparin (H1027, Sigma-Aldrich), 100 U/100 ug mL"1
penicillin/streptomycin (15140-122, Invitrogen Corporation, Grand Island, NY, USA), 10%
foetal bovine serum (FBS, F1051, Sigma-Aldrich), 2 mM L-glutamine (25030149, Invitrogen
Corporation), and 15 ug mL"1 endothelial growth factor supplement (ECGS, 356006, BD
Biosciences, San Jose, CA, USA). HUVEC between passages 2 and 6 were used in all
experiments. Skin fibroblasts extracted from human foreskin and purchased from PromoCell
(C-12350, Heidelberg, Germany) were cultured in Ml99 supplemented with 10% FBS and
100 U/100 ug mL"1 penicillin/streptomycin at 37°C and 5% CO2. Fibroblasts between
passages 6 and 15 were used in all experiments.
4.3.3 Culture system and fibrin preparation
PET fibres were fixed on polycarbonate frames to be precisely distanced from each other [24].
Frames bearing fibres were cleaned in RBS detergent 35 (cat 27952, Pierce Biotechnology,
Rockford, IL), sonicated for 15 min, rinsed with a flow of Milli-Q gradient water (Millipore
Canada, Nepean, Canada) with a resistivity of 18.2 Mii-cm, and finally blow-dried with 0.2-
um filter-sterilized air. Frames bearing fibres were then immersed in 2 mL of sterile PBS
containing 10% antibiotics (100 U/100 ug mL"1 penicillin/streptomycin) and exposed under
UV light for 30 min, as the final sterilization step.
In 6-well plates, fibrin gels were prepared to be used as the attachment bench using 1 mL/well
of fibrinogen solution (4.0 mg mL"1) made in HBSS and supplemented with 350 KIU mL"1 of
aprotinin. This solution was directly mixed with 1 mL of a thrombin solution (2 U mL"1 in
HBSS) for the polymerization process of fibrinogen into fibrin, 5 min at room temperature
followed by 20 min at 37°C and 5 % C02 .
4.3.4 Cell adhesion and angiogenesis assay
After the polymerization process, sterile frames bearing the fibres were transferred and
deposited on the top of a fibrin gel prepared as described in Section 2.3. Then, 100,000
HUVEC were directly seeded over it and cultures were run for 2 days to allow cell attachment.
Two (2) mL of supplemented Ml99 medium were finally added to each well to cover the
frames [24].
86
After the attachment phase, frames bearing fibres were transferred to a new fibrin gel with the
same composition as described above, and then covered with a second layer of fibrin that
contained 100,000 HUVEC per milliliter suspended in heparin-free Ml99 medium. The
fibrinogen solution was allowed to clot for 5 min at room temperature and then, at 37°C and
5% CO2 for 30 min. After the polymerization process, 2 mL of M199 culture medium (with or
without growth factors) were poured over it. Culture media were changed daily and the culture
process was evaluated after 2 and 4 days. For the co-culture experiment, 300,000 of human
foreskin fibroblasts were mixed directly with Ml99 culture media and added on the top of the
fibrin. In another set of experiments, HUVEC were fed with M199 supplemented with either
VEGF or bFGF (2, 5 or 20 ng/mL) with no fibroblasts.
4.3.5 Visualization of microvessels and lumen formation
To observe endothelial cell behaviour, pictures were daily taken with a phase contrast
microscope. After 2 and 4 days, the fibrin gels containing the frames were fixed for further
examination. For cell staining, cell-seeded fibres were gently washed with PBS (3 times) and
fixed in a formaldehyde solution (3.75%, wt/v) in PBS for 20 min. Following three washes
with PBS, cells adhered on fibres were permeabilized with a Triton X-100 solution (0.5% v/v
in PBS) for 15 min. Samples were finally rinsed three times in PBS, incubated 1 h in a PBS
solution containing 2% (wt/v) BSA, and rinsed three more times in PBS. Samples were
incubated in a solution containing a mixture of rhodamine-phalloidin (1:300 dilution, R415,
Molecular Probes, Eugene, OR) for F-actin labelling and Hoechst 33258 (1:10,000 dilution,
cat. # B2883, Sigma-Aldrich) for nuclei staining, made in a blocking buffer solution
containing BSA (2% (wt/v) in PBS) for 1 h at room temperature, in the dark. Finally, after
three washes with PBS, samples were conserved with 3 mL per well of PBS.
Samples were imaged and recorded as high-resolution files (*.tif). To improve quality, images
were then processed with Image-Pro-Plus Software. The number of cell-cell connections was
manually quantified after 2 and 4 days of culture by counting the numbers of sprouts and
branch points. Sprout is an elongated structure where a connection starts, whereas branch
87
points are single trunk structures that give rise to two divergent outgrowths. The numbers of
cells as well as cell-cell connections per fibre length are reported.
To assess the formation of multi-cellular lumen, human fibrinogen conjugated with Alexa
Fluor 546 (F-13192, Molecular Probes, Eugene, OR) was combined with unlabelled human
fibrinogen at a mass ratio of 1:10 (conjugated to unconjugated fibrinogen) to produce the fibrin
gel, as in Section 2.3, in which the HUVEC-covered fibres were sandwiched. In this
experiment, HUVEC were labelled with the CellTracer™ CFSE staining (C34554, Molecular
Probes, Eugene, OR) for 30 min prior to the experiment. The fibrin degradation by HUVEC
could then be examined using confocal microscopy.
In another series of experiments, fibres covered with unlabelled HUVEC were sandwiched
between fluorescent fibrin (see the above paragraph for composition) and after polymerization,
fibroblasts were subsequently added on the top of the fibrin gel. The cultures were carried out
for a period of 4 days. Following fixation and permeabilization, as described above, cells were
stained with Alexa Fluor 488 green phalloidin (1:300 dilution, A12379, Molecular Probes,
Eugene, OR) for F-actin and Hoechst for nuclei. Layer-by-layer images were taken with a
Confocal Laser Scanning Biological Microscope (Olympus Fluoview FV1000, Olympus
Optical Co. Ltd., Tokyo, Japan).
4.3.6 Histological section
Fibrin gels containing fibre holders were fixed overnight in 4% neutral buffered formalin.
Fibres embedded in fibrin were gently removed from their frame, and processed for paraffin
sections according to standard protocols [27]. Six-um-thick sections were then prepared for
hematoxylin and eosin staining (both from Sigma-Aldrich). Pictures were taken with the
bright field mode of the Biolmaging Navigator fluorescence microscope (Olympus FSX100,
Olympus Optical Co. Ltd., Tokyo, Japan).
4.3.7 Imaging microvessels by confocal microscopy
To investigate microvessel formation between adjacent fibres, frames bearing fibres embedded
in fibrin were gently washed with PBS (3 times) and fixed 20 min in a formaldehyde solution
88
(3.75%, wt/v, PBS). Following three washes with PBS, cells were permeabilized with a Triton
X-100 solution (0.5% v/v in PBS) for 5 min. Samples were rinsed three times in PBS,
incubated 1 h in a PBS solution containing 2% (wt/v) BSA, and rinsed three more times in
PBS. Samples were then incubated in a solution of rhodamine-phalloidin red (1:300 dilution)
made from a blocking buffer containing BSA (2% (wt/v) in PBS) for 1 h at room temperature,
in the dark, and rinsed three times with PBS. Finally, samples were incubated with SYTOX
Green Nucleic Acid Stain (cat. S7020, 1:10,000 dilution in Milli-Q water, Invitrogen) for 20
min and washed three times with PBS.
Images of microvessels were taken with a Confocal Laser Scanning Biological Microscope
(Olympus Fluoview FV1000, Olympus Optical Co. Ltd., Tokyo, Japan) equipped with an
Olympus 1X70 camera and recorded as high-resolution layer-by-layer files. Images were edited
with Olympus FV10-ASW 2.0 viewer and a complete image reconstruction was made to
provide a 3D view of the microvessels.
4.3.8 Statistical analysis
The results are expressed as mean values ± standard deviations for each group of samples.
Data were statistically analyzed by using the analysis of variance (ANOVA).
4.4 Results and Discussion
This three-dimensional cell culture system allows to precisely controlling the fibre-to-fibre
distance to 100|im [24]. When HUVEC-covered fibres were embedded in fibrin also
containing HUVEC, cells oriented and elongated along the fibre axis at the fibre-gel interface,
allowing the patterning of microvessels. When using HUVEC-free monofilaments embedded
in fibrin containing HUVEC, neither tube-like structures nor cell-cell connections were
observed (Supplementary Figure 4.i). The step-by-step process of microvessel development
occurring in the culture system described here has been presented elsewhere [24].
Compared to our previous study in which lower HUVEC concentration was used in the fibrin
gel [24], the numbers of cells as well as cell-cell connections per fibre length after 2 and 4
days of culture found in this present study were higher. In the present study, the cell density
89
was increased, as we found that it plays a significance role in the microvessel development.
This is in agreement with another study revealing that when cell number was smaller than a
certain threshold value, the formation of tube-like cell connections, sprouting and branching
were not observed [28].
After sandwiching the HUVEC-covered fibres in fibrin and adding fibroblasts, some cells that
cover the fibres migrate along the fibre axis and some into the fibrin (Fig. 4.1a) in order to
interact with the cells located inside the gel, finally forming cell-cell connections after 1 day of
culture. During this phase, cell cytoskeleton shape became more flatten and elongated to form
tube-like structures (discontinued arrows, Fig. 4.1b) and even some cells were able to bridge
between two fibres to build a cell connection (thick white arrows, Fig. 4.1b). When HUVEC
were cultured in the absence of fibroblasts, many HUVEC migrated away from the fibres and
failed to build cell-cell connections, and even though connections were formed after 2 days
(Fig. 4.1c), the number of connections was lower when compared to the system in which
fibroblasts were added (Fig. 4.Id).
Lower magnification image of samples with fibroblasts reveals the tube-like structures seen
between adjacent fibres, almost completely covering the area between fibres after 4 days (Fig.
4.1e). Also, a complex tube-like structure network occurred between adjacent fibres (black
and white arrows, Figs. 4.If and 4.1g).
When HUVEC-covered fibres were co-cultured with fibroblasts for a period of 4 days,
confocal microscopy pictures (Figs. 4.2a-4.2e) confirmed the better maturation of the
microvessels, compared to control samples (Supplementary Figure 4.ii). A lower magnification
of the reconstructed confocal image (Fig. 4.2f) as well as a higher magnification image (Fig.
4.2g) also revealed that the fibrin between adjacent fibres, where cells formed connections,
was degraded over time. In these experiments (Figs. 4.2f and 4.2g), fibrin was prepared using
fibrinogen conjugated with a red fluorescent probe mixed with regular fibrinogen. HUVEC
were labelled with CFSE CellTracer™ (green).
90
The effect of VEGF and bFGF over the formation of cell-cell connections and micro vessel
development was dose-dependent (Table 4.1). Numbers of cell-cell connections between
fibres as well as cell numbers per fibre length are listed in Table 4.1. The mean numbers of
cell-cell connections between fibres as well as cell numbers per fibre length between controls
and systems with either VEGF or bFGF at three concentrations (i.e. 2, 5 and 20ng/mL) were
compared for a culture period of 2 days (Figs. 4.3a-4.3d and Table 4.1). The effect of VEGF
was stronger at 2 and 5ng/mL (Fig. 4.3a), when compared to the highest concentration tested
here (20ng/mL, Fig. 4.3b) and to the control system with no added growth factors. The
addition of bFGF yielded higher numbers of cell-cell connections and cells per fibre length at
a concentration of 20ng/mL (Fig. 4.3d), when compared to those with bFGF concentrations of
2 or 5ng/mL (Fig. 4.3c) and control.
Our results (Fig. 4.3 and Table 4.1) showed that VEGF and bFGF supported cell proliferation
and migration, thus enhancing the formation of cell-cell connections in a dose-dependent
manner. This finding is in good agreement with other studies [17,22,29-31]. The use of VEGF
at a concentration of 20ng/mL resulted in lower cell numbers and cell-cell connections
compared to control cultures. This phenomenon is referred to as the biphasic effect of VEGF,
as also reported by others [30,32].
Following 2 and 4 days of culture, the number of cell-cell connections per fibre length
(connections/mm of fibre) was statistically significantly higher when fibroblasts were added
compared to those of control and samples with bFGF (20ng/mL) and VEGF (2ng/mL). Similar
findings were found when the numbers of cells per fibre length (cells/mm of fibre) were
compared for the different samples (Table 4.2).
Cell-cell connections and evidence of the degradation of fibrin are presented in Figure 4.4.
Examining the layer-by-layer confocal pictures, multiple cell-cell connections were observed
(see left sides of Figs. 4.4a-4.4d). At time zero, cells were rounded with a strong uniform
signal from the fluorescent fibrin (Fig. 4.4f). Cell-cell connections (discontinued lines) were
associated to areas where fibrin was degraded (stars, see right sides of Figs. 4.4a-4.4d). We
believe that these cell-cell connections are made of multiple cells resulting in lumen, as
91
reported by others [33,34]. The complete image reconstruction (Fig. 4.4e) also reveals that the
multi-cellular connections covered almost all the area in between the adjacent fibres.
Histological sections also confirmed that HUVEC were forming multi-cellular lumen (star,
Fig. 4.4g). Lumen were guided by the PET fibre (discontinued arrow, Fig. 4.4h) and made up
of multiple cells (arrows, Fig. 4.4i).
These findings reveal that fibroblasts enhance microvessel formation, development and
maturation. This is in agreement with several studies available in the scientific literature. For
example, fibroblasts were found to stabilize endothelial cell-lined tube formation [34], and
provide important signalling that lead to the maturation of capillaries made of HUVEC
[32,35]. Fibroblasts are known to release many growth factors including VEGF, FGF, platelet-
derived growth factor (PDGF), and angiopoietin-1 [12,16-18,35]. When comparing the effects
of VEGF and bFGF and fibroblasts over the formation of cell-cell connections and subsequent
microvessel maturation, our results show that fibroblasts can provide sufficient pro-angiogenic
factors to promote vessel-like formation (Figs. 4.2a-4.2e). The use of fibroblasts yielded
higher numbers of cells and cell-cell connections, when compared to experiments involving
VEGF or bFGF. However, there are still some debate on how VEGF and bFGF concentration
affects angiogenesis and microvascularization (see Table 4.3 for some examples of results
from the scientific community).
In samples with fibroblasts, they were deposited on top of the upper fibrin gel. In this way,
they were from a distance of approximately 2 to 3 mm from the endothelial cells and the
fibres. To ascertain that fibroblasts were not migrating into the fibrin and so would never come
in direct contact with HUVEC up to 4 days of culture, in some experiments fibroblasts were
labelled with the CFSE CellTracer to follow their localization (Supplementary Figure 4.iii).
These experiments reveal that fibroblasts remain on the top of the fibrin and thus the effect of
fibroblasts over microvessel development was not a result of a direct contact between
fibroblasts and HUVEC.
To carry out longer cultures (more than 4 days), we believe that the recruitment of fibroblasts
or pericytes at the site of microvessel formation will be necessary in order to stabilise and
92
completely close the walls of the microvessels formed between adjacent fibres, as also
postulated by some researchers [34,36-38]. For instance, many studies have demonstrated the
ability of pericytes, an immature cell type that can differentiate into fibroblasts, smooth
muscle cells or macrophages, to initiate vascular sprouts, to stabilize vessels and to support
vessel maturation [18,39,40].
4.5 Conclusions
The development of a functional micro-vasculature is a crucial factor for the survival as well
as the development and function of engineered vascularised tissue substitutes. Here, the effects
of fibroblasts and two growth factors (VEGF and bFGF) over microvessel development were
studied using a system in which human umbilical vein endothelial cells (HUVEC) are guided
by polymer monofilaments to form oriented vessel-like structures. VEGF and bFGF were
shown to enhance HUVEC proliferation and migration, thus improving the development of
cell-cell connections and subsequent microvessel development. When HUVEC-covered fibres
were co-cultured with fibroblasts, the HUVEC responses were more pronounced compared to
control and to cultures in which VEGF and bFGF were used. The use of fibroblasts resulted in
more mature microvessels containing more cell-cell connections. These microvessel-like
structures were also shown to contain a lumen. This culture process in which microvessels
with multi-cellular lumen are patterned can find applications in cancer and tissue engineering
studies.
Acknowledgements
This work was supported by the Canadian Foundation for Innovation/Ministere de 1'Education
du Quebec through an On-going New Opportunities Fund (project no.7918), the National
Sciences and Engineering Research Council of Canada (NSERC), and by the Universite de
Sherbrooke. Mr. Sukmana was supported by the IDB Merit Scholarship Program. The authors
are grateful to Johanne Dussault and Dr. Sameh Geha, for their help in histology, and to Gilles
Grondin and Dr. Leonid Volkov for their assistance in confocal microscopy analysis. Also, the
authors would like to thank Evan Dubiel and Georges Sabra for their critical review of the
manuscript.
93
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Table 4.1: Effects of VEGF and bFGF concentrations over the numbers of cells and cell-cell
connections following 2 days of culture.
Cell number and cell-cell connection*
Number of cell-cell connections per fibre length (connections/mm) Number of cells per fibre length (cells/mm)
Growth factor concentrations (ng/mL)
Control
6.6d
±0.9
52.6d
±7.7
VEGF
2
9.1a
±0.4
83.9a
±2.3
5
7.2" ±1.1
82.2a
±6.3
20
5.1e
±1.0
53.0 d
±12. 3
bFGF
2
7.0C
±1.1
68.8C
±4.4
5
8.1b
±0.9
78.6b
±5.8
20
9.8a
±1.0
85.8a
±5.4
CD value
p<0.05
0.95
7.07
p<0.01
1.43
10.60
*Means in a row followed by different letters are significantly different.
Table 4.2: Comparison of the effects of fibroblasts, VEGF and bFGF over the numbers of cells
and cell-cell connections.
Cell number and cell-cell connection*
Number of cell-cell connections per fibre length (connections/mm) Number of cells per fibre length (cells/mm)
Culture time
(days)
2
4
2
4
Parameters
Control
6.6C
±0.9 12.5C
±1.6 52.6b
±7.7 92.7C
±18.5
Fibroblasts
12.4a
±0.9 20.9a
±3.8 89.5a
±4.6 124.9a
±12.1
VEGF (2ng/mL)
9.2b
±0.4 13.2C
±2.2 83.9b
±2.3 89.5b
±5.7
bFGF (20ng/mL)
9.8b
±1.0 16.4b
±1.5 85.8b
±5.4 97.9" ±5.3
CD value
p<0.05
0.87
2.64
5.78
12.62
p<0.01
1.35
4.11
8.99
19.65
*Means in a row followed by different letters are significantly different.
98
Table 4.3: Review of some studies on the effects of VEGF and bFGF on endothelial cells and
angiogenesis.
Cell type/culture method
HUVEC cultured in fibrin and collagen gel containing growth factors and sphingosine-1-phosphate(SlP). EC were seeded on microcarrier beads and then embedded in fibrin containing growth factors.
Mouse aortic rings (contained EC and mural cells) were cultured in a collagen-based matrix.
HUVEC were cultured on microcarrier beads and embedded in fibrin, then covered with Ml99 containing growth factors.
EC cultured on microcarrier beads embedded in a serum-free fibrin with bFGF. Human umbilical artery endothelial cells were cultured in plastic wells pre-coated with growth factors. HUVEC were cultured on microcarrier beads, then embedded in fibrin gels and covered with Ml99 containing growth factors. Canine jugular vein EC (CJEC) cultured in fibrin glue then embedded in fibrin gel and covered with Ml99 containing FGF-1. Bovine aorta endothelial cells cultured in fibrin glue, then embedded in fibrin and covered with Ml99 containing 1 or 2 growth factors.
Growth factors
bFGF and VEGF (40ng/mL)
VEGF (lOOng/mL) and bFGF (30ng/mL) VEGF and bFGF (5, 20, and 50ng/mL)
VEGF and bFGF at several concentrations (l-35ng/mL) bFGF (30ng/mL)
VEGF165 and VEGF189 at 4ug/mL
VEGF12i and VEGF165 at several concentrations (l-35ng/mL) FGF-1 1, 10, 100 ng/mL
FGF-1 and VEGF at concentrations between 1 to lOOng/mL.
Results
S IP combined with either VEGF or FGF stimulated EC to invade gels and form tube-like structures and not in the absence of SIP. The use of either bFGF or VEGF stimulated capillary formation from EC.
Both growth factors dose-dependently stimulated angiogenesis, with bFGF showing a stronger effect than VEGF. The stronger stimulatory dose for both factors was 20 ng/mL. VEGF enhanced sprout formation from beads but not bFGF. VEGF was optimal at low concentrations (2.5-5ng/mL) and decreased at higher concentrations (15-35ng/mL). bFGF played an important role in the regulation of guided EC migration during angiogenesis. Immobilized VEGF supported cell migration and survival. VEGF 189 enhanced EC adhesion when compared to VEGF165. Low concentrations of both VEGF12i and VEGFi65 promoted the growth of long, thin vessels, whereas higher concentrations of VEGF remarkably enhanced vessel diameter. Low concentration of FGF-1 (1 ng/mL) maintained a persistent microvascular network, whereas higher concentrations resulted in an abnormal CJEC phenotype and vessel regression. Significant effect of FGF-1 started at 1 ng/mL while VEGF at 2ng/mL. The optimum effect was at higher concentration (lOOng/mL) for both growth factors. When combining bFGF and VEGF at concentration of lOng/mL, the effect was greater than either bFGF or VEGF alone.
Ref.
[7]
[9]
[12]
[17]
[22]
[29]
[30]
[41]
[42]
EC: endothelial cells.
99
Figure 4.1: Effect of fibroblasts over microvessel development.
After being sandwiched in fibrin, HUVEC migrated along the fibre axis and some toward the gel (a). After 1 day of culture, some cells were able to build cell-cell connections along the fibre axis (discontinued arrows) and even able to bridge between fibres (arrows) (b). In the absence of fibroblasts, many cells failed to establish cell-cell connections after 2 days of culture (arrows) (c). Comparing to the control, the use of fibroblasts produced more tube-like cell-cell connections after 2 days (d). After 4 days of culture, low magnification images of samples with fibroblasts reveal that cell-cell connections covered almost all the area between adjacent fibres (e). With the presence of fibroblasts, phase contrast microscopy (f) as well as immunofluorescence (F-actin and nucleus staining) (g) confirmed the presence of a complex tube-like structure network after 4 days of culture. Scale bars correspond to 100 um.
100
Figure 4.2: Immunofluorescence picture illustrating the effect of fibroblasts on
microvessel development.
After 4 days of culture, cells were stained for F-actin using rhodamine-phalloidin (red) and counterstained with SYTOX Green to highlight the position of nuclei during confocal analyses. Confocal pictures were recorded as layer-by-layer images from the top to the bottom of samples (a-d). Complete image reconstruction is presented in (e). To confirm the degradation of fibrin after 4 days, HUVEC were stained with the CFSE Cell Tracer (green) prior to be cultured in fibrin made of a mixture of fluorescent (red) and non-conjugated fibrinogen. A complete image reconstruction is shown at low (f) and high magnification (g). Scale bars correspond to 100 urn.
101
Concentration (ng/mL)
Low (2-5) High (20)
VEGF
bFGF
Figure 4.3: Immunofluorescence picture (F-actin and nuclei staining) illustrating the effect of
VEGF and bFGF on microvessel development. Scale bars correspond to 100 um.
102
\ .
fibre
AS-
^
t "-.
Figure 4.4: Illustration of multi-cellular lumen formation for cultures carried out for a period
of 4 days with fibroblasts.
HUVEC were stained for F-actin (green) and nuclei (blue). Fibrin was made using a mixture of fibrinogen conjugated with Alexa Fluor 546 (red) and non-conjugated fibrinogen. Microvessel structure and fibrin degradation are illustrated in the layer-by-layer pictures (a-d) as well as in the complete image reconstruction (e). Image of the culture system at time zero (f). Examination of histological sections stained using hematoxylin and eosin reveals HUVEC lining (arrows) and lumen formation (star) (g). Lumens were guided by the fibre (discontinued arrow) (h). Higher magnification reveals that HUVEC are lining (i). Scale bars correspond to 30 um (a-f), to 100 um (g and h), and to 10 um (i).
103
Culture time
8h
48 h
72 h
Uncovered PET fibres
PET fibres pre-covered with HUVEC
^ « ^ | » ! W « i « « * « * » ^ ' # i * | S ' Jffi*s4
wwnipiiiim mn'tfiii'.'imr •aa>***i*,«<t,*m&<c^&i
man, fmtiwi* <am mmxiiMen^mf.i^*^ jfflfc «as*. trsM • R W vwwmijmmmw «•***«*
t a^ f^p^te^ l
I d
••-• - « • -^- rt
f
• i i f f • » « • - • a t ••»•
^ * - ^ t *4 . « , • » • • •
.. * : i . <*uf--<*->* v . •*»
. - * • I , • • • . •• •
• . * » •
.» " *
-^g».*IWi.-Aff4*W :«i.JW'» * f • < * '•#*- * * l
• * * • . ; ! " 7 . . . . - . -V <f * •- ' I - - <
PtTt-hie
! ' *v i5 i ' "it,-- .•ta-.fc.iit.i.'.yi -; ' * . *.*••******-ar-tim. :'*s .9**.ri»i.-"ram
Supplementary Figure 4.i: Comparison between systems using cell-free fibres (left) and
HUVEC-covered fibres (right) over cell-to-cell connections and microvessel development.
Scale bars correspond to 100 um.
104
P6T fibre
Supplementary Figure 4.ii: Immunofluorescence confocal pictures of a sample cultured with
no fibroblasts.
After 4 days, cells were stained for F-actin using rhodamine-phalloidin and counterstained with SYTOX Green to highlight the position of nuclei. Confocal pictures were recorded as layer-by-layer images from the top to the bottom of samples (a-d). Complete image reconstruction is presented in (e). Scale bars correspond to 100 u.m.
105
Supplementary Figure 4.iii: Fibroblasts stained with the CFSE CellTracer.
106
CHAPTER 5
Conclusions and Suggestions
5.1 Conclusions
This thesis project aimed at developing a tridimensional cell culture system for the guidance of
microvessel formation. It was the initial part of a larger research program aiming to create an
oriented functional blood microvessel network within three-dimensional scaffolds. The
development of mature and oriented blood microvessels should facilitate oxygen and nutrient
transport as well as waste removal within tissue constructs and can be applied to culture
vascularised tissue substitutes.
Overall, the results presented in this thesis reveal that this novel culture system can be used to
successfully orient the development of microvessels. However, to reach the goals targeted in
this research program, some further investigations should be carried out in the future. For
example, investigating microvessel functionality should be done in future studies, as suggested
by others [Chrobak et al. 2006]. Testing the culture method presented in this thesis in a
dynamic culture system [Chrouinard et al. 2009] to achieve more mature microvessels should
be further studied.
Briefly, this thesis presents the development and validation of a novel cell adhesion method to
increase HUVEC adhesion and spreading on untreated polymer fibres. Further tridimensional
culture experiments using these HUVEC pre-coated fibres sandwiched between fibrin gels
revealed that this system can induce formation of tube-like structures in a directional fashion.
The optimum fibre-to-fibre distance to promote oriented microvessels was found to be IOOUMTI.
The use of growth factor (i.e., Vascular Endothelial Growth Factor (VEGF) and Basic
Fibroblast Growth Factor (bFGF)) was applied to improve microvessel development. Also, co-
culturing these HUVEC-covered fibres with fibroblasts significantly increased the maturation
107
of the formed microvessels. Further histology and fluorescent fibrin experiments also
confirmed that microvessel-like structures, which were developing between polymer
monofilaments embedded in fibrin, contained a lumen.
We believe that the well oriented microvessels reported in this thesis, which consisted of
multi-cellular lumens, should facilitate flow induction within these lumens. Also, this assay
will find applications in cancer research and tissue engineering.
5.2 Conclusion (en francais)
Ce projet de these visait a developper un systeme tridimensionnel de culture cellulaire pour
orienter la formation de micro-vaisseaux. II represente la partie initiale d'un programme de
recherche plus large qui a pour objectif de creer un reseau de micro-vaisseaux sanguins
fonctionnels a l'interieur d'echafaudages tridimensionnels. Le developpement de micro-
vaisseaux sanguins matures devrait faciliter le transport de l'oxygene et des nutriments ainsi
que l'enlevement des dechets dans les masses tissulaires.
Dans l'ensemble, les resultats presentes dans cette these montrent que ce systeme de culture
novateur peut etre utilise avec succes pour orienter le developpement des micro-vaisseaux.
Cependant, pour atteindre les objectifs vises dans ce programme de recherche, des
investigations complementaires devraient etre menees dans le futur. Par exemple, 1'etude de la
fonctionnalite des micro-vaisseaux devrait etre realisee tel que suggere par d'autres [Chrobak
et al. 2006]. Tester la methode de culture presentee dans cette these dans un systeme de culture
dynamique [Chrouinard et al. 2009] pour atteindre des micro-vaisseaux plus matures devrait
etre fait.
En bref, cette these presente le developpement et la validation d'une nouvelle methode pour
augmenter 1'adherence et la migration de cellules endotheliales humaines provenant de la
veine ombilicale (HUVEC) sur des fibres de polymere non-modifiees. D'autres experiences de
culture tridimensionnelle en utilisant ces fibres pre-recouvertes d'HUVEC et « sandwichees »
entre des gels de fibrine ont revele que ce systeme peut induire la formation de structures
108
tubulaires de facon directionnelle. La distance optimale fibre-a-fibre pour promouvoir la
formation de micro-vaisseaux orientes est d'environ lOOum.
L'utilisation du facteur de croissance (i.e., Vascular Endothelial Growth Factor (VEGF) et le
basic Fibroblast Growth Factor (bFGF)) a ete appliquee pour ameliorer le developpement des
micro-vaisseaux. De plus, dans une autre serie d'experiences, la co-culture des fibres
recouvertes de HUVEC avec des fibroblastes augmente significativement la maturation des
micro-vaisseaux formes. L'histologie et des experiences menees avec de la fibrine
fluorescente ont egalement confirme que ces pseudo-micro-vaisseaux qui se developpent entre
les fibres de polymere incorporees dans la fibrine possedaient un lumen.
Nous croyons que les micro-vaisseaux bien orientes rapportes dans cette these, qui possedent
des lumens multicellulaires, devraient faciliter l'induction d'un flux au sein de ces lumens. En
outre, cette methode de culture et son utilisation pourrait trouver des applications dans la
recherche sur le cancer et en ingenierie tissulaire.
5.3 Suggestions
The proof-of-concept of this tridimensional culture system opens the door to other
experiments either in static or dynamic cultures using a perfusion bioreactor system in order to
achieve the final goal of this research program. The following future works and techniques are
suggested:
5.3.1 Further studies of the angiogenesis assessment
Angiogenesis is a multi-factorial process in which new blood vessels are formed from existing
vessels [Carmeliet 2000; Folkman and Haudenschild 1980]. The process of angiogenesis
occurring during tumour progression involves, in many respects, events found during
morphogenesis and development [Salcedo et al. 2005]. In tissue engineering, models of
angiogenesis development were adapted for the vascularization of tissue constructs. In this
thesis, the emphasis was to demonstrate the proof-of-concept that microvessel formation can
109
be guided by polymer monofilaments. Studies to further assess angiogenesis development
using our novel system can include:
• To investigate the involvement of integrins during microvessel formation. Some
integrins, particularly av[33, av[35 and cc5(3l, have been identified as markers for ligand-
based and functional mediators of tumour angiogenesis [Hwang and Varner, 2004; Davis
et al. 2002]. For example, Max et al. (1997) have reported the use of
immunohistochemistry to show that normal tissue gives a lower expression of ocvP3
integrin when compared to tissues undergoing angiogenesis. Also, RGD-containing
peptides, which bind to the ocvP3 integrin, have been proposed as tracer for tumour
targeting and angiogenesis imaging [Haubner et al. 2001].
• To study the expression of growth factors and proteolytic enzymes during
angiogenesis process. The assessment of angiogenesis can also be done by investigating
growth factor expression from endothelial cells (EC), such as Vascular Endothelial
Growth Factor (VEGF), Basic Fibroblast Growth Factor (bFGF), and Platelet-derived
Growth Factor (PDGF) [Nakatsu et al. 2003b; Yancopoulos et al. 2000]. For example,
using Western blot, the level of VEGF expression was compared between control cells and
those undergoing angiogenesis. Also, the expression of proteolytic enzymes (i.e., matrix
metalloproteinases (MMP)) that can be related to fibrin degradation during capillary
formation can indicate when endothelial cells undergo angiogenesis [Koh et al. 2009].
• Proteins as markers for angiogenesis. The occurrence of angiogenesis process can also
assessed by characterizing the expression of other proteins related to endothelial cell
function, such as endothelial nitric oxide synthase (eNOS), vascular cell adhesion
molecule-1 (VCAM-1), von Willebrand Factor (vWF), inter-cellular adhesion molecule-1
(ICAM-1), platelet endothelial cell adhesion molecule-1 (PECAM-1), and E and P-
selectins [Bramfeld et al. 2010; Davis et al. 2002; Montano et al. 2009], only to name
a few.
• Gene expression during the development of an oriented microvessel. Transfection
techniques (i.e., adenoviral and siRNA transfection) have been used to silence (either
knock-out or knock-down) some genes (i.e., Tie-2, delta 4, notch 1 and HESR-1) that are
related to angiogenesis [Nakatsu et al. 2003a; Taylor et al. 2002]. In this study, HUVEC
110
were firstly treated in order to knock-down the gene which is related to angiogenesis,
before using these cells. Therefore, the proof of angiogenesis process occurring in our 3D
method presented in this thesis can be further investigated.
• To study expression of the extra-domain B (EDB domain) of fibronectin. The EDB
domain is a sequence of 91 amino acids that can be inserted into the fibronectin molecule
[Halin et al. 2001] and is one of the best markers for angiogenesis [Alessi et al, 2004]. In
tumour progression, fibronectin containing EDB accumulates around neovascular
structure, which appears to be found only in tissues undergoing angiogenesis [Halin et al.
2001]. To evaluate the expression, radio-labelled single-chain Fv (scFv) antibody which
would recognise the same epitope of EDB domain of fibronectin can be used. For
example, Viti et al. (1999) have concluded that scFv expression of tumour vasculature was
higher compared to that in normal tissues [Viti et al. 1999].
5.3.2 The effect of dynamic culture system over microvessel development and
proof of microvessel functionality
Dynamic cell culture experiments can be carried out using a simple set-up (see Figure 5.1) as
well as using an automated bioreactor system, which has been previously described
[Chrouinard et al. 2009]. Using a dynamic culture system, below are further suggested works
to study the development of oriented microvessels.
• Effect of dynamic culture over microvessel development. Endothelial cells are
experiencing fluid shear stress and dynamic flow conditions in the entire circulatory system
[Aird 2007]. The use of dynamic culture parameters (i.e., flow rate and pulsation) can allow
different mechanical stimulation [Levenberg et al. 2005]. A simple dynamic culture (see
Figure 5.1) can be used to study the effect of flow rate, while to investigate the influence of
pulsation an automated bioreactor system such as the one available in Prof. Vermette's
laboratory can be used. In this study, the effect of mechanical stimulation on cellular
guidance and microvessel development could be further investigated in order to find the
optimum parameter.
• Assessment of angiogenesis process during microvessel development in a dynamic
culture environment. For further validation of the occurrence of angiogenesis, several
111
characterizations listed in the above section could be used. For example, the level of VEGF
and PDGF-BB expression on EC cultured with flow were found to be higher than those in
static cultures [Sumpio et al. 1998].
• Study of endothelial nitric oxide (NO) production and eNOS expression in dynamic
culture. As previously described, expression of some proteins can used to follow the state
of the EC involved in the angiogenesis process. Among them eNOS expression and NO
production have been found to increase when endothelial cells were exposed to flow
conditions [Li et al. 2003].
• Smooth muscle oc-actin (SMocA) to assess microvessel maturation. During the
formation of new microvessels in vivo, the expression of SMocA is indicative of the
presence of smooth muscle cells (SMC), which are essential to blood microvessel
maturation [Jain 2003]. Also, dynamic culture with flow and pulsation has been found to
improve the structural organization of SMC [Akhyari et al. 2002; Holtorf et al. 2005].
Therefore, when using co-culture systems of EC with SMC, the level of SMocA expression
at the site of microvessel formation is of significance in the overall angiogenesis process
[Kofidis et al. 2002].
• Assessment of microvessel functionality. After the microvascular tube is being mature
i.e., the vessel wall is completely covered by endothelial cells and pericytes, further
assessment of its functionality is important. Testing its functionality can be adapted from
techniques published by others [Chrobak et al. 2006].
112
Figure 5.1 Simple set-up for dynamic cell cultures.
A custom-made cell culture chamber (a) was prepared to fit the fibre-frame (b). The simple dynamic culture system consists of a power supply (c) (located outside a C02 incubator) and a microperistaltic pump connected to a bottle containing culture medium; air filter, silicon tubing and culture chamber can be placed within the incubator (d). An image of HUVEC cultured with a flow rate of 80 mL/min after 3 days is presented at low (e) and high (f) magnifications, illustrating healthy cells.
113
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115
APPENDIX A
Protocols for cell staining
Basic Solutions
• Phosphate buffered saline solution (PBS IX, pH 7.4).
• Formaldehyde solution (3.75%, wt/v) in PBS for fixation.
• Triton X-100 solution (0.5% v/v in PBS) for permeabilization.
• PBS containing 20% (wt/v) BSA (referred to as PBS/BSA below) as a blocking buffer
solution.
A . l Protocol for F-Actin staining
One of the common markers for F-actin filament is TRITC-phalloidin (cat. #P1951, Sigma-
Aldrich). Phalloidin is isolated from Aminata phalloides mushroom and binds in the same site
of F-actin (see Appendix B for detail information).
Step-by-step staining procedure for F-Actin:
• Samples were gently washed with PBS (3 times) then fixed in a formaldehyde solution
for 20 min.
• Following three washes with PBS, samples were then permeabilized with a Triton X-
100 solution for 5 min at room temperature.
• After three rinsing steps in PBS, samples were incubated 1 hour in a PBS/BSA
solution for a blocking process then rinsed three times in PBS.
• Samples were then incubated in a solution containing a mixture of TRITC-phalloidin
(1/300) and Hoechst (1/10,000) made in PBS/BSA for 1 hour at room temperature, in
the dark.
• After two final rinsing steps with PBS, samples were conserved in PBS.
116
A.2 Protocol for Di l -Ac-LDL uptake
Dil-Ac-LDL (Acetylated Low Density Lipoprotein) is a marker to distinguish endothelial cells
from other cell types (i.e., fibroblasts, smooth muscle cells, epithelial cells and pericytes).
When cells were labelled with Dil-Ac-LDL, the lipoprotein is degraded by lysosomal enzymes
and the fluorescent probe (Dil) accumulates in the intracellular membranes of endothelial
cells. Dil-Ac-LDL used in this project was purchased from Biomedical Technologies Inc.,
Stoughton, MA, USA (cat. # BT-902).
Step-by-step staining procedures:
• Dilute Dil-Ac-LDL to lOug/mL in supplemented Ml99 media.
• Add this diluted staining solution over the endothelial cells monolayers then incubate
at 37°C and 5% C0 2 for 4 hours.
• Remove the Dil-Ac-LDL staining solution and wash twice with Ml99 medium with no
Dil-Ac-LDL.
• After incubation at 37°C and 5% CO2 for 30 minutes, cells are ready to be observed
with fluorescence microscopy using a standard filter for Alexa 546 or rhodamine.
117
A.3 Protocol for von Willebrand Factor staining
Von Willebrand Factor (vWF) is a large multimeric glycoprotein secreted by megakaryocytes
and endothelial cells. vWF participates fundamentally in the hemostasis process in the
association with thrombogenic Factor VIII (FVIII) (i.e., blood clotting factor or pro-cofactor).
During the hemostasis, vWF and FVIII have been found to be synthesized and released by the
endothelium into the bloodstream. Therefore, vWF is known as a marker for endothelial cells.
Goat anti-vWF was purchased from Haematologic Technologies Inc., USA (cat.# PAHVWF-
G) and diluted 1/500 in PBS/BSA. Secondary antibody FITC rabbit anti-goat was purchased
from Chemicon (cat.# AP106F) diluted 1/100 in PBS/BSA mixed with Hoechst (cat. # H-
33258 Sigma-Aldrich) (1/10000).
Below is the detailed protocol:
• Samples were gently washed with PBS (3 times) then fixed in a formaldehyde solution
for 20 min.
• Following three washes with PBS, samples were permeabilized with a Triton X-100
solution for 5 min at room temperature.
• After three rinsing steps in PBS, samples were saturated in rabbit serum (Biodesign,
USA; cat# N66010R) diluted 1/20 in PBS for 30 minutes, then rinsed three times in
PBS.
• Incubate cells with primary antibody goat anti-vWF in PBS/BSA for 1 hour, then rinse
three times in PBS.
• Incubate samples with secondary antibody FITC rabbit anti-goat and Hoechst in
PBS/BSA for 1 hour at room temperature and protect from light.
• Finally, rinse three times in PBS.
118
APPENDIX B
Cell cytoskeleton
The cytoskeleton is a framework structure inside the cytoplasm of eukaryotic cells. Its
function is mainly to give a cell its distinctive shape and high level of internal organization.
The major structural elements of cytoskeleton are: microtubules, intermediate filaments, and
microfilaments. These three major elements play important roles to support cell movement
and motility.
Microtubules (MT) are the largest structures and are composed of a hollow structure with
outer and inner diameters of 25nm and 15nm, respectively. MT are responsible for motility of
the cell and also support cell shape and distribution of micro and intermediate filaments.
Intermediate filaments (IF) have a diameter about 8-12nm and are the most stable and least
soluble constituents compared to the two other major cytoskeleton components. IF are thought
to have a tension-bearing role since they are often observed in areas of mechanical stress. In
contrast with other components, they differ in size and chemical compositions between one
tissue to another.
Microfilaments (MF) are the smallest structures, with a diameter of approximately 7nm. They
play important role in cell contraction and cell locomotion. They also contribute in the
maintenance of cell shape. MF are composed of the protein actin, called G-actin, which can
form double-helical strands of F-actin filaments (F stands for filamentous). To date, F-actin
filaments are known as sites for cell-cell and cell-ECM interactions.
119