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Instructive Matrix Cues for Tissue Engineering Applications
by
Yun Xiao
A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy
Department of Chemical Engineering and Applied Chemistry
Institute of Biomaterials and Biomedical Engineering University of Toronto
© Copyright by Yun Xiao 2016
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Instructive Matrix Cues for Tissue Engineering Applications
Yun Xiao
Doctor of Philosophy
Department of Chemical Engineering and Applied Chemistry
Institute of Biomaterials and Biomedical Engineering
University of Toronto
2016
Abstract
Tissue engineering holds the promise of generating functional tissues to replace, or regenerate
impaired native tissues by combining knowledge from cell biology, material science, and
engineering. Designing instructive biomaterials that harness regeneration potential of native
tissue requires incorporation of biochemical and biophysical cues. Besides, micro-tissues
analogues are generated by emerging microfabrication techniques with precise control. This
thesis aims to facilitate cell-matrix interactions by biochemical and topographical cues in
collagen-based matrix for tissue regeneration in vivo with a focus on cardiac tissue engineering
and wound healing applications, and creating cardiac micro-tissues with better physiological
relevance in vitro.
Chapter 2 introduces the key topics on designing instructive biomaterials for tissue engineering
applications, with an emphasis on cardiac tissue engineering and wound healing applications.
Chapter 3 describes covalent immobilization of peptides or angiogenic growth factors on
collagen scaffolds. Immobilization efficiency and release profile were characterized and showed
sustained releases. The modification did not affect the porous structure and tensile strength of the
collagen scaffolds.
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Chapter 4 describes the study on protective effect of QHREDGS peptide on primary human
keratinocytes and its efficacy in promoting diabetic wound healing. In vitro studies on normal
and diabetic human keratinocytes showed that immobilized QHREDGS promoted keratinocyte
attachment, collective migration, and survival against H2O2 stress. QHREDGS immobilized in
chitosan-collagen hydrogel accelerated wound healing in diabetic mice by promoting re-
epithelialization in vivo.
Chapter 5 describes the design of a microfabricated bioreactor providing topographical cues to
generate cardiac micro-tissues recapitulating native cardiac bundles. The micro-tissues were
perfusable with micro-tubing in the center, mimicking capillaries. We demonstrated the utility of
this platform by investigating the effects of nitric oxide on electrophysiological properties of the
perfusable cardiac biowires.
We finally conclude with remarks on the future prospects for biochemical and topographical cues
in instructive biomaterials design for tissue engineering applications.
iv
Acknowledgments
First of all, I would like to thank my supervisor Dr. Milica Radisic for giving me the opportunity
to work on these studies and her support and guidance over the years. Milica has inspired me to
be the best scientist I could be with challenges and encouragement throughout my thesis, and her
enthusiasm and vision in science. I am thankful for every opportunity she provided for me to
grow as a graduate student.
I would also like to thank my committee members, Dr. Michael Sefton and Dr. Alison McGuigan,
for their valuable feedbacks on my thesis. Thanks also to Dr. Edgar Acosta for being my
departmental examiner and to Dr. Todd Hoare for being my external examiner. The invaluable
advices from these experts from different research area made my thesis the best it could be.
It is my greatest pleasure to work with previous and present members of the Laboratory for
Functional Tissue Engineering since 2010: Rohin Iyer, Loraine Chiu, Anne Hsieh, Lewis Reis,
Katherine Chiang, Hannah Song, Nimalan Thavandiran, Larry Meng, Iran Rashedi, Boyang
Zhang, Sara Nunes, Jason Miklas, Lan Dang, Aarash Sofla, Mark Li, Lara Fu, Kujaany Kana,
Nicole Feric, Carol Laschinger, Dario Bogojevic, Miles Montgomery, Yimu Zhao, Aric Pahnke,
Locke Davenport Huyer, Genna Conant, Erica Knee, Anastasia Korolj, Stasja Drecun, Junhao
Gu, Shuwen Cao, Samad Ahadian, and Ben Lai. All of you have provided great support and help
over the years and I am ever so grateful.
I also acknowledge the opportunities to work with and learn from my collaborators: Dr. Kang
Kai and Dr. Lu Sun from Dr. Ren-Ke Li’s group, Haijiao Liu from Dr. Yu Sun and Dr. Craig
Simmons’ group, Dr. Mark Gagliardi from Dr. Dordon Keller’s group, Dr. Lindsay Fitzpatrick
from Dr. Michael Sefton’s group, and Camila Londono from Dr. Alison McGuigan’s group. I
would also like to thank A.J. Wang from the animal facility for his help with my animal studies.
To all the animals who were sacrificed in my studies, I acknowledge your contribution in the
name of science.
I would like to thank my friends here who made Toronto my second home and the others who
have always been there for me despite the thousands of miles between us. To my boyfriend,
Haijiao Liu, without whom I could not have come this far. I am so thankful that you came into
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my life at the beginning of my graduate study and changed it throughout. You have been a great
support for me through the stressful times, the person I can share everything with, a great
company when I work overtime, an inspiring young scientist to have a discussion with, and an
example of self-motivation and determination to me.
Finally, I dedicate this thesis to my parents, who love me unconditionally and always allow me
to pursue my dreams. I feel blessed to have such loving and supportive parents and grandparents,
and sorry that I was not always there to be with you.
vi
Declaration of Co-Authorship
The original scientific content of the thesis is comprised of three previously published, peer
reviewed articles in internationally recognized journals and a fourth article submitted. The
literature review is from one published review paper, one review paper in preparation for
submission, and two published book chapters, the transcribed writing being that of Yun Xiao.
The contributions of co-authors are stated in the thesis, in conformity with the requirements for
the degree of Doctor of Philosophy.
vii
Abstracts of Published Articles Appearing in the Thesis
Modifications of biomaterials with immobilized growth factors or peptides for tissue
engineering applications.
Xiao Y, Reis L, Zhao Y, Radisic M. Methods 2015;84:44-52.
In order to provide an instructive microenvironment to facilitate cellular behaviors and tissue
regeneration, biomaterials can be modified by immobilizing growth factors or peptides. We
describe here our procedure for modification of collagen-based biomaterials, both porous
scaffolds and hydrogel systems, with growth factors or peptides by covalent immobilization.
Characterizations of the modified biomaterials (immobilization efficiency, release profile,
morphology, mechanical strength, and rheology) and in vitro testing with cells are also discussed.
Contributions: Y.X.-concept and design, performed experiments and data analysis, manuscript
writing; L.R.-concept and design, performed experiments and data analysis, manuscript writing;
Y.Z.-concept and design, manuscript writing; M.R.-concept, data interpretation, final approval of
manuscript.
viii
Aged human cells rejuvenated by cytokine enhancement of biomaterials for surgical
ventricular restoration.
Kang K*, Sun L*, Xiao Y, Li SH, Wu J, Guo J, Jiang S, Yang L, Yao TM, Weisel RD, Radisic
M, Li RK. J Am Coll Cardiol 2012;60:2237–49. (* equal contribution)
Objectives: This study investigated whether cytokine enhancement of a biodegradable patch
could restore cardiac function after surgical ventricular restoration (SVR) even when seeded with
cells from old donors.
Background: SVR can partially restore heart size and improve function late after an extensive
anterior myocardial infarction. However, 2 limitations include the stiff synthetic patch used and
the limited healing of the infarct scar in aged patients.
Methods: We covalently immobilized 2 proangiogenic cytokines (vascular endothelial growth
factor and basic fibroblast growth factor) onto porous collagen scaffolds. We seeded human
mesenchymal stromal cells from young (50.0 ± 8.0 years, N = 4) or old (74.5 ± 7.4 years, N = 4)
donors into the scaffolds, with or without growth factors. The patches were characterized and
used for SVR in a rat model of myocardial infarction. Cardiac function was assessed.
Results: In vitro results showed that cells from old donors grew slower in the scaffolds.
However, the presence of cytokines modulated the aging-related p16 gene and enhanced cell
proliferation, converting the old cell phenotype to a young phenotype. In vivo studies showed
that 28 days after SVR, patches seeded with cells from old donors did not induce functional
recovery as well as patches seeded with young cells. However, cytokine-enhanced patches
seeded with old cells exhibited preserved patch area, prolonged cell survival, and augmented
angiogenesis, and rats implanted with these patches had better cardiac function. The patch
became an elastic tissue, and the old cells were rejuvenated.
Conclusions: This sustained-release, cytokine-conjugated system provides a promising platform
for engineering myocardial tissue for aged patients with heart failure.
ix
Contributions: K.K., L.S.-concept and design, performed in vitro and in vivo experiments and
data analysis, manuscript writing; Y.X.-design, scaffold preparation and characterization,
manuscript writing; S.H.L., J.W., J.G., S.J., L.Y., T.M.Y.-performed surgical work and
evaluation, R.D.W., M.R., R.K.L.-concept, data interpretation, final approval of manuscript.
x
Diabetic wound regeneration using peptide-modified hydrogels targeting the epithelium
Xiao Y, Feric N, Knee EJ, Gu J, Cao S, Laschinger CA, Londono C, McGuigan AP, Radisic M.
Submitted to Proceedings of the National Academy of Sciences.
There is a clinical need for new, more effective treatments for chronic wounds in diabetic
patients. Lack of epithelial cell migration is a hallmark of non-healing wounds and diabetes often
involves endothelial dysfunction. Therefore, targeting re-epithelialization, which mainly involves
keratinocytes, may improve therapeutic outcomes of current treatments that mostly focus on
angiogenesis. In this study, we present an integrin-binding prosurvival peptide derived from
angiopoietin-1, QHREDGS, as a novel therapeutic candidate for diabetic wound treatments by
demonstrating its efficacy in promoting human primary keratinocytes attachment, survival,
collective migration, and Akt and MAPKp42/44 activation. The QHREDGS peptide, both as a
soluble supplement and when immobilized in a substrate, protected keratinocytes against
hydrogen peroxide stress in a dose dependent manner. Collective migration of both normal and
diabetic human keratinocytes was promoted on chitosan-collagen films immobilized with the
QHREDGS peptide. The clinical relevance was further demonstrated by assessing the
QHREDGS-immobilized chitosan-collagen hydrogel in full-thickness excisional wounds in a
db/db diabetic mouse model, which showed accelerated wound closure compared to peptide-free
hydrogel and blank wound controls. Furthermore, the accelerated wound closure was primarily
due to faster re-epithelialization and increased granulation tissue formation. There were no
observable differences in blood vessel density or size within the wound. Together, these findings
indicate that QHREDGS is a promising candidate for new wound-healing interventions that
enhance re-epithelialization and granulation tissue formation.
Contributions: Y.X. designed and performed experiments, analyzed data and prepared the
manuscript. N.F. analyzed data and prepared the manuscript. E.J.K. analyzed data. J.G. and S.C.
preformed peptide conjugation and analyzed data. C.A.L. performed Western blotting and
analyzed data. C.L. performed a preliminary migration experiment. A.P.M. provided feedback on
the design of the collective migration experiments, suggested appropriate controls and helped
with data presentation. M.R. supervised the work and wrote the manuscript.
xi
Microfabricated perfusable cardiac biowire: a platform that mimics native cardiac bundle.
Xiao Y, Zhang B, Liu H, Miklas JW, Gagliardi M, Pahnke AQ, Thavandiran N, Sun Y,
Simmons C, Keller G, Radisic M. Lab Chip 2014;14:869–82.
Tissue engineering enables the generation of three-dimensional (3D) functional cardiac tissue for
pre-clinical testing in vitro, which is critical for new drug development. However, current tissue
engineering methods poorly recapitulate the architecture of oriented cardiac bundles with
supporting capillaries. In this study, we designed a microfabricated bioreactor to generate 3D
micro-tissues, termed biowires, using both primary neonatal rat cardiomyocytes and human
embryonic stem cell (hESC) derived cardiomyocytes. Perfusable cardiac biowires were
generated with polytetrafluoroethylene (PTFE) tubing template, and were integrated with
electrical field stimulation using carbon rod electrodes. To demonstrate the feasibility of this
platform for pharmaceutical testing, nitric oxide (NO) was released from perfused sodium
nitroprusside (SNP) solution and diffused through the tubing. The NO treatment slowed down
the spontaneous beating of cardiac biowires based on hESC derived cardiomyocytes and
degraded the myofibrillar cytoskeleton of the cardiomyocytes within the biowires. The biowires
were also integrated with electrical stimulation using carbon rod electrodes to further improve
phenotype of cardiomyocytes, as indicated by organized contractile apparatus, higher Young's
modulus, and improved electrical properties. This microfabricated platform provides a unique
opportunity to assess pharmacological effects on cardiac tissue in vitro by perfusion in a cardiac
bundle model, which could provide improved physiological relevance.
Contributions: Y.X.-concept and design, performed experiments and data analysis, manuscript
writing; B.Z.-design, device fabrication; H.L.-performed AFM experiments and data analysis;
J.W.M., M.G., A.Q.P.-cardiomyocyte differentiation from human embryonic stem cells (hESCs);
N.T.-hydrogel composition development; Y.S., C.S., K.G., M.R.-concept, data interpretation,
final approval of manuscript.
xii
Table of Contents
Abstract ........................................................................................................................................... ii
Acknowledgments .......................................................................................................................... iv
Declaration of Co-Authorship ........................................................................................................ vi
Abstracts of Published Articles Appearing in the Thesis ............................................................. vii
Table of Contents .......................................................................................................................... xii
List of Tables .............................................................................................................................. xvii
List of Figures ............................................................................................................................ xviii
List of Abbreviations .................................................................................................................... xx
Chapter 1 ......................................................................................................................................... 1
1 Introduction ................................................................................................................................ 1
1.1 Overview ............................................................................................................................. 1
1.2 Hypothesis ........................................................................................................................... 2
1.3 Specific aims ....................................................................................................................... 3
Chapter 2 ......................................................................................................................................... 5
2 Literature review ........................................................................................................................ 5
2.1 Instructive biomaterials for tissue engineering ................................................................... 5
2.1.1 Motivation for instructive biomaterials .................................................................. 5
2.1.2 Naturally derived biomaterials ................................................................................ 7
2.1.3 Instructive biochemical cues provided by biomaterials ........................................ 10
2.1.4 Biomechanical instructions provided by biomaterials .......................................... 12
2.1.5 Mesenchymal stromal cells ................................................................................... 13
2.2 Cardiac tissue engineering ................................................................................................ 15
2.2.1 Motivation for cardiac tissue engineering ............................................................. 15
xiii
2.2.2 Cell sources ........................................................................................................... 15
2.2.3 Biomaterials .......................................................................................................... 20
2.3 Tissue engineering for wound healing .............................................................................. 23
2.3.1 Wound healing process ......................................................................................... 23
2.3.2 Diabetic wound healing ........................................................................................ 26
2.3.3 Current tissue engineering products for topical wounds ....................................... 27
2.3.4 Instructive biochemical cues for wound healing .................................................. 28
2.4 Cardiac tissue engineering in vitro ................................................................................... 36
2.4.1 Motivation for generating cardiac micro-tissues .................................................. 36
2.4.2 Cardiac micro-tissues as research platform .......................................................... 39
Chapter 3 ....................................................................................................................................... 42
3 Collagen patches immobilized with growth factors or peptides for cardiac regeneration ....... 42
3.1 Introduction ....................................................................................................................... 42
3.2 Materials and methods ...................................................................................................... 44
3.2.1 Materials ............................................................................................................... 44
3.2.2 Covalent immobilization of growth factors and peptides on collagen scaffolds .. 44
3.2.3 Quantification of growth factor immobilization efficiency .................................. 45
3.2.4 Quantification of QHREDGS peptide immobilization efficiency ........................ 46
3.2.5 Characterization of release profile ........................................................................ 47
3.2.6 Scanning electron microscopy .............................................................................. 47
3.2.7 Tensile testing of porous collagen scaffolds ......................................................... 48
3.3 Results and discussion ...................................................................................................... 50
3.4 Conclusion ........................................................................................................................ 54
3.5 Acknowledgments ............................................................................................................. 54
Chapter 4 ....................................................................................................................................... 55
xiv
4 Diabetic wound regeneration using peptide-modified hydrogel targeting the epithelium ....... 55
4.1 Introduction ....................................................................................................................... 55
4.2 Materials and methods ...................................................................................................... 56
4.2.1 Primary human keratinocytes cell culture ............................................................. 56
4.2.2 Evaluation of soluble QHREDGS in vitro ............................................................ 57
4.2.3 Proliferation assay ................................................................................................. 57
4.2.4 H2O2 treatment on HEKs with soluble QHREDGS peptide ................................. 57
4.2.5 Conjugation of QHREDGS to chitosan ................................................................ 58
4.2.6 Solvent casting of chitosan-collagen films ........................................................... 58
4.2.7 Coating validation ................................................................................................. 58
4.2.8 Keratinocyte attachment on chitosan-only films .................................................. 59
4.2.9 H2O2 treatment on keratinocytes on the chitosan-collagen films ......................... 59
4.2.10 EarlyToxTM Cell Integrity assay ........................................................................... 59
4.2.11 Western blotting .................................................................................................... 60
4.2.12 Migration assay ..................................................................................................... 60
4.2.13 Immunostaining .................................................................................................... 61
4.2.14 Animals, wound model, and treatment ................................................................. 61
4.2.15 Histology analysis ................................................................................................. 62
4.2.16 Microvessel analysis algorithm ............................................................................. 63
4.2.17 Statistical analysis ................................................................................................. 63
4.3 Results ............................................................................................................................... 63
4.3.1 QHREDGS peptide prevents H2O2-induced apoptosis in human primary
keratinocytes and upregulates Akt and MAPKp42/44 signaling .............................. 63
4.3.2 Immobilized QHREDGS peptide promotes human primary keratinocytes
attachment, survival and migration in vitro .......................................................... 66
xv
4.3.3 Immobilized QHREDGS peptide promotes diabetic human primary
keratinocytes attachment, survival and migration in vitro .................................... 71
4.3.4 QHREDGS-immobilized hydrogel promotes wound healing in db/db diabetic
mice ....................................................................................................................... 74
4.3.5 Accelerated QHREDGS-induced diabetic wound healing does not involve
changes in the extent of angiogenesis of the granulation tissue ........................... 79
4.4 Discussion ......................................................................................................................... 82
4.5 Conclusion ........................................................................................................................ 86
4.6 Acknowledgments ............................................................................................................. 87
Chapter 5 ....................................................................................................................................... 88
5 Microfabricated perfusable cardiac biowire: a platform that mimics native cardiac bundle ... 88
5.1 Introduction ....................................................................................................................... 88
5.2 Materials and methods ...................................................................................................... 90
5.2.1 Biowire bioreactor design and fabrication ............................................................ 90
5.2.2 Perfusion system design and fabrication ............................................................... 90
5.2.3 Cell culture ............................................................................................................ 90
5.2.4 Generation of cardiac biowires ............................................................................. 91
5.2.5 Quantification of compaction rate ........................................................................ 92
5.2.6 Immunostaining and Fluorescent Microscopy ...................................................... 92
5.2.7 Quantification of nuclei elongation and alignment ............................................... 92
5.2.8 Characterization of perfusable biowires ............................................................... 92
5.2.9 Quantification of NO perfusion ............................................................................ 93
5.2.10 NO treatment of human cardiac biowires ............................................................. 93
5.2.11 Electrical stimulation ............................................................................................ 94
5.2.12 Atomic force microscopy (AFM) ......................................................................... 95
5.2.13 Statistical analysis ................................................................................................. 95
xvi
5.3 Results ............................................................................................................................... 96
5.3.1 Generation and characterization of cardiac biowires ............................................ 96
5.3.2 Generation and characterization of perfusable cardiac biowires ........................ 100
5.3.3 NO treatment of human cardiac biowires by perfusion ...................................... 101
5.3.4 Electrical stimulation of cardiac biowires ........................................................... 103
5.4 Discussion ....................................................................................................................... 105
5.5 Conclusion ...................................................................................................................... 109
5.6 Acknowledgments ........................................................................................................... 109
Chapter 6 ..................................................................................................................................... 110
6 Discussion and conclusions .................................................................................................... 110
6.1 Discussion ....................................................................................................................... 110
6.2 Significant contributions ................................................................................................. 114
6.3 Conclusion ...................................................................................................................... 116
Chapter 7 ..................................................................................................................................... 117
7 Recommendations for future work ......................................................................................... 117
7.1 Investigate cardiac regeneration by collagen patch immobilized with QHREDGS
peptide ............................................................................................................................. 117
7.2 Determine the mechanism of accelerated keratinocyte collective migration promoted
by QHREDGS peptide .................................................................................................... 117
7.3 Improve the perfusable biowire for drug candidates with high molecular weight ......... 118
7.4 Investigate the synergy between biochemical cues and topographical cues ................... 118
References ................................................................................................................................... 120
Appendices List of publications and contributions ..................................................................... 161
Copyright Acknowledgements .................................................................................................... 165
xvii
List of Tables
Table 2-1 Principal tissue distribution and cells of origin for different collagen types in human
body ................................................................................................................................................. 8
Table 2-2 Principal properties of chitosan in relation to its use in biomedical applications ........ 10
xviii
List of Figures
Figure 2-1 Chemical structure of chitosan, comprising N-acetyl-D-glucosamine (right) and D-
glucosamine (left) units. .................................................................................................................. 9
Figure 2-2 Three classic stages of wound healing. ....................................................................... 25
Figure 2-3 Different biochemical cues provided by matrix to regulate the native cells. .............. 29
Figure 2-4 Engineering heart tissue for replacement therapeutics and in vitro models by physical
and electrical control of cells and microenvironment. .................................................................. 38
Figure 2-5 Strategies for generating 2D and 3D cardiac tissue in vitro. ....................................... 41
Figure 3-1 Reaction diagram for immobilization of growth factors or peptides on collagen
sponges using 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide HCl (EDC) and N-
hydroxysulfosuccinimide (Sulfo-NHS). ....................................................................................... 44
Figure 3-2 Characterization of scaffolds. ...................................................................................... 51
Figure 3-3 Characterisation of peptide immobilization. ............................................................... 53
Figure 4-1 Soluble QHREDGS peptide prevents H2O2-induced cell death in human primary
keratinocytes with up-regulation of Akt and MAPK phosphorylation. ........................................ 65
Figure 4-2 The presence of soluble QHREDGS peptide does not accelerate HEKs migration on
collagen coated surfaces. .............................................................................................................. 66
Figure 4-3 Immobilized QHREDGS peptide in chitosan-collagen films promotes human neonatal
primary keratinocytes survival and migration. ............................................................................. 68
Figure 4-4 The presence of the immobilized QHREDGS peptide promotes HEK attachment on
chitosan-only films. ....................................................................................................................... 69
Figure 4-5 HEKs form calcium-induced adherens junctions during migration and the accelerated
migration is not associated with a difference in cell density. ....................................................... 70
xix
Figure 4-6 Immobilized QHREDGS peptide in chitosan-collagen films promotes diabetic adult
human primary keratinocyte survival and migration. ................................................................... 72
Figure 4-7 The presence of immobilized QHREDGS peptide promotes DHEK attachment on
chitosan-only films. ....................................................................................................................... 73
Figure 4-8 DHEKs form adherens junctions during migration and the accelerated migration is not
associated with a difference in cell density. .................................................................................. 74
Figure 4-9 QHREDGS-immobilized hydrogel promotes wound healing in db/db diabetic mice. 77
Figure 4-10 Thickness of the unwounded epidermis. ................................................................... 78
Figure 4-11 An example of wound re-epithelialized after two weeks with a single treatment of
QHREDGS peptide in the chitosan-collagen hydrogel. ............................................................... 78
Figure 4-12 The improvements in the diabetic wound healing process induced by the QHREDGS
peptide are not associated with increased angiogenesis within the granulation tissue. ................ 79
Figure 4-13 QHREDGS peptide does not affect microvessel number and size within granulation
tissue. ............................................................................................................................................ 81
Figure 5-1 Cardiac bundles in native myocardium. ...................................................................... 96
Figure 5-2 Generation of cardiac biowires with microfabricated bioreactor. ............................... 97
Figure 5-3 The suture template provides topographical cues in the biowires for the
cardiomyocytes to elongate and align. .......................................................................................... 99
Figure 5-4 Generation of perfusable cardiac biowires. ............................................................... 100
Figure 5-5 Nitric oxide (NO) treatment on human tubing-templated biowires. ......................... 102
Figure 5-6 Electrical stimulation and perfusion of cardiac biowires. ......................................... 104
xx
List of Abbreviations
2D Two-dimensional
3D Three-dimensional
ABTS 2,2'-Azinobis [3-ethylbenzothiazoline-6-sulfonic acid]-diammonium salt
AFM Atomic Force Microscopy
Ang1 Angiopoietin-1
bFGF Basic fibroblast growth factor
BMP-2 Bone morphogenetic protein-2
BrdU Bromodeoxyuridine
BSA Bovine serum albumin
CABG Coronary artery bypass grafting
cDNA Complimentary DNA
CPCs Cardiac progenitor cells
cTnT Cardiac troponin T
DAPI 4',6-diamidino-2-phenylindole
DHEKs Diabetic human adult epithelial keratinocytes
DTMRI Diffusion tensor magnetic resonance imaging
EB Embryoid body
ECM Extracellular matrix
xxi
EDC 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide HCl
EDGS EpiLife Defined Growth Supplement
ELISA Enzyme-linked immunosorbent assay
eNOS Endothelial nitric oxide synthase
EPCs Endothelial progenitor cells
ESCs Embryonic stem cells
ET Excitation threshold
FDA Food and Drug Administration
GAG Glycosaminoglycan
HBOT Hyperbaric oxygen therapies
HEKs Human epithelial keratinocytes
HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)
HGF Hepatocyte growth factor
HIF Hypoxia-inducible factor
ID Inner diameter
IL-1β Interleukin 1β
iNOS Induced nitric oxide synthase
iPSCs Induced pluripotent stem cells
KGF Keratinocyte growth factors
xxii
LDH Lactate dehydrogenase
LQT3 Long QT syndrome type 3
MAPK Mitogen-activated protein kinase
MCR Maximum capture rate
MEA Microelectrode arrays
MEFs Mouse embryonic fibroblasts
MES 2-(N-morpholino)ethanesulfonic acid
MI Myocardial infarction
miRNA Micro RNA
MMP Matrix metalloproteinase
mRNA Messenger RNA
MSCs Mesenchymal stromal cells
nNOS Neuronal nitric oxide synthase
NOS Nitric oxide synthase
OD Outer diameter
PBAE Poly(β-amino esters)
PBS Phosphate buffered saline
PCL Polycaprolactone
PDGF Platelet derived growth factor
xxiii
PDMS Poly(dimethysiloxane)
PEG Poly(ethylene glycol)
PGA Polyglycolic acid
PGS Poly(glycerol sebacate)
PHD2 Prolyl hydroxylase domain 2
pHEMA Poly(2-hydroxyethyl methacrylate)
PLLA Poly(L-lactide)
PNIPAAm Poly-N-isopropylacrylamide
POC Poly(1,8-octanediol-co-citric acid)
PTFE Polytetrafluoroethylene
PU Polyurethane
ROS Reactive oxygen species
SDF Stromal-derived factor
siRNA Small interfering RNA
SMA Smooth muscle actin
SNP Sodium nitroprusside
Sulfo-NHS N-hydroxysulfosuccinimide
SVR Surgical ventricular restoration
TCP Tissue culture polystyrene
xxiv
TGF-β1 Transforming growth factor-β1
TOT Topical oxygen therapy
VEGF Vascular endothelial growth factor
α-SMA α-smooth muscle actin
1
Chapter 1
1 Introduction
1.1 Overview
Tissue engineering holds the promise of restoring or regenerating functional tissues by
combining knowledge from cell biology, material science, engineering, and medicine [1]. The
focus of tissue engineering studies has evolved from replacing damaged tissues or organs with
functional tissues generated in vitro, into creating an instructive microenvironment to regenerate
the impaired tissue in situ [2]. Meanwhile, empowered by recent advances in microfabrication
technologies, a variety of functional micro-tissue constructs were generated in vitro and have
been proposed to be cogent platforms for pre-clinical drug screening studies with better
physiological relevance compared with animal models. Not surprisingly, both tissue regeneration
in vivo and micro-tissue generation in vitro can be facilitated by the biochemical and biophysical
cues in the microenvironment. In this thesis, we aimed to create instructive microenvironments to
facilitate cell-matrix interactions in tissue engineering applications with a particular focus on
cardiac tissue engineering and wound healing applications.
Native cardiac tissue has limited regeneration capacity and can be assisted by delivery of
external cells [3]. Mesenchymal stromal cells (MSCs) is the leading candidate for cell therapies
but MSCs from aged patients exhibit limited regeneration potential compared with MSCs from
young patient [4]. We propose that collagen scaffolds immobilized with angiogenic growth
factors (vascular endothelial growth factor (VEGF) and basic fibroblast growth factor (bFGF))
can rejuvenate MSCs from aged donors and improve their cardiac regeneration potential for
surgical ventricular restoration (SVR).
Cutaneous wounds are generally capable of regeneration by themselves through a series of well-
orchestrated events [5]. However, diabetic patients suffer from chronic wounds due to disruption
of the well-coordinated cellular events involved in wound healing [6]. Impaired angiogenesis has
been the main target of current therapeutic interventions in diabetic chronic wounds yet their
clinical outcomes have been limited. A short peptide sequence derived from angiopoietin-1
(ang1), QHREDGS, has been reported to activate pro-survival pathways through integrin
2
interaction in various cell types [7-13]. Therefore, it is of our interest to investigate the pro-
survival effect of QHREDGS peptide on human keratinocytes and its efficacy in promoting
diabetic wound healing in an angiogenesis-independent manner.
Cardiovascular related toxicity is one of the leading causes of marketed drug withdrawal and
current pre-clinical cellular drug screening platforms poorly recapitulate the three-dimensional
(3D) structure of native myocardium. Emerging microfabrication techniques offer the generation
of micro-tissues with precise control and greater complexity that recapitulates native tissue [14].
We proposed that perfusable cardiac micro-tissues can be generated under guidance of a
suspended template in microfabricated bioreactors and would better recapitulate the native
myocardium with a lumen in the center mimicking the capillary. This platform would be more
physiologically relevant for drug screening applications as the pharmaceutical agents are
circulated in the capillaries in vivo.
1.2 Hypothesis
The overarching hypothesis of this thesis is that topographical and biochemical cues provided by
the matrix can facilitate cell-matrix interaction and thus regulate cell assembly, cell functions,
and tissue morphogenesis. Specifically, we hypothesized that:
1. The QHREDGS peptide can be covalently immobilized onto collagen scaffold by 1-ethyl-
3-(3-dimethylaminopropyl) carbodiimide HCl (EDC) chemistry.
2. Collagen scaffolds immobilized with angiogenic growth factors (VEGF and bFGF) will
rejuvenate MSCs from aged donors and improve their regeneration potential for cardiac
remodeling.
3. The QHREDGS peptide will promote keratinocyte survival in vitro with up-regulated Akt
and MAPK phosphorylation and improve diabetic wound closure in vivo by accelerating
re-epithelialization.
4. A suspended template will guide tissue remodeling of cardiac microtissues generated in
microfabricated bioreactors and the microtissues can be perfused with a micro-tubing
template.
3
1.3 Specific aims
Our overall goal is to design matrix systems that provide appropriate biophysical and/or
biochemical cues to assist tissue remodeling and regeneration in vivo or generate micro-tissues
with higher complexity that recapitulate the native tissue in vitro. The specific aims of the work
include:
1. Immobilize QHREDGS peptide or angiogenic growth factors onto collagen scaffold.
a) Covalently immobilize VEGF and bFGF onto collagen scaffold and characterize their
immobilization efficiency and release profile.
b) Covalently immobilize QHREDGS peptide onto collagen scaffold and characterize its
immobilization efficiency and release profile.
c) Characterize the physical properties of the collagen scaffold before and after growth
factor and peptide immobilization.
2. Investigate the pro-survival effect of QHREDGS peptide on human keratinocytes in vitro
and evaluate its efficacy in promoting diabetic wound healing in vivo.
a) Using normal human primary keratinocytes, investigate the effect of soluble
QHREDGS peptide on keratinocyte survival against H2O2 stress and collective
migration.
b) Using normal human primary keratinocytes, investigate the effect of immobilized
QHREDGS peptide on keratinocyte attachment, survival against H2O2 stress and
collective migration.
c) Using diabetic human primary keratinocytes, investigate the effect of immobilized
QHREDGS peptide on keratinocyte attachment, survival against H2O2 stress and
collective migration.
d) Evaluate the efficacy and the potential mechanism of immobilized QHREDGS peptide
in promoting wound healing in db/db diabetic mice.
4
3. Develop perfusable cardiac micro-tissues recapitulating the 3D structure of native
myocardial fibers.
a) Design a bioreactor with suspended template to guide the self-remodeling of collagen-
based hydrogel seeded with cardiac cells.
b) Characterize the cardiac cell alignment under topographical guidance from the
template.
c) Develop a perfusable biowire system with micro-tubing in the center of the cardiac
micro-tissues.
d) Evaluate the efficacy of the perfusable biowires by perfusing nitric-oxide-releasing
reagent and investigating its effect on electrophysiological properties of the cardiac
micro-tissues.
5
Chapter 2
2 Literature review1, 2, 3, 4
2.1 Instructive biomaterials for tissue engineering
2.1.1 Motivation for instructive biomaterials
Tissue engineering is defined as “an interdisciplinary field that applies the principles of
engineering and life sciences toward the development of biological substitutes that restore,
maintain, or improve tissue function.” [1] The classical tissue engineering approach to develop
biological substitutes involves incorporation of living cells into a scaffold, and cultivation of the
construct in a bioreactor. As potential alternative candidates for organ transplantations, tissue
constructs created in vitro have been used in clinical studies as biological substitutes for tissues
including skin [15], cartilage [16], bone [17], blood vessels [18, 19], heart valve [20], nerve [21],
bladder [22], trachea [23], urethra [24], and vaginal organs [25]. Moreover, modern tissue
engineering approaches have demonstrated great promises for treating and curing debilitating
health conditions including myocardial infarction [26], spinal cord injury [27, 28], osteoarthritis
[29], osteoporosis [30], diabetes [31], liver cirrhosis [32] and retinopathy [33].
1 Copyright © 2013 BioMed Central. Contents of this chapter have been published in Stem Cell Res Ther:
Thavandiran N, Nunes SS, Xiao Y, Radisic M. Topological and electrical control of cardiac differentiation and
assembly. Stem Cell Res Ther. 2013;4:14. Reuse with permission from BioMed Central. A link to the published
paper can be found at: http://www.stemcellres.com/content/4/1/14
2 Copyright © 2015 Elsevier. Contents of this chapter have been published in Methods: Xiao Y, Reis LA, Zhao Y,
Radisic M. Modifications of collagen-based biomaterials with immobilized growth factors or peptides. Methods.
2015;84:44–52. Reuse with permission from Elsevier. A link to the published paper can be found at:
http://www.sciencedirect.com/science/article/pii/S1046202315001723
3 Copyright © 2014 Cambridge University Press. Contents of this chapter have been published in: Chiu LLY, Zhang
B, Xiao Y, Radisic M. Cardiac tissue regeneration in bioreactors. Biomaterials and Regenerative Medicine,
Cambridge University Press; 2014, p. 640-668. Reuse with permission from Cambridge University Press. A link to
the published chapter can be found at:
http://ebooks.cambridge.org/chapter.jsf?bid=CBO9780511997839&cid=CBO9780511997839A046
4 Copyright © 2015 IOP Publishing. Contents of this chapter have been published in Biomedical Materials:
Davenport Huyer L, Montgomery M, Zhao Y, Xiao Y, Conant G, Korolj A, Radisic M. Biomedical Materials.
2015;10:034004. Reuse with permission from IOP Publishing. A link to the published chapter can be found at:
http://iopscience.iop.org/article/10.1088/1748-6041/10/3/034004
6
Although the health benefits are obvious, the tissue engineering industry finds itself on a “roller
coaster ride” [34-39]. After its emergence and soaring development with enthusiasm from
business community in the 1990s, tissue engineering industry entered a dark period in the early
2000s, with the capital value of publicly traded tissue engineering companies reduced by 90%
between 2000 and 2002 [38]. Organogenesis (Canton, MA) and Advanced Tissue Sciences (ATS;
La Jolla, CA), two leading companies that brought the first commercially-produced tissue
engineering products to the market, engineered skin substitutes, declared bankruptcy in 2002
[40]. The dismal performance resulted in the reassessment of tissue engineering products, which
are typically limited by their long preparation time, challenging quality control, complex
distribution chains, and short shelf-life.
After the devastating years, tissue engineering industry has rebounded alongside with recent
advances in pluripotent stem cells, including embryonic stem cells (ESCs) and induced
pluripotent stem cells (iPSCs) [41]. Meanwhile, the market of tissue engineering products
expanded significantly. The capital value for publicly traded tissue engineering companies
increased over 10-fold from 2003 to 2008 with new products entering the market [39]. The sale
of products exceeded $1.3 billion in 2007, and half of this was contributed by the Medtronic
Infuse®, a recombinant bone morphogenic protein product that is acellular [42]. During this time,
the focus of tissue engineering approach has undergone considerable evolution from replacement
to regeneration in situ because it was recognized that instead of recreating the complexity of
living substitutes for transplantation [43], we should aim to develop instructive materials that
harness the body’s innate powers of self-repair [2] as the latter products may have a faster route
to the market. In these scenarios, the matrix not only serves as a scaffold that provides
mechanical support and defines the shape of tissue constructs, but also provides a multitude of
complex stimuli to support a range of cell functions and promote tissue remodeling.
Advances in biomaterials science combined with increasing knowledge of cell biology and cell-
extracellular matrix (ECM) interactions have led to the development of biomaterials tailored to
provide appropriate biological and mechanical guidance for tissue regeneration in vivo. More
importantly, the complexities in matrix design and fabrication were achieved by innovative
technologies with precise control and improved reproducibility [44]. Here, we discuss key
aspects of designing instructive biomaterials for tissue engineering applications, under the
7
principle of facilitating the cell-ECM interactions with biochemical and mechanical cues
reminiscent of native ECM to support or induce tissue regeneration.
2.1.2 Naturally derived biomaterials
Biomaterials for tissue engineering applications can be categorized into naturally derived and
synthetic biomaterials with unique advantages of each group. Naturally derived biomaterials,
including polypeptides (e.g. collagen) and polysaccharides (e.g. chitosan), are found in many
products approved by the Food and Drug Administration (FDA). Their extensive use leads to
thorough characterization and small likelihood of side effects in new applications [45]. A key
advantage associated with naturally derived biomaterials is their general capacity to support cell
attachment, proliferation, and differentiation [46]. The inherent composition and structure
properties of naturally derived biomaterials enable biological recognition, including presentation
of receptor-binding ligands and susceptibility to cell-triggered matrix degradation and
remodeling [47]. On the other hand, synthetic biomaterials provide attractive alternatives with
greater control over material properties (e.g. biochemical cues, mechanical properties,
topography, structure etc.), simplified purification process, and reduced possibilities of
immunogenicity and pathogen transmission [47].
In the scope of this thesis, instructive biomaterials were designed based on naturally derived
biomaterials because of the orientation towards clinical transition. Specifically, biomaterials
based on collagen and chitosan have been approved by the FDA and widely used in clinical
tissue engineering applications such as wound dressings. It is our major interest to investigate the
modification of the biomaterials with immobilized biochemical cues such as angiogenic growth
factors and QHREDGS peptide. However, we think our results from naturally derived
biomaterials can be translated and will contribute to our general knowledge of instructive
biomaterials design including synthetic biomaterials as well.
8
2.1.2.1 Collagen
Table 2-1 Principal tissue distribution and cells of origin for different collagen types in human body
(Reproduced from reference [48])
Collagen Type Principal Tissue Distribution Cells of Origin
I
Loose and dense connective tissue;
collagen fibers
Fibroblasts and reticular cells; smooth
muscle cells
Fibrocartilage
Bone Osteoblasts
Dentin Odontoblasts
II Hyaline and elastic cartilage Chondrocytes
Vitreous body of eye Retinal cells
III
Loose connective tissue; reticular fibers Fibroblasts and reticular cells
Papillary layer of dermis Smooth muscle cells; endothelial cells
Blood vessels
IV Basement membranes Epithelial and endothelial cells
Lens capsule of eye Lens fibers
V
Fetal membranes; placenta Fibroblasts
Basement membranes
Bone
Smooth muscle Smooth muscle cells
VI Connective tissue Fibroblasts
VII Epithelial basement membranes Fibroblasts
Anchoring fibrils Keratinocytes
VIII Cornea Corneal fibroblasts
IX Cartilage
X Hypertrophic cartilage
XI Cartilage
XII Papillary dermis Fibroblasts
XIV (undulin) Reticular dermis Fibroblasts
XVII P170 bullous pemphigoid antigen Keratinocytes
Collagen is the most abundant component of mammalian ECM and exists in tissues including
skin, bone, cartilage, cornea and blood vessels (Table 2-1) [48]. Native collagen has unique
9
triple helix fibril structure and type I, II, III, V and XI collagen are known to form collagen fibers
[49]. Three α chains, each of which are based on the sequence -Gly-X-Y-, assemble into collagen
molecule. While 29 distinct collagen types have been characterized, type I collagen is most
widely used for tissue engineering applications [49]. Different forms of type I collagen products,
including freeze-dried sheets, pastes, pads, powder and hydrogels, have been derived using
techniques such as direct decellularization of ECM and chemical extraction in acid solutions [50],
neutral salt solutions [51], or proteolytic solutions [52]. For tissue engineering applications,
physical, chemical, and enzymatic crosslinking techniques have been developed to improve
mechanical properties and enzymatic resistance of collagen biomaterials [49]. Collagen scaffolds
have been applied as skin grafts (e.g. INTEGRA™ Matrix Wound Dressing), vascular implants
(e.g. Artegraft®), orthopaedic filings (e.g. Foundation®) and nerve guides (e.g. NeuraGen®).
Furthermore, micro-tissue analogues have been generated in vitro based on self-assembly of
collagen-based hydrogels encapsulating cells and serve as platforms for research in cell biology
and pharmaceutical development [53].
2.1.2.2 Chitosan
Figure 2-1 Chemical structure of chitosan, comprising N-acetyl-D-glucosamine (right) and D-
glucosamine (left) units.
Chitosan is a cationic polysaccharide derived from chitin, the second most abundant natural
polymer, by replacing at least 60% acetyl groups along chitin chain with amino groups (which
corresponds to a deacetylation degree of 60) (Figure 2-1) [54]. Due to these amino groups,
chitosan can be protonated in acidic solutions and become soluble. Indeed, chitosan is the only
positively charged naturally occurring polysaccharide [55]. Besides, the amino groups may form
complex with metals thus chitosan is used for waste water treatment [54]. More importantly, the
10
amino groups may be quaternized or reacted with aldehyde groups under mild conditions through
reductive amination [54]. Therefore, various chitosan derivatives have been developed for a
multitude of applications in agriculture, waste treatment, food industry, cosmetics, and
biopharmaceutics [54].
For tissue engineering applications, chitosan possesses unique advantages such as antimicrobial
activity [56], mucoadhesion [57], analgesic effects [58], haemostatic properties [59], and
stronger mechanical properties compared to other naturally derived biopolymers [60]. Moreover,
chitosan can be biodegraded into non-toxic residues mainly by lysozymes [61]. Thus, chitosan
and its derivatives have been applied extensively in biomedical applications such as wound
dressings [62], drug delivery [63], bone grafts [64], and medical coatings [65] (Table 2-2).
Table 2-2 Principal properties of chitosan in relation to its use in biomedical applications
(Reproduced with permission from Elsevier, Rinaudo M. 2006 [54])
Potential biomedical applications Principle characteristics
Surgical sutures Biocompatible
Dental implants Biodegradable
Artificial skin Renewable
Rebuilding of bone Film forming
Corneal contact lenses Hydrating agent
Time release drugs for animals and human Nontoxic, biological tolerance
Encapsulating material Hydrolyzed by lysozyme
Wound healing properties
Efficient against bacteria, viruses, fungi
2.1.3 Instructive biochemical cues provided by biomaterials
The bioactivity of biomaterials is primarily conferred by the molecular information from the
basic scaffold material together with any embedded macromolecules, such as growth factors [2].
Growth factors are potent regulators that facilitate a multitude of cell activities including
migration, proliferation, differentiation, and survival. Since their mass production became
available through recombinant protein technology [66], growth factors have been extensively
applied to regulate different stages of tissue regeneration process in preclinical and clinical
studies.
11
For example, basic fibroblast growth factor (bFGF) was administrated in a canine myocardial
infarct model to enhance neovascularization and improve left ventricular function [67]. Another
potent angiogenic growth factor, vascular endothelial growth factors (VEGF), was administrated
both intra-arterially and intra-muscularly in a rabbit model of chronic hindlimb ischemia and
significantly augmented revascularization [68, 69]. Platelet derived growth factor (PDGF) was
the first growth factor approved by the FDA (Regranex approved in 1997) as an adjunct to
proper ulcer care in the treatment of lower extremity diabetic neuropathic ulcers [70]. Bone
morphogenetic protein-2 (BMP-2) is another growth factor that has been approved by the FDA
for clinical applications (Infuse® bone graft) to promote interbody spinal fusion (approved in
2002) [71], tibial fracture recovery (approved in 2004) [72], and sinus augmentation (approved in
2007) [73, 74].
Peptides are short functional amino acid sequences that are derived from primary receptor-
domains of specific proteins. Different peptides have been identified in numerous primary
proteins and some of them were designed synthetically to enable novel properties such as stimuli
responsiveness [75] and self-assembly [76]. The most well-characterized peptide is the integrin-
binding Arg-Gly-Asp (RGD) sequence, which can be found in the sequence of many ECM
proteins, including fibronectin, collagen type IV, and laminin [77-79]. RGD sequence has been
extensively applied in modifications of synthetic biomaterials to provide cell-adhesion sites on
bio-inert materials [47]. Other peptide sequences have been derived from various growth factors
[80], bringing the advantage of cost-effectiveness in scalable chemical synthesis compared to
recombinant growth factors productions. While retaining comparable bioactivities, peptides are
more stable than native growth factors, which may simplify the preparation and distribution of
biomaterials [2]. Moreover, the bioactivity of peptide may be more specific compared to the
native growth factors, which are potent regulators of a multitude of cellular activities and
sometimes associated with increased risks of systematic side effects, such as carcinogenesis [81].
To overcome the short half-life and enzymatic degradation of free growth factors in solution,
controlled-release strategies are frequently adopted [2]. Dynamic release of different growth
factors with independent release profile might provide more leverage over cell behavior than
indiscriminate delivery, recapitulating the dynamic regulation in developmental pathways [82,
83]. Moreover, localized and sustained delivery of growth factors would decrease the dosage
12
required while supraphysiological dosing may lead to systemic side effects such as hypotension
(VEGF) and nephrotoxicity (bFGF) [84]. Synergistic signaling of growth factors and ECM
proteins has been described in recent studies [85], which indicates necessities of sequestering
growth factors within the matrix, preventing their degradation and presenting them to cell-
surface receptors. Various growth factor immobilization methods have been used when
designing instructive biomaterials with growth factors to localize, enhance and sustain their
bioactivities [85-88].
2.1.4 Biomechanical instructions provided by biomaterials
There is growing recognition of the biomechanical cues provided by biomaterials that regulate
cell activities and tissue morphogenesis, with our expanding knowledge of mechanobiology and
emergence of technologies such as microfabrication and 3D printing. Incorporation of
topographical cues brings another dimension in biomaterials design, providing unique
opportunities in building tissue analogues involving multiple cell types, and could potentially
simplify biomaterials preparation.
Coupled by focal adhesions between the cytoskeleton and ECM, cells can pull on the matrix not
only to generate the traction forces, but also to sense the mechanical properties and structure of
the ECM, potentially through the bonding dynamics of integrin adhesion receptors [89]. The
importance of mechanical properties of ECM is first demonstrated on stem cell differentiation
with the first study showing different lineage and phenotype commitment of mesenchymal stem
cells seeded on substrates with varied rigidity [90]. Specifically, stiff substrates were myogenic,
soft substrates were neurogenic, and comparatively rigid substrates were osteogenic [90]. Recent
studies showed that stem cell fate can also be influenced by past mechanical environments,
suggesting a “mechanical memory” on stem cell differentiation [91]. Moreover, nano- and
micro-scale topographic features have been recently controlled to regulate stem cell renewal [92,
93]. Taken together, translation of biomechanical regulation from in vitro to in vivo becomes
more addressable with the wide range of emerging materials and analytical technologies and
future instructive biomaterial design may require synergistically providing both biomechanical
and biochemical factors [94].
13
Cell alignment is critical for tissue-scale functionality in muscular tissues and can be regulated
by topographical guidance from substrates as well. Heart tissue possesses complex structural
organization on multiple scales. On macro-scale, native myocardium contains elongated
myofibers aligned in parallel; the structure enables coordinated contraction of the ventricle and
expulsion of blood. On micro-scale, adult cardiomyocytes are rod shaped and contain registries
of sarcomeres that enable cell contraction in response to electrical signals. On nano-scale, each
sarcomere contains precisely organized sarcomeric proteins (e.g. sarcomeric α-actin/α-actinin
and myosin heavy chain) that enable coordinated contractions of sarcomeres. Cardiomyocytes
alignment was first achieved by confining cell attachment on bio-inert substrates patterned with
cell-adhesive sites, such as fibronectin [95, 96]. Larger scale patterning aided with high-
resolution diffusion tensor magnetic resonance imaging (DTMRI) has created cardiomyocytes
monolayer that recapitulates cross-sections of native cardiac tissue [97]. Micro-and nano-scale
patterned grooves were designed to guide cardiomyocyte alignment as well and the tissue
constructs displayed anisotropic action potential propagation and contractility characteristic of
native heart tissue [98].
3D cardiac micro-tissues have been created by microfabrication technology and demonstrated 3D
alignment of heart cells more closely resemble native myocardium [99]. Constructs containing
both parallel channels and micro-pores were designed to guide multicellular organization with
cardiomyocytes predominantly occupying the parallel channels and grouping into bundles and
micro-pores remained acellular for mass transfer [100]. Other templates have been integrated to
guide the formation of 3D micro-tissues formed by hydrogel compaction driven by contractile
cells [44]. The anisotropic cell organization within 3D micro-tissues is critical for
electromechanical properties and better recapitulates the native heart tissue.
2.1.5 Mesenchymal stromal cells
In some tissues, such as the heart and brain, the native tissue has intrinsically limited
regeneration potential while the other tissues age with a declining progenitor population, thus
requiring the aid from external cell sources to regenerate [2]. The primary criteria for choosing
the external cells include easy access to autologous source, large population available after short
ex vivo expansion, capability of replenishing native cells or regenerating the native tissue. MSCs,
currently defined by their fibroblast-like morphology, adherence to plastic, expression of a
14
specific set of surface antigens (CD105+, CD90+, CD73+), and capacity for osteogenic,
chondrogenic, and adipogenic lineages in vitro [101], are the leading candidate for cell therapy
and have demonstrated considerable promise in tissue regeneration in pre-clinical and clinical
studies [102].
A significant advantage of using MSCs in clinical application is that they can be readily obtained
from a variety of tissues including bone marrow [103], adipose tissue [104], placenta [105], skin
[106], umbilical cord blood [107], umbilical cord perivascular cells [108], umbilical cord
Wharton’s jelly [109], dental pulp [110], amniotic fluid [111], synovial membrane [112], and
breast milk [113]. Moreover, MSCs proliferate rapidly in vitro with up to 18-fold increase after 2
weeks of culture [114]. Therefore, a large number of MSCs may be available readily to meet the
dose requirement for clinical trials (up to millions of cells/kg body weight).
Although the exact mechanism of how MSCs exert their regenerative benefits remains to be fully
defined, several potential mechanisms have been suggested by recent studies. (1) MSCs may
transdifferentiate into specific cell type and replenish the damaged tissue, which has been
reported in clinical trials to treat bone and cartilage defects [115, 116]. (2) Fusion of MSCs with
endogenous cells may improve their regeneration capacity, which has been mainly described in
cardiac tissue [117, 118]. (3) MSCs modulate both adaptive and innate immuno systems by
suppressing T cells and modulating inflammatory cytokines expression [119, 120]. (4) The
secretome from MSCs, the spectrum of regulatory and trophic factors secreted by MSCs,
including growth factors, cytokines, and chemokines, exhibit paracrine effects that stimulate
regeneration capacity of endogenous cells [121, 122].
Clear understanding of the mechanism is important to optimize the design of MSCs clinical
studies to maximize the therapeutic benefits. For example, acellular approach would be more
cost-effective than MSCs transplantation if the therapeutic benefit of their secretome is well-
defined and could be further enhanced by targeted preconditioning and genetic manipulation
[121]. For MSCs transplantation, poor retention at target location necessitates engraftment within
delivery vehicle and various biomaterials have been used for different tissue engineering
applications [123-126]. With a growing body of literature reporting conflicting results, the
interactions between MSCs and tumor cells requires special attention for the safety concerns
related to MSCs clinical applications [127]. Also, limited regeneration potential associated with
15
MSCs from aged patients compared to the ones from younger patients raises reevaluation
between autologous and allogenic cell sources for aged patients [128].
2.2 Cardiac tissue engineering
2.2.1 Motivation for cardiac tissue engineering
Myocardial infarction (MI) leads to the death of cardiomyocytes, and the infarct area becomes
replaced by a fibroblastic scar tissue that has no contractile function. This reduces the pumping
ability of the heart and the cardiac output. In addition, the scar tissue thins due to the lack of
vasculature to provide oxygen and nutrients to the infarct site, thus leading to high wall stress
and cardiac dilatation, which may ultimately lead to heart failure. Heart failure triggered by MI is
a leading cause of death globally [129].
The adult heart has a limited regenerative capacity. The shortage of donor organs further
suggests a need to develop new treatment strategies for cardiovascular diseases. Cardiac tissue
regeneration can be achieved through several strategies, including (1) gene therapy, (2) cell
transplantation, and (3) implantation or injection of biomaterials or engineered cardiac tissues.
The goal of these cardiac tissue regeneration strategies is to repair the damaged myocardium
through supporting vascularization and cell survival, in turn reducing wall thinning and
preventing dilatation and heart failure.
Since the damaged myocardium has limited capacity to regenerate, cell transplantation can
replace the damaged and lost cells, thus attenuating pathological remodeling. However, cell
transplantation is limited by the washout of cells from the injection site and the inability of
injected cells to integrate with the native tissue. Solutions include delivering cells within a
scaffold or hydrogel to support cell engraftment, and growing functional cardiac tissues that can
be implanted and would integrate with the native tissue.
2.2.2 Cell sources
Generally, cells for cardiac tissue engineering should be 1) expandable to achieve high numbers
of cells; 2) compatible with the host without causing immune reactions; and 3) able to survive
and maintain function in vitro and in vivo [130]. Cells for cardiac tissue engineering can be
16
categorized into primary cardiac cells, ESCs, iPSCs, resident cardiac progenitor cells, and adult
progenitor cells from other native tissues.
2.2.2.1 Fetal and neonatal cardiac cells
Adult cardiac cells are the target cell source in cardiac regeneration since they are differentiated
cardiomyocytes with developed contractile apparatus and can integrate with host cardiomyocytes
through gap junctions and intercalated disks. However, they are present in low numbers and have
little proliferative and developmental potential. The advantages of using fetal and neonatal
cardiac cells include their greater proliferative and developmental potential than that of adult
cardiac cells. These cells also showed efficacy in cardiac regeneration, since the injection of fetal
or neonatal cardiomyocytes attenuated pathological ventricular remodeling by forming viable
grafts, increasing the ventricle thickness, and improving left ventricular function [131, 132]. By
contrast, injected adult cardiomyocytes were unable to survive both in acutely cryoinjured
myocardium and in granulation tissue. Contractile cardiac tissues have also been engineered
using fetal or neonatal rat cardiomyocyte-enriched cell populations [133-135]. However, fetal or
neonatal cardiac cells are not available in large numbers to provide the millions of
cardiomyocytes required for cardiac tissue regeneration, thus motivating the search for
alternative cell sources. Moreover, while fetal or neonatal cardiac cells from animals can readily
be used, a direct translation to human studies will not be possible due to obvious ethical
considerations involving the use of human cells.
2.2.2.2 Embryonic stem cells
ESCs have been investigated intensively due to their ability to undergo indefinite self-renewal
without losing the capacity to differentiate into all cell types. ESCs are commonly cultivated and
differentiated in 3D aggregates known as embryoid bodies (EBs). Quantification of “beating”
EBs is commonly applied to assess cardio-genicity in ESCs. However, despite robust cardiomyo-
genicity in EBs, only 1%–5% of their total cell number consists of cardiomyocytes under
standard culture conditions (i.e. plated on 0.1% gelatin-coated culture dishes and cultured in 80%
knockout Dulbecco’s Modified Eagle’s Medium with 20% fetal bovine serum, 1 mM l-glutamine,
0.1 mM mercaptoethanol, and 1% non-essential amino acid stock [136]). The most efficient and
reproducible protocols to date for differentiation of cardiomyocytes from pluripotent stem cells
17
are those that have replicated the signaling pathways that regulate lineage commitment in the
early embryo. With this approach, the earliest stages of cardiovascular development in ESC
differentiation cultures were mapped, identifying a multipotent cardiovascular progenitor that
displays the capacity to generate cardiac and vascular progeny [137, 138]. In mouse Flk1 and in
humans KDR expression can be used to enrich for cardiac-specified mesoderm [139]. When
isolated from the differentiated EBs and cultured as a monolayer, these progenitors generate
contracting cardiomyocytes [137, 138]. As these progenitors differentiate, they progress through
the developmental stages thought to be involved in the establishment of the cardiovascular
lineages in vivo, for which specific cytokines are required. The combination of activin A and
BMP4 on days 1–4 of EB differentiation induces a primitive-streak-like population and
mesoderm development. Subsequent application of WNT inhibitor DKK1 and KDR ligand
VEGF165 significantly enhances the differentiation of KDR+ progenitors into cardiomyocytes
[137, 138], while bFGF is added to support continued expansion of cardiovascular lineages.
ESC-derived cardiomyocytes have been used to engineer cardiac tissues. Guo et al. [140] seeded
mouse ESC-derived cardiomyocytes into circular molds with collagen type I and Matrigel to
produce engineered heart tissues. Stevens et al. [141, 142] tricultured human ESC-derived
cardiomyocytes with mouse embryonic fibroblasts and ESC-derived endothelial cells or human
umbilical vein endothelial cells to generate scaffold-free cardiac patches.
2.2.2.3 Induced pluripotent stem cells
The recent advent of iPSCs has been considered as a solution for the ethical issues and
immunological concerns associated with the usage of ESCs. Briefly, fully differentiated cells can
be reprogrammed to have pluripotency like ESCs by introducing a set of transcription factors
including Oct3/4, Sox2, c-Myc, and Klf4 [143]. Although the efficiency of current protocols for
derivation of iPSCs is still low, iPSCs can serve as a novel cell source for cardiac tissue
engineering. Previously, Mauritz et al. [144] generated functional cardiomyocytes from mouse
iPSCs. Importantly, human iPSCs, which were derived from reprogramming of adult fibroblasts,
were differentiated into cardiomyocytes [145, 146]. The derivation of cardiomyocytes from
iPSCs allows the generation of autologous human cardiomyocytes necessary for cardiac tissue
regeneration [146]. Human embryonic stem cell and human iPSC-derived cardiomyocytes were
previously cultured in collagen matrices using uniaxial mechanical stress conditioning and the
18
addition of endothelial cells and stromal supporting cells [147]. The engineered human
myocardium was transplanted onto hearts of athymic rats. The grafts resembled host
myocardium and contained microvessels that were perfused by the host coronary circulation.
Fibroblasts, with their demonstrated ability to be transdifferentiated into skeletal muscle cells
[148], have been considered as an attractive cell source for cardiac tissue engineering. Efe et al.
[149] reprogrammed mouse embryonic fibroblasts (MEFs) directly into cardiomyocytes through
over-expression of Oct4, Sox2, Klf4 and c-Myc. Reprogramming fibroblasts into cardiomyocytes
is a very appealing approach since “on-site” therapy would be possible if the reprogramming
could be done directly in the infarct area.
However, it should also be noted that clinical applications of iPSCs are still under development
and are currently challenged by the possibility of tumor development after transplantation [150,
151]. Specifically, carcinogenesis can be caused by the integrated oncogenes used in
reprogramming, c-Myc in particular [152], by insertional mutagenesis from viral vectors [153],
by disruption of tumor suppressor genes [154], and by genetic and epigenetic abnormalities in
the reprogrammed cells [155].
2.2.2.4 Cardiac progenitor cells
Recent studies have revealed evidence for DNA synthesis and an increase in the number of
cardiomyocytes in diseased human hearts [156, 157]. Although it occurs at a very slow rate, the
regeneration of cardiomyocytes in the heart seems to be an appealing solution for myocardial
infarction. Endogenous regeneration of the myocardium may be due to cardiomyocytes re-
entering the cell cycle and dividing, or the proliferation and differentiation of resident
populations of cardiac progenitor cells (CPCs) [158]. Genetic manipulation or application of
bioactive molecules can be used to alter the cell cycle control in adult cardiomyocytes in order to
promote cell proliferation.
Urbanek et al. [159] reported that the increase in cardiac mass after human aortic stenosis is a
result of combined myocyte hypertrophy and hyperplasia. The number of cells expressing stem
cell markers and telomerase was increased in aortic stenosis. The cell clusters containing these
stem cells made the transition to cardiogenic and myocyte precursors. There was also evidence of
primitive myocytes turning into terminally differentiated myocytes. In a separate study, Urbanek
19
et al. [160] found a cardiac stem cell pool in the human heart that is capable of promoting
myocardial regeneration after infarction. The number of cardiac stem cells increased both in
acute and in chronic infarcts. These cells were telomerase-competent dividing stem cells that
were committed to the myocyte, smooth muscle, and endothelial cell lineages. This suggests the
activation of cardiac stem cells in response to ischemic injury. However, chronic infarcts had
fewer functionally competent cardiac stem cells in the viable myocardium than were present in
acute cases, as indicated by the higher expression of markers of cellular senescence (e.g. p16 and
p53), shorter telomeres, and greater apoptosis in cardiac stem cells of chronic infarcts. This
underlies the progressive cardiac deterioration that leads to terminal cardiac failure in chronic
infarcts. It was later reported that the myocardium has interstitial structures with stem cell niches
that contain resident cardiac stem cells and lineage-committed cells [161]. These cells are
connected through gap and adherens junctions to myocytes and fibroblasts, which act as
supporting cells. The cardiac stem cells divide both symmetrically and asymmetric- ally, with a
dominance of asymmetric division in which the cells give rise to one daughter cardiac stem cell
and one daughter committed cell. This preserves primitive cells while generating myocyte
progenitors, endothelial cells, and smooth muscle cells.
Laugwitz et al. [162] reported the identification of isl1+ cardiac progenitors in post-natal rat,
mouse, and human myocardium. When co-cultured with neonatal myocytes, these isl1+ cells
underwent highly efficient conversion into the mature cardiac phenotype. Subsequent studies
have identified CPCs in post-natal hearts with the expression of surface markers such as c-kit and
Sca-1, and physiological properties such as the ability to efflux fluorescent dye and to form
multicellular spheroids [163, 164]. However, these cell subpopulations are insufficient for
cardiac repair without appropriate signaling. Bioactive molecules and biomaterials can be used to
recruit these cells to the infarct region for in situ cardiac repair. In addition, these cells can be
isolated and then expanded in vitro for further use in cardiac tissue regeneration.
Different resident stem cell populations have been used for cardiac repair. The injection of lin−,
c-kit+ cells into the ischemic heart improved the regeneration of the myocardium [165]. In
addition, cardiomyocytes were derived from the isl1+ cardiac progenitor population [166]. These
cells were then used to generate beating thin films [159]. Adult cardiac progenitors can also be
isolated from explant cultures of human endomyocardial biopsies, and can be expanded as
20
cardiospheres in vitro [167, 168]. These cardiosphere-derived cells improved cardiac
regeneration in mice through direct differentiation and the secretion of angiogenic growth factor
to improve cell survival [169]. However, it remains a challenge to generate high numbers of
cardiac progenitor cells for cardiac tissue engineering.
2.2.2.5 Bone marrow cells
Bone-marrow-derived stem cells have elicited considerable interest in cardiac tissue engineering
because there were reports of hematopoietic stem cells regenerating into cardiomyocytes [170].
Adult bone marrow cells have been used to regenerate and improve the functional properties of
the myocardium [171-174]. Marrow-derived stromal cells, also referred to as mesenchymal stem
cells, have been applied in clinical trials, and this resulted in a significant improvement in
ejection fraction [175]. However, subsequent studies showed that the improvement brought about
by bone marrow cells was more likely due to their secretion of soluble factors and to their
vasculogenic potential after transplantation rather than their differentiation into cardiomyocytes.
2.2.2.6 Adipose-derived cells
Besides bone marrow cells and cardiac progenitor cells, the adipose tissue is another source of
autologous adult stem cells. A significantly higher density of MSCs is present in the adipose
tissue than in the bone marrow (5% compared with 0.01%) [176]. Adipose-derived cells have
been shown to improve cardiac tissue regeneration by direct differentiation and secretion of
paracrine factors [177-179].
2.2.3 Biomaterials
The cells described above hold great promise in cardiac regeneration yet face the challenge from
poor engraftment at the target site. Delivery within a supporting matrix can improve the
engraftment of cells by preventing cell loss during delivery, and providing a protective
environment afterwards. Moreover, the matrix provides mechanical support to alleviate the
increased wall stress caused by functional cardiomyocytes loss. The biomaterials applied can be
categorized into porous scaffolds and hydrogels and each of them have advantages over the other.
Porous scaffolds provide stronger mechanical support during cell delivery and thereafter while
hydrogel systems can be delivered in a minimally invasive manner.
21
Many natural and synthetic biomaterials have been fabricated into scaffolds for cardiac tissue
engineering by techniques such as freeze-drying, salt casting, and electro-spinning. Scaffolds can
be made of naturally derived biomaterials such as collagen [180, 181], gelatin [135], and alginate
[133]. Synthetic biomaterials have also been extensively investigated such as polyglycolic acid
(PGA) [182], polycaprolactone (PCL) [183], poly(L-lactide) (PLLA) [134, 184], polyurethane
(PU) [185], poly(1,8-octanediol-co-citric acid) (POC) [186], and poly(glycerol sebacate) (PGS)
[99]. More recently, decellularization technique provides an attractive method of generating
naturally derived scaffolds (mainly composed of collagen) with proper cardiac architecture [187].
Different hydrogel systems have been developed to deliver cell therapies for cardiac regeneration.
Naturally derived hydrogels have been investigated such as collagen [188, 189], fibrin [190, 191],
chitosan [192, 193], alginate [194, 195], and Matrigel [196, 197]. Hydrogels based on synthetic
polymers including poly(ethylene glycol) (PEG) [198], poly-N-isopropylacrylamide (PNIPAAm)
[199, 200], and poly(2-hydroxyethyl methacrylate) (pHEMA) [199, 201] have been developed to
deliver cells for cardiac regeneration as well.
In cardiac tissue engineering, biomaterials serve predominately as scaffolds for tissue formation
and vehicles for the delivery of engineered tissues [202-207]. Scaffolds for cardiac tissue
engineering require a number of criteria to be carefully considered to allow for optimal tissue
function including: physical properties of the polymer (e.g. strength and elasticity), degradation
rates, and host response [204]. Furthermore, these properties help to dictate the body's elicited
immune response.
To satisfy the dynamic nature of heart function and myocardial remodelling post-MI, the ideal
cardiac biomaterial should account for several design parameters. Matching the mechanical
properties of the myocardium is an important cardiac biomaterial property [208]. The Young's
modulus of the adult human myocardium ranges non-linearly from 10–20 kPa (start of diastole)
to 200–500 kPa (end of diastole) [209-212]. A rigid and inelastic patch placed on the heart will
impede contraction. A scaffold should not be constructed too soft as pathological cardiac dilation
can be reduced by mechanically reinforcing the myocardium [213]. In addition, materials
capable of achieving tissue-like compliance (e.g. hydrogels) must allow for easy
handling/suturing. An ideal biomaterial should also comply with the changes in strain
experienced by the myocardium of approximately ±15% [214-216]. The anisotropic
22
(directionally dependent) stiffness of the heart tissue [216-220] is an important design parameter
that the scaffold should replicate. Mathematical modelling and in vivo experiments on dogs
demonstrated that the patches with anisotropic mechanical properties (Dacron with slits), placed
onto an infarcted myocardium resulted in improved functionality compared to isotropic patches
[221, 222]. The cyclic contraction of the myocardium necessitates a biomaterial that is
elastomeric. The patch should biodegrade over the desired period, matching the remodelling
process of the heart post MI, that usually takes 6–8 weeks, to avoid fibrous capsule formation
and a chronic inflammatory response in the myocardium. Ensuring that the material degrades at
the same rate as the heart heals is crucial for success [223].
Comprehending the immune response to an implanted biomaterial is required to determine the
clinical relevance. The immediate response of the host to implantation of a foreign material is
known as the tissue response [224-227]. Upon implantation, host proteins adhere to the
implanted biomaterial surface. These proteins signal monocytes and other leukocytes to begin the
foreign body cascade [227-229]. Macrophage recruitment and differentiation dictates the
formation of foreign body giant cells and fibrous capsule formation [227-229]. The way a
biomaterial influences macrophage polarization has been found to influence the host immune
response to implanted devices significantly. Specifically, M1 macrophages induce an
inflammatory response in the body, while M2 macrophages encourage tissue repair [227-229].
Material design looks to minimize the inflammatory and immune response as well as the fibrous
capsule formation. Formation of a fibrous capsule inhibits the efficacy of a tissue scaffold, as
cells are not able to integrate and interact with the surrounding cells, and the apparent stiffness of
the material is increased [230]. Furthermore, this limits vascularization, which is important as it
facilitates growth through delivery of blood, as well as ensures the viability of exogenously
applied cells [231]. Many designs of cardiac biomaterials aim to encourage the polarization of
macrophages to the M2 phenotype, in order to minimize fibrous capsule formation and promote
incorporation into host tissue [227, 229, 232]. Minimization of fibrous capsule formation, as well
as reduction in the foreign body reaction and chronic inflammation, are considered benchmarks
to biocompatibility assessment [203, 226, 233, 234].
23
2.3 Tissue engineering for wound healing
2.3.1 Wound healing process
Wound healing is a complex process involving a highly regulated cascade of biochemical and
cellular events coordinated to achieve restoration of tissue integrity following tissue damage or
loss [5, 235]. The most extensively studied wound healing process is cutaneous wound healing as
skin is the largest organ of the human body and the outermost barrier. The skin is composed of
two main layers, the actual barrier, epidermis, and the underneath supporting layer, dermis. The
epidermis is mainly composed of hard, cornified keratinocytes, which maintain skin integrity and
serve as the physical barrier against external environment. The dermis is the residence of various
cell types, including dermal fibroblasts, macrophages, lymphocytes, and mast cells. The cells
within dermis, together with the fibrous and amorphous ECM between them, provide tensile
strength, support, and moisture retention to the skin. Moreover, the dermis is where blood vessels
are located, which provides oxygen and blood to the skin.
Clinically, cutaneous wound healing process can be characterized as repair by first, second and
third intention [236]. Primary intention healing occurs when tissue is incised in a clean manner
and re-approximated (i.e. after surgical procedures) and generally results in less scarring because
the incisional defect re-epithelizes rapidly and is sealed by matrix deposition. Secondary
intention healing occurs in open wounds where the wound edges are not approximated and the
defect heals by formation of granulation tissue, wound contraction and migration of epithelial
cells. Third intention healing (also called delayed primary intention healing) occurs when a
wound is allowed to heal open for a few days (usually due to gross contamination), and then
closed as in primary healing.
Cutaneous wound healing consists of a symphony of well-orchestrated events that can be divided
into three overlapping phases: inflammation, proliferation and remodeling. Right after injury, a
fibrin clot (a meshwork of mainly fibrin with platelets embedded) forms to stop bleeding
(hemostasis) and protect the wound. Meanwhile, platelets release a wide range of growth factors
such as PDGF to recruit inflammatory cells, among which neutrophils are the first cell type
coming from the bloodstream to the wound. At this time, the wound is hypoxic due to the
damage of the blood vessels immediately after injury. This seemingly deleterious situation
24
actually has beneficial effects as it increases keratinocytes migration, early angiogenesis,
fibroblasts proliferation, and the transcription and synthesis of crucial growth factors and
cytokines, including PDGF, VEGF, and transforming growth factor-β1 (TGF-β1) (Figure 2-2a).
25
Figure 2-2 Three classic stages of wound healing. (a) Inflammation. The wound is characterized by a
hypoxic (ischemic) environment in which a fibrin clot has formed. Bacteria, neutrophils and platelets are
abundant in the wound. (b) New tissue formation. An eschar (scab) has formed on the surface of the
wound. Most cells from the previous stage of repair have migrated from the wound, and new blood
vessels form (angiogenesis). The migration of keratinocytes can be observed under the eschar. (c)
Remodelling. Disorganized collagen has been laid down by fibroblasts in the wound. Re-epithelialization
and contraction contributed to the final closure of the wound. (Reproduced with permission from Nature
Publishing Group, Gurtner G. et al. 2008 [5])
The second phase is proliferation and is marked by the appearance of granulation tissue, a robust,
dark pink, granular-appearing tissue that consists of fibroblasts, macrophages, newly forming
ECM, and developing blood vessels. Fibroblasts are recruited to the wound by PDGF (secreted
by platelets and macrophages) and deposit new matrix, including collagen, elastin, reticulin,
proteoglycans, and glycosaminoglycans (GAGs). Monocytes from circulation take up residence
at the wound site as tissue macrophages, which serve a dual role in wound healing process:
between inflammatory and proliferation phase, macrophages shift from a proinflammatory
secreting phagocyte that clears debris to an anti-inflammatory initiator of repair. Macrophages
also serve a dual role in angiogenesis: they first respond to hypoxia by promoting angiogenesis,
and later, they respond to interferons by inhibiting angiogenesis. Some fibroblasts differentiate
into a tension-generating phenotype known as myofibroblasts induced by TGFβ1.
Myofibroblasts express α-smooth muscle actin (α-SMA) and participate in wound contraction
through their extensive cell-matrix contacts (supermature focal adhesions) which are linked to
their intracellular stress fibers. Keratinocyte migration is mainly mediated by keratinocyte
growth factors (KGF-1 and KGF-2), which are produced by fibroblasts. Importantly,
keratinocytes do not bind to the fibrin clot, but to the dermal matrix, therefore providing an
efficient debridement of eschar from the dermal layer while re-epithelializing. Coverage by
keratinocytes over the wound site marks the final step of the repair phase as these cornified
epithelial cells provide the outermost cellular barrier to the environment. (Figure 2-2b)
The last phase of wound healing is remodeling, which mainly involves collagen maturation and
reorganization. Over time, the immature dermal matrix is reorganized to a stronger, more mature
scar mainly due to realignment of the collagen fibers from a random orientation to specific
structure according to function. The remodeling is usually accompanied by regression of blood
vessels and degradation of excess ECM synthesized at earlier stages. (Figure 2-2c)
26
2.3.2 Diabetic wound healing
Optimum cutaneous wound healing process requires a well-orchestrated integration of the
complex biological and molecular events of cell migration and proliferation, and of ECM
deposition and remodelling. This complicated process can be described as several overlapping
phases: inflammation, new tissue formation (angiogenesis and re-epithelialization), and
remodeling. The transition between phases is mediated by appropriate and precise regulation
from inflammatory mediators, growth factors, cytokines, and mechanical forces [5, 235].
However, this orderly healing process can be impaired by systemic complications such as
diabetes mellitus, and stalled in a specific phase resulting in chronic wounds that fail to heal [6].
Chronic wounds are pervasive worldwide, affecting more than 70 million people, with senior
population especially vulnerable [237]. Many face harsh medical and social consequences.
Specifically, diabetic foot ulcer affects 15% of people with diabetes and is the leading cause of
nontraumatic amputation [238].
Denervation of local sympathetic nerve system is a feature of diabetic neuropathy and results in
sensory deficits, so that the diabetic patients do not respond to external insults including pressure
and heat [239]. Absence of protective symptoms against the persistent insults leads to further
deformities and increased infection rates [6]. Meanwhile, diabetic wound healing is challenged
by vasculopathy including impairments in both vasculogenesis and angiogenesis. Vasculogenesis
is impaired in diabetic wounds due to insufficient mobilization of circulating endothelial
progenitor cells (EPCs) from the bone marrow [240] and impaired homing to the wound site with
diminished expression of the chemokine stromal derived factor-1α (SDF-1α) [241]. Impaired
endothelium function also involves a reduction of nitric oxide synthesis and faster nitric oxide
degradation due to the presence of excess reactive oxygen species (ROS) [242]. Moreover,
excessive glycosylation of matrix proteins induced by hyperglycemia leads to crosslinks between
molecules in the basement membrane of ECM and impairs angiogenesis [243]. Excessive
activation of some matrix metalloproteinases (MMPs) such as MMP9 can impair cell migration
and lead to breakdown of some critical matrix proteins and growth factors [244].
On the cellular level, the resident cells in diabetic chronic wounds have been reported to be
associated with abnormalities compared to those from normal wounds. Fibroblasts isolated from
diabetic chronic ulcers are senescent with decreased responses to various growth factors
27
including transforming growth factor-β1 (TGF-β1), platelet derived growth factor (PDGF),
epidermal growth factor (EGF), insulin-like growth factor-1 (IGF-1), basic fibroblast growth
factor (bFGF) [245-248]. Macrophages in diabetic wounds show a decrease in release of
cytokines, including tumor necrosis factor α (TNF-α), interleukin 1β (IL-1β), and vascular
endothelial growth factor (VEGF) [249]. Keratinocytes attachment and migration, the critical
events in re-epithelialization, are impaired in diabetic chronic wounds due to altered ECM
composition and an enhanced ECM degradation rate [250, 251]. Meanwhile, all the resident cells
in the wound site are stressed by excessive production of reactive oxygen species (ROS) from
macrophages and neutrophils due to consistent inflammation, coupled with an impaired
antioxidant defense capability in response to hyperglycemia [252-254].
2.3.3 Current tissue engineering products for topical wounds
Wound dressing was the first category of FDA-approved tissue engineering products and has
seen numerous innovations since then. In general, wound dressings can be categorized into
acellular dressing and dressings with cells. Based on how acellular dressings interact with the
native wound environment, they can be further grouped into bio-inert dressings and bioactive
dressings.
Bio-inert dressings mainly serve as a physical barrier and keep the wound environment moist.
Winter et al. first described that moisture-retaining dressings speed epithelialization of acute,
superficial wounds in pigs compared with air-exposed wound [255] and similar results were
observed in humans later [256]. Moreover, some conformable bio-inert dressings can absorb
exudate in the draining wounds [257].
A group of bioactive acellular wound dressings were developed to deliver bioactive agents to the
local wound environment. Antibiotic drugs have been delivered in wound dressings that include
streptomycin [258], minocycline [259], vancomycin [260], neomycin [261] and ciprofloxacin
[262, 263]. Antiseptic agents have been extensively applied such as iodine-releasing agents (e.g.
Hyiodine®), silver-releasing agents (e.g. SilverSeal®), iodine (e.g. Iodozyme™), chlorhexidine
(e.g. BACTIGRAS◊), and nitric oxide [264]. Having been applied in cutaneous wound treatment
for decades [265], natural honey received a lot of attention recently as the underlying
28
mechanisms of honey-promoted wound healing became better understood, which mainly
involves antimicrobial and anti-inflammatory effects [266].
Another important function of bioactive wound dressing is providing the matrix for wound
regeneration, which is critical for angiogenesis and re-epithelialization. Type I collagen is used
most often for this purpose due to its close similarity to the native ECM [267]. Scaffolds based
on chitosan [62], cellulose [268], fibrin [269], gelatin [270, 271], silk [270], and alginate [272]
have been investigated as well. More recently, a bioelectric dressing that generates physiologic
levels of micro-current (2-10 μA) (Procellera®) has been reported to accelerate wound healing by
promoting re-epithelialization [273].
Living skin substitutes consist of bioactive dressing hosting dermal cells and/or epidermal cells.
With current techniques, epidermal grafts capable of covering the entire surface area of the body
can be generated from a 3-cm2 biopsy from autologous tissue [274]. However, these living skin
substitutes are still limited by their cost in terms of time and money needed for preparation, short
shelf-life and difficulties in storage [15]. More recently, MSCs from various sources have also
been reported to promote both normal and diabetic wound healing and many clinical trials are
under way [275].
2.3.4 Instructive biochemical cues for wound healing
The importance of biochemical cues on cellular behavior and tissue morphogenesis has been well
recognized and utilized in tissue engineering applications. When designing biomaterials,
biochemical property is often the first to consider as the bioactivity of biomaterials is primarily
conferred by the molecular information from the basic scaffold material together with any
embedded macromolecules, such as growth factors [2]. For wound healing applications, growth
factors and derivatives, bioactive matrix materials (e.g. collagen and chitosan), small bioactive
molecules (e.g. oxygen and nitric oxide), and genetic regulators (e.g. cDNA, siRNA, and miRNA)
have been implemented in instructive matrix designed for regulating wound healing (Figure 2-3).
29
Figure 2-3 Different biochemical cues provided by matrix to regulate the native cells. Growth factors
and derivatives can interact with native cells through their specific receptors. The composition of ECM
proteins are usually recognized by integrins. Small bioactive molecules including oxygen and nitric oxide
can diffuse into the cells and mainly affect mitochondria activities. Genetic regulators including
complimentary DNA (cDNA), small interfering RNA (siRNA) and micro RNA (miRNA) can be
delivered by non-viral vehicles and facilitate gene transcription and translation.
2.3.4.1 Growth factors and derivatives
Various growth factors have been delivered within wound dressings, including epidermal growth
factor (EGF) [276], VEGF [277], bFGF [278], transforming growth factor-β (TGF-β) [279] and
the FDA-approved PDGF [70]. Conventional delivery methods often apply the growth factors as
soluble form which result in burst release that can be affiliated with severe systematic side
effects such as hypotension (VEGF) and nephrotoxicity (bFGF) [84]. Growth factor variants
have been developed to render desirable properties including solubility [280], improved retention
[281], ECM affinity [85]. Moreover, short peptide sequences have been developed to recapitulate
the bioactivity of target growth factors by chemical synthesis with better cost-effectiveness.
Recently, novel delivery strategies other than providing growth factors as soluble supplements
have been explored. Synergistic signaling between growth factor receptors and integrin ligands
30
has been proposed, which motivates immobilizing the growth factors in close affinity to ECM
proteins [282]. Hubbell et al. demonstrated that the growth factors delivered by this method can
be effective at lower dosage, which is important for both eliminating potential side effects by
supraphysiological dosage and improving cost effectiveness of growth factors therapies [282].
2.3.4.2 Small bioactive molecules
2.3.4.2.1 Oxygen
It is with no doubt that among all the chemicals, oxygen is the most vital to the human body due
to its involvement in cellular respiration and energy production. Upon injury, the wound site
immediately suffers from ischemia due to the disruption of vasculature and the tissue oxygen
tension in chronic wounds has been transcutaneously measured to be lower than normal tissues
[283]. Historically, hyperbaric oxygen therapies (HBOT) have been applied exposing the body
intermittently to pure oxygen under pressure in a stationary pressure chamber [284]. HBOT
delivers oxygen through systematic circulation therefore its efficacy is limited in tissues with
poor circulation. Since the early 1960s, topical oxygen therapy (TOT) has been developed, which
typically involves applying pure oxygen with sealing around wound tissue for a mean of 90 min,
once a day at an absolute pressure slightly above atmospheric pressure [285]. More recently,
Oxygen-releasing wound dressings (e.g. Oxyzyme™) have been developed to promote wound
healing by addressing cellular hypoxia after tissue damage [284]. Instead of attaching a bag filled
with pure oxygen to the wound, Oxyzyme™ system generates oxygen by chemical reaction.
The importance of oxygen in wound healing is well documented as increased energy is
demanded in the granulation tissue for cellular activities throughout wound healing including
bacterial defense, cell proliferation, and cell migration. Once recruited to the wound site, the
bactericidal activity of leukocytes is positively correlated to local oxygen concentration [286].
HBOT has been recently shown to reverse the diabetic defect in endothelial progenitor cell
mobilization from bone marrow, which is critical for angiogenesis [287]. During remodeling
phase, fibroblast proliferation and collagen synthesis are oxygen dependent [288, 289]. Other
than directly supplying energy for cellular metabolism, oxygen has been found critical for
growth factor signaling as well [290].
31
The metabolically more active cells in the wound consume large amount of oxygen and this,
together with interrupted blood supply, contributes generation of a hypoxia gradient in the
wound tissue. Recent discovery of hypoxia induced factor α (HIF 1-α) highlighted the
importance of hypoxia gradient in wound healing process [291]. This hypoxia gradient is
important and sometimes prerequisite for effective wound healing and stabilized HIF 1-α
expression is critical to improved diabetic wound healing [292]. However, it should be noted that
the hypoxia gradient only benefits wound healing process with transient presence while chronic
hypoxia would result in wounds that fail to heal.
2.3.4.2.2 Nitric oxide
Ever since establishing its important role in physiological regulation as the endothelial relaxing
factor [293], nitric oxide (NO) has been intensively studied and its extensive physiological
impact has been revealed in virtually all organ and tissue systems under both normal and
pathological conditions [294]. Specifically, NO has been reported to be involved throughout the
three phases of cutaneous wound healing and function as an important diffusible, gaseous
regulator of angiogenesis, inflammatory response, and collagen deposition [295]. Over the past
two decades, many NO delivery devices and vehicles have been developed to transit the
therapeutic potential of NO to improve cutaneous wound healing.
NO is generated by the enzyme nitric oxide synthase (NOS), which catalyzes the conversion of
amino acid L-arginine to L-citrulline. Three NOS isoforms have been identified: two
constitutively expressed isoforms (endothelial NOS (eNOS) and neuronal NOS (nNOS)) and one
inducible isoform (iNOS) [296-298]. The expression of iNOS is induced by a variety of
cytokines, growth factors, and inflammatory stimuli on target cells which lead to high expression
levels and NO output compared with eNOS and nNOS [299]. All three NOS isoforms have been
found in skin tissue: nNOS has been observed in keratinocytes and melanocytes [300, 301];
eNOS have be detected in keratinocytes of the basal epidermal layer, dermal fibroblasts,
endothelial capillaries, and eccrine glands [302, 303]; and iNOS have be induced in keratinocytes
[304, 305], dermal fibroblasts [302], and endothelial cells [306].
The implication of NO in wound healing process has received a lot of attention since studies
showing correlation between cutaneous wound healing process and increased levels of NO
32
metabolites (e.g. nitrite and nitrate) [307], mRNA and protein expression of all three NOS
isoforms [308-310]. Moreover, inhibition of NOS by competitive inhibitors decreases collagen
deposition and breaking strength of wounds and impairs healing [307]. The critical role of NO in
wound healing is further proved by both eNOS and iNOS knockout mice, which showed serious
deficit in cutaneous wound healing [311, 312]. Importantly, there are also strong correlations
between impaired wound healing in diabetic animals and decreased NOS expression and activity,
and/or NO levels [309, 313, 314].
It is evident that NO participates throughout the three phases of wound healing. First of all, NO
has been applied as an antimicrobial agent [264]. During the early inflammatory phase, NO
regulates infiltration of monocytes and neutrophils by activating pro-inflammatory cytokines (e.g.
IL-8 and TGF-β1) and also serving as chemoattractant by itself [315-317]. NO is involved in
proliferative phase as it has been reported to promote angiogenesis and keratinocytes migration.
Importantly, NO is vital to the activity of VEGF as blockade of NOS prevents VEGF-induced
endothelial cell proliferation and mitogen-activated protein (MAP) kinase [318]. NO also
mediates angiogenesis directly by promoting ECs proliferation and migration [319]. Moreover
NO has been found to promote proliferation of keratinocytes and inhibits their apoptosis [320,
321]. At the final phase of cutaneous wound healing, NO predominantly regulates collagen
synthesis in fibroblasts. Treatments with NO donors, dietary L-arginine, or iNOS overexpression
all enhance collagen deposition in the wound [322-324].
The wide-ranging functionalities of NO in the wound healing process motivated numerous
attempts to implement NO as therapeutic agent to improved wound healing outcomes. However,
as a gas molecule NO is extremely difficult to handle owing to its instability and potential to be
oxidized into toxic nitrogen dioxide molecule. Therefore, NO delivery devices vehicles including
S-nitrosothiols [325, 326], diazeniumdiolates (NONOates) [327, 328], and nano particles[324,
329, 330] have been designed and demonstrated improved wound healing outcomes.
2.3.4.3 Genetic regulators
Genetic therapies have been developed originally using viral vectors and successfully applied in
promoting wound healing [331]. The general principle is to deliver nucleic acids that are
developed either for blocking harmful genes or for restoring the activity of defective genes.
33
However, the use of viral vectors in humans faces safety challenges such as immune reactions.
Therefore genetic regulators including naked DNA [332] and RNA interference [333] delivered
by non-viral vectors have been developed. Efficient delivery and targeted release are the main
challenges for non-viral based genetic therapies and recent advances such as electroporation have
significantly improved the efficiency [334]. Here we review recent advances in developing
genetic regulators delivered by non-viral vectors with a specific focus on wound healing
applications.
2.3.4.3.1 Complementary DNA (cDNA)
Complementary DNA (cDNA) is a DNA copy synthesized from the target messenger RNA
(mRNA) that encodes specific protein via reverse transcription. Non-viral vectors including
cationic polymers, cationic liposomes, and naked plasmids have been designed to deliver cDNAs
encoding for peptides (e.g. LL-37 [335], secretoneurin [336]) and growth factors (e.g. VEGF
[337], KGF [338]) to regulate different wound healing phases and promote skin regeneration.
Sonic hedgehog is a prototypical morphogen that plays essential roles during embryonic
development and it has been shown to regulate postnatal tissue remodeling and regeneration by
promoting angiogenesis [339]. Asai et al. demonstrated that topical sonic hedgehog gene therapy
delivered by plasmid vector accelerated wound healing in type 2 diabetic mice by promoting
microvascular remodeling [340]. Park et al. delivered sonic hedgehog DNA intradermally as
polyplexes formed with biodegradable cationic poly(β-amino esters) (PBAE) and reported
similar angiogenic benefits and improved wound healing [341].
The safety of delivering cDNA using naked plasmids has been demonstrated in a number of
recent clinical studies using intramuscular injections of hepatocyte growth factor (HGF) DNA
plasmids to treat critical limb ischemia [342-345]. Importantly, local delivery of HGF plasmids
did not cause peripheral edema and did not increase systemic HGF protein level [343]. Moreover,
therapeutic benefits including reduced pain, increased ankle-brachial index and improved wound
healing have been demonstrated in these studies and some ischemic ulcers healed completely
[342-345].
cDNA transfection usually results in transient expression of exogenous genes and methods have
been developed to prolong the gene expression. Kulkarni et al. encapsulated lipoplexes carrying
34
eNOS encoding gene in fibrin microspheres and formed a “fibrin in fibrin” temporal release
system [346]. Electrospun fibers have been immobilized with plasmids to deliver growth factors
such as bFGF [347] and EGF [348].
2.3.4.3.2 Small interfering RNA (siRNA)
While cDNA transfection enables production of a functional protein, small interfering RNAs
(siRNAs) can be delivered to silence deleterious genes by complimentary binding to the mRNA
sequences of the corresponding target genes. Therefore, cDNA and siRNA can serve as controls
for each other in recovery experiments designed for specific pathways [349, 350]. More
importantly, siRNA-based post-transcriptional modification methods have been applied in
wound healing applications to achieve transient local functional ablation of malignant genes at
different phases of wound healing including angiogenesis and ECM remodeling. Overview of
siRNA-based therapies for other diseases such as cancer can be found in other excellent review
papers [351, 352].
Chen et al. identified INT6/eIF3e as regulator of hypoxia-inducible factor 2α (HIF2α) activity
and they reported that a single siRNA-Int6 application promoted neoangiogenesis by
accumulating HIF2α and downstream transcription of angiogenic factors in a hypoxia-
independent manner [353]. Using a previously developed agarose matrix topical delivery system,
Nguyen et al. achieved near-complete local knockdown of p53, a cell cycle regulator, and
accelerated wound healing in diabetic mice with increased vasculogenic cytokines including
VEGF and SDF-1 [354]. Wetterau et al. delivered prolyl hydroxylase domain 2 (PHD2) siRNA
with agarose matrix in diabetic mice and suppression of PHD2 increased the expression of
HIF1α and subsequent angiogenic regulators [355].
Remodeling phase is another therapeutic target of wound healing as dysregulated remodeling
could result in hypertrophic scars that are unaesthetic. Lee et al. first delivered Smad3 siRNA to
inhibit skin fibrosis induced by radiation [356]. Similarly, Wang et al. inhibited activation of
TGF-β/Smad signaling cascade by applying TGF-β type I receptor siRNA and reported reduced
hypertrophic scarring [357]. To sustain the release of these TGF-β/Smad targeting siRNAs,
delivery matrices have been designed including trimetyl chitosan [358] and a pressure-sensitive
35
hydrogel [359]. MMP-responsive nanofibrous matrix was developed for diabetic wound
remodeling to control release MMP-2 siRNA and restored the wound recovery rates [360].
More recently, Charafeddine et al. developed nanoparticles encapsulating siRNA to deplete
Fidgetin-like 2 in vivo and found accelerated cell migration potentially caused by increased
directional cell motility modulated by microtubule growth [361]. In another study, Randeria et al.
demonstrated that delivery of siRNA that suppress ganglioside-monosialic acid 3 synthase, a
critical mediator of insulin resistance, in spherical nucleic acid gold nanoparticle conjugates was
able to reverse impaired diabetic wound healing [362].
2.3.4.3.3 MicroRNA (miRNA)
Besides the genetic information that is transacted by proteins, recent evidence shows that the
majority of the genome is actually transcribed into noncoding RNAs. MicroRNAs (miRNAs) are
a major group of these noncoding RNAs, and are important regulators of the coding genes by
posttranscriptional gene regulation. They are small (approximately 22-nuleotide long)
endogenously formed repressors that usually bind to the 3’-untranslated region of the target
mRNAs. Importantly, their interactions are non-complimentary and a single microRNA can
regulate multiple mRNAs simultaneously and a single mRNA can be regulated by various
microRNAs [363]. Recent studies suggest that miRNAs play important roles in dermal wound
healing including regulating angiogenesis, re-epithelialization and wound remodeling [364, 365].
Angiogenesis is an important step in wound healing and microRNAs have been shown to be
important regulators [366]. Bonauer et al. first discovered that miR-92a targets mRNAs
corresponding to several important angiogenic proteins and systematic inhibition of miR-92a
improves angiogenesis during myocardial tissue recovery [367]. In diabetic wounds, Chan et al.
showed that downregulating miR-200b supports angiogenesis and accelerates healing by
desilencing GATA binding protein 2 and VEGF receptor 2 [368]. In another study, they
described that downregulation of miR-199a-5p promotes angiogenesis both in vitro and in vivo
by inducing Ets-1 and MMP-1 expression [369]. Other studies have demonstrated the promise of
pretreatment of angiogenic cells (e.g. MSCs) with microRNAs to improve wound healing
outcomes after cell implantation [370].
36
Re-epithelialization is another essential step of wound healing and has been a target for
microRNA interventions. Biswas et al. found that hypoxia induces miR-210 expression and
down-regulate the cell-cycle regulatory protein E2F3, resulting in impaired wound closure with
limited keratinocyte proliferation [371]. miR-21 has been reported to be induced by TGF-β [372]
and promote keratinocytes migration in vitro and re-epithelialization in vivo [373, 374]. However,
Pastar et al. reported contradicting results showing that miR-21 inhibits wound healing with
suppressed leptin receptor (LepR) signaling [375]. miR-203 has been discovered as an important
regulator of mRNAs responsible for both keratinocyte proliferation and migration, including
RAN and RAPH1 [376]. Sundaram1 et al. identified miR-198 as an important regulatory switch
in controlling multiple gene expressions to facilitate re-epithelialization [377]. Li et al. identified
miR-31 as another key regulator to promote keratinocyte proliferation and migration with
epithelial membrane protein 1 as its direct target [378]. In another study, they identified miR-132
as a regulator to facilitate transition from inflammation to proliferation phase and its implication
in chronic wounds, which are stalled in the inflammation phase with impaired re-epithelialization
[379].
Another important phase of wound healing is remodeling and it is essentially related to scar
formation that is usually caused by dysregulated collagen production and remodeling [380]. Kato
et al. first identified miR-192 as a regulator of TGF-β-induced collagen expression in diabetic
kidney glomeruli [381]. miR-29 has been identified as a key regulator of collagen expression in
systemic sclerosis [382] and collagen deposit in skin fibroblasts [383]. Yang et al. reported that
downregulation of miR-155 at wound sites does not accelerate wound closure but leads to a
reduced fibrosis with less collagen and α-SMA expression [384].
2.4 Cardiac tissue engineering in vitro
2.4.1 Motivation for generating cardiac micro-tissues
Cardiac tissue engineering in vitro aims to manipulate the microenvironment cells interact within
in order to facilitate cell assembly and build functional tissue with the goal of providing
replacements for diseased or damaged native tissues. Additionally, engineered heart tissue may
serve as increasingly accurate in vitro model for studies in normal and diseased heart physiology,
as well as drug discovery, validation, and toxicology [385-387]. With the advent of serum-free
37
cardiac differentiation protocols [388-392] comes the ability to generate large quantities of
cardiomyocytes derived from human pluripotent stem cell sources for engineered heart tissue.
Additionally, cardiomyocyte-specific surface markers have been identified and microfluidic cell
separation methods have been advanced which can be used to purify heterogeneous populations
[393-395].
The adult mammalian heart is composed of a complex and well-integrated mosaic of anatomical
modules. The contractile muscle (atria, and ventricles) positioned between the supporting epi-
and endocardium, the conduction system (pacemaker nodes, and Purkinje fiber network), and the
highly dense vasculature (endothelial and smooth muscle cells) constitute the key elements of the
cardiac system, which is the engine for the larger cardiovascular system. In the heart, the many
cell types form specific integrated structures which contribute to their individual cell and overall
organ function. To engineer these cells in the appropriate positions and to temporally give them
the correct biochemical, physical, and electrical cues is the overarching goal (Figure 2-4).
38
Figure 2-4 Engineering heart tissue for replacement therapeutics and in vitro models by physical
and electrical control of cells and microenvironment. Depiction of current methods used to
manipulate heart cells to develop, mature, and assemble into functional heart tissue. Tuning the cell
microenvironment by means of geometry and electrical control exhibits upstream effects on adhesion,
cell-cell and cell-extracellular matrix interactions, growth and differentiation, cellular and tissue
alignment via cytoskeletal organization, and electrical and contractile apparatus. The small dark arrows in
the flow diagrams indicate the sequence by which the specific method of microenvironmental control
effectively manifests downstream. These end changes in the cardiac cells include changes in gene/protein
expression, electrical properties, and mechanical properties. Top: during development pluripotent stem
cells differentiate into mesodermal progenitors, then cardiovascular progenitors that give rise to various
cell types in the heart (cardiomyocytes, fibroblasts, endothelial and smooth muscle cells). Cell
differentiation and assembly into a highly organized structure is governed by biochemical, mechanical
and electrical stimuli in vivo. Tissue engineering aims to recapitulate some of these environmental factors
in vitro. Middle: control of substrate topography and stiffness affects cell orientation and, as a result,
functional properties. Bottom: control of electrical properties is achieved by use of conductive
biomaterials, electrical stimulation bioreactors or changes in gene expression of key ion channels. The
large green arrows (middle and bottom) depict the span of current techniques used in the field and link
them to the regimes of cardiac differentiation and assembly where they have been applied (top). CM,
cardiomyocyte; CVP, cardiovascular progenitor; E-C, excitation-contraction; EC, endothelial cell; ECM,
extracellular matrix; ET, excitation threshold; FB, fibroblast; MCR, maximum capture rate; PSC,
pluripotent stem cell; SMC, smooth muscle cell.
(Reproduced with permission from BioMed Central, Thavandiran N. et al. 2013 [44])
39
2.4.2 Cardiac micro-tissues as research platform
Serving as models to discover novel therapeutics or drug toxicities in vitro, single
cardiomyocytes cultured in vitro have been well-characterized by various methods, such as
immunostaining and patch-clamping [396-398]. However, single cardiomyocytes are not
necessarily an appropriate model for all toxicology studies, and do not lend themselves to in vivo
applications. Therefore, two-dimensional (2D) and 3D tissue constructs have been developed to
better recapitulate the higher functions of tissues containing a multitude of cardiomyocytes.
Studies that have focused on the development of different tissue constructs are outlined below.
Cardiomyocytes cultured as monolayers enable cell-cell coupling by gap junctions, which plays
an important role in the action potential propagation in cardiac tissue. Moreover, the alignment
and cell-cell interaction in the monolayer could be controlled by micro-patterning methods. Karp
et al. patterned photocrosslinkable chitosan on glass and tissue culture polystyrene to create cell-
repellent regions and cardiomyocytes on the cell-adhesive regions were able to beat
spontaneously after one week [399]. Nima et al. cultured cardiomyocytes following micro- and
macroscopic guidance of fibronectin coating and recapitulated the directions of native cardiac
fibres characterized by high-resolution diffusion tensor magnetic resonance imaging [97].
Microelectrode arrays (MEA) have been developed to culture cardiomyocyte monolayers and
provide insight on cardiac field potentials by impedance measurements [400-402]. Natarajan et
al. created cardiomyocyte monolayers aligned with MEAs by patterning fibronectin to guide
cellular attachment thus controlling action potential propagation [403] (Figure 2-5A). Creating
topographical cues on culture substrates guides the alignment of cardiomyocytes, which results
in the desired characteristics of anisotropy and improved physiological properties [404, 405].
Moving from 2D culture of cardiomyocyte monolayers, researchers from Parker’s group
developed contractile cardiomyocyte thin films on elastomers to measure contractile forces
combined with a quantification of action potential propagation (Figure 2-5B) [406]. This high-
throughput platform has been used to provide new insights into the pathophysiology underlying
the cardiomyopathy of Barth syndrome [407].
Native cardiac tissue has a 3D structure where it works to generate force against a load. This
structure is replete with extracellular matrix and cellular interactions on all geometric faces of
40
cardiomyocytes, as well as a topography and stiffness which 2D substrates have a difficult time
replicating. 3D scaffolds made of natural or synthetic polymers can create an environment on
which dense cardiac tissue can be seeded that has properties comparable to the human
myocardium [387]. More recently, technologies including stereolithography and femtosecond
laser scanning have been applied to precisely control the geometry and structure of scaffolds. Ma
et al. seeded cardiomyocytes derived from long QT syndrome type 3 (LQT3) IPS-CMs onto
synthetic filamentous matrices fabricated by femtosecond laser and studied the cardiac
contractility in response to a panel of drugs [408].
Gel compaction has been a widely-applied method to generate cardiac tissue constructs as the
self-assembled constructs produce increased force of contraction due to the higher cell density
after the gel compaction [409]. During cultivation, the cells in the hydrogel contract and generate
pulling forces that lead to hydrogel compaction and this process highly depends on the cell
population and the hydrogel stiffness [410]. Importantly, placing microfabricated constraints to
guide the gel compaction process can control the geometries of the final cardiac microtissue.
Moreover, during cultivation, the cells developed cell-cell and cell-matrix interactions to
compact the cell-gel mixture and cell alignment was then mediated by the stress due to resistance
of constraints. Hansen et al. developed a technique to construct a large series of fibrin-based mini
engineered heart tissues that developed high yield and reproducibility (Figure 2-5C) [411].
Boudou et al. generated arrays of cardiac microtissues using small cell number (~5000 cardiac
cells) per construct with potential in high-throughput drug screening (Figure 2-5D) [412].
Thavandiran et al. showed that appropriate population percentage (25%) of nonmyocytes played
critical roles in the tissue remodeling dynamics and enhanced final structure and function
properties of cardiac microtissues [410]. The biowire platform (Figure 2-5E) used suspended
suture templates to create longitudinal microtissues that remained stable for weeks and generated
more matured cardiac tissues with electrical stimulation after the gel compaction stage [413].
Xiao et al. further developed this platform by replacing the suture template with micro-tubing to
mimic the capillaries in cardiac bundles. This created a novel method to introduce
pharmaceutical agents to cardiac tissue by perfusion and the negative inotropic effect of nitric
oxide has been demonstrated as an example [414].
41
Figure 2-5 Strategies for generating 2D and 3D cardiac tissue in vitro. (A) Cardiomyocyte
monolayers cultured on patterned MEA for guided action potential propagation. (B) Cardiomyocyte
monolayers cultured on flexible elastomer films for contractile force measurement. (C) High-throughput
platform for monitoring contractile activities of an array of fibrin-based engineered heart tissues. (D)
Cardiac micro-tissues around micro-cantilevers to measure contractile properties. (E) Cardiac biowires
set-up with a suspended template to guide tissue formation and cell alignment. The phenotype of the
cardiomyocytes were matured under external electrical stimulation and the drug candidates were applied
by perfusion through micro-tubing within the biowire, providing improved physiological relevance.
(Reproduced with permission from IOP Publishing, Davenport Huyer L. et al. 2015 [53])
42
Chapter 3
3 Collagen patches immobilized with growth factors or peptides for cardiac regeneration5, 6
3.1 Introduction
Tissue engineering aims at generating tissues and organs to regenerate and restore the structure
and function of their pathologically altered native counterparts [1]. Biomaterials for tissue
engineering do not merely serve as a mechanical support, but rather provide an instructive
microenvironment that facilitates cellular behaviors and tissue regeneration. General principle of
designing these biomaterials is to mimic the biological and physical properties of native ECM.
Bioactive agents such as growth factors and peptides have been incorporated in biomaterials to
promote or enhance the tissue regeneration process [415].
Growth factors are potent regulators that modulate many cellular functions including migration,
proliferation, differentiation, and survival. Various growth factors have been applied to facilitate
different stages of specific tissue regeneration process. For example, basic fibroblast growth
factor (bFGF) was administrated in a canine myocardial infarct model to enhance
neovascularization and improved left ventricular function [67]. Another potent angiogenic
growth factor, VEGF, was administrated both intra-arterially and intramuscularly in a rabbit
model of chronic hindlimb ischemia and significantly augmented revascularization [68, 69].
PDGF was the first growth factor approved by US FDA (Regranex approved in 1997) as an
adjunct to proper ulcer care in the treatment of lower extremity diabetic neuropathic ulcers [70].
However, clinical translations of growth factors have been limited, partly because of
5 Copyright © 2015 Elsevier. Contents of this chapter have been published in Methods: Xiao Y, Reis LA, Zhao Y,
Radisic M. Modifications of collagen-based biomaterials with immobilized growth factors or peptides. Methods.
2015;84:44–52. Reuse with permission from Elsevier. A link to the published paper can be found at:
http://www.sciencedirect.com/science/article/pii/S1046202315001723
6 Copyright © 2012 Elsevier. Contents of this chapter have been published in J Am Coll Cardiol: Kang K, Sun L,
Xiao Y, Li SH, Wu J, Yao TM, Weisel RD, Radisic M, Li RK. Aged human cells rejuvenated by cytokine
enhancement of biomaterials for surgical ventricular restoration. J Am Coll Cardiol. 2012;60:2237–2249. Reuse
with permission from Elsevier. A link to the published paper can be found at:
http://www.sciencedirect.com/science/article/pii/S0735109712043677
43
supraphysiological uses of them risk systemic side effects such as hypotension (VEGF) and
nephrotoxicity (bFGF) [84]. In 2008, Regranex received boxed warning from FDA as patients
treated with three or more tubes of it had increased rate of mortality secondary to malignancy
[416]. Recently, we and others have demonstrated that controlled release of growth factors by
immobilization onto biomaterials localizes, enhances and sustains their bioactivities [85-87]. As
a result, the growth factors could be delivered locally and remain effective at a lower total dose,
addressing the issues for their clinical transitions.
Peptides are short amino acid sequences that are derived from primary receptor-domains of
specific proteins. The adhesion-promoting peptide derived from fibronectin, RGD, is the most
intensively-investigated peptide and has been used to promote cell attachment on different
surfaces and materials [77]. Our group has recently described a novel ang1fibrinogen-like
domain based peptide, QHREDGS [7]. It has demonstrated pro-survival benefits on cardiac cells
[7-9], endothelial cells [10], osteoblasts [12], and human iPSCs [11]. The advantages of using
small peptides is that they are more stable than their growth factor counterparts and also less
susceptible to conformational changes during immobilization or encapsulation processes [2].
Moreover, peptide sequences can be synthesized and purified in more cost-effective manner
compared to recombinant human proteins.
Traditional biomaterials for tissue engineering applications could be categorized into porous
scaffolds and hydrogels and both of them have merits that are exclusive of the other. Porous
scaffolds provide strong mechanical support and guidance for cell growth with the overall
architecture and porous structures tunable by different fabrication methods [417]. Hydrogel
systems can be delivered in a minimally invasive manner with or without encapsulating cells to
promote tissue regeneration [418]. With recent advances in microfabrication, hydrogel systems
have also been applied to generate micro-tissues in vitro for drug screening applications [410,
413, 419, 420]. Here, we discuss modification and characterization of collagen-based
biomaterials in the forms of both porous sponge and hydrogel systems with growth factors or
peptides by covalent immobilization using 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide
(EDC) chemistry.
44
3.2 Materials and methods
3.2.1 Materials
Avitene® Ultrafoam™ (2 mm thick, 8 cm × 12.5 cm) was purchased from Davol Inc. Ultrapure
PROTASAN™ chitosan salt (UP G 213, viscosity = 20-200 mPa·s, Mw = 200-600 kDa, 75-90%
DA) was purchased from NovaMatrix®. 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide
hydrochloride (EDC, Cat# 22980) and N-hydroxysulfosuccinimide (Sulfo-NHS, Cat#24510)
were purchased from Thermo Fisher Scientific Inc.
3.2.2 Covalent immobilization of growth factors and peptides on collagen scaffolds
The carboxyl groups (-COOH) in collagen sponge react with N-hydroxysulfosuccinimide (Sulfo-
NHS) in the presence of EDC, resulting in a semi-stable Sulfo-NHS ester intermediate, which
may then be reacted with primary amine groups (-NH2) in growth factors or peptides to form
stable amide crosslinks (Figure 3-1). The semi-stable intermediate enables us to perform step
immobilization, which has shown enhanced immobilization efficiency compared to bulk
immobilization [87]. We use phosphate buffered saline (PBS) as a buffer rather than distilled
water or 2-(N-morpholino)ethanesulfonic acid (MES) according to previous studies [87].
Figure 3-1 Reaction diagram for immobilization of growth factors or peptides on collagen sponges using 1-
ethyl-3-(3-dimethylaminopropyl) carbodiimide HCl (EDC) and N-hydroxysulfosuccinimide (Sulfo-NHS).
1. Using sterile tools, cut the 2 mm thick collagen sponge sheet into desired shape (2 cm × 2
cm).
2. For each collagen sponge (2 cm × 2 cm), prepare 1 mL EDC reaction solution composed of
45
24 mg/mL EDC and 60 mg/mL Sulfo-NHS in PBS (Lonza).
3. Filter-sterilize the EDC reaction solution using sterile 0.2 μm syringe filters (VWR) and use
it promptly.
4. Immerse the collagen sponge into the EDC reaction solution in a 6-well plate and allow the
activation to proceed for 20 minutes at room temperature.
5. Prepare 600 μL growth factor solution composed of 1 µg/mL VEGF (PeproTech) and 1
µg/mL bFGF (PeproTech) or peptide solution composed of 1 µg/mL QHREDGS (Biomatik)
in PBS in another 6-well plate.
6. Remove the activated collagen sponge from EDC reaction solution and drain excess liquid
by dabbing to the wall.
Note: The NHS-activated collagen sponge should be proceeded immediately to step 7.
7. Immerse the activated collagen sponge into prepared growth factor solution or peptide
solution and allow the reaction to proceed for 2 hours at room temperature.
8. Wash the sponge with fresh PBS for 8 times of at least 5 minutes each time, to wash away
the uncrosslinked EDC, sulfo-NHS, and growth factors or peptides.
9. Keep the sponge in PBS at 4°C and use within 24 hours after preparation.
3.2.3 Quantification of growth factor immobilization efficiency
In order to investigate the amount of growth factors immobilized onto collagen sponge, modified
sponges were prepared freshly (immobilization group). Sponges were also prepared following
the protocol without adding the EDC reaction solution (physical adsorption group) or without
adding the growth factor solution (control group). All the washes in step 8 were collected
(labelled as supernatant). After the washes, all the collagen sponges were digested by 0.276
mg/mL collagenase type IA (Sigma Aldrich) for 1.5 hours at 37 °C and the digestion solution
was collected. All the samples were stored at -80 °C if not used immediately.
Enzyme-linked immunosorbent assay (ELISA) utilizes antigen-antibody interaction and color
developing enzyme to analyze amount of specific proteins in solution. Here we used ELISA
Development Kit (PeproTech) to quantify the amount of immobilized VEGF and bFGF on
collagen sponges and the unbounded amount in the supernatant in all three groups
(immobilization, physical adsorption, and control). To obtain accurate results, all samples and
standards were arranged in the same ELISA plate.
46
1. Reconstitute all the components from the kit, aliquot and store at recommended temperature.
2. Dilute capture antibody with PBS to 0.5 μg/mL and add 100 μL to each ELISA plate well.
Seal the plate with parafilm and incubate overnight at room temperature. Aspirate and wash
the plate 4 times.
Note: After each wash, it is easier to remove liquid by blotting on clean paper towel.
3. Add 300 μL blocking buffer to each well and incubate for 2 hours at room temperature.
Aspirate and wash plate 4 times.
4. Prepare standards (from the ELISA kit) and samples on ice (dilute with PBS when necessary)
and add 100 μL to each well in triplicate. Incubate at room temperature for at least 2 hours.
Aspirate and wash the plate 4 times.
5. Dilute detection antibody in 0.05% Tween-20, 0.1% BSA to the final concentration of 0.25
μg/mL and add 100 μL to each well. Incubate at room temperature for 2 hours. Aspirate and
wash plate 4 times.
6. Dilute avidin-HRP conjugate 1:2000 in 0.05% Tween-20, 0.1% bovine serum albumin (BSA)
and add 100 μL to each well. Incubate 30 minutes at room temperature. Aspirate and wash
the plate 4 times.
7. 2,2'-Azinobis [3-ethylbenzothiazoline-6-sulfonic acid]-diammonium salt (ABTS) liquid
substrate should be equilibrated at room temperature. Add 100 μL of substrate solution to
each well. Shake the plate gently and monitor color development with SpectraMax i3
(Molecular Device) at 405 nm at 5-minute intervals for 30 minutes.
8. The built-in software (SoftMax Pro 6.4) calculates the concentration of target growth factor
in sample solutions automatically by comparing with standard solutions based on absorbance
readings.
3.2.4 Quantification of QHREDGS peptide immobilization efficiency
Fluorescently labelled peptide, FITC-QHREDGS was used to assess the conjugation efficiency
and true final concentration of peptide attached to chitosan post dialysis. This then determined
the final concentration of peptide in the hydrogels. FITC-QHREDGS (Biomatik) was substituted
for regular peptide in the protocol above with the critical addendum that all steps in the method
were protected from light to prevent photobleaching of the fluorophore. As the molecular weight
cut-off of the dialysis membrane is at most a tenth that of chitosan it can be safely assumed that
47
all of the chitosan is retained and recovered, and the peptide present is only that which was
successfully attached to the chitosan.
1. Standards of the FITC-QHREDGS in PBS should be made ranging from 0.0005 to 0.01
mg/mL.
2. Both the standards and the reaction solutions recovered post dialysis should have their pH
adjusted to 7 using 0.1N NaOH and 0.1N HCl, as fluorescence is greatly affected by pH
[421].
3. Test samples (reaction solutions recovered post dialysis) should be diluted 1:10 and 1:100.
4. Pipette 100μL of sample or standards, in triplicate, into wells of a 96-well black plate.
5. Run the plates through a fluorometer (Spectra Max Geminin EM, Molecular Devices) at an
excitation wavelength of 490 nm and emission of 520 nm.
6. The true final concentrations of the peptide and conjugation efficiency are calculated by
comparing the fluorescence of the samples to the standards, correcting for the dilution factor
and for the volume recovered post dialysis.
3.2.5 Characterization of release profile
It is important to investigate the release profile of the growth factors or peptides immobilized on
the modified biomaterials to confirm that they are being released gradually over extended time
(e.g. 2-3 weeks). We immersed freshly-prepared sponges in 1 mL PBS in 24-well plates and
incubated at 37 °C for 28 days. The supernatant was collected on day 1, 3, 7, 14, 21, and 28 and
stored at -80 °C. The collagen sponge was digested after collecting the supernatant on day 28 by
methods described in 3.2.2. All the samples (all the supernatant samples and digestion samples)
were characterized for VEGF and bFGF amount by ELISA as described before. Similarly, in the
case of peptide immobilization, the supernatant was collected and analyzed as described in 3.2.3.
3.2.6 Scanning electron microscopy
Porous scaffolds provide mechanical support and guidance for the cell infiltration and it is
necessary to confirm that our modification does not change their porous structures (e.g. clotting
the pores). We used environmental scanning electron microscopy (Hitachi S-3400 N) to examine
both collagen sponges and collagen-chitosan hydrogel samples. A filter paper was used to gently
remove the excess water from the samples. The chamber was closed and the temperature of the
48
chamber decreased to -20°C. The samples were imaged under variable pressure mode at 70 Pa
and15 kV.
3.2.7 Tensile testing of porous collagen scaffolds
The collection of human samples was approved by the Research Ethics Board of University
Health Network. Each patient provided informed consent. Bone marrow aspirates were obtained
from the sternum of patients undergoing coronary artery bypass grafting (CABG) at Toronto
General Hospital. “Young” hMSCs were isolated from patients age ≤57 years (50.0 ± 8.0 years,
N = 4); “old” hMSCs were isolated from patients age ≥66 years (74.5 ± 7.4 years, N = 4). Four
groups of scaffolds were tested with and without growth factors: fresh (scaffolds right after
preparation), blank (scaffolds incubated in media for 3 days), young (scaffolds seeded with
young hMSCs and incubated in media for 3 days), and old (scaffolds seeded with old hMSCs and
incubated in media for 3 days).
We used a protocol modified based on ASTM D412-06a to perform the tensile testing of
modified collagen sponges using ElectroForce 5200 BioDynamic Test Instrument (Bose) with a
22 N load sensor. Please note that “displacement” in following protocol is defined by the
ElectroForce program as the distance between the two grips.
1. After modification (Step 2.2), carefully cut the collagen sponge into straight specimen (2 cm
× 1 cm) and briefly dry with Kimwipes (50 ± 5% moisture).
2. Start the machine and set the displacement to be 2 mm.
3. Place the specimen in both grips to the same depth, carefully adjust the grips to align the
specimen with the direction of pull. Make sure the specimen is not stretched initially and
reset the force reading to 0.
4. Set the displacement rate at 1 mm/min and displacement to 3 mm (approximately 50%
strain). Start recording both displacement and force reading.
5. Start pulling and record the distance L0 where force reading starts increasing. Wait until
displacement stops at 3 mm.
6. Set the displacement to 2 mm (original distance between the two grips) and start again. The
force reading should decrease to 0 at displacement L0, indicating that the sponge behaves
like elastomer within 50% strain range.
49
Note: We aim for 50% strain range based on final application for heart implantation. The elastic
behavior would be better verified by plotting the stress-strain curve.
7. Set the displacement to 10 mm and start. Release the specimen after rupture.
Calculations:
1. Calculate the strain as the elongation of specimen in percentage:
ε = the strain of the specimen,
L = observed displacement,
L0 = initial displacement when the specimen is not extended.
2. Calculate the tensile stress at any specified strain as follows:
T(x) = tensile stress at (x) % elongation,
F(x) = force at specified elongation,
A = cross-sectional area of unstrained specimen.
3. Calculate the ultimate tensile strength as follows:
UTS = ultimate tensile strength, the stress at rupture,
Ft = the force at rupture,
A = cross-sectional area of unstrained specimen.
4. Young’s modulus should be calculated from the elastic region of the stress-strain curve as
follows:
E = Young’s modulus
Tx = tensile stress at (x) % elongation,
εx = the strain on the specimen at (x) % elongation.
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3.3 Results and discussion
It is important to characterize the mechanical properties of the modified porous scaffolds to
ensure it will withstand tensile force for adequate performance in certain tissue engineering
application. For example, scaffolds for myocardium implantation should have Young’s modulus
close to the native myocardium (10-150 kPa in the physiological regime [422]) and remain
elastic within the deformation of myocardial segments during beating (about 20% in longitudinal
direction and up to 60% in radial direction [423, 424]). We characterized the scaffolds to ensure
that cytokine immobilization did not adversely affect the stiffness, strength, or porosity of the
scaffolds and, therefore, their usefulness for surgical ventricular restoration (SVR). Stress–strain
curves demonstrated that cytokine-free and -enhanced scaffolds exhibited similar behaviors
during the breaking process (Figure 3-2A). Young’s modulus, a measure of stiffness, did not
differ with the addition of growth factors to the scaffold (Figure 3-2B). Ultimate tensile strength,
the maximum stress a material can withstand while being stretched, was lower in cytokine-
enhanced scaffolds than in cytokine-free scaffolds, with the exception of the Old group (Figure
3-2C). However, the ultimate tensile strength was calculated at the final breaking point, at
approximately 300% strain. Because this strain is not applicable in vivo, the Young’s modulus is
more suitable for characterizing the mechanical property of the scaffolds for our application.
Scanning electron microscopy showed no significant differences in the porosity or pore
structures of the scaffolds with or without cytokines (Figure 3-2D).
To ensure covalent immobilization of the growth factors to the collagen scaffolds (and not just
physical attachment), we used ELISA to determine the immobilization efficiencies for VEGF
and bFGF, which were 42% and 24%, respectively, and the physical bonding efficiencies, which
were 2% and 5% (Figure 3-2E), demonstrating that both cytokines were chemically immobilized
to the scaffolds. The scaffolds were prepared to provide a slow release of cytokines over time
after SVR. The release study, based on ELISