fabrication of a 3-dimensional cardiac tissue using a ... · fabrication of a 3-dimensional cardiac...
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Fabrication of a 3-dimensional cardiac tissue using a modular tissue engineering approach
by
Brendan Martin Pue-Bun Leung
A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy
Institute of Biomaterials and Biomedical Engineering
University of Toronto
© Copyright by Brendan Martin Pue-Bun Leung 2011
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Fabrication of a 3-dimensional cardiac tissue using a modular
tissue engineering approach
Brendan Martin Pue-Bun Leung Doctor of Philosophy
Institute of Biomaterials and Biomedical Engineering University of Toronto, 2011
Abstract
Implantation of engineered cardiac tissue may restore lost cardiac function to damaged
myocardium. We propose that functional cardiac tissue can be fabricated using a modular,
vascularized tissue engineering approach developed in our laboratory. In this study, rat aortic
endothelial cells (RAEC) were coated onto sub-millimetre size modules embedded with
cardiomyocyte-enriched neonatal rat heart cells (CM) and assembled into a contractile,
macroporous sheet-like construct.
Cell morphologies, contractility and responsiveness to electrical stimulus were examined
to evaluate the function of the resulting modular construct. CM embedded modules contracted
spontaneously at day 7 post-fabrication and remained viable in vitro at day 14. Modules cultured
in 10% bovine serum were more contractile and responsive to external stimulus compared to
10% FBS medium cultured modules. VE-cadherin staining showed a confluent layer of RAEC
on CM embedded co-culture modules at day 7. Co-culture modules were also contractilie, but
when compared to CM only modules their electrical responsiveness was slightly diminished.
Modules assembled into macroporous sheets retained their characteristics at day 10 post-
assembly. Micrographs from histological sections revealed the existence of muscle bundles near
the perimeter of modules and at inter-module junctions.
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The fate of modular cardiac tissues in vivo was examined using two implantation
strategies based on a syngeneic animal model. Co-culture modules (CM and EC) were either
injected into the peri-infarct zone of the heart, or fabricated into a patch form and implanted
over a right ventricular free wall defect. In both models, donor EC were involved in the
formation of blood vessels-like structures, which appeared to have connected with the host
vasculature. Co-culture implants had a higher overall vessel density compared to CM-only
implants, but only in the absence of MatrigelTM. Moreover, donor CM organized into striated
muscle-like structures, at least when MatrigelTM was removed from the matrix. Together these
results suggest that modular cardiac tissue can survive and develop into native-like structures
when implanted in vivo and the potential of the modular approach as a platform for building 3-D
vascularised cardiac tissue should be explored in greater depth.
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Acknowledgements
I’d like to begin by expressing my most sincere gratitude to my supervisor, Dr. Michael
Sefton, for his guidance, patience and understanding during my graduate training. I feel very
blessed to have had Dr. Sefton as my mentor. His genuine interest in my personal and
professional development has shaped me into the researcher that I am today. I will cherish all
the lessons that I have learned from our numerous discussions in his office and our countless
chit-chats in front of the espresso machine; I thank you from the bottom of my heart.
I am also very fortunate to have had a wonderful group of professors who served on my
advisory committee. Thank you Dr. Simmons, Dr. Backx and Dr. Rocheleau for all your
insightful comments and suggestions. In particular, I’d like to thank Dr. Radisic for helping me
get the project off the ground and for all her expert advice in the field of cardiac tissue
engineering.
I’d like to thank everyone in the Sefton lab and the 4th floor of CCBR for all the laughter
and support. A special thanks goes out to Chuen Lo for all his technical assistance and
encouraging words. To Omar Khan, Mark Butler and Rohini Gupta, thank you for making my
graduate experience memorable both inside and outside of the lab.
I am forever indebted to my parents, Robert and Nancy Leung. Thank you for giving me
the inspiration and opportunity to pursue my dreams. Also, to my brother Brian, thank you for
your support and good times.
Finally, to my loving wife Helen, thank you for believing in me. Your love and
encouragement have helped me see the positive side of everything and have made me a better
person. To you, I dedicate this thesis.
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Table of Contents
ABSTRACT .................................................................................................................... II
ACKNOWLEDGEMENTS ............................................................................................. IV
TABLE OF CONTENTS ................................................................................................. V
LIST OF FIGURES ...................................................................................................... VIII
LIST OF TABLES ........................................................................................................... X
ABBREVIATIONS ......................................................................................................... XI
CHAPTER 1: RESEARCH INTRODUCTION, HYPOTHESES AND OBJECTIVES ..... 1
1.1 The Need for Engineered Cardiac Tissue ................................................................................................... 1
1.2 Research Overview and Hypothesis ............................................................................................................ 2
1.3 Research Objectives ........................................................................................ Error! Bookmark not defined. 1.3.1 Overall Objectives ...................................................................................................................................... 3 1.3.2 Specific Objective 1: Fabrication of CM-EC co-culture modules ............................................................. 3 1.3.3 Specific Objective 2: Characterization of CM/EC co-culture modules ..................................................... 3 1.3.4 Specific Objective 3: Fate of CM/EC co-culture modules in vivo ............................................................. 4
1.4 Thesis Organization ...................................................................................................................................... 5
CHAPTER 2: CURRENT METHODS AND CHALLENGES IN CARDIAC TISSUE ENGINEERING ............................................................................................................... 7
2.1 Engineered Cardiac Muscles for Functional Cardiac Regeneration ................................................................ 7
2.2 General Considerations for in vitro Tissue Engineering ................................................................................... 8
2.3 Cell Sourcing for Cardiac Tissue Engineering ................................................................................................... 9 2.3.1 Cardiomyocytes ............................................................................................................................................. 10 2.3.2 Endothelial cells ............................................................................................................................................ 13
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2.3.3 Cardiofibroblast ............................................................................................................................................. 17
2.4 Biomaterials for Cardiac Tissue Engineering ................................................................................................... 18 2.4.1 Properties and Functions of Cardiac Extracellular Matrix ............................................................................ 18 2.4.2 Collagen ......................................................................................................................................................... 19 2.4.3 Elastin and Other ECM Proteins ................................................................................................................... 21 2.4.4 Synthetic Scaffold and Materials ................................................................................................................... 22
2.5 Fabrication and Culture Platforms for Cardiac Tissue Engineering ............................................................. 23
2.6 Angiogenesis and Cardiac Tissue Vascularization Strategies ......................................................................... 25
2.7 In vivo models for evaluating engineered tissue ............................................................................................... 28 2.7.1 Allogeneic and immunosuppressed animal model ........................................................................................ 29
CHAPTER 3: A MODULAR APPROACH TO CARDIAC TISSUE ENGINEERING .... 49
3.1 Abstract ................................................................................................................................................................ 49
3.2 Introduction ......................................................................................................................................................... 50
3.3 Materials and Methods ....................................................................................................................................... 52 3.3.1 Cardiomyocyte isolation and module fabrication .......................................................................................... 52 3.3.2 Endothelial cell seeding ................................................................................................................................. 53 3.3.3 Module sheet .................................................................................................................................................. 54 3.3.4 Electrical response assessment and field stimulation .................................................................................... 54 3.3.5 Immunofluorescence staining ........................................................................................................................ 55 3.3.6 Statistical analysis .......................................................................................................................................... 56
3.4 Results ................................................................................................................................................................... 57 3.4.1 Contractility of CM embedded modules ....................................................................................................... 57 3.4.2 Behavior of RAEC seeded CM embedded modules ..................................................................................... 62 3.4.3 Module sheets ................................................................................................................................................ 64
3.5 Discussion ............................................................................................................................................................. 70 3.5.1 Electrical characterization ............................................................................................................................. 70 3.5.2 Co-culture model ........................................................................................................................................... 71 3.5.3 Sheet formation .............................................................................................................................................. 72
3.6 Conclusion ............................................................................................................................................................ 74
CHAPTER 4: FATE OF MODULAR CARDIAC CONSTRUCTS IN VIVO ................... 79
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4.1 Abstract ................................................................................................................................................................ 79
4.2 Introduction ......................................................................................................................................................... 80
4.3 Materials and methods ........................................................................................................................................ 81 4.3.1 Isolation of rat primary CM and EC .............................................................................................................. 81 4.3.2 Modular cardiac patch model ........................................................................................................................ 81 4.3.3 Peri-infarct module injection model .............................................................................................................. 82 4.3.4 Immunostaining and Histological Analysis ................................................................................................... 83 4.3.5 Statistical analysis .......................................................................................................................................... 84
4.4 Results ................................................................................................................................................................... 85 4.4.1 Characterization of CM embedded modules and rat heart EC ...................................................................... 85 4.4.2 Modular patch and injection implants ........................................................................................................... 88 4.4.3 Host immune response and the removal of MatrigelTM ................................................................................. 97
4.5 Discussion ........................................................................................................................................................... 103 4.5.1 Implantation strategies ................................................................................................................................. 103 4.5.2 Fates of implanted cells and host immune response .................................................................................... 105
4.6 Conclusion .......................................................................................................................................................... 107
CHAPTER 5: FUTURE PERSPECTIVES AND CONCLUSIONS .............................. 112
5.1 Conclusions ........................................................................................................................................................ 112
5.2 Recommendations ............................................................................................................................................. 114 5.2.1 Improving embedded cell viability .............................................................................................................. 114 5.2.2 Using autologous cell source ....................................................................................................................... 116 5.2.3 Characterization of host-module coupling .................................................................................................. 117 5.2.4 Assessing the functional benefits of modular cardiac tissue ....................................................................... 119
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List of Figures Figure 1-1: Schematic of modular construct and modular sheet………………………………...5 Figure 2-1: A schematic diagram showing the major components in the coagulation cascade…………………………………………………………………………………………..15 Figure 2-2: Illustration of components found in endothelial cell tight junctions and adherens junctions…………………………………………………………………………………..……..16 Figure 2-3: Schematic diagram of the three major cardiac tissue engineering strategies……………………………………………………………………...…………………25 Figure 2-4: Changes in the endothelium during angiogenesis…………………………………27 Figure 3-1: Viability (live/dead assay) of cardiomyocytes embedded in modules…………….58 Figure 3-2: Fractional area change of a single module when stimulated by external signal…………………………………………………………………………………………….59 Figure 3-3: Electrophysiological differences between BS and FBS cultured modules……………………………………………………………………………………….…60 Figure 3-4: Electrophysiological differences between modules with and without MatrigelTM……………………………………………………………………………………….61 Figure 3-5: Electrophysiological characteristic of CM/EC co-culture modules……………….63 Figure 3-6A: Schematic diagram of the testing chamber used to condition module sheets……66 Figure 3-6(B-E): Characterization of CM/EC co-culture modular sheets………………… …..67 Figure 3-7: Troponin I expression in cardiac modular sheets………………………………….68 Figure 3-8: Troponin I, vimentin and Cx-43 expression in cardiac modular sheets…………...69 Figure 4-1: Immunofluorescence staining of CM embedded in small diameter modules supplemented with MatrigelTM………………………………………………………………….86 Figure 4-2: Immunoflorescence micrograph of rat cardiac EC isolated from transgenic GFP+ Lewis rats………………………………………………………………………………………..87 Figure 4-3: VE-cadhering expression in EC coated modules before and after injection through a 28G needle………………………………………………………………………………………88
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Figure 4-4: Schematic diagrams and tissue morphology at explant for modular patch implant module injection implant………………………………………………………………………..90 Figure 4-5: Masson’s trichrome stain of patch implants ……………………………….……...91 Figure 4-6: Masson’s trichrome stain of module injection implants…………………………...92 Figure 4-7: Immunohistochemical staining of GFP+ donor cells in explants…………………..93 Figure 4-8: GFP and CD31 expression in patch implants...........................................................94 Figure 4-9: GFP and CD31 expression in module injection implants.........................................95 Figure 4-10: GFP and MHC expression in patch and module injection implants……………………………………………………………………………………….....96 Figure 4-11: Immunohistochemical staining of CD68+ macrophages in patch and module injection implants………………………………………………………………………………..98 Figure 4-12: Immunohistochemical staining of T cells in patch and module injection implants……………………………………………………………………………………….....99 Figure 4-13: GFP and CD31 expression in MatrigelTM-free module injection implants………………………………………………………………………………………...100 Figure 4-14: GFP and MHC expression in MatrigelTM-free module injection implants………………………………………………………………………………………...101 Figure 4-15: Vessel density count in patch and modules injection implants…………………102
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List of Tables Table 2-1: Sbtypes of collagen in human ……………………………………………………...20 Table 2-2: A summary of recent implantation studies using various cardiac tissues engineering strategies………………………………………………………………………………………...31 Table 3-1: Medium compositions (presumed key differences) and serum types used for in vitro study……………………………………………………………………………………………..53
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Abbreviations ASC : Adipose stem cell
Ang-1 : Angiopoietin 1
AP : Action potential
BMP-2: Bone morphogenic protein 2
CM : Cardiomyocyte
CF : Cardiofibroblast
Cx43: Connexin 43
d-CMP : Dynamic cardiomyoplasty
EC : Endothelial cell
ECM : Extracellular matrix
EPC : Endothelial progenitor cell
ESC : Embryonic stem cell
FN : Fibronectin
b-FGF : Basic fibroblast growth factor
GAG : glycosaminoglycan
GFP : Green fluorescent protein
IPN : Interpenetrating network
iPS: Induced pluripotent stem cell
MHC-β : Myosin heavy chain beta
MMP : Matrix metalloproteinase
MI : Myocardial infarction
bmMSC : Bone marrow derived mesenchymal stem cell
MSC : Mesenchymal stem cell
NO : Nitric oxide
PGA : Polyglycolic acid
PLA : Polylactic acid
PCL : Polycaprolactone
PDGF : Platelet derived growth factor
PDMS : Polydimethylsiloxane
PFA : Platelet activating factor
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SMC : Smooth muscle cell
TFPI : Tissue factor pathway inhibitor
TF : Tissue factor
TGF-β : Transforming growth factor beta
TnI : Troponin I
VEGF : Vascular endothelial growth factor
vWF : von Willebrand factor
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Chapter 1: Research Introduction, Hypotheses and Objectives
1.1 The Need for Engineered Cardiac Tissue
The heart is a complex organ with a simple function: to pump blood throughout the body
without interruption over an individual’s lifespan. However, certain diseases including coronary
artery disease and cardiomyopathy can cause cell death in the myocardium. Unlike skin or liver
tissues, adult myocardium has limited regeneration capacity. Thus, any loss of cells usually
translates to a permanent loss of cardiac function. On a tissue level, cell debris draws in
inflammatory cells and contributes to the formation of scar tissue. Ultimately, the combination
of tissue loss and scar tissue formation will further decrease the output of the heart and cause
more damage to the rest of the myocardium.
The vicious cycle of functional loss due to cell death leading to further tissue damage
could be stopped, and possibly reversed, by replacing diseased myocardium with functional
cardiac tissue. Tissue engineering has made it plausible to create tissues in vitro for transplant
and replacement of scar tissue. To date, several promising methods have been devised for
building cardiac tissue in vitro, including cell seeded scaffolds, cell-embedded hydogel, or
matrix-free cell sheets. However, many of these engineered cardiac tissues lack a functional
vasculature to sustain a high cell density. The challenges in creating cardiac tissue with
physiologically relevant thickness and function remain to be overcome before they can achieve
widespread clinical significance.
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1.2 Research Overview and Hypothesis
The overall goal of this work was to create functional, implantable cardiac tissue from a
modular tissue engineering construct embedded with neonatal rat cardiomyocytes (CM). We
hypothesized that the modular tissue engineering construct developed in our laboratory1 would
serve as a useful platform for producing vascularized cardiac tissue in vivo. A module is a small
cylindrical collagen gel with a length of approximately 1500 µm and diameter of 500 µm.
Modules were embedded with a CM-rich cell mixture composed of CM, fibroblast, cardiac
progenitor stem cells, and endothelial cells (EC). We coated the surface of each module with a
monolayer of EC with the view that such design would provide the means to control initial cell
seeding density, reduce thrombogenicity and improve scaffold vascularization in vivo. We
examined the responses of CM-only as well as cardiomyocyte-endothelial cell (CM-EC) co-
culture modules to external electrical stimulation. In addition, we assessed the effect of surface
coated EC on the maturation and function of embedded CM.
The fate of the cardiac modular construct in vivo was examined using syngeneic animal
models. Cells from transgeninc eGFP Lewis rats were used to construct modules and then
implanted into host Lewis rats. Cell survival, maturation and immune responses were
monitored. We hypothesized that the presence of EC on CM-EC co-culture modules would
increase implant vascularity and lead to subsequent improvements in overall cell survival and
tissue morphology compared to CM only modules. Also, we expected the cardiac modular
construct, with or without EC, to be well tolerated by their syngeneic host with minimal immune
response.
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1.3 Research Objectives
1.3.1 Overall Objectives
The overall objective of this project was to create cardiac tissue using a modular tissue
engineering construct. To achieve this we had to 1) devise a method of fabricating CM-EC co-
culture modules and higher order structures, 2) characterize modular cardiac tissue using
functional in vitro assays, and 3) investigate the fate and remodeling process of cardiac tissues
using in vivo models. These three objectives will be covered in the following section.
1.3.2 Specific Objective 1: Fabrication of CM-EC co-culture modules
Modular co-culture constructs (CM-EC) were fabricated using a collagen based modular
engineering construct developed in our laboratory. Each module was embedded with a CM-rich
cell mixture harvested from neonatal rat heart and the surface was coated with a monolayer of
rat EC (Figure 1-1A).
We have also devised a method to assemble modules into a single, sheet-like structure
with thickness and pores the size of a module (Figure 1-1B). Modules embedded with CM
underwent spontaneous agglomeration and fusion when placed in close proximity to one
another. By immobilizing a single layer of modules in alginate gel, intermodular junctions were
fused together while alginate, a non-cell adherent natural polymer, acted as a physical and
biochemical barrier to prevent the collapse of interstitial spaces between modules and to
maintain sufficient permeability for nutrient transport. An interlinking, perfuseable,
macroporous modular network was obtained by dissolving the alginate gel with a citric acid
buffer.
1.3.3 Specific Objective 2: Characterization of CM/EC co-culture modules
Earlier studies have demonstrated that collagen modules can support and maintain
surface coated HUVEC and embedded HepG2 cells2. However, the functionalities of embedded
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cells have not been investigated. In this project, both CM and RAEC were assessed for the
presence of functional phenotypes. Electrical responsiveness and contractility were measured as
primary indicators of embedded CM functionality. While, the expression of cardiac specific
markers, including troponin I, troponin T and myosin heavy chain on immunohistological
sections served as secondary markers. For surface coated EC, VE-cadherin expression was used
as a quiescent EC marker.
1.3.4 Specific Objective 3: Fate of CM/EC co-culture modules in vivo
The interactions between engineered tissues and their hosts can significantly impact the
viability and functionality of the implant. In this project, we examined the fate of the modular
cardiac construct in vivo using a syngeneic rat model. Cells harvested from neonatal eGFP
transgenic Lewis rats were incorporated into modular constructs and implanted into the hearts of
Lewis rats. We used two separate implantation models to determine the best method of
deploying the modular construct into the heart and also to observe their impacts on host-graft
interactions. Our first model involved the construction of a cardiac patch, where macroporous
cardiac modular sheets were sandwiched between two pieces of porous gelatin foam, also know
as GelFoam. These patches were then implanted over a transmural wound on the right
ventricular free wall of a healthy Lewis rat heart and explanted after 2 weeks. The second model
attempted to determine the fate of cardiac modules when implanted as individual microtissues in
the heart. An acute myocardial infarct (MI) was induced in a Lewis rat by ligation of the left
descending coronary artery. After 7 days, small diameter modules embedded with CM were
injected into the peri-infarct zone and explanted after 21 days. Explanted tissues were assessed
for cell viability and morphologies by histological analyses (H&E, Mason Trichrome, eGFP,
TnI, TnT, MHC, CD31, CD68, T-cell receptor, and other relevant immunohistochemical
stainings). Blood vessel density (both GFP positive and GFP negative) were measured as a
surrogate marker for angiogenesis.
In both models, we expected CM-embedded modules implanted into the rat heart to
develop into more native-like morphology compared to CM-modules in vitro. Also, we expected
the presence of EC in co-culture modules to improve overall cell survival and illicit a more
pronounce angiogenic response. Finally, in the injection model, we expected the modular
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approach to yield better cell engraftment and improve survivability compared to conventional,
matrix-free cell injection therapy.
Figure 1-1: Schematic diagrams of a CM embedded EC coated module (A) and a single layer,
porous modular sheet (B). Modular sheet were fabricated by immobilizing a single layer of
modules on a nylon mesh with alginate gel. After the intermodular junctions have fused, the
alginate was removed using a citrate buffer wash, leaving behind a porous modular sheet.
Details about the fabrication and characterization of modular cardiac construct will be covered
in Chapter 3 and 4.
1.4 Thesis Organization
The thesis is organized into self contained chapters. The first chapter outlines the scope and
objectives of the thesis. While, chapter 2 goes on to review and present the current methods and
challenges in cardiac tissue engineering as well as relevant background literature. In chapter 3
and 4, we discuss the fabrication of modules and their behavior in vitro. Specifically, chapter 4
focuses on the fate of modular constructs in vivo. In the last chapter, we summarize the lessons
learned from the project and evaluate the usefulness of the modular construct in the field of
cardiac tissue engineering. We also discuss the future direction of the project in chapter 5.
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Reference List
1. A. P. McGuigan and M. V. Sefton, Design criteria for a modular tissue-engineered
construct. Tissue Eng 13, 1079-1089 (2007).
2. A. P. McGuigan and M. V. Sefton, Vascularized organoid engineered by modular
assembly enables blood perfusion. Proc. Natl. Acad. Sci. U. S. A 103, 11461-11466
(2006).
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Chapter 2: Current Methods and Challenges in Cardiac Tissue Engineering
2.1 Engineered Cardiac Muscles for Functional Cardiac Regeneration
Similar to other organs, the heart owes its specialized functions to the unique complement
of cells and their complex arrangements. Its contractile function derives primarily from
cardiomyocytes (CM). These myocytes are tightly packed with neighboring fibroblasts, which
act as supporting cells by maintaining the extracellular matrix (ECM) as well as guiding
electrical current propagations throughout the heart. On the other hand, the myocytes’ demand
for oxygen and nutrients are satisfied by a dense endothelial cell (EC) lined vasculature.
Together these three cell types form a mechanically and electrically coupled unit that generates
the force necessary to pump blood throughout the body.
Until the recent discovery of resident cardiac progenitor cells1,2, the myocardium was
thought to be composed entirely of terminally differentiated cells. However, in their native state,
these progenitor cells do not contribute sufficiently to the repairing of major cardiac trauma.
Thus, from a clinical perspective, the adult heart has little capacity for regeneration after injury.
A constant supply of oxygen and nutrient is required to sustain the high metabolic demand of
CM. For this reason, any disruption of circulation to the heart muscle, such as that seen during a
myocardial infarction (MI), will lead to rapid cell death and tissue necrosis. In most cases, the
cell debris triggers an inflammatory response that is characterized by the infiltration of
monocytes and macrophages. The necrotic tissue is then replaced by granulation tissue
consisting mainly of fibroblasts and collagen. Finally, this tissue is remodeled to form a stiff and
non-contractile scar tissue.
Recent studies have demonstrated that injection of cells into the myocardium, including
bone marrow derived mesenchymal stem cells (bmMSC)3,4, skeletal myoblasts5-7 and CM
derived from embryonic stem cells8,9, may prevent scar tissue formation and restore cardiac
function in patients with acute MI. It has also been reported that circulating endothelial
progenitor cells (EPC) from the patients’ bone marrow may become mobilized and participate in
angiogenesis in the peri-infarct zone10. A comprehensive review of current theories on cell-
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based therapies for MI has been published by Laflamme et al.11 and hence will not be covered
within this chapter.
In many cases, the presence of scar tissue and extensive tissue remodeling caused by
chronic ischemic cardiomyopathy requires the complete replacement of damaged tissue with
functional ones in order to achieve significant clinical benefits. Early attempts such as dynamic
cardiomyoplasty (d-CMP) aimed to replace cardiac function by implanting an autologous
skeletal muscle flap taken from the patients’ body. Although the implanted muscle flap
improved cardiac performance, the progressive degeneration of the implant and the lack of
electrical coupling with host tissue limited its clinical usefulness12. While d-CMP failed as a
replacement therapy to prevent total heart transplantation, it sparked the notion that one could
fabricate implantable cardiac tissue in vitro by combining CM and other supporting cell types in
an appropriate scaffold. The resulting engineered cardiac tissue should possess similar
mechanical and functional properties as native cardiac tissue. The presence of functional cells
should have the capability to secrete ECM and growth factors to encourage beneficial host
remodeling, which will significantly contribute to the long term integration of the implant13.
2.2 General Considerations for in vitro Tissue Engineering
In vitro engineered tissues could provide the means to rapidly replace tissue functions that
are lost due to trauma or acute diseases. These tissues may be biomimetic in nature, or they
could be designed to functionally imitate the target tissue. However, there are many obstacles
facing the clinical implementation of these therapies. Strategies for engineering tissue in vitro
must address the following elements:
1. Providing a mechanically and biochemically suitable extracellular environment to support
cellular function.
2. Finding the appropriate cell source and/or incorporating bimolecular cues into the scaffold
to promote recruitment of functional endogenous cells.
3. Providing sufficient perfusion of oxygen and nutrients to embedded cells.
4. Providing the necessary environmental stimuli to promote proper tissue development.
5. Integrating engineered tissue structurally and functionally with the host tissue.
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The biological and structural functions of engineered tissue are intimately related to the
phenotypes of the incorporated cells. Besides the obvious differences among cell types, it is also
important to consider the subtle phenotypic differences of cells from different origins and
lineages. For example, while most EC in various organs share common EC markers, there are
heterogeneous subpopulations of EC within the endothelium and even within the organ.
Functional differences between EC found in the brain, kidney, liver and gastrointestinal tract
manifest as different EC phenotypes. Mechanical forces in large diameter vessels versus those
associated with capillary blood flow contributes to functional and morphological differentiation
of EC14. Moreover, paracrine signaling by cytokines often causes local differential EC gene
expression15.
While achieving the necessary mechanical properties are essential, the design for
engineered tissues must also satisfy the basic requirements for supporting cellular growth.
Among the most crucial criterion is the need to provide adequate mass transport to and from the
implanted cells so that nutrients and waste products can be exchanged freely. Under desirable
circumstances, adequate blood supply into the tissue also ensures that the secreted therapeutic
molecules from the tissue can reach the systemic circulation.
Beyond the basic survival requirements of nutrient delivery, cells also need a suitable
extracellular environment to engage in cellular functions such as proliferation, migration and
differentiation. Selection of a tissue engineering scaffold that resembles in vivo ECM
architecture not only enhances the functionality of embedded cells, but may also encourage the
migration of endogenous cells into the implant and peri-implant region. Natural ECM
components possess many of the desirable qualities as a building block for tissue engineering
scaffolds. Considerations of cell sourcing, vascularization and ECM interaction in the modular
tissue engineering scaffold are discussed in detail within the following sections.
2.3 Cell Sourcing for Cardiac Tissue Engineering
Cell sourcing is perhaps the most important consideration when designing tissue
regeneration strategies. The choice of cell can directly impact host response, ECM remodeling,
and ultimately the functioning of the regenerated tissue. The cells selected should also be
matched with suitable scaffolds, culture conditions and delivery methods in order to maximize
its therapeutic potential. In cardiac tissue engineering, the cells of interest are primarily
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cardiomyocytes, cardiofibroblasts, and endothelial cells. The following sections will outline the
features of these cell types and their most common sources.
2.3.1 Cardiomyocytes
The most striking feature of the heart is its ability to contract continuously and
synchronously during a person’s lifespan. Heart muscle cells, called cardiomyocytes (CM), are
the primary cell type responsible for the generation of contractile force. The developmental
origin of CM can be traced back to the mesoderm layer of the gastrulated embryo16. The heart
itself is formed by two separate waves of myogenesis known as the primary and secondary heart
fields. The primary heart field originates from the anterior lateral mesoderm and give rise to the
left ventricle of the mature heart, while the secondary heart field develops into the right ventricle
inflow and out flow tracks17. The secondary heart field also give rise to some of the non-
myocyte component of the heart including cardiofibroblast (CF), smooth muscle cells (SMC)
and endothelial cells (EC).
CM share many similarities with skeletal muscle cells; both are striated, multinucleated,
and excitable cells. In comparison to skeletal muscle cells, CM are more resistant to fatigue,
contain more mitochondria, and are more adapted to aerobic respiration. Nevertheless, perhaps
the most distinct anatomical feature of CM are the intercalated discs found between adjacent
cells. Intercalated discs are areas of cell membrane rich in desmosomes and gap junctions.
Desmosomes are cadherin family transmembrane proteins that are associated with intermediate
filaments. Through homophilic binding, desmosomes also confer mechanical cell adhesion and
integrity between CM. On the other hand, intercellular communications are mediated by gap
junctions. Transmembrane proteins called connexins form hemi-channels on the membranes of
adjacent cells. These connexons can align and form a direct channel between the cytoplasm of
these adjacent cells, allowing membrane depolarization to propagate from one cell to the next,
thus facilitating action potential propagation over the myocardium.
The ultimate goal of cell sourcing in tissue engineering is to find a cell type that can
differentiate into all the necessary cells found in an organ in order to recapitulate organogenesis
during development. Perhaps, the cells that carry the most potential toward this goal are
embryonic stem cells (ESC) derived from the inner cell mass of the blastocyst. These cells are
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pluripotent and have strong proliferative capacity compared to adult stem cells. However,
pluripotency is not without its downfalls. Direct injection of undifferentiated ESC can lead to
teratoma formation18. Therefore, the phenotype of ESC and ESC derived cells must be carefully
regulated. Selective differentiation of ESC have been shown to be a promising path towards a
steady source of CM19-21. Laflamme et al. demonstrated in a rat model that the injection of
differentiated human ESC maintains cardiac phenotypes and continues to proliferate over 4
weeks22. In addition, the same group has demonstrated the utility of human ES derived CM in
tissue engineering application by fabricating contractile cardiac constructs23.
Although ESC derived CM improved cardiac function in vivo, their allogeneic nature
poses a significant obstacle towards clinical application. The recent discovery of induced
pluripotent stem (iPS) cells derived from adult fibroblast may be the answer for an autologous
and pluripotent cell source. The Yamanaka group pioneered the technique in restoring
pluripotency of somatic cells to the same level as ESC by ectopically expressing Oct3/4, Sox2,
Klf4 and c-Myc24. It can be assumed that current methods being used to drive ESC toward CM
differentiation can similarly be applied to iPS cells. Like other stem cells, the in vivo
tumorgenicity and immune response toward iPS cells will ultimately determine their clinical
usefulness. Further studies are needed to answer these questions.
From a clinical therapy perspective, the ideal CM for tissue engineering should retain
contractile phenotype indefinitely, cause minimal immune response in the host, and participate
in functional remodeling by organizing into contractile bundles25. Much of these criteria can be
satisfied by using CM differentiated from stem or progenitor cells. Studies in stem cell lineages
have identified populations of adult stem cells in tissues, such as adipose26, skin, dental pulp27
and amniotic fluid28,29, some of which are capable of differentiating into beating CM30.
However, the most studied population for cardiac regeneration in clinical application to date is
the bone marrow derived messenchymal stem cells (MSC). These cells inhabit the stromal space
of bone marrow. They can be obtained through direct bone marrow aspiration or from peripheral
blood. In both cases, the MSC reside as a subpopulation of cells within the mononuclear cell
fraction31. Under the right condition, these MSC can be induced to express cardiomyogenic
markers32 and play a cardioprotective role to nearby cardiomyoblasts33. In clinical studies,
intravenous or intramyocardial injections of MSC in acute MI have been shown to increase
ejection fractions and vascularity around the infarct area34,35. However, few MSC differentiate
into CM lineage and those that are committed have little to do with the functional improvement
12
that are observed36. Therefore, it appears that much of the benefits of MSC in acute MI are
derived from their cytoprotective and angiogenic roles37,38.
Another potential source of autologous CM can be found in adipose-derived stem cells
(ASC). Like MSC, these cells can be kept undifferentiated over long periods of time in culture
and they are capable of differentiating into muscle, bone and fat cells in vitro. Even though the
immunophenotype of ASC closely resemble that of MSC39, they behave very differently in
culture. ASC are capable of differentiating spontaneously into beating CM through vascular
endothelial growth factor (VEGF) paracrine signalling40. A similar phenomenon can also be
observed when ASC are cultured with cell extract of neonatal rat CM41, with TGF-β alone30, or
in the presence of explanted cardiac tissue42. Moreover, the induction of ASC into myogenic
phenotype by surrounding CM can be observed in vivo when ASC are injected intracardially in
an acute MI model43. Due to their strong VEGF paracrine signaling, ASC can promote
angiogenesis and stimulate vessel growth when cultured with neonatal hearts containing CM
and EC44. Intramyocardially injected ASC were also shown to significantly improved cardiac
function after MI in animal models45,46. These results have demonstrated the therapeutic
potential of ASC in cell therapy as well as a source of autologous CM for cardiac tissue
engineering.
Although not a viable CM source for therapeutic applications, primary rat neonatal
ventricular cardiomyocytes have served as a useful cell model that has greatly contributed to the
understandings of CM physiology and development. These CM differ from adult ventricular
CM in its ability to grow and differentiate during the transitional phase from neonatal to adult
myocytes in culture47. Structurally, neonatal CM lack a well developed T-tubule system, the
deep invagination of the sarcolemma, found in adult CM48. This leads to a more heterogeneous
cytosolic Ca2+ concentration compared to adult myocytes and contributes to its unique
excitation-contraction coupling characteristic.
13
2.3.2 Endothelial cells
Endothelial cells (EC) are found on the luminal surface of all blood carrying vessels. A
monolayer of EC, known as the endothelium, has been accepted as a separate organ weighting 1
kg in an average human that lines the entire circulatory system49. The lineage of EC is closely
related to hematopoietic cells. It is known that both cells differentiate from a common precursor
referred to as the hemangioblast. Although EC are considered a terminally differentiated cell
type, they are also believed to transdifferentiate into intimal smooth muscle cells (SMC)50-52.
Physiologically, EC are rarely present without cellular interaction with supporting cell types
such as pericytes and SMC. In a large diameter vessel, a layer of intimal SMC underlies the
endothelium, and together controls vascular tone by various paracrine and autocrine pathways.
At the capillary level, the endothelium is associated with pericytes, SMC-like cells. The
endothelium has long been considered as an inert barrier between blood and the underlying
tissues. However, recent advances in basic EC biology and cardiovascular research have
identified numerous biological and metabolic processes involving the endothelium. The
endothelium serves as a haemostatic organ by dynamically balancing pro-thrombotic and anti-
thrombotic signals. It is actively involved in controlling transendothelial transport of molecules,
gating the migration of various immune cells from circulating blood into interstitial space, and
may act with or against inflammatory responses. The endothelium is also an integral part of
establishing and maintaining vascular tone by sensing and responding to changes in shear stress
induced by blood flow. From a tissue engineering perspective, EC provide the building block for
enabling functional blood vessel to carry nutrients into the seeded cells. Consequently, the
following sections will cover the attributes of EC that are of primary interest, namely their
haemostatic and angiogenic (Ch 2.6) properties.
It is the haemostatic property of the EC that makes it an important component in a tissue
engineering scaffold vascularization strategy. The endothelium actively regulates thrombosis
and thrombolytic factors in response to platelet adhesion, blood chemistry, vasomotor tone and
blood flow regime53. The endothelium can be described in one of two states: (1) a basal or
quiescent state, where EC are non-proliferative and display low platelet adhesion, or (2) an
activated state, where EC phenotype change to a pro-thrombotic, proliferative state. The
endothelium can be triggered to switch from a quiescent to an activated phenotype by
pathogens, injury signals (tissue factor, collagen), or chemical cues (thrombin).
14
The basal state of the endothelium actively suppresses intravascular thrombosis by
secreting tissue factor pathway inhibitor (TFPI), which binds with factor Xa and tissue factor-
VIIa to form an inactive complex54. While TFPI is primarily bound to the EC membrane, it may
also be released into the circulation in a soluble form. Factors such as heparin, thrombin and
blood flow increases the production and/or release of TFPI55-57. EC also secrete
glycosaminoglycans (GAGs), namely heparan sulfate, that catalyze the inhibition of factor Xa
and thrombin by antithrombin58. Moreover, basal state endothelium mediates protein C (PC)
activation and inhibits thrombosis by expressing thrombomodulin (TM). TM is a membrane
bound protein that binds thrombin. The formation of TM-thrombin complex has two
anticoagulation effects. First, it prevents thrombin from activating fibrinogen, platelets, factor V,
VIII and XIII59. Secondly, the TM-thrombin complex activates protein C via the endothelial
protein C receptor (EPCR), which inactivates factor Va and VIIa and disables the synthesis of
thrombin60.
One of the major procoagulation proteins that are expressed by EC is tissue factor (TF).
Although TF is known to be constitutively expressed by subendothelial vascular cells, it is only
expressed by EC in an activated state61. TF forms a complex with plasma serine protease factor
VII/VIIa and in turn activates IX and X to initiate the blood coagulation cascade. In quiescent
endothelium, TF transcription is actively suppressed62. The endothelium can be activated by
many agonists including vascular injury, alteration of blood flow regime, and infection. Bacteria
derived endotoxins have been used routinely to activate EC in culture. In its activated state, EC
also express platelet activating factor (PAF), which together with P-selectin causes the adhesion
of platelet to the endothelium49. Platelet adhesion is often viewed as a critical step in the
initiation of intravascular thrombosis. The activated endothelium may also facilitate thrombosis
by releasing von Willebrand factor (vWF) to act as a stabilizer for factor VIII and promote its
binding to ECM components.
15
Figure 2-1: A schematic diagram showing the major components in the coagulation cascade.
The cascade is based on the sequential conversion of clotting factor precursors into active
proteases. The ultimate result is the generation of a fibrin, which then polymerizes and
crosslinks to form a clot. (Adapted from NCBI tutorial: Mutation and blood clot, CC)
The endothelium is one of a handful of non-structural organs that is capable of sensing
and responding to mechanical changes in its environment. EC from different regions of the
circulatory system are known to have different responses to external stimuli, including their
sensitivity toward hemodynamic forces63-65. Platelet-endothelial cell adhesion molecule
(PECAM or CD31) is an EC specific transmembrane adhesion protein that plays a major role in
mechanosignal transduction. The extracellular domain of PECAM participates in cell-cell
adhesion through homophilic association while the intracellular domain is associated with
various adaptor proteins through tyrosine phosphorylation and is ultimately associated to the
cytoskeleton. Integrins are another class of transmembrane proteins that are responsible for
mechanosignal transduction. Conformational activation of integrins induced by fluid shear is
responsible for adaptive EC morphogenesis including cytoskeletal alignment. However, the
16
extracellular domains of integrin are mainly associated with ECM components and hence they
are not considered as a mechnosensory components to fluid induced shear stress66,67.
As an organ that controls mass transport across the blood-tissue barrier, it is imperative for
EC to have strong and tight intercellular adhesion. Junctional components found between EC are
associated with one of the two specialized regions on the plasma membrane, namely tight
junctions and adherens junctions. Each type of junction contains its own unique membrane
proteins (Figure 2-2).
Figure 2-2: Illustration of components found in endothelial cell tight junctions and adherens
junctions68.
Tight junctions are located on the apical (or luminal) side of the EC cell-cell junction.
Their main function is to seal off the space between EC to give a continuous luminal surface in
order to facilitate blood flow. The main proteins involved in tight junction architecture are
occludins, claudins, and junction adhesion molecules (JAM)69. Their proximity to the lumen
allows them to serve as mechnosignaling sensors70.
Unlike tight junctions, adherens junctions are responsible for the mechanical integrity of
cell-cell junctions and intercellular signaling. A family of cell specific proteins known as
cadherins maintains adherens junctions. In EC, two types of cadherin are expressed: vascular-
endothelial cadherin (VE-cadherin or CD144) that localize to adherens junctions only and N-
17
cadherin that are found dispersed over the plasma membrane69. The function and structure of
VE-cadherin are well characterized and it is believed to be the major component of adherens
junctions. The extracellular domain of VE-cadherin exhibits homophilic, calcium dependent
associations. On the other hand, its intracellular domain associates with plakoglobin and β-
catenin, which ultimately connects to the actin cytoskeleton71. Similar to PECAM, the
intracellular domain of VE-cadherin can undergo tyrosine phosphorylation, which has been
shown to destabilize adherens juctions72 and may be used as a pathway to modulate endothelium
permeability. For this reason, the expression and localization of VE-cadherin has been used as a
vessel maturation and endothelium stability marker73. One of the receptors for VEGF, VEGFR-
2, mediates VE-cadherin tyrosine phosphorylation upon VEGF binding and is believed to
potentiate the sprouting response in early stage of angiogenesis74. The importance of VE-
cadherin as a structural component in the adherens junctions is further substantiated by attempts
to develop VE-cadherin knockout mice, which was demonstrated to result in embryonic
lethalaty75 due to impaired vessel development.
2.3.3 Cardiofibroblast
Cardiofibroblast (CF) of messenchymal origin constitutes the bulk of non-myocyte cell
population in the heart76. The main function of CF is to secrete and maintain ECM components,
including collagen, fibronectin, and laminin. Unlike CM and EC, CF is found natively in the
stromal space and lack basement membranes. This feature gives CF its unique ability to migrate
and populate injury sites, such as a myocardial infarct, and quickly restore tissue volume and
ECM proteins77. In the developing fetal heart, CF contributes to ECM rich structure including
valves and atrial ventricular walls. As the heart matures, a 3D network of collagen and CF,
known as the cardiac skeleton, begins to take shape. This network allows CF to exert forces on
the myocytes, as well as, to respond to external stimuli through degradation and synthesis of
ECM. These processes help maintain the mechanical integrity of the heart through cell-cell and
cell-ECM interactions. More importantly, CF forms heterotypic gap junctions with neighboring
CM or CF, whereby the conductivity of these junctions can be modulated by the differential
expression and coupling of connexins isoforms, including Cx45, Cx43 and Cx4078,79. The cell-
18
cell interaction between CF and CM helps to ensure the long range synchronization of
myocardium contraction.
CF also plays a pivotal role during cardiac repair. Residence CF can mobilize in
response to tissue damage and differentiate into smooth muscle-like cells called myofibroblast80.
These are not true smooth muscle cell, but they are more contractile and active compared to
fibroblast. The ability of myofibroblast to contract collagen and produce new ECM helps
stabilize wound site and form scar tissue. In myocardial infarcts, these cells are responsible for
the formation of granulation tissue. However, their eventual loss through apoptosis results in the
weakening of the scar and the ultimately leads to heart wall thinning77,81. Therefore, the dynamic
balance between fibroblast and myofibroblast phenotype is a critical factor in the wound healing
outcome.
2.4 Biomaterials for Cardiac Tissue Engineering
2.4.1 Properties and Functions of Cardiac Extracellular Matrix
Extracellular matrix (ECM) refers to the non-cellular components found in tissue. It is a
complex mixture of molecules including proteins such as collagen, elastin, laminin, fibronectin
and polysaccharides such as glycosaminoglycan (GAG). The composition and architecture of
ECM varies depending on the native tissue. The primary purpose of ECM is to provide
structural integrity to tissue. However, it is now known that ECM components such as collagen
and laminin also play a crucial role in functional cell signaling through ECM receptors, such as
the integrins, on cell membranes82-85. ECM of load-bearing tissues such as tendons and bones
may persist for long periods of time and have a much lower turnover rate compared to ECM in
skin and mucosal membrane, which are constantly degraded and re-synthesized to accommodate
cellular functions such as cell migration and proliferation. The majority of ECM components in
the heart are secreted by cardiofibroblasts and myofibroblasts. They provide a wide range of
ECM molecules for various functions, including fibrillar collagen and elastin for structural
integrity, laminin and collagen IV for basement membrane maintenance, and proteoglycans for
cell signaling modulation86,87. Moreover, these ECM components are able to transmit external
stimuli to the myocytes and fibroblast and trigger intracellular signaling that can alter cellular
19
functions. Recent work by Ott et al. demonstrated that decellularized native cardiac tissue can
be repopulated with CM and CF and give rise to functional tissue that possesses the
microarchitecture of native tissue88. This suggests that both the composition and organization of
ECM molecules are important for proper tissue development. The following section will cover
the properties of these major ECM molecules.
2.4.2 Collagen
It is the most abundant protein in the body and accounts for 20 to 30% of total body
weight89. Collagen has a simple amino acid sequence consisting of tripeptide repeats. This leads
to a high degree of homology among collagens from different species. For example, type I
collagen from bovine and porcine skin has been demonstrated to be highly compatible with
human cells90. The abundance and compatibility of collagen makes it an ideal matrix substrate
for tissue engineering applications.
The widespread presence of collagen in the ECM is attributed to its unique micro and
macro structure. Collagen is secreted mainly by fibroblasts, smooth muscle cells and EC, and it
is present only in the ECM. The basic primary structure of collagen consists of repeating tri-
peptide units of -Gly-X-Y-, where the amino acid X and Y are often represented by proline and
hydroxyproline90 respectively. The space and flexibility provided by the small hydrogen residue
on glycine is essential for the assembly of collagen polypeptide into more complex, superhelical
secondary structures, while the ring structure of proline stabilizes the superhelix.
Each collagen polypeptide chain forms an extended left-handed helix. Three helices self
assemble to form a trimeric right-handed superhelix91. The triple helix is held together through
hydrogen bonds among the three associated chains. These triple chain molecules comprise the
basic units for larger collagen structures, such as collagen fibrils, which are parallel, staggered
bundles of collagen superhelices. Polypeptides of different collagen isoforms, or α-chain, may
also associate to form heterotrimers; thus, yielding a large variety of collagen fibrils of different
mechanical properties. More than 20 different collagens α-chains have been identified and each
has their unique primary and secondary protein structure. Based on amino acid sequence,
microstructure and functional attributes, collagen can be subdivided into several types. Type I
collagen is the most abundant collagen. Togethertype I, II, III and V collagen share the same
triple-helix structure and are referred to as fibrous forming collagens. In cardiac tissue, over
20
90% of collagen belongs to either type I or III92. Other types of short fibrous collagen (type IX,
XI, XII and XIV) serve as anchor fibers to secure cells and other ECM components to the larger,
continuous collagen fibers. Type IV collagen is found in basement membrane and has unique
globular structures along the triple-helix backbone93 resulting in the formation of sheet-like
structure. Common collagen subtypes and their tissue distributions are listed in Table 2-1.
Table 2-1: Subtypes of collagen in human (adapted from Table 19-5, Molecular Biology of Cell,
4th edition).
There are many characteristics that make collagen a desirable material for tissue
engineering scaffolds. Collagen is an excellent substrate for cell attachment. Certain amino acid
sequences found in collagen are targets for integrin receptor binding. For example, native type I
collagen expresses the tetrapeptide domain -Arg-Gyl-Asp-Ala- (RGDA), which binds the
ubiquitously expressed α1β1 and α1β2 integrin66.
Collagen can be readily remodeled by matrix metalloproteinase (MMP) secreted from
implanted cells or native tissues surrounding the implant. Various experimental and
21
commercially available collagen products such as IntergraTM (Intergra Lifesciences Inc.) are
designed to gradually degrade and be absorbed by peri-implant tissues. As a natural matrix
component in bone, collagen and collagen-elastin matrix with embedded cytokines such as bone
morphogenic protein-2 (BMP-2) have been used as a osteogenic matrix for bone regeneration94-
96. Unfortunately, collagen is a natural agonist for thrombosis via platelet activation97.
Therefore, it is a poor choice for blood contacting applications.
In the adult myocardium, the five most common types of collagen, in descending order
of abundance, are type I, III, IV, V and VI. Collagen plays several essential roles in cardiac
ECM. It mediates the fibrotic response of CF by forming complexes with nonadhesive signaling
protein, such as osteopontin, through the integrin receptors98. Type I collagen is the major
component of the perimysium, a sheath-like connective tissue which surrounds and
interconnects groups of myocytes to help maintain the ordered laminar structure of muscle
bundles in the myocardium99,100. Each laminar layer consists of several fiber bundles stacked
together. The layers are organized in varying directions depending on their trasmural locations.
Fibers located on the epicardial and endocardial surfaces run in a base-apex orientation, whereas
fibers in the mid-wall run in a circumferential orientation101. This structure gives rise to the
anisotropy of the myocardium and also magnifies the shorting of each fiber bundle into gross
ventricular wall thickening that is much greater than anticipated102.
From a biomaterial perspective, collagen is a highly versatile material. It can be easily
extracted from a wide variety of sources and can be processed into many different forms from
sheets to discs, and hydrogel sponge to electrospun mesh. The amino acid residues on the
collagen polypeptide backbone can be readily crosslinked or modified with synthetic polymers
to modulate its mechanical and biological properties. For example, an interpenetrating network
(IPN) made from poloxamine, a synthetic hydrogel, and natural collagen fiber has been shown
to improve the mechanical strength of the resulting composite hydrogel while maintaining
cytocompatability103.
2.4.3 Elastin and Other ECM Proteins
Elastin fibers provide resilience and elasticity to tissues such as blood vessels, skin and
cardiac tissue. In arteries, elastin fibers constitute up to 50% of its dry weight and is secreted
22
mainly by vascular smooth muscle cells104. Moreover, elastin fibers are long-lasting and have a
low turnover rate, thus making it an important structural ECM component.
The basic molecular unit of elastic fibers is elastin. Like collagen, elastin is synthesized
in a soluble pro-protein form known as tropoelsatin. Tropoelastin is bound by a 67 kD
galatolectin chaperone protein on the cell membrane to prevent premature coacervation105.
Tropoelastin consist of alternating Lys-Ala rich hydrophilic and Val-Pro-Gly rich hydrophobic
domains106. Unlike collagen, no proteolytic processing of tropoelstin is required for
elastogenesis. Instead, the lysine/alanine rich domain takes on the α-helix structure, where the
lysyl residues are readily deaminated by extracellular lysyl oxidase to form translysyl
crosslinkages107. On the other hand, hydrophobic domains from adjacent tropoelastin molecules
form loose interpenetrating chain structures that are responsible for the elasticity of elastin
fibers108. Once crosslinked, elastin forms an insoluble elastin fiber.
Besides collagen and elastin, fibronectin (FN) is the next most abundant ECM
component. FN molecules form homodimers and consist of several distinctive functional
domains. FN exists in both insoluble and soluble forms. In its insoluble ECM form, FN
associates with collagen fibers with its collagen-binding domain, while interacting with cell
surface adhesion molecules such as integrin through its Arg-Gly-Asp (RGD) domain. This
bifunctional nature allows FN to serve as a biological crosslinker between cells and the ECM109.
Also, by using integrin as a transmembrane adapter protein, the actin cytoskelekton exerts forces
on extracellular FN fibrils to aid in its polymerization and organization104. Soluble FN circulates
through the blood stream and participates in thrombosis and wound healing.
Another bifunctional ECM component is laminin (LM). Laminin is found associated
with type IV collagen in the basal lamina. Each laminin molecule consists of three polypeptide
subunits (α−, β− and γ− chains) stabilized by disulfide bridges. Like FN, laminin can bind to
both ECM components and cell surface receptors and it is believed to influence cell attachment,
migration and differentiation104.
2.4.4 Synthetic Scaffold and Materials
A wide range of synthetic polymeric materials can be fabricated into scaffolds suitable
for cardiac tissue engineering. These scaffolds can be porous or fibrous in nature. By altering
the polymeric structure, the mechanical properties and biological responses of scaffolds can be
23
fine tuned. Polyglycolic acid (PGA) and its copolymers with polylactic acid (PLA) and
polycaprolactone (PCL) form biodegradable scaffolds that have been used extensively in tissue
engineering applications. Rat neonatal CM seeded on PGA scaffold remain viable and
contractile when cultured in a spinner flask in vitro110. To better control the orientation of cell,
alignment polymer can be electrospun such that fibers are directionally arranged111.
Furthermore, bioactive molecules that enhance cell attachment112 and biodegradability113 can be
incorporated into the polymer structures during synthesis. Therefore, the resulting material can
inherit the mechanical advantages of synthetic polymers while gaining a more natural-like
cellular interface that is more suited for tissue development.
2.5 Fabrication and Culture Platforms for Cardiac Tissue Engineering
The interaction between cells and biomaterials on a molecular level is one of many
factors that will contribute to the functionality of the engineered tissue. To maximize the
regenerative potential of the cells, one must consider the impact of tissue fabrication and culture
strategies on the cells’ behavior.
Most biomaterial-cell combinations are fabricated by either seeding cells onto a pre-
fabricated substrate, or by mixing cells with the biomaterial prior to scaffold casting. Both
methods are currently used in the fabrication of cardiac tissue, and each has its benefits. The use
of pre-formed scaffolds allows the fabrication process to be uncoupled from the cell seeding and
permits the usage of processes that are not compatible with cells. Examples of these processes
include salt leaching, free radical polymerization, and plasma surface modifications. The wide
array of methods available for cell-free scaffold fabrication means that scaffold can have a wider
range of biological and mechanical properties. The disadvantage of this method is that cells can
only be seeded on the surface and must actively migrate into the core of the scaffolds. This may
lead to uneven cell seeding throughout the depth of the scaffold.
One way to improve the uniformity of seeding cells within the scaffold is to combine
cells with compatible hydrogel prior to casting and gelation. The Zimmermann group has
demonstrated that CM embedded in a collagen and Matrigel based hydrogel can be casted into
sheet114, ring115 or pouch116 forms; each with their unique structures and applications.
Nonetheless, cells embedded near the core of the gel still suffer from diffusion limitation and
tend to have low viability. Therefore, recent studies have shifted their focus towards the
24
fabrications of microtissue such as the modular construct approach described within this thesis.
By shrinking the size of the scaffold down to several hundred micrometers in diameter, cells
embedded in the core may receive sufficient nutrients to survive. Kelm et al. have created
microtissues (~100 µm) containing CM in a hang drop culture system that are capable of
supporting core tissue viability117,118.
A variation on the modular scaffold design is cell sheet assembly. Many cells, including
fibroblast and CM, secret their own ECM components and form monolayer sheets when
cultured in vitro. Using a thermoresponsive poly N-Isopropylacrylamide coating to control the
growth of cell sheets, Shimizu et al. were able to cultivate contractile CM sheets and assemble
them into multilayer tissues119. The same group has also stacked sheets of fibroblast and EC
with CM sheets in alternating layers to improve angiogenesis when implanted in a rat model120.
Schematic diagrams of these cardiac tissue engineering strategies can be found in Figure 2-3.
Achieving adequate tissue perfusion and proper ECM architecture are essential for a
culture platform, but they are not enough to ensure the maintenance of phenotype and
functionality of cells 121. It is well known that primary cells can be successfully cultured and
expanded in vitro, but over time they lose cell-specific metabolic functions and cell-cell
interactions, this limiting their ability to organize into native-like, functional microstructures. In
native cardiac muscle, CM and CF are subjected to high level of mechanical stress during each
contraction cycle. Moreover, they are constantly generating and transmitting electrical impulses
over the myocardium. It is known that exposure to subtle mechanical stresses such as tensional
and shear on engineered cardiac tissues induces cell alignment, myosin fiber expression and
microstructure organization in vitro resembling native tissue 122. Electrical field stimulation
elicits a similar response in CM, and the expression of Cx43 corresponds to the direction of cell
alignment. Mechanical and electrical conditioning also results in a stronger contractile force,
although the magnitude is still much smaller than compared to native tissue123,124. These
evidences have illustrated that the delivery of physiologically relevant stimulus is crucial for
cardiac tissue development.
25
Figure 2-3: Schematic diagram of the three major cardiac tissue engineering strategies. Cardiac
construct could be fabricated by seeding functional cells, such as CM and EC, onto pre-
fabricated scaffold. Alternatively, cells could be embedded into hydrogel and molded into the
desired geometry. An emerging technique for fabricating 3-D tissue is to stack 2-D cell sheets
together into a scaffold free constructs.
2.6 Angiogenesis and Cardiac Tissue Vascularization Strategies
The capacity to delivery sufficient nutrients to cells is one of the important determinants
to the viability and functionality of implanted tissue. The challenge in solving the mass transport
problem is obvious, considering that the first generation of successfully commercialized
artificial tissue comprise mainly of thin structures, such as skin grafts and corneas (in the
26
millimeter thickness range), or avascular tissues like cartilage and tendons. There have been
continual clinical demand and interest in fabricating large, 3-dimensional tissues containing
metabolically functional cells. In particular, studies of angiogenesis and vasculogenesis in the
fields of tumor biology and developmental biology have been instrumental in the development
of novel strategies in scaffold vascularization.
Angiogenesis is the sprouting and growth of existing blood vessels into areas of poorly
vascularized tissues. This phenomenon has gained significant interest after tumor researchers
observed the in-growth of new vasculature into developing tumors. The process of angiogenesis
is well characterized and can be described in the following steps125. First, the parent blood
vessel undergoes vasodilation in order to disrupt interendothelial cell junctions. The sub-
endothelium basement membrane is then digested by secreted matrix metalloproteinases
(MMPs), followed by EC mobilization and the formation of a migration front of the new
capillary. The luminal space is formed by the morphological change of EC into tubules like
structure. Maturation of the new vessel is achieved through basement membrane synthesis and
recruitment of pericytes. In large caliber vessels, this pericyte layer is eventually replaced by
smooth muscle cells. On the other hand, the process of vasculogenesis does not require
sprouting from existence blood vessels. Instead, endothelial progenitor cells (EPC) known as
angioblasts begin to aggregate and establish cell-cell contact. The cell mass then gives rise to
nascent endothelial tubes that mature through the recruitment of pericytes. Although
vasculogenesis is the major mechanism of embryonic blood vessel formation, it has
demonstrated that EPC exist in the adult blood stream and may be involved in postnatal blood
vessel formation.
In both angiogenesis and vasculogenesis, the initiating events are driven by the presence
of angiogenic factors, usually in the form of growth factor such as vascular endothelial growth
factor (VEGF), basic fibroblast growth factor (b-FGF), transforming growth factor beta (TGF-β)
and platelet-derived growth factor (PDGF). Chemical species such as nitric oxide (NO) may
also serve as angiogenic factors. Some of the growth factors act directly on EC motility and
proliferation (VEGF and b-FGF). Others act indirectly (TGF-β and PDGF) on supporting cell
types that are involved in vessel maturation. In either case, the spatial and temporal distributions
of angiogenic factors are crucial to the growth rate, degree of branching and quality of new
blood vessels formed. Angiogenic factors are known to function synergistically via receptor
tyrosine kinase mediated downstream signaling pathways crosstalk126.
27
Figure 2-4: Changes in the endothelium during angiogenesis. Cell-cell integration is maintained
by adherens junction proteins (VE-cadherin and PECAM), while the basal surfaces are anchored
to ECM composed of laminin and type IV collagen. In response to stimulus such as VEGF,
adherens junctions between EC are disassembled and EC begin to sprout toward the angiogenic
signals. At the same time, the underlying basement membrane is dissolved by proteolytic
enzymes and a provisional ECM is secreted by underlying cells to facilitate EC migration.
(Adapted from Stupack et al.127, 2004)
Early 3D cardiac scaffold designs relied on hypoxic environment and growth factor
secreted by embedded cells as angiogenic signals114,128,129. The rationale for this approach was
based on what was observed during tumor angiogenesis, where hypoxia induced growth factors
secretions were able to induce sufficient new blood vessel growth to sustain the tumor.
28
However, this method is largely incompatible in tissue engineering application, since the time
frame of angiogenesis is usually measured in weeks, whereas most cells would not survive for
more than a day or two without sufficient perfusion.
One way to enhance the rate of vascularization is to incorporate EC into cardiac scaffolds
with the view that they can help initiate vasculogenesis and secrete growth factor to drive
angiogenesis. Variations of EC seeding strategies include static surface seeding and
homogenous EC embedding into 3D scaffolds. Seeding of EC on scaffolds is often combined
with some form of growth factors delivery (Ang-1, VEGF, b-FGF, TGF-β and PDGF) that are
designed to locally affect the angiogenic kinetics of implanted EC130. Controlled release of
VEGF has been shown to induce EC tubule formation131. A variation to embedding bioactive
molecules is to include naked plasmid DNA encoding VEGF or b-FGF in the scaffold. The idea
is to have seeded cells incorporate the naked plasmid and express the gene of interest thus
delivering the desired angiogenic effect132. However, this method is not widely used due to the
variability of VEGF expression. Besides DNA and growth factor embedded scaffold, transgenic
Chinese hamster ovary (CHO) cells expressing exogenous VEGF or b-FGF have been
encapsulated and co-implanted with cell seeded scaffold126. An obvious advantage of the
cellular protein expression method over bolus injection and controlled polymer release of
growth factor is the longevity and consistency of the growth factors delivered. This is especially
critical considering most growth factors and biological signaling molecules have a short half
life.
The rate of scaffold vascularization depends not only on growth factor and cells, but also
the material and design of the scaffold. Scaffolds made out of natural biomaterials can be easily
remodeled by cell and can also promote cell mobility through integrin binding. Therefore, the
ability of cells to remodel their surrounding may have an effect on their survival by influencing
the speed at which vascularization occurs.
2.7 In vivo models for evaluating engineered tissue
While in vitro assays provide a good starting point for the development of engineered
tissue, it is by no mean a good predictor of their in vivo behavior and performance.
Discrepancies between in vitro and in vivo observations are primarily due to the complex
interaction found in whole animals, including host immune responses, mechanical forces and
29
extracellular matrices remodeling. Aside from species, animal models can be classified
according to their level of immune response toward the implanted tissue. A summary of recent
in vivo studies of engineered cardiac constructs is presented in Table 2-2. The choices of animal
models in these studies reflect the research question being addressed. Their features will be
discussed in the following section.
2.7.1 Allogeneic and immunosuppressed animal model
An allograft is an implant derived from a different individual of the same species. This is
by far the most common type of transplantation in human patients. The immune response
involved in allograft rejection is mediated by histocompatibility antigen on the donor cells, and
the most significant ones are the class I and II major histocompatibility complex (MHC). Class I
MHC is present on all cell types, while class II MHC is expressed on antigen presenting cells
(APC), namely dendritic cells, B cells and macrophages. Variations of MHC antigen between
individuals allow the host’s immune system to recognize self from non-self tissue. In general,
the severity of an allograft immune response is proportional to the divergence between the MHC
antigens.
An immune response can be triggered by MHC antigens via the binding of T cell
receptors and the subsequent activation of T cells. The process of T cell activation is further
regulated by co-stimulatory signaling molecules expressed on APCs. Once activated, T cells can
take on either CD4+ or CD8+ phenotype. T cells expressing CD4 molecule, also known as
helper T cells, function as a mediator of the immune response by promoting the proliferation
and cytotoxicity of macrophages and other T cells. The other type of T cells expressed CD8
molecule are known as cytotoxic T cell, and they are capable of causing cell death by releasing
cytokines or via direct Fas ligand binding with Fas receptors on target cells.
Even though allograft rejection is most prominent in whole organ transplants with intact
vasculature, the likelihood of initiating an immune response in engineered tissue derived from
allogeneic cells is significant enough to raise concerns when trying to achieve a controlled in
vivo model for studying tissue remodeling. In recent years, the development of genetically or
pharmacologically induced immunecompromised animal models allows researchers to
circumvent some of the challenges in host-versus-graft reactions. Genetically altered animals
such as athymic nude rats and severe combined immuno-deficient (SCID) mice lack the specific
30
organ and cell types to effectively mount an adaptive immune response toward non-self tissues.
However, their disadvantage is that their immune system mutation may affect other organ
system and lead to other undesirable phenotypes, such as their increased susceptibility toward
tumor formation and shorter life span.
Another method of creating immunecompromised animal is by using pharmacological
agents to inhibit the activation and proliferation of host T cells and B cells. Immunosuppressants
such as azathioprine and mycophenolate mofetil act as antimetabolites to T and B cells. Others
such as cyclosporine and tacrolimus exert their action by inhibiting calcineurin mediated IL-2
expression by T cells. While long-term side effects commonly seen in human patients are
unlikely to be present during the typical implantation period of 1 to 4 weeks in animal models,
they should be taken into consideration and closely monitored to ensure that the observations are
not affected. Animals that are receiving immunosuppressants generally do not suffer from
aforementioned systemic problems that plague other immunocompromised animals. The uses of
pharmacological agents also allow researchers to titrate the desire amount of immune response
in the final model. This may be a useful way to study the effects that different immune
components have on the outcome of the implants. Finally, the use of immunosuppressant in
allotransplantation represents a model that closely resembles current clinical practice. The
results generated from these types of studies can be more easily translated into bedside
applications.
31
Cel
l typ
e Im
plan
t typ
e A
nim
al M
odel
E
ndpo
int
Key
resu
lts
Neo
nata
l rat
CM
Pe
ri-in
farc
t inj
ectio
n
Rat
- sy
ngen
eic
4 w
eeks
Im
plan
ted
CM
exp
ress
CX
-43
(200
4)4
Neo
nata
l rat
CM
C
olla
gen
+ M
atrig
el ri
ng-
type
EH
T R
at -
syng
enei
c 2
wee
ks
Imm
unos
uppr
esse
nt n
eces
sary
for E
HT
surv
ival
in v
ivo.
EH
T he
avily
va
scua
lrize
d an
d tis
sue
mat
urat
ion
obse
rved
in v
ivo
(200
2)13
3 . N
eona
tal r
at C
M
Alg
inat
e sc
affo
ld
Rat
– a
llogr
aft,
4 w
eeks
Pr
evas
cual
rized
scaf
fold
s stru
ctur
ally
and
ele
ctric
ally
inte
grat
ed in
to
host
myo
card
ium
. Als
o th
icke
n sc
ar ti
ssue
and
pre
vent
s fur
ther
LV
di
latio
n (2
009)
134
Skel
etal
myo
blas
t C
olla
gen
+ M
atrig
el p
atch
ov
er M
I R
at -
syng
enei
c 4
wee
ks
Stro
ng im
mun
e re
spon
se. I
mpr
oved
syst
olic
func
tion,
but
no
indi
catio
n of
impl
ante
d ce
ll su
rviv
al (2
008)
135
hESC
-CM
, em
bryo
nic
FB a
nd
HU
VEC
PLLA
/PLG
A sc
affo
ld
Rat
– a
llogr
aft,
imm
unos
uppr
esse
d
2 w
eeks
So
me
mat
urat
ion
of C
M in
to st
riate
d st
ruct
ure
and
expr
esse
s CX
-43.
Fo
rmat
ion
of d
onor
and
hos
t der
ived
blo
od v
esse
ls (2
010)
136
hESC
der
ived
CM
, FB
and
EC
M
atrix
-fre
e sh
eets
ove
r he
alth
y he
art
Rat
– a
llogr
aft,
athy
mic
hos
t 1
wee
k Fo
rmat
ion
of b
lood
ves
sel-l
ike
stru
ctur
es. M
HC
and
Nkx
-2.5
pos
itive
C
M fo
und
in p
atch
(200
9)13
7 N
eona
tal r
at C
M
Col
lage
n m
esh
into
skel
etal
m
uscl
e po
uch
Rat
– a
llogr
aft,
imm
unos
uppr
esse
d
2,3
and4
w
eeks
C
M su
rviv
ed a
t 4 w
eeks
, but
did
not
form
stria
ted
bund
les.
Hos
t ve
ssel
aro
und
perip
hery
at 4
wee
ks (2
008)
138 .
Neo
nata
l rat
CM
and
H
UV
EC
ECM
-fre
e m
icro
tissu
e in
ject
ed in
to p
eric
ardi
al
cavi
ty
Rat
- al
logr
aft
1 w
eek
Mic
rotis
sue
inte
grat
ed in
to h
ost m
yoca
rdiu
m a
s wel
l as t
issu
e el
onga
tion
by d
ay 7
(200
6)13
9
Neo
nata
l rat
CM
C
olla
gen
+ M
atrig
el b
and-
like
EHT
over
MI
Rat
– a
llogr
aft,
imm
unos
uppr
esse
d
4 w
eeks
Fo
rmat
ion
of h
ighl
y di
ffer
entia
ted
card
iac
mus
cle
over
MI s
car.
Ves
sel s
truct
ures
foun
d w
ithin
EH
T. Im
prov
e sy
stol
ic a
nd d
iast
olic
fu
nctio
n (2
006)
140
Neo
nata
l rat
CM
C
olla
gen
+ M
atrig
el p
ouch
-lik
e EH
T ov
er h
ealth
y he
art
Rat
– a
llogr
aft,
imm
unos
uppr
esse
d
2 w
eeks
A
ggre
gatio
n of
CM
nex
t to
host
myo
card
ium
but
sepa
rate
d by
cel
l-fr
ee g
ap. C
M fo
rm lo
ose
by d
iffer
entia
ted
mus
cle.
Abu
ndan
t CX
43
expr
essi
on (2
007)
116 .
Neo
nata
l rat
CM
C
olla
gen
+ M
atrig
el
inje
ctio
n R
at –
allo
graf
t, im
mun
osup
pres
sed
4 w
eeks
C
M su
rviv
ed a
nd e
xpre
sses
CX
43, b
ut n
o st
riate
d tis
sue
form
atio
n (2
006)
141 .
Rat
CM
and
EC
m
atrix
-fre
e sh
eets
R
at –
allo
graf
t, at
hym
ic h
ost
4 w
eeks
Im
prov
ed v
esse
l den
sity
in c
o-cu
lture
pat
ch14
2 Im
plan
ted
EC in
filtra
ted
into
hos
t myo
card
ium
and
form
ves
sels
(2
008)
142
Tab
le 2
-2: A
sum
mar
y of
rece
nt im
plan
tatio
n st
udie
s usi
ng v
ario
us c
ardi
ac ti
ssue
s eng
inee
ring
stra
tegi
es.
32
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Chapter 3: A modular approach to cardiac tissue engineering Leung BM and Sefton MV Tissue Engineering Part A, 2010 June 29 PMID: 20504074
3.1 Abstract
Functional cardiac tissue was prepared using a modular tissue engineering approach with
the goal of creating vascularized tissue. Rat aortic endothelial cells (RAEC) were seeded onto
sub-millimetre size modules made of type I bovine collagen supplemented with MatrigelTM
(25% v/v) embedded with cardiomyocyte (CM)-enriched neonatal rat heart cells and assembled
into a contractile, macroporous sheet-like construct. Modules (without RAEC) cultured in 10%
bovine serum were more contractile and responsive to external stimulus (lower excitation
threshold, higher maximum capture rate and greater en face fractional area changes) than
modules cultured in 10% fetal bovine serum. Incorporating 25% Matrigel in the matrix reduced
the excitation threshold and increased the fractional area change relative to collagen only
modules (without RAEC). A co-culture medium, containing 10% bovine serum, low Mg2+
(0.814 mM) and normal glucose (5.5 mM) was used to maintain RAEC junction morphology
(VE-cadherin) and CM contractility, although the responsiveness of CM was attenuated with
RAEC on the modules. Macroporous, sheet-like module constructs were assembled by partially
immobilizing a layer of modules in alginate gel until day 8, with or without RAEC. RAEC/CM
module sheets were electrically responsive, however, like modules with RAEC this
responsiveness was attenuated relative to CM only sheets. Muscle bundles co-expressing cardiac
troponin I and connexin 43 were evident near the perimeter of modules and at inter-module
junctions. These results suggest the potential of the modular approach as a platform for building
vascularised cardiac tissue.
50
3.2 Introduction
Congestive heart failure (CHF) is a major cause of death in both the developed and
developing world. Although the pathophysiologies relating to CHF are complex, the condition is
ultimately caused by the weakening of heart muscle tissue resulting in lowered cardiac output.
The decrease in cardiac output causes chronic systemic ischemia that leads to decompensation
responses such as ventricular hypertrophy, increased blood volume, and increased blood
pressure, which in turn increases cardiac load and leads to further damage to cardiac muscle
tissue. As a result, there is considerable interest in fabricating a cardiac tissue substitute using
cardiomyocytes in vitro. The resulting tissue engineered cardiac muscle would be expected, at a
minimum, to possess similar mechanical and functional properties as native cardiac tissue, but
also have the capability to be integrated into native tissue thus preventing long term cardiac
tissue remodeling1.
Attempts to fabricate functional cardiac tissues in vitro have yielded promising results2,3.
A crucial factor in creating viable 3-dimensional tissue in vitro is to achieve adequate perfusion
throughout the tissue. This is especially true for cardiac tissue due to its high metabolic rate and
oxygen demand. With a view to improving engineered tissue perfusion, Radisic et al.4
incorporated oxygen carriers and flow channels into neonatal rat cardiomyocyte (CM) seeded
scaffolds. Aside from nutrients, environmental factors including mechanical5 and electrical6
stimuli are also needed for the proper development of cardiac tissue. Many of these concepts
have been incorporated into recent culture systems7 in which CM are incorporated with other
materials to form functional tissues 8. For example, a 3-dimensional model was used by
Zimmermann et al.5, where they embedded a neonatal rat CM enriched cell population (also
containing fibroblasts and endothelial cells, EC) into a MatrigelTM/collagen gel. In their model,
the EC in culture facilitated angiogenesis and host vasculature coupling. Other 3-dimensional
culture systems rely on cell secreted extracellular matrix (ECM) as the major matrix component.
Kelm et al.9,10 created microtissues containing CM in a hanging drop culture system. The scale
of these microtissues (~100 µm) has been shown to support core tissue viability. In addition,
contractile CM sheets have been cultivated and assembled into multilayer tissues by Shinizu et
al.11. These strategies improve tissue perfusion by promoting either angiogenesis of surrounding
blood vessels, or neovasculogenesis of embedded endothelial cells (part of the CM mixed cell
population) via cytokine and chemokine signaling.
51
The overall goal of this work is to create functional, cardiac tissue using a modular tissue
engineering approach with neonatal rat CM embedded in a collagen/MatrigelTM gel (the module)
and seeded with rat aortic endothelial cells (RAEC). Modules had a post-contraction length of
approximately 500 µm and diameter of 300 µm. When assembled into a construct, the resulting
interconnected channels are perfuseable, mimicking the required vasculature. The small
diameter of each module ensures sufficient nutrient delivery to the centre by diffusion. Modules
may be assembled into larger constructs in vitro followed by implantation, or they may be
deployed in vivo and allowed to be remodeled by host tissue. We hypothesized that the modular
tissue engineering approach 12,13 can be used to produce vascularized cardiac tissue by
assembling modules into a macroporous sheet, where EC in the intermodular space can form
vessel-like structures. This approach results in uniform, scalable and vascularized constructs14
and is being explored elsewhere for its utility in pancreatic islet transplantation and fat
reconstruction.
In work reported here, we have characterized the in vitro phenotype and functional
response of a CM/RAEC co-culture system comprised of both individual modules and modules
in the form of a contractile sheet. We examined the responses of CM-only and RAEC/CM co-
culture modules to external electrical stimulation, as well as the effect of surface coated RAEC
on selected features of the embedded CM. Key aspects of this study was the definition of a
suitable medium for co-culture of CM and EC and a means of preparing sheet-like structures
from the modules.
52
3.3 Materials and Methods
3.3.1 Cardiomyocyte isolation and module fabrication
The protocol for cell isolation was similar to that reported by Radisic et al6. Hearts from
neonatal rat pups (1-2 days old) were harvested and kept in ice cold HBSS (Hanks balanced salt
solution, Gibco, Mississauga, ON). The aortas were trimmed and the remaining heart tissues
were quartered followed by serial digestion with bovine pancreas trypsin (0.6mg/mL in HBSS,
Sigma, Mississauga, ON) and collagenase II (1 mg/mL in HBSS, Worthington Biochemical,
Lakewood, NJ). The extracted cell mixture was pre-plated onto tissue culture polystyrene for 1
hour to enrich its CM content. Harvested cells were maintained in native CM medium consisting
of high glucose (25 mM) DMEM medium (Gibco) supplemented with 10% fetal bovine serum
(FBS; or 10% bovine serum, BS, Sigma), 1% HEPES buffer (10 mM, Gibco), and 1%
penicillin/streptomycin (Gibco). Each CM preparation consisted of cells from an entire litter (10
to 13 neonatal rats) to enhance consistency. Over the course of the study several litters were
used in each set of data. The number of experiments reflects the use of these replicate litters.
The CM enriched cell mixture was loaded onto a Cytospin® slide and stained for
troponin I and prolyl-4-hydroxylase (see below) to identify CM and fibroblasts respectively: the
mixture consisted of 50% cardiomyocytes and 40% fibroblasts. The remaining 10% was
presumed to consist of other cardiac cell types, including EC. The CM-rich cell mixture was
embedded in a type I bovine collagen matrix supplemented with 25% MatrigelTM (v/v, BD
Bioscience), and cast into modules using methods as described previously 15. Viability of
embedded cells was determined using Live/DeadTM assay kit (Molecular Probe, Eugene, OR).
53
3.3.2 Endothelial cell seeding
RAEC were purchased and cultured in an MCDB-131 based native EC medium (VEC
Technologies, Rensselaer, NY), supplemented with 10% FBS. In some experiment, RAEC were
transduced with a lentiviral eGFP vector16 to enable tracking over time (in collaboration with
Dr. J. Medin, Ontario Cancer Institute).
CM-embedded modules were cultured in native CM medium supplemented with 10%
BS for 3 days prior to EC seeding. To coat the surface of modules, 2-3 million RAEC were
suspended in a seeding medium consisting of a 50:50 mixture of native CM (10% BS) and
native EC (10% FBS) medium, and mixed with one mL of CM-embedded modules inside a 15
mL centrifuge tube. The tube was agitated for 30 minutes on a uniaxial rocker at low speed. The
mixture was transferred to a non-tissue culture treated 6-well plate and kept in seeding medium
overnight in order to promote RAEC attachment. The seeding medium was used to transition the
CM and EC from their separate native media to the co-culture medium, described in Table 3-1.
After one day, modules were transferred into a new non-tissue culture treated 6-well plate to
remove any RAEC that did not attach to modules, and the seeding medium was changed to the
co-culture medium. Modules were maintained in this medium until assayed.
Table 3-1: Medium compositions (presumed key differences) and serum types used in the
study.
54
3.3.3 Module sheet
Four days after fabrication, CM embedded modules were assembled into a monolayer
sheet. For RAEC/CM modular sheets, CM embedded modules (3 days after fabrication) were
seeded with RAEC and assembled into sheets the next day. Using a 10 mL serological pipette,
modules, with or without RAEC, were placed over a sterile nylon mesh (pore size = 100 µm,
Millipore, Cork, Ireland) to form a single layer. A dilute alginate solution (1.2% w/w, Sigma)
was dripped over the modular sheet followed by a few drops of 50 µM calcium chloride to
crosslink the alginate. Once the alginate formed a soft gel (approximately 5 to 10 seconds), the
sheet was rinsed in PBS (1.06 mM KH2PO4, 155.17 mM NaCl, 2.97 mM Na2HPO4-7H2O) and
transferred into CM or co-culture medium. These alginate embedded sheets were stimulated (see
below) for 10 days and then the alginate gel was removed prior to assay by brief incubation in
100 µM citrate buffer at room temperature.
3.3.4 Electrical response assessment and field stimulation
CM embedded modules were equilibrated in 1x Tyrode’s salt solution (Sigma) at 37oC
for 1 hour prior to measurement. Electrical responses of modules were assessed using custom
made testing chambers. Modules were paced between a pair of graphite electrodes kept 1 cm
apart by a polycarbonate bracket. Using a 6-well plate made from tissue culture polystyrene,
modules and electrodes were placed in an individual well with 1.5 mL of Tyrode’s salt solution,
which was enough to partially submerge the electrodes. The small chamber volume ensured that
most of the modules were located between the electrodes.
Module sheets were assayed using a PDMS/glass testing chamber. A pair of graphite
electrodes were kept 1 cm apart by a PDMS bracket and placed over a layer of PDMS poured
into a 60mm glass Petri dish. This allowed the module sheets to be fixed in position relative to
the electrode.
For both systems, a signal generator (S88X stimulator, Grass Technologies, Brossard,
QC) connected to the electrodes provided the desired electrical stimulation (biphasic, 2 ms
square pulse, up to 12 V at various frequencies). The samples were kept at 37oC during the
experiment using a solid-state microscope stage heater. Contractions of modules and module
sheets were recorded with a CCD camera (Hitachi Denshi, model KP-M1U) on an inverted
55
phase contrast microscope (Olympus CX2). The image sequences were analyzed using ImageJ
(NIH, version 1.38x).
Excitation threshold (ET) was defined as the minimum voltage required for synchronous
contraction at 1 Hz. Maximum capture rate (MCR) was defined as the maximum rate of
synchronous contraction under an electric field equivalent to twice the ET. The contractility of
modules was defined as the fractional en face area change during one contraction cycle using
frame-by-frame video analysis in ImageJ.
Module sheets were electrically conditioned prior to assay. Briefly, CM-embedded
modules were maintained in native CM medium with 10% BS for 3 days. Then the modules
were fabricated into sheets (see above) and transferred into a well, fitted with a pair of graphite
electrodes as above, and stimulated continuously with biphasic pulses at 5 V/cm and 1 Hz for 10
days. Medium was supplemented with 10 µM of ascorbic acid and changed daily to minimize
the effect of reactive oxygen species generated at the electrodes. Individual modules were not
electrically conditioned because they could not be fixed in position relative to the electrical field
over the course of an experiment.
3.3.5 Immunofluorescence staining
Freshly harvested, CM enriched cell mixtures were analyzed by cytospin. Slides were
stained with rabbit anti-rat troponin I (Santa Cruz Biotechnology, Santa Cruz, CA) and mouse
anti-rat prolyl 4-hydroxylase (Sigma, Oakville, ON) antibodies to identify cardiomyocytes and
fibroblasts, respectively.
RAEC coverage on modules was determined by whole mount immunoflourescence
staining of goat anti-rat VE-cadherin primary antibody (Santa Cruz Biotechnology, Santa Cruz,
CA) followed by rabbit anti-goat IgG AlexaFluorTM 488 secondary antibody (Molecular Probes,
Eugene, OR). Modules were fixed in 4% paraformaldehyde solution and permeabilized using
0.2% Triton-X100 solution. On immunohistochemical sections, CMs were stained using rabbit
anti-rat troponin I (Santa Cruz Biotechnology, Santa Cruz, CA) followed by goat anti-rabbit IgG
secondary antibodies conjugated with AlexaFluorTM 488 (Molecular Probes, Eugene, OR).
Cardiac fibroblasts were stained with mouse anti-rat vimentin antibody (Sigma, Oakville, ON)
followed by goat anti-mouse IgG AlexaFluor 568 (Molecular Probes, Eugene, OR). In sections
that were triple stained (troponin I, vimentin and connexin-43), connexin-43 was stained prior to
immunofluoresence staining using rabbit anti-rat Cx43 antibody (Chemicon) coupled by a three-
56
step IHC chromogeninc method. In some experiments, surface seeded RAEC were tracked by
eGFP, where cell nuclei were counterstained with Hoechst nuclear stain (Sigma, Oakville, ON).
Samples were visualized using a Zeiss LSM510 confocal microscope.
3.3.6 Statistical analysis
Statistical analysis including calculations of standard error of means and mean
comparisons (t-test or one-way ANOVA) were performed using Prism (Version 5.0, Graphpad
Software, La Jolla, CA)
57
3.4 Results
3.4.1 Contractility of CM embedded modules
Cardiomyocytes embedded in modules remained viable for at least 14 days in either type
1 collagen or type 1 collagen supplemented with 25% MatrigelTM. However, higher viability
was obtained with an initial cell seeding of 107 cells/mL compared to 106 cells/mL (Figure 3-1).
Modules without RAEC were cultured for 7 days in cardiomyocyte medium containing either
10% FBS or 10% BS and then assayed for electrical responsiveness. In both cases, modules
displayed synchronous contractions corresponding to the external stimulation frequency (Figure
3-2). A small proportion of the modules (< 10%) contracted spontaneously even in the absence
of external electrical stimulation and continued to contract when stimulated. Modules cultured
in 10% BS had a lower excitation threshold but similar maximum capture rate compared to
modules cultured in 10% FBS (Figure 3-3A). Moreover, modules cultured in 10% BS had better
defined peaks with higher maximum contraction amplitudes, compared to modules cultured in
10% FBS; a representative sequence of a single module (from a group of >30) is shown in
Figure 3-3B. Histological sections of modules cultured in 10% BS showed higher troponin I
expression and elongated cell morphology compared to modules cultured in 10% FBS (Figure
3-3C). These results suggest that bovine serum enhanced the electrical responsiveness and
contractile performance of CM-embedded modules, through an effect on the expression or
organization of the contractile apparatus within the cells. For this reason, all subsequent
experiments were carried out in 10% BS supplemented medium.
The effect of extracellular matrix on the function of embedded CM (10% BS
supplemented medium) was tested by embedding CM in either collagen (type 1) or collagen
supplemented with MatrigelTM; the latter choice was based on the work of Zimmerman et al5.
CM embedded in MatrigelTM supplemented matrix exhibited lower excitation thresholds (Figure
3-4A, p<0.05) and higher contractile amplitudes (Figure 3-4C) compared to CM embedded in
collagen only modules. However, the maximum capture rates were found to be similar for the
two groups (Figure 3-4B).
58
Figure 3-1: Viability (live/dead assay) of cardiomyocytes embedded in modules, cultured in
10% BS, at 14 days post fabrication. Matrix composition did not affect embedded cell viability,
while increasing seeding density from 106 to 107 cell/mL dramatically improved viability.
(Magnification = 100x)
59
Figure 3-2: Fractional area change of a single module when stimulated by external signal. CM-
embedded modules (2 x 107 cell/mL collagen supplemented with 25% v/v MatrigelTM) were
cultured in 10% BS medium for 7 days prior to assay. At a voltage twice the excitation
threshold, modules displayed synchronous contraction corresponding to the external signal
frequency: 1 Hz from 0 to 7 sec, 2 Hz from 7 to 13 sec, and 3 Hz from 13 to 17 sec.
60
Figure 3-3: (A) Excitation thresholds (ET) and maximum capture rates (MCR) of CM (2 x 107
cell/mL) modules (collagen with 25% MatrigelTM) cultured in medium containing 10% FBS
(solid) or 10% BS (hatched) for 7 days post fabrication. Maintaining modules in medium with
10% BS decreased ET (*, p<0.05, n=3), but did not affect MCR. (B) Fractional area change of
single modules (taken from an aliquot of >30) on day 7. Modules cultured in 10% BS or 10%
61
FBS were stimulated at 200% excitation threshold voltage (2ms, 1 Hz) and recorded on video.
Fractional area change is normalized against a reference frame in each sample. (C) CM cultured
in 10% BS expressed more cardiac troponin I (green) and assumed a more elongated
morphology than those cultured in 10% FBS when examined using immunofluorescent
microscopy.
Figure 3-4: At day 7 post fabrication, CM (2 x 107 cell/mL) embedded in MatrigelTM (25% v/v)
supplemented collagen modules, cultured in 10% BS, exhibited a lower excitation threshold (A,
*, p<0.05, n=3) and higher contractility (C, single modules taken from an aliquot of >30)
compared to collagen only control. However, both collagen only and collagen+MatrigelTM
modules had similar maximum capture rates (B).
62
3.4.2 Behavior of RAEC seeded CM embedded modules
Primary RAEC (2 x 107 cell/mL) were seeded onto modules embedded with CM 3 days
after fabrication to form endothelialized modules; approximately 25% of the module surface
was covered at initial seeding. After 24 hours in a seeding medium consisting of a 50:50 mixture
of the two native media, the RAEC/CM modules were placed in a co-culture medium that had a
lower glucose concentration (5.5 mM) than the native CM medium and a much lower Mg2+
concentration than native EC medium (Table 3-1). These factors, among other perhaps less
crucial medium differences, helped maintain the contractility of embedded CM and promoted
EC adherent junction formation. After 14 days in co-culture medium, the surface of the modules
were almost completely covered by RAEC with well-developed adherent junctions, confirmed
by junction localized VE-cadherin staining (Figure 3-5A), similar to that seen with RAEC
seeded on empty collagen modules and cultured in native EC medium. In contrast, RAEC
seeded on modules cultured in native cardiomyocyte medium showed no morphologically
distinguishable junction features (Figure 3-5B).
When stimulated by an external electric field, endothelialized modules were able to
contract synchronously, but they had a higher excitation threshold (p<0.001) and lower
maximum capture rate (p<0.05) compared to CM-only modules (Figure 3-5C). In fact their
electrical response profile resembled that of CM-only modules cultured in 10% FBS. However,
both endothelialized modules and CM-only modules had similar contraction amplitudes (Figure
3-5D).
63
Figure 3-5: (A) Immunofluorescence staining for VE-Cadherin on RAEC/CM modules after 14
days in co-culture medium indicated the presence of a confluent layer of RAEC. (B) In contrast,
RAEC seeded on CM modules cultured in high glucose native CM medium displayed
disorganized junctions. (C) Surface coated RAEC attenuated the electrical responsiveness of
modules, as seen by an increase in excitation threshold (***, p<0.0001, n=3) and decrease in
maximum capture rate (*, p<0.05, n=3). (D) With external field stimulation, CM-embedded
modules coated with RAEC displayed similar contractility when compared to CM-only modules
(single modules taken from an aliquot of >30). Contractility was assessed by comparing the en
face fractional area change.
64
3.4.3 Module sheets
The modular approach mimics the multilayered organization of cardiac muscle by first
fusing sub-millimeter modules into a sheet-like construct with a view to subsequently stacking
these “sheets” together to yield a perfuseable, macroporous, endothelialized three dimensional
tissue. One of the challenges of assembling multiple cell embedded modules into a sheet is to
control the rate and degree of agglomeration caused by remodeling. Modules embedded with
contractile cell types, such as smooth muscle cells17 and CM, tend to clump and fuse into a large
cell mass spontaneously under static conditions. The size of these non-porous aggregates is
expected to impair diffusion and ultimately lead to necrosis of the core. To minimize this, the
modules were partially embedded in a thin layer of alginate gel, a non cell adhering matrix, to
prevent the spaces between modules from collapsing, while at the same time allowing a limited
amount of intermodule fusion to occur (Figure 3-6A). In as little as five days, a macroporous
module sheet was produced, and then the alginate was dissolved, releasing the fragile, yet
physically stable sheet from the supporting nylon mesh (Figure 3-6B).
Both CM only as well as RAEC/CM module sheets were characterized. At 4 days post
fabrication, modules were assembled into sheets and stimulated for an additional 10 days.
RAEC/CM modules were made by seeding eGFP tranduced RAEC onto modules one day prior
to sheet assembly (i.e., on day 3). After 10 days of stimulation, RAEC were tracked and found
to be evenly distributed over the surface of module sheets (Figure 3-6D). Moreover, the
attachment density was enhanced by removing the alginate gel and reseeding the sheet with
more RAEC (Figure 3-6E), after 10 days of stimulation. When placed in an electrical field
(Figure 3-6C), both RAEC/CM and CM-only module sheets contracted synchronously.
RAEC/CM sheets exhibited a higher excitation threshold (p<0.01), but a roughly similar
maximum capture rate compared to CM-only sheets (Figure 3-6C). When RAEC/CM sheets
were not stimulated for the 10 days of culture, they did not contract when placed in an electric
field.
Module sheets with or without RAEC were stained for cardiac troponin I, a protein
subunit associated with the troponin-tropomyosin complex in cardiac muscle. Troponin I
positive cells were found throughout the modules. However, most of the muscle bundle-like
structures were found at the perimeter of the modules or near the inter-module junctions (Figure
3-7). Most of these structures were no more than 5 cell layers thick and covered 12 ± 2% (n = 4)
65
of the cross sectional area in both RAEC/CM or CM-only sheets (Figure 3-7B). When triple-
stained with vimentin, troponin I and connexin-43, we saw a fibroblast rich layer at the
periphery of the modules (Figure 3-8A). We also found connexin-43 co-localized with troponin
I positive structures along the perimeter of modules and inter-modular space, just below the
fibroblast layer (Figure 3-8C, D).
66
Figure 3-6A: (A) Schematic diagram of the testing chamber used to condition module sheets.
Modules were partially embedded in alginate gel on top of a nylon mesh and conditioned
electrically for 10 days.
67
Figure 3-6(B-E): (B) After 10 days of continuous stimulation, macroporous RAEC-CM module
sheets were removed from the alginate and nylon mesh and assessed by field stimulation. (C)
Excitation thresholds and maximum capture rates of RAEC/CM sheets and CM-only sheets;
excitation threshold was significantly higher in the presence of RAEC (**, p<0.001, n=3). (D)
eGFP transfected RAEC were uniformly distributed, at a relatively low density, over RAEC/CM
modular sheets, 10 days after stimulation. (E), RAEC/CM sheets were reseeded with 3 x106
RAEC, using the same protocol as in the first seeding, after removing the alginate gel. With this
protocol, RAEC surface coverage was increased.
68
Figure 3-7: (A) Immunofluorescence staining of cardiac troponin I (green) shows a high density
of CM near the perimeter of modules and at inter-module junctions (dash lines indicate edges of
modules) in a module sheet after 10 days of stimulation. (B) At higher magnification, the CM
appeared to form muscle like bundles, similar to that seen in native cardiac tissue, but such
bundles constituted only ~12% of the field, shown in A.
69
Figure 3-8: (A-B) Triple-stained (vimentin, troponin I and connexin-43) modular sheets at after
10 days of stimulation indicate that surfaces of module sheets were covered by a layer of
fibroblasts with an underlying layer of CM. (C-D) Connexin-43 positive structures were also
observed. (E-F) Co-localization of connexin-43 and troponin I was confirmed by overlaying
high magnification images suggesting the CM were electrically coupled.
70
3.5 Discussion
We have characterized sub-millimeter sized modules containing cardiomyocytes and
endothelial cells and examined the feasibility of building larger constructs with this modular
approach. We focus here on characterizing the individual modules and the assembled sheet-like
structures. Future reports will describe the fate of these modules and sheets in cardiac infarct
models.
3.5.1 Electrical characterization
CM-embedded modules responded to external field stimulation after 7 days in culture.
Addition of MatrigelTM (25% v/v) to the collagen matrix lowered the excitation threshold and
improved the contractility of the modules, but did not affect the maximum capture rate. This
benefit is attributed to the presence of basal lamina proteins that are found in MatrigelTM,
including laminin and collagen type IV, as well as the various growth factors that are found in
MartigelTM. The presence of MatrigelTM is presumed to provide an extracellular matrix
environment that is more similar to the native myocardium than collagen type I alone, resulting
in improved cell organization and tissue remodeling6. For further development of this approach,
an alternative is required since the tumour derived Matrigel has limited utility in clinical
applications.
Culturing modules in 10% BS instead of 10% FBS also resulted in improved electrical
performance and contractility, and this was accompanied by an increase in troponin I expression
and cell elongation. This observation is consistent with that of Schwarzkopf et al.18 who showed
that medium supplemented with adult serum (same species) promoted viability and maturation
of cardiomyocytes compared to fetal calf serum. While we did not investigate the effect of adult
rat serum on rat CM, we saw a similar improvement in CM maturation with adult bovine serum
in comparison to fetal bovine serum. It was beyond the scope of this study to elucidate the
underlying mechanism. Similarly, further optimizations of the matrix or the medium are left for
the future although the value of this for rat CM or EC may not be warranted.
We acknowledge that the current method for assessing electrical performance is highly
dependent on the materials used and the geometry of the testing chamber, and therefore cannot
be viewed as a true analog for representing in vivo electrical performance. Of particular
71
importance is the inability to assess regional differences in contractility and whether action
potentials propagated throughout the thickness of the construct or just merely on the surface.
Nevertheless, the current method allowed us to compare the overall effects exerted by different
environmental signals and enabled us to optimum culture conditions.
3.5.2 Co-culture model
One of the main objectives of this study was to develop a means of maintaining a
functional RAEC layer on the surface of contractile CM-embedded modules. By adjusting the
composition of the medium (Table 1) and lowering the glucose level to that of native EC
medium, primary RAEC remained viable and showed typical VE-cadherin expression after 14
days, suggesting the endothelial cells were in a quiescent state19 in the co-culture medium.
Using the same medium we were also able to generate viable RAEC/CM co-culture modular
sheets.
To our knowledge, this is the first study to investigate the effects of surface coated
endothelial cells on the electrical performance and responsiveness of underlying CM in a
modular construct. A 24 hour transition period and seeding medium (a 50:50 mixture of the
native media) was used to enable RAEC seeding and coverage. Without this seeding medium,
RAEC attachment and coverage was poor. We found that the presence of a quiescent RAEC
layer in the co-culture medium, on either modules or modular sheets, permitted the contractile
response of embedded CM, provided the Mg2+ concentration was at the low level of CM
medium and not the high value of the native EC medium. However, the RAEC reduced the
module and module sheet electrical responsiveness, manifested as an increase in excitation
threshold and decrease in maximum capture rate. These changes in electrical performance were
not seen when CM-only modules were cultured in the same co-culture medium. We believe that
these changes were most likely caused by the presence of the RAEC layer which may have
changed the surface conductance of the modules. Such changes would cause the excitation
threshold to increase compared to CM-only modules. Differences in membrane protein
expression (e.g. connexin proteins), may also attenuate the propagation of electrical impulses
from the surface to the underlying CM, resulting in a lower maximum capture rate.
Judging from VE-cadherin staining, the native cardiac EC embedded in the modules did
not form vessel-like structures, nor did they appear to interact with surface seeded RAEC.
72
However, these cardiac EC may still play an important role in CM maturation, as demonstrated
in other multi-culture cardiac constructs20-22. For example, Iyer et al. showed the presence of
fibroblasts and EC in cardiac tissue constructs enhanced CM elongation and excitability23,
possibly by spatially orienting CM into muscle-like bundles.
Our initial aim was to use the RAEC/CM modules as a vehicle to uniformly deliver
viable EC through out an assembled construct containing multiple modules. In a separate study
involving RAEC modules (without embedded cells) implanted in a rat omental pouch, we
observed that RAEC migrate from the modules and form chimeric (host-donor vessels)
functional blood vessels in the intermodular space after 21 days24. We expect the RAEC on the
co-culture modules to migrate and form vessels in a similar fashion, when implanted. Plans to
evaluate the modular construct in a rat implant model are expected to reveal the fate and
remodeling process of RAEC/CM modules in vivo; how this then changes the in vivo electrical
properties and morphology of CM remains to be defined.
3.5.3 Sheet formation
A major challenge in creating clinically relevant, thick (1 to 10 mm) cardiac tissues in
vitro is the lack of tissue vasculature. Specifically, the formation of blood vessels, both host and
graft derived, can be quite slow and insufficient to meet the high nutrient demands of
cardiomyocytes. There are several approaches designed to circumvent this limitation, including
the use of oxygen carriers in culture medium25, incorporating vessel-like flow channels in
preformed scaffolds26, and preloading scaffolds with various growth factors and/or functional
cell types that promotes angiogenesis27,28.
We devised a method to produce a sheet-like structure from an assemblage of modules.
A thin layer of modules was laid onto a nylon mesh and partially immobilized in alginate gel. At
least some of the CM in these modules matured into muscle-like bundles and formed a
physically integrated sheet capable of synchronous contraction when stimulated. Moreover, the
CM at inter-modular spaces expressed connexin-43, suggesting that the module sheet was
electrically coupled. However, these muscle-like bundles, as observed on histological sections,
are relatively thin and only occupy a fraction of the total volume of the module. The lack of
troponin I positive CM near the core of the module suggests that the CM may have been
crowded out by the faster growing fibroblasts. However this may also be the result of active CM
73
migration and maturation. Further studies at different time points are needed to reveal the
mechanism of the remodeling process. Since we assume that these bundles are responsible for
generating the contractile forces, it is crucial to increase the proportion of these bundles for the
modular construct to produce physiologically relevant force. For instance, in future
implementations it may be beneficial to reduce the diameter of modules so that the surface to
volume ratio is increased, thus increasing the contractile force on a per module basis. It is also
conceivable that over a longer incubation period in vitro or when implanted in vivo, more CM
may migrate to the surface thus increasing the thickness of this layer. These issues will be
explored in future studies.
In recent years, there has been interest in building cardiac constructs using small
functional sub-units: e.g., micro-tissues9, cardiac sheets11 or other components29. In the approach
most conceptually similar to modules, Kelm et al. demonstrated that human umbilical vein
endothelial cell (HUVEC) coated micro-tissue (a “spheroid” without collagen gel) with
embedded CM were able to agglomerate to form a patch with functional HUVEC lined
capillaries10,30. In all cases, the goal is to create uniform, scalable, vascularized and functional
cardiac tissue and the merits of one approach relative to the other will be determined by the
success of meeting these goals based on in vivo performance. Of particular significance in this
work is that co-culture conditions that maintained both CM function and RAEC junction
morphology were devised. We found, for example, that RAEC/CM sheets that were not
stimulated during culture did not contract when subject to external field stimulation at the time
of analysis (after 10 days of stimulation).
We chose to embed modules in alginate not only because of its ease of application and
removal by changing local calcium concentration, but also because it is cell compatible and non
cell adherent. A single-module thickness across the sheet ensured that sufficient nutrients could
diffuse into the core of each module, at least in vitro. These macroporous module sheets will
serve as intermediate components for eventually building thicker tissues in a scalable fashion,
where the tissue maturation process can be monitored. When the modules were assembled into
sheets, there was a small decrease in the electrical responsiveness relative to modules alone.
This is likely the result of differing geometries of the stimulation apparatus resulting in
differences in the field perceived by the CM. It is also conceivable that intermodular interactions
of surface seeded RAEC may have lead to a change in tissue conductance.
Fabrication of functional cardiac tissue in vitro depends on many parameters, including
medium composition, extracellular matrix, geometry and cell seeding sequence. Native
74
myocardium consists of cardiomyocytes that form mechanically and electrically coupled
bundles. These bundles are encased in ECM secreted by cardiac fibroblasts, and maintained by
capillaries lined with endothelial cells. The three cells types exist in their respective
microenvironments. The RAEC/CM co-culture system described in this study consists of more
than two cell types, since the primary CM enriched cell population contained a significant
quantity of fibroblasts and cardiac EC. While in this study, the electrical responsiveness of CM
and the adherent junction morphology of RAEC were chosen as the primary endpoints, we must
also consider the contribution of cardiac fibroblasts and EC to these measurements. The data
presented here suggest that cardiac fibroblasts actively segregate and migrate to the surface of
the modular sheet, and potentially play an important role in the formation and remodeling of the
modular sheet since the spatio-temporal relationships between CM and the various supporting
cell types has been shown to enhance CM maturation and the overall functionality of
construct23,31. Therefore, aside from being a viable platform for in vitro tissue formation, the
module system may also serve as a cell delivery vehicle that is capable of providing a supportive
environment to multiple cell types in a pre-determined seeding ratio and geometry.
3.6 Conclusion
We have created rat aortic endothelial cell coated, neonatal rat cardiomyocyte embedded
modules as well as module sheets. In both cases, embedded cells formed troponin I positive,
native muscle-like structures (albeit to a limited extent), and contracted with external field
stimulation. In the presence of endothelial cells in an appropriate co-culture medium, the
construct became less responsive, although it continued to be excitable and contractile. While an
in vivo study will be needed to elucidate the fate of these modules, this study has demonstrated
the feasibility of creating functional cardiac tissue using the modular approach.
75
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Chapter 4: Fate of modular cardiac constructs in vivo
4.1 Abstract
Cardiac tissue engineering holds the promise of restoring cardiac function by replacing
scar tissue with functional cells. The modular tissue engineering approach has been
demonstrated in a previous study to be a useful method for generating functional cardiac tissue
in vitro. However the effects of host immune response and remodeling on modular cardiac
tissues remain unclear. In this study, the fates of modular cardiac tissues in vivo were examined
in two implantation strategies based on a syngeneic animal model. CM-only or co-culture (CM
and EC) modules were either injected into the peri-infarct zone of the heart, or fabricated into
patch form and implanted over a right ventricular free wall defect. After 2 weeks (3 weeks for
patch implants), donor EC developed into blood vessels-like structures, which appeared
functional and connected with host vasculature. However, no donor CM was found within the
implant sites. Also, host immune cells including macrophages and T-cells infiltrated extensively
into implantation site in both models. To lessen host immune response, MatrigelTM was omitted
from matrix and modules were rinsed with serum-free medium prior to implantation. The
removal of xenoprotein attenuated host immune cell infiltration, while donor EC appeared
unaffected and formed functional blood vessels. Co-culture implants had a higher overall vessel
density compared to CM-only implants, but only in the absence of MatrigelTM. Most
importantly, donor CM matured into striated muscle-like structures throughout MatrigelTM-free
implants. In this study we have demonstrated that modular cardiac tissues can be implanted into
the myocardium in multiple configurations and are able to develop into native-like structures in
vivo. The results also suggest that host immune response play a crucial role in the survival and
maturation of implanted modular cardiac tissue.
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4.2 Introduction
Myocardial infarction (MI) is caused by the interruption of blood supply to the heart and
can lead to massive cell death. The extracellular matrix and cell debris that are left behind
undergo extensive remodeling mediated by immune cells and resident myofibroblasts. The
inflammatory region eventually stabilizes into collagen-rich, acellular and non-contractile scar
tissue. The scar tissue reduces the overall contractile force of the heart, and at the same time
their inelastic nature increases overall cardiac load. Therefore there is a need to develop
therapeutic strategies that aim to replace scar tissue with functional tissue and ultimately restore
cardiac function.
Current research focuses on two major strategies to achieve this goal. The first is by
injecting cells, either differentiated myocytes1-4 or undifferentiated cardiac progenitor cells
(embryonic5,6 or mesenchymal7-9 stem cells) into or around the infracted region in an attempt to
repopulate the site with functional cells. The second strategy aims to replace the scar tissue with
engineered cardiac muscles consisting of myocytes (CM10-12 or skeletal myoblast13,14) and other
supporting cells types, including EC and fibroblast15,16. One of the obstacles in these strategies is
low cell survival caused by a combination of poor cell engraftment and lack of functional
vasculature within the implant sites. A novel vascularization technique was demonstrated by our
laboratory when EC seeded collagen modules formed functional blood vessels in a rat omental
pouch model after 21 days17. It is proposed that a similar method can create a scalable cardiac
construct with uniformly distributed EC that may accelerate graft vascularization in vivo.
The goal of this study is to devise methods to implant the modular cardiac construct and
to assess their fates in vivo using a syngeneic rat model. Two implantation methods were
investigated. In the first method, macroporous modular sheets containing CM and EC were
sandwiched between two pieces of commercially available porous gelatin foam (GelFoam) to
create an implantable patch. This patch was implanted over a transmural defect on the right
ventricular free wall of a healthy Lewis rat heart for 2 weeks. Using this model we were able to
elucidate the biological interaction, such as ECM remodeling, cell maturation and tissue
integration between implanted cardiac modular construct and host tissue. The second method
investigated the delivery of modules into the peri-infarct zone via intracardial injection.
Modules were injected (CM-only or CM/EC co-culture) into the peri-infarct zone of an acute MI
in a Lewis rat. Using this approach we expected to see better cell engraftment and improved
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viability compared to matrix-free cell injection therapy. We also expected that CM-embedded
modules implanted into the rat heart would develop more native-like morphology compared to
CM-modules cultured in vitro.
4.3 Materials and methods
4.3.1 Isolation of rat primary CM and EC
Primary CM and EC were isolated from green fluorescent protein (GFP) transgenic
neonatal Lewis rats (Lewis-Tg [EGFP] F455) obtained from the Rat Resources and Research
Center (RRRC, University of Missouri, Columbia, MO). The aorta and atrium were trimmed
away and the remaining tissues were cut into small pieces. The tissues were incubated in bovine
trypsin (0.6mg/mL in HBSS) overnight at 4oC, followed by serial digestion in collagenase type
II (1 mg/mL) at 37oC. CM-rich cell suspensions were pre-plated onto TCPS flasks for 1 hour to
enrich the CM content. The CM enriched cell mixture was loaded onto a Cytospin® slide and
stained for troponin I and prolyl-4-hydroxylase (see below) to identify CM and fibroblasts
respectively: the mixture consisted of 50% cardiomyocytes and 40% fibroblasts. The remaining
10% was presumed to consist of other cardiac cell types, including EC.
Primary rat cardiac EC were isolated from the cells attached on to the TCPS flasks in the
previous step. The CM-rich cell suspensions were replaced with native EC-medium (MCDM-
131 with 10% FBS). After 2 days, attached cells were trypinized and the EC were labeled with
mouse anti-rat CD31 FITC conjugated antibody followed by anti-FITC magnetic beads
(Mylteny Biotech), and labeled cells were separated using a magnetic cell sorting column.
Isolated EC were maintained with native EC medium in fibronectin coated flasks. EC phenotype
was confirmed by Dil-Ac-LDL uptake assay (10µg/mL Dil-Ac-LDL, 4 hours at 37oC).
4.3.2 Modular cardiac patch model
Modular cardiac patch was prepared by placing CM-embedded modular sheets within a
GelFoam pocket. CM embedded modular sheets were fabricated as described in Chapter 3.3.3.
The sheets were stimulated continuously for 5 days (1 Hz, 5V, 2ms, biphasic square pulse). At
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the end of day 5, three modular sheets were placed on a piece of GelFoam, measuring 2mm
thick, 6mm wide and 12mm long. The GelFoam was then folded in half length-wise and sutured
at the open corners to create an implantable patch. For CM/EC co-culture patches, CM modular
sheets were seeded with EC (passage 3-5) at the end of the five-day stimulation period, and the
sheets were culture for 1 day in co-culture medium before being fabricated into patches similar
to CM-only sheets.
The modular patch was implanted over a transmural defect on the right ventricular free
wall using methods described the Li group18. Wildtype Lewis rats (12-20 weeks old) were
sedated, intubated and ventilated (Harvard Apparatus, Holliston, MA, USA), and maintained
with 1.5-2.5% isoflurane. The heart was visualized through a median sternotomy. A 5-6 mm
diameter section of the right ventricular (RV) free wall was isolated with a purse-string suture
(with 7-0 polypropylene monofilament suture) to create a “neck” of the bulging RV free wall, as
described before18-20. A 16-guage angiocatheter was passed over the stitch to create a tourniquet
to prevent bleeding when a full thickness circular segment of the RV free wall was removed.
Patches containing modular sheets were laid on top of the resected ventricular wall and then
sutured to the margin of the defect with a purse-string stitch with 7-0 polypropylene
monofilament stitch using an over-and-over running suture technique to close the defect in the
RV free wall. Then the tourniquet was released and the purse-string stitch was removed. The
chest wall was closed with 5-0 silk suture. Patches were explanted after 14 days and fixed in 4%
paraforamldehyde for histological analysis.
4.3.3 Peri-infarct module injection model
Small diameter CM-only modules and CM/EC co-culture modules were cast in
polyethylene tubes with an inside diameter of 540µm and cut to 1mm in length. The mean post
contraction diameter was approximately 220µm. Other than the difference in diameter, these
modules were fabricated in the same way as the ones used for making the modular patch. CM-
only modules were maintained in native CM-medium until implantation at day 7. Co-culture
modules were made by seeding EC (passage 3-5) onto CM embedded modules at day 3 post-
fabrication and kept in co-culture medium from day 3 to 7 in culture before implantation.
MatrigelTM-free modules were made with type I bovine collagen only. Prior to implantation,
they were rinsed and incubated for 1 hour in defined serum-free medium as describe by Kessler-
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Icekson et al.21. Briefly, the medium was composed of 1:1 mixture of Ham’s F12 and DMEM
supplemented with 10 mM HEPES at pH 7.3, additional 1mg/mL glucose, 0.1 µM
hydrocortisone, 25 µg/mL insulin, 25 µg/mL transferrin, 1 mg/mL fetuin (all from Sigma).
An acute MI rat model was created by ligating the left descending coronary artery of
wild type Lewis rats (12-20 weeks old) for 7 days prior to modules implantation. Small diameter
modules were injected into the peri-infarct zone using a 28G needle (3 sites per animal, equal
distance apart). Approximately 50 modules were injected per site for a total of 150 modules per
animal. Animals were sacrificed at 21 days post injection and the hearts were fixed in 4%
paraformaldehyde for histological analysis.
4.3.4 Immunostaining and Histological Analysis
Modules cultured in vitro for 7 days were fixed with 4% paraformaldehyde and
permeabilized using 0.2% Triton-X100 solution. Samples were then triple stained for troponin I,
F-actin and cell nucleus. Embedded CM were identified using rabbit anti-rat troponin I
(Chemicon) antibody followed by Alexa FluorTM 488 goat anti-rabbit secondary antibody
(Molecular Probes). Alexa FluorTM 568 phalloidin (Molecular Probes) and Hoechst (Sigma)
were used to stain for actin cytoskeleton and cell nucleus, respectively. Samples were visualized
using a Zeiss LSM510 confocal microscope.
For in vivo analysis, animals were sacrificed at 2 week (patch model) or 3 week (module
injection model) after implantation. Explanted tissues were fixed in 4% paraformaldehyde for
48 hours at 4oC, followed by incubation in 30% sucrose solution for 24 hours. Tissue samples
were rinsed in 70% ethanol before being embedded in paraffin. Tissue sections were stained
with H&E and Masson’s trichrome for tissue morphology assessment. Macrophages and T-cells
were stained with monoclonal mouse anti-rat CD68 (Serotec) and mouse anti rat T cell receptor
(BD Pharagen) antibodies respectively. Implanted cells were tracked by staining tissue sections
with polyclonal rabbit anti-GFP antibody (Abcam), or double staining with GFP and CD31 (or
myosin heavy chain, MHC). For MHC staining tissue section were sequentially stained with
polyclonal rabbit anti-GFP antibody (Abcam) visualized with DAB, followed by monoclonal
mouse anti-rat MHC (α- and β- isoforms, Abcam) antibody visualized with Vector Red. Donor
derived blood vessels were detected by sequentially staining tissue sections with polyclonal goat
anti-rat CD31 antibody (Santa Cruz) visualized with DAB, followed by polyclonal rabbit anti-
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GFP antibody (Abcam) visualized with Vector Red. Vessel density for each sample was
determined by averaging the number of target structures in three hot spots under 100x
magnification.
4.3.5 Statistical analysis
Statistical analysis including calculations of standard error of means and mean
comparisons (t-test or one-way ANOVA) were performed using Prism (Version 5.0, Graphpad
Software, La Jolla, CA)
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4.4 Results
4.4.1 Characterization of CM embedded modules and rat heart EC
We examined the morphology and viability of CM embedded small diameter modules
after 7 days in culture to establish baseline conditions prior to implantation. Cells on the
perimeter appeared elongated in collagen-only modules as well as in collagen+MatrigelTM
module. However, troponin I positive cells near the core appear rounded (Figure 4-1) compared
to cells on the perimeter.
Primary cardiac microvascular EC from transgenic GFP+ Lewis rats were isolated and
concentrated, using CD31 antibody conjugated magnetic beads after CM harvest. To verify their
phenotype, EC were incubated with Dil-Ac-LDL and visualized by confocal microscopy (Figure
4-2). Over 90% of cells are EC, as indicated by their uptake of Dil-Ac-LDL.
We were interested in determining if rat EC adhesion and junction morphology were
affected during the module injection procedure. To answer this question, small diameter
modules embedded with CM were coated with non-GFP rat aortic EC using similar methods as
before22. At 7 days post seeding, CM/EC modules were loaded into a syringe then ejected
through a 28G disposable needle, similar to the ones used for in vivo injection. Immediately
after injection the modules were fixed and stained for VE-cadherin (Figure 4-3A). Confocal
microscopy showed no significant loss of EC in modules which passed through the needle when
compared with modules that did not pass through the needle (Figure 4-3B). However, EC
junctions did appear to be disturbed.
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Figure 4-1: Immunofluorescence staining of CM embedded small diameter modules
supplemented with MatrigelTM after 7 days in static culture containing 10% BS (green - troponin
I, blue - cell nucleus, red - F-actin). Cells on the perimeter appear elongated in both collagen-
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only (A,B,C) and collagen+MatrigelTM (D,E,F) modules. In both groups, cell density decreases
toward the core of modules.
Figure 4-2: Immunoflorescence micrograph of rat cardiac EC isolated from transgenic GFP+
Lewis rats. EC were isolated using CD31 conjugated magnetic beads. After 2 passages, EC were
seeded at 50% confluence onto fibronectin coated glass chambered slides and stained with Dil-
Ac-LDL (10µg/mL, 6 hours). EC stained with Dil-Ac-LDL (red) maintained GFP expression
(green).
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Figure 4-3: EC coated modules before and after injection through a 28G needle. Matrigel
supplemented CM modules made with small diameter tubing (I.D. = 500µm) were coated with
RAEC and static cultured for 7 days. Modules were loaded into a 1 mL syringe and ejected
through a diposible ½ inch, 28G needle. Modules were fixed and stained immediately. VE-
cadherin staining (green) show no significant cell loss compared to modules before injection,
however, EC junctions appeared more disorganized (100x magnifications).
4.4.2 Modular patch and injection implants
In this study we explored two different implantation strategies to deliver modular cardiac
constructs. In the first strategy an implantable version of the modular sheet was fabricated by
stacking modular sheets between two pieces of collagen foam. Three modular sheets, containing
collagen, MatrigelTM, and cells harvested from transgenic GFP+ Lewis rats, were placed
between collagen foams and the corners were sutured to form a pouch (Figure 4-4A). The pouch
was implanted over the right ventricular free wall of wildtype syngeneic Lewis rats. At 14 days
the right ventricular wall was excised and sectioned for histological analysis.
In the second implanation model, small diameter CM-only and CM/EC co-culture
modules containing MatrigelTM were injected through a 28G needle into the peri-infarct zone of
a 7-days old acute left ventricular infarct. Modules were seen around the injection site, and they
remained visible when explanted after 3 weeks (Figure 4-4B).
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Patch and module injection implants were retrieved after 2 week and 3 week,
respectively. In both models, implanted modules were extensively remodeled by host cells, and
in most cases, cannot be identified by histological staining (Figure 4-5). Moreover, there were
significant amount of collagen deposit in the implantation sites by host tissue. Histological
sections also indicate thinning of the heart wall caused by the remodeling of the infarct in the
injection model. (Figure 4-6).
In both implantation models GFP+ cells were found though out the implant sites (Figure
4-7). The presence of EC in the co-culture implant did not lead to a substantial change in GFP+
cell density. Cells that were GFP+/CD31+ formed blood vessel-like structures within the co-
cultured patch (Figure 4-8E and F) and injection (Figure 4-9E and F) implants. In some cases,
blood vessel-like structures were lined with a combination of GFP+/CD31+ and GFP-/CD31+
cells (Figure 4-8F and 8F). Furthermore, red blood cells were found inside the lumen of these
vessels, which suggest that these vessels may have connected with host vasculature and were
functional. Similar structures were also found in CM-only implants in both models. However,
these vessels were lined with GFP-/CD31+ cells only, suggesting that the EC were host-derived
(Figure 4-8B,C and 4-9B,C).
Lastly we stained explanted tissues for GFP and myosin heavy chains (MHC, α- and β-
isoforms) to evaluate the fate of implanted CM. MHC+ structure was found in neither the CM-
only or co-culture patch implants (Figure 4-10A and D). In module injection implants, GFP+/
MHC+ cells in muscle like bundles were found in CM-only implants (Figure 4-10B and C), but
not in co-culture implants (Figure 4-10E and F).
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Figure 4-4: Schematic diagrams and tissue morphology at explant for modular patch implant
module injection implant. (A) Patch implants were created by placing 3 modular sheets between
two pieces of GelFoam and were sutured over a transmural defect on the right ventricular free
wall of a wildtype Lewis rat. (B) In a different model, individual small diameter modules were
injected into the peri-infract zone (3 sites, 50 modules/site) of an 7 day old MI (white dash line)
in a wildtype Lewis rat using a 28G needle. Modules (white arrow) can be found around the
border zone (yelow dotted line) outside of the infarcted area (white dotted line) when explanted
after 3 weeks.
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Figure 4-5: Masson’s trichrome stain of patch implants after 2 weeks. MatrigelTM supplemented
collagen modules in all groups underwent extensive host remodeling. Large deposits of collagen
and granulation tissues were found within the GelFoam matrix in CM-only (A and B) and co-
culture (C and D) modular patch implants. In some cases, implanted modules could be identified
morphologically (D, white dotted line).
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Figure 4-6: Masson’s trichrome stain of module injection implants after 3 weeks. MatrigelTM
supplemented collagen modules injected into the peri-infarct region underwent extensive host
remodeling.
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Figure 4-7: Immunohistochemical staining of explants revealed the presence of GFP+ donor
cells scattered within the implant site in patch implants (A and C) and module injection implants
(B and D). The expression of GFP suggests implanted cells remained viable. The presence of
EC in co-culture implants did not translate into higher GFP+ cell density (slides shown at 100x
magnification).
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Figure 4-8: Blood vessel-like structures can be found in both CM-only (A,B and C) and co-
culture (D, E and F) modular patch implants. The presence of red blood cells suggests vessels
are functional. Blood vessels were evenly distributed within the modular cardiac construct. High
magnification (400x) revealed blood vessels in co-culture patches are lined entirely (E) or
partially (F) with GFP+/CD31+ donor derived EC (pink and brown). In CM-only modular patch
implants, most cells lining the vessels are CD31+ but GFP-, suggesting that they are derived
from host EC.
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Figure 4-9: Blood vessel-like structures can be found in both CM-only (A,B and C) and co-
culture (D, E and F) injection implants. The presence of red blood cells suggests vessels are
functional. Higher magnification (400x) revealed blood vessels were lined entirely (E) or
partially (F) with GFP+/CD31+ donor derived EC (pink and brown) in co-culture injection
implants. In CM-only implants (B and C), most vessels are lined with cells that are CD31+ but
GFP-, suggesting they are derived from host EC.
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Figure 4-10: Double staining for GFP (brown) and MHC (pink) revealed donor derived CM
developed into striated muscle like structure in CM-only injection implant (B, 100x and C,
400x). These double stained, GFP+/ MHC+ structures were not found in other groups (co-culture
patch – D, CM-only patch – A, and co-culture module injection – E,F)
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4.4.3 Host immune response and the removal of MatrigelTM
Host responses towards implanted constructs can significantly affect construct function
and survival. In this study we examined these responses by staining for macrophages and T-cell.
The presence of MatrigelTM in the patch implants and the module injection implants elicited a
strong macrophage response (Figure 4-11A, B, D and E) as well as T-cell infiltration (Figure 4-
12A, B, D and E). We attempted to attenuate these immune responses in the module injection
implant model by removing MatrigelTM from the matrix and rinsing the modules in serum-free
medium prior to implantation. After 3 weeks these implants were removed and examined by
histology. There were substantially less macrophage and T-cell infiltration into the implants in
the absence of MatrigelTM (Figure 4-11C, F and 4-12C, F). Blood vessel like structures lined
with GFP+/CD31+ cells were found through out CM/EC co-culture implants. In contrast, similar
structures found in CM-only implants were lined with GFP-/CD31+ cells (Figure 4-13).
When quantified, patches containing CM-only modules and co-culture modules showed
no difference in total vessel density (Figure 4-15A). But when the vessel are grouped based on
cell phenotype, we found an increased number of vessels lined with GFP+/CD31+ cells in the co-
culture patch implants compared to CM-only patches (Figure 4-15B). Also, co-culture implants
without MatrigelTM have significantly higher vessel density counts compared to co-culture
implants with MatrigelTM and CM-only MatrigelTM-free implants (Figure 4-15A). However, the
removal of MatrigelTM did not alter the proportion of GFP+ vessels found in the implants
compared to those with MatrigelTM (Figure 4-15B).
The removal of MatrigelTM also affected muscle bundle formation. In both CM-only and
co-culture implants, we found an abundance of muscle-like structures throughout the implants.
Unlike implants with MatrigelTM, many of these bundles contained GFP+/MHC+ cells (Figure 4-
14).
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Figure 4-11: Immunohistochemical staining of CD68 revealed large number of macrophage
infiltration in all implantation models with MatrigelTM (A,B,D and E, 100x magnification).
MatrigelTM-free CM-only (C) and co-culture (F) injection implants lead to a significant
reduction in macrophage infiltration after 3 weeks, as shown by the reduction of CD68+ cells.
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Figure 4-12: Immunohistochemical staining of T cell receptor (TCR) revealed large number of
T-cell infiltration in all implantation models with MatrigelTM (A,B,D and E, 100x
magnification). It was not known whether these T-cells were activated or not. MatrigelTM-free
CM-only (C) and co-culture (F) injection implants lead to a significant reduction in T-cell
infiltration after 3 weeks, as shown by the reduction of TCR+ cells.
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Figure 4-13: Blood vessel-like structures were found in MatrigelTM-free CM-only (A and B)
and co-culture (C and D) injection implants after 3 weeks. Under higher magnification (400x),
blood vessels comprise partially of GFP+/CD31+ donor derived EC (pink and brown) were found
in co-culture injection implants (D). In CM-only patches (B), vessels are lined with GFP-/CD31+
cells (brown), suggesting the vessels arise from host derived EC. The presence of red blood cells
in the lumens suggests the vessels connected to host vasculature and are functional.
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Figure 4-14: Donor CM in MatrigelTM-free CM-only (A and B) and co-culture (C and D)
injection implants formed striated muscle-like structures after 3 weeks. Under higher
magnification (400x) striated muscle bundles were found. These structures contain both host
derived CM (GFP-/MHC+, pink) as well as donor derived CM (GFP+/ MHC+, pink and brown).
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Figure 4-15: Total vessel density counts show no significant difference in vessel densities
between CM-only constructs and their co-culture counterparts (A). In MatrigelTM-free injection
implants (CM-Injection-MF and CO-Injection-MF), however, the presence of EC significantly
increased vessel density compared to CM-only implants and co-culture injection implants with
MatrigelTM (*, p<0.05, n=3). In all cases, donor cells participated in vessel formation, as shown
by the increased proportion of GFP+ and mixed vessel when counted based on their phenotypes
(GFP+ EC only, mixed EC, or GFP- EC only).
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4.5 Discussion
4.5.1 Implantation strategies
In previous studies we have demonstrated that modules consisting of MatrigelTM
supplemented collagen were able to maintain the viabilities and functionalities of embedded CM
and surface coated EC. We have also shown that the modules could be used to form
macroporous sheets in vitro22. In this study our goal was to create implantable constructs using
the modular approach and determine their fates in vivo. Our first approach was to assemble
module sheets into an implantable patch. However, these sheets did not posses the necessary
mechanical strength to be surgically handled as a stand alone patch; therefore we had to devise a
way to support the module sheets. The method described in this study involved sandwiching
module sheets between two pieces of GelFoam, a commercially available sponge-like substrate
made of gelatin. The GelFoam layers provided mechanical stability to the patch construct so that
it could be sutured onto the right ventricular free wall defect. Histological analysis revealed that
a significant number of host cells including macrophage, T-cells and fibroblast infiltrated into
the GelFoam after 2 weeks. These cells resorbed and remodeled much of the GelFoam similar to
what was observed in other studies involving collagen or gelatin foam scaffolds19,23. We found
most of the GFP+ cells clustered in module-shaped areas within a collagen rich deposit, and
almost none of them were found near the host-implant interface, where the tissue type is
predominantly host derived. The presence of EC on co-culture modules did not seem to have a
noticeable effect on the gross tissue remodeling outcome.
Aside from using the module sheets to create constructs to cover a full thickness defect,
we also explored the possibility of deploying modules as individual functionalized microtissues
to repopulate damaged cardiac tissue. To test this approach CM-only or co-culture modules
were injected into the peri-infarct region of an acute infarct in the left ventricle. The peri-infarct
zone was chosen due to its location between the infarct zone and healthy myocardium. At this
location, the modules will be more likely to survive, being adjacent to oxygenated tissue, while
remaining close to the infarct zone to exert its therapeutic effects. Histological analysis after 3
weeks showed heart wall thinning and decellularization within the infarct zone. Modules were
found in the peri-infarct zone with most of their collagen matrix still intact. When compared to
the patch implant model, the injection model has a much smaller host-tissue transition zone, and
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the modules are separated from the host tissue by only a thin layer of collagen. In the absence of
GelFoam, the host tissue remained organized and few myofibroblasts infiltrated into implants.
Like the patch implants a large amount of macrophage and T-cells infiltrated into the implants
sites, suggesting that the use of GelFoam in the patch used may not be sole factor for triggering
the larger than expected host immune response.
These two implantation strategies were intended to highlight the potential applications of
the modular cardiac construct. The patch implant was initially designed to cover full thickness
defect, and as such it should possess the necessary mechanical strength. In its current form the
module sheet construct cannot easily be handled or sutured into native myocardium. The
addition of a GelFoam layers solved this problem, but at the same time it may act as a barrier for
proper implant-host integration by separating the modular sheets from the wound edge of the
RVOT defect. Moreover, the addition of GelFoam decreased the overall cell-to-scaffold ratio
when compared to constructs consisting of modules alone, thus potentially reducing the
theoretical force that could be generated by such implant. Therefore, more work is still needed
to fabricate the modules into a clinically useful cardiac patch implant.
Alternatively we deployed loose CM-embedded modules as pre-fabricated tissue unit
into the peri-infarct zone in an MI; an area that has undergone an ischemic event and
experienced heavy cell loss. The rationale here is similar to that of cell therapies based on
intramyocardial cell injection, where the goal is to rapidly repopulate functional cell types
including CM and EC. The potential advantages of injecting modules over cell suspension may
be better cell retention and cell survival. Indeed, the modules were physically lodged within the
injection site and we saw no modules escaping through the needle puncture hole after implant.
Histology sections confirmed the presence of module and viable cells after 3 week. On the other
hand, when compared with scaffold free cell injection methods, the presence of collagen may
impede with the integration and assembly of CM into muscle bundles. Whether the modular
injection method of cell delivery into the myocardium is superior to conventional cell injection
remains to be determined by functional assays in future studies.
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4.5.2 Fates of implanted cells and host immune response
The ultimate goal of cardiac tissue engineering is to restore lost function to the heart.
Whether or not this goal is achieved depends on the survival and function of implanted cells as
well as their interactions with host tissue. Specifically, the new tissue must contain contractile
CM and the vasculature to support them. The fates of implanted cells were examined by
histological analysis.
Previous studies have demonstrated that allogeneic EC coated on collagen modules were
capable of forming functional blood vessels in an immunosuppressed animal after 3 weeks17. In
this study we saw similar vessel-like structures within the modular patch implants and injected
modules implants. It is likely that these vessels have anastamosed with host vasculature and are
functional, as erythrocytes were found within their lumens. In both cases, we found vessels lined
with GFP+/CD31+ cell when co-culture modules were implanted, which suggests that these
vessels were derived from implanted EC. Also, vessels consisted of host derived EC alone or a
mixture of host and implanted EC were found within the vicinity of implants. These
observations suggest that the implanted EC not only form vascular structures, but may also be
involved in creating and maintaining an angiogenic gradient to attract host EC migration and
play a role in stabilizing new blood vessels formation around the implant site10. The factors
responsible for the observed angiogenic events remain to be determined in future studies.
We hypothesized that the presence of EC in co-culture implants would lead to an
increase in overall vessel density. Indeed this was the case in the modular injection implant
model, but only in the absence of MatrigelTM. It is possible that host immune response against
xenoprotein may have masked the difference in vessel density between CM-only and co-culture
implants. Alternatively, the presence of growth factors in implants containing MatrigelTM may
have mobilized host EC and overwhelmed the angiogenic effect of donor EC, resulting in
similar overall vessel density in both implants.
The presence of xenoprotein also had an effect on the overall vessel density amongst
similar implants. MatrigelTM-free co-culture implants had a higher overall vessel density
compared to co-culture implants with MatrigelTM. This suggests that the host immune response
against implants with MatrigelTM may have diminished the angiogenic potential of donor EC.
The main motivation in building a functional vasculature in engineered cardiac tissue is
to support the metabolic needs of CM, which are essential for their development and
106
organization into differentiated muscles, as several studies have highlighted this point15,16,24. In
this study, we looked at the expression of MHC as the primary marker of tissue development.
With the exception of CM-only injected module, few implanted CM survived. However, those
that did survive developed into striated muscle like structures in vivo in modules containing
MatrigelTM. This suggests that the maintenance of CM survival shortly after implantation may
be the key for successful tissue formation.
The low CM survival rate was accompanied by a strong macrophage and T-cell
infiltration. It is important to recognize that the recruitment of macrophage and the associated
inflammatory response are crucial for the proper wound healing and revascularization process.
Not only do macrophages remove cell debris from the trauma site, certain subpopulation of
macrophages, namely the M2 sub-type, are known to secrete VEGF and other factors to promote
angiogenesis25. However, an overactive inflammatory response brought on by non-self proteins
may impair cell survival. For this reason, we have chosen to perform all implantation studies
using syngeneic animals in order to minimize the immune response caused by major
histocompatibility complex mismatch between donor and host. Even though all cells were
harvested from syngeneic donors, the MatrigelTM and bovine serum used during culture may
have aggravated the inflammatory response and triggered the activation of T-cells, leading to a
cytotoxic environment that contributed to CM death. Other studies have shown that the presence
of xenogeneic protein in MatrigelTM and bovine serum used during culture can provoke a strong
host immune response13,26. In particular, a study by Zimmermann et al. showed that the
immunogenicity of MatrigelTM elicited an immune response that can significantly reduce
implanted CM survival12. It is likely that the implanted CM in MatrigelTM+collagen modules
suffered the same fate. Not surprisingly, the removal of MatrigelTM and pre-implantation rinse
of modules in serum-free medium substantially reduced the amount of macrophage and T-cell
infiltration. Along with the reduction in host immune response was an increase in CM survival
and the presence of GFP+/MHC+ striated muscle structure in both CM-only and co-culture
injection implants. However, it appeared that the presence of EC in co-culture injection implants
did not lead to a significant difference in CM survival after 3 weeks. Perhaps the benefit of EC
will be more pronounced in later time points.
Beside improved CM survival, the removal of xenoproteins also improved blood vessel
density. Module injection with collagen-only, co-culture modules had a higher vessel density
compared to the CM-only group. In fact, the difference in vessel density was only present in the
collagen-only implants. It is possible that host immune response triggered by MatrigelTM and
107
bovine serum have also lead to the death of implanted EC, similar to the fate of implanted CM.
These results highlight the issue of using xenomaterial as part of the cardiac scaffold for clinical
applications. Development in synthetic scaffolds and hydrogel based on autologous ECM may
mitigate this problem27.
4.6 Conclusion
In this study we have demonstrated that modular cardiac tissues could be
implanted successfully in its basic form or as a patch implant in a syngeneic animal model. In
both implantation strategies, donor EC formed blood vessels-like structures and appeared to
have anastamosed with host vasculature. In co-culture implants, these vessels were lined with a
mixture of donor and host derived EC. In contrast, vessels found in CM-only implants only
contain host derived EC. The presence of MatrigelTM and bovine serum in CM-only and co-
culture implants triggered significant host immune and inflammatory responses which may be
responsible for the loss of implanted CM in both the patch and injection model. The subsequent
removal of foreign protein substantially reduced macrophage and T-cell infiltration into the
implants. Co-culture implants had a higher overall vessel density compared to CM-only
implants, but only in the absence of MatrigelTM. It also led to an increase in striated muscles
bundles containing donor CM. These results will serve as the basis for future optimization
studies toward the realization of the modular cardiac construct as clinically relevant method to
replace damaged cardiac tissue.
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Chapter 5: Future Perspectives and Conclusions
5.1 Conclusions
The overall goal of this project was to apply the modular tissue engineering construct to
create functional cardiac tissues. To begin, we fabricated and characterized a co-culture modular
cardiac tissue in vitro containing neonatal CM and EC. Modules made of type I collagen or
MatrigelTM supplemented collagen were able to support CM growth and function for 14 days in
culture. Modules contracted synchronously in response to external electrical stimulation. The
presence of MatrigelTM increased contraction amplitude and decreased excitation threshold.
Similar functional improvements could be achieved when using medium supplemented with
10% bovine serum instead of 10% fetal bovine serum. Culture conditions for co-culture modules
were determined by fine tuning medium compositions to satisfy the need of both embedded CM
and surface seeded EC so that both cells types maintained their phenotype while in culture.
After 14 days in culture, surface seeded EC covered the entire surface of the module and
expressed VE-cadherin while embedded CM exhibited decreased electrical responsiveness
(excitation threshold, maximum capture rate), but there was no reduction in contraction
amplitude compared to CM-only modules. These studies helped to establish the baseline
conditions for fabricating functional cardiac modules.
The next step was to build on the modular concept and fabricate larger constructs with
an eventual goal of achieving the scale and complexity of native myocardium. We created
macroporous modular sheets by immobilizing co-culture modules in alginate gel in vitro for 10
days, while simultaneously subjecting the sheet to electrical stimulation during culture. Over
this period, adjacent modules fused with one another to form an integrated sheet, while
intermodular spaces were preserved by the alginate gel. When released from the alginate gel,
modular sheets contracted synchronously in response to external electrical stimulation.
Histological analysis revealed many embedded CM expressing troponin I and connexion-43
around the perimeter of modules as well as at intermodular junctions. Morphologically, CM
around the perimeter of modules appeared aligned and elongated. Their resemblance to native
muscle bundles suggested that these structures may be responsible for the generation of the
observed contractile force. Overall, these results demonstrated the feasibility of building higher
113
order structures (albeit small ones) using the modular approach as well as the structural and
functional integration of modules into a modular sheet.
In the last study, we investigated the fate of modular cardiac tissue in the myocardium.
Two syngeneic implantation models, namely patch and injection models, were used to study the
remodelling of cardiac modules and their interactions with host tissue. Co-culture modules
containing MatrigelTM and collagen were either injected into the peri-infarct zone of the heart or
fabricated into a patch form and implanted over a defect on the right ventricular free wall.
Explanted tissues indicated that donor cells remained viable (2 weeks for patch and 3 weeks for
injection), while host remodelling of collagen matrix was observed. In both models, donor EC
along with other cells, developed into blood vessel-like structures that appeared to connect with
host vasculature. In CM-only implants, blood vessels lined with host derived EC were found
throughout the implant. Therefore, the presence of donor EC in co-culture implants did not lead
to an overall increase in implant vascularity compared to CM-only implants. Interestingly, no
donor CM were found in either model, which suggested that they may have died. Also, host
immune/inflammatory cells such as macrophages and T-cells were found in or near the implant.
To lessen the host response, collagen-only co-culture modules were rinsed in serum-free
medium prior to peri-infarct implantation. Removal of xenoproteins attenuated host immune
response by reducing macrophage and T-cell infiltration into implants. Unlike previous
observations, co-culture implants had a significantly higher vessel density compared to CM-only
implants in the absence of MatrigelTM. In fact, MatrigelTM-free co-culture implants had a higher
overall vessel density compared to co-culture implants with MatrigelTM. Most importantly, in
vivo, donor CM matured into striated, muscle-like structure throughout the implant. This was a
significant improvement over implants with MatrigelTM and the results suggested that early host
response plays a significant role in the survival and maturation of modular cardiac constructs.
The studies presented in this thesis demonstrated the utility of the modular construct as a
means for combining EC and CM in a spatially defined and functional subunit. The results
highlighted the novelty and challenges of building scalable cardiac-tissue equivalence using the
modular approach. Finally, data from in vivo studies showed that cardiac modular tissue
implanted into the myocardium survives and undergoes extensive remodelling and maturation;
events that are mediated by host immune response. These studies have laid the groundwork for
the development of modular cardiac tissue and their use as a therapeutic platform.
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5.2 Recommendations
The modular tissue engineering construct was able to solve some of the challenges in
creating large 3-D tissue. However, many obstacles remain to be overcome. Some of these
issues relate specifically to the modular construct, while others are more general and apply to
most tissue engineering strategies. These issues, along with some recommended studies to
address them, are discussed in the following sections.
5.2.1 Improving embedded cell viability
The maintenance of cell viability has been a major challenge in designing engineering
tissue constructs. In many cases, the lack of a vasculature in constructs limits both the size and
complexity of in vitro engineered tissue. The modular approach to engineering tissue constructs
was designed primarily to address this issue by reducing the maximum distance between
embedded cells and the perimeter of the construct to within the theoretical nutrient diffusion
limit of 100-200 µm. Nevertheless, we saw a decline in the density of viable cells towards the
core of modules (diameters ≤ 400 µm). In culture, embedded CM near the core of modules
expressed troponin I, but did not mature into elongated and striated muscle bundles like those
found near the perimeter. This observation may imply that the effective diffusion limit in CM
embedded modules may be much less than expected.
One of the major factors that causes CM death in tissue engineering constructs is
hypoxia. Cells embedded near the module core may suffer from hypoxia due to diffusion
limitation. Core hypoxia can be confirmed by monitoring the expression of hypoxia inducibled
factor 1α (HIF-1α), either by immunohistochemistry or PCR. HIF-1α is a transcription factor
expressed under reduced oxygen tension1. Its expression has been linked to the up regulation of
genes including erythropoietin2, VEGF3, HO14 and i-NOS5; all of which are involved in
promoting systemic O2 absorption and transportation6. However, these genes exert their effects
through distant effecter cells and tissues, such as EC and hematopoietic stem cells. Therefore,
they do little to restore the O2 homeostasis of tissue engineered constructs when cultured in
vitro. The reduction of ATP production rate causes the cytoplasm to accumulate ions that are
normally expelled via the cation pumping ATPase and eventually contributes to CM necrosis7.
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The most effective way to correct hypoxia is to increase the flux of O2 diffusion into the
engineered cardiac construct. This could be achieved by decreasing the average distance of CM
from culture medium and by increasing the O2 tension of culture medium. As covered in
previous chapters, numerous studies have attempted to vascularize tissue engineering constructs
with the view of reducing the distance of embedded cell from the culture medium. A recent
study published by the Sefton group indicates that the viability of embedded HepG2 cells can be
maintained in small diameter collagen modules, with a post-contraction diameter of 400 µm and
cell density of up to 8x107 cells/mL8. At the same time, HpeG2 cells embedded in large
diameter modules (700 µm, post-contraction) developed a necrotic core. In comparison, CM
embedded in similar size modules displayed core cell death at a cell density of 2x107 cells/mL,
which is likely due to their higher oxygen demand compared to HepG2 cells. Nevertheless,
these results suggest that the maximum density of viability cells supported by the modular
construct is inversely proportional to the diameter of the modules. For cardiac module, this may
represent a further reduction in module diameters, perhaps down to less than 100µm from the
current 200-300µm range, in order to maintain core viability at the current cell density.
Reduction of module diameter may be an adequate solution for improving O2 diffusion
in applications where the modules are meant to be used in their unassembled form, such as the
injection implant model. One of the potential issues when trying to create higher order structures
with smaller modules compared to larger ones is the increased number of intermodular junctions
per unit volume in the final assembled constructs. Assuming that these intermodular junctions
are mechanically weaker than the modules themselves, this would translate into an overall
decrease in mechanical strength of the assembled constructs. Indeed, during physical handling,
we found that the intermodular junctions in cardiac modular sheets tended to be the weaker
spots. Histological data suggested that cells in one module do not assemble into muscle bundles
with other cells in neighbouring modules. Therefore, increasing the number of intermodular
junctions using the current generation of cardiac modules will further decrease the mechanical
strength of the assembled constructs.
Other ways to increase O2 transfer into modules include tissue perfusion9 and increasing
O2 tension in the culture medium. Both methods operate by maintaining the concentration
gradient of oxygen across the module-medium interface. The amount of oxygen carried by
culture medium at saturation is governed by temperature and the partial pressure of O2 in the gas
phase above the liquid. In contrast, most of the oxygen carrying capacity of blood is handled by
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haemoglobin. Specifically, the total concentration of O2 from plasma and haemoglobin in
arterial blood can reach 8600 µM compared to 220 µM found in standard cell culture medium.
To compensate for the lack of haemoglobin, synthetic oxygen carriers have been formulated as
substitutes for haemoglobin in vitro10,11. Radisc et al. have found that the addition of
perfluorocarbon (PFC) emulsion in culture medium can increase the total amount of oxygen
delivery in a perfused cardiac bioreactor12. A similar strategy could be used to alleviate hypoxia
in the modular cardiac constructs.
5.2.2 Using autologous cell source
One of the crucial factors in successfully creating functional engineered tissue is to
select the correct cell source that complements the tissue engineering strategy. In this study, we
used neonatal rat CM as a model cell type to demonstrate the usefulness of the modular
construct. However, their allogeneic nature and scarcity in clinical settings severely limit their
potential as a viable treatment option for human patients. Recent advances in stems cell research
have gathered much attention as they hold the promise of generating the various cell types that
are necessary for cardiac tissue engineering with minimal immune barrier. For example,
autologous adult stromal stem cells can be readily harvested from patients’ own skin, fat, blood
or bone marrow and expanded ex vivo. Among these cell types, adipose stem cells (ASC) and
bone marrow derived mesenchymal stromal cells (bmMSC) are of particular interest. When
implanted into the myocardium, they have been shown to differentiate into myocytes and EC
respectively; two major cell types that are necessary to create cardiac tissue. In the context of
modular tissue engineering, ASC can be embedded into modules in their undifferentiated
progenitor state and allowed to mature in vitro into functional CM using biochemical factors,
including TGF-β113, VEGF14 and CM extracts15,16. Work by Tandon et al. demonstrated that
differentiated ASC respond to external electrical stimulation by upregulating connexin-43
expression as well as aligning and elongating perpendicular to the electric field17. Moreover,
ASC have been shown to differentiate into CM when injected directly into the myocardium18,19.
Together, these studies illustrate the potential of adult derived stem cell for generating
functional cardiac tissues.
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5.2.3 Characterization of host-module coupling
In order to maximize the functional benefits of modular cardiac tissue in the host’s
myocardium, the interface between host tissue and implanted modules must be electrically and
mechanically coupled with each other. Mechanical integration can be assessed by staining for
adherens junction components, including cardherins and desmosomes. Intermodular junction
must also be mechanically coupled to transmit force over the entire assembled construct. This
could be assessed by measuring the twitch force of the modular sheets in vitro. Ultimately, the
establishment of mechanical coupling allows the contractile forces generated by the implant to
be effectively transmitted to the myocardium.
The synchronous contraction of myocardium is achieved by the rapid and anisotropic
propagation of action potentials (AP) across the myocardium. This is mediated by gap junction
proteins known as connexins that are present on the surface of CM. Although the expressions of
connexins can be detected by immunohistological methods, only functional assays can confirm
that the connexins are organized into hemi-channels and that they are aligned with other hemi-
channels on neighbouring CM. The physical establishment of intercellular gap junction can be
detected using dye-transfer assay20. However, the diffusion of dye from one cell to another is
generally a much slower process compared to the propagation of AP. Therefore, the data
obtained will not carry sufficient spatial-temporal resolution to fully describe AP propagation on
the myocardium.
A more physiologically relevant assay to determine the functional status of gap junctions
is to optically or electrically measure the propagation velocity of AP across the modular sheet.
Changes in membrane potential during depolarization can be detected directly using ANEPPS
dyes, a family of fluorescence membrane dyes that varies their fluorescence output relative to
membrane potential21. Alternatively, CM depolarization can be detected indirectly by
monitoring intracellular calcium concentration using calcium sensitive fluorescent dyes. The
changes in fluorescence output in both systems can be mapped optically to indicate the
propagating front of AP across the modular sheet as a mean to assess the status of electrical
coupling between modules21,22. This method may also be applied to study electrical coupling
between the modular cardiac tissue and the host myocardium in explanted hearts. A similar
approach was used by Dumas et al., where the AP on a rabbit heart mounted in a Langendorff
perfusion apparatus was monitored using two-photon excitation of di-4-ANEPPS dye.
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Optical mapping of AP propagation across the cardiac modular sheets were attempted in
preliminary studies, but have achieved limited success. Upon further analysis, we have
identified several issues that may have hindered our ability to observe AP propagation across the
cardiac modular sheet. First and for most, the plane of AP propagation must be parallel to the
focus plane of the microscope in order to achieve accurate ratiometric fluorescence
measurements. This is mainly due to the drastic fall off of fluorescence signals in out-of-focus
planes. Although, this could be easily achieved in 2-D cultures, the curvatures of individual
module as well as the modular sheet have made it difficult to fit the entire depth of the modular
sheet within the focal plane. One way to correct this would be to mechanically fix the sheet on a
substrate, such as PDMS. Another solution would be to decrease the physical size of the
aperture of the lens in order to increase its depth of view, thereby increasing the depth of tissue
that would be in focus. However, in doing so, the number of photons reaching the camera would
be reduced and either the exposure times or camera sensitivity would have to be increased to
compensate for the decrease in signal. Of course, AP propagation could also be mapped using
an electrode array, whereby an electrical pulse is sent out to the test tissue by an input electrode.
The resulting changes in membrane potential are recorded using an array of recording electrodes
and the propagation velocity of AP is calculated based on the time delay and location of the
recorded signals. Since this modality relies on physical contact, it should solve some of the
issues being faced in optically mapping AP on modular cardiac tissues.
Results from these experiments could help elucidate the fate of modular cardiac
constructs in several ways. First, the electrophysiological properties of the implants can be
characterized. Studies have shown that engineered cardiac construct may be susceptible to
arrhythmia due to incompatible electrophysiological characteristics21,23. By mapping AP
propagation, any beneficial or adverse effects caused by the implant on the electrical conduction
in the host’s myocardium can be detected and studied. Overall, these data would provide
insights into the remodelling process that occurs during host-implant coupling.
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5.2.4 Assessing the functional benefits of modular cardiac tissue
The ultimate goal of engineered cardiac tissues is to replace and restore lost cardiac
function. Animal injury models like the ones described in this thesis serve as a useful platform
to assess the functional benefit of implanted tissue. It is necessary to quantify the functional
benefit of these implants in order to evaluate their efficacies and improve upon their designs.
From a clinical stand point, cardiac function refers primarily to the ability of the heart to pump a
sufficient volume of blood at the necessary pressure over each cycle. This can be translated to
measurable parameters, including end-systolic and end-diastolic ventricular pressures, ejection-
filling interval, stroke volume and ejection fraction. Many of these parameters can be
determined directly using a pressure-volume catheter inserted into the left ventricle24, or
measured using echocardiography as a non-invasive alternative.
Functional assays such as the one described above would be well suited for detecting the
restoration of function after implantation in an animal injury model, such as the peri-infarct
module injection model presented in this thesis. Moreover, control groups must be carefully
selected in order to obtain meaningful results. For example, in the module injection model, there
should be three control groups; one with CM-only modules, one with cell-free modules, and one
sham (saline injection). The differential changes in functional parameter between each group
would yield clues regarding the importance of each component in restoring cardiac functions.
These results would provide critical feedback for improving the therapeutic benefits of modular
cardiac tissues.
120
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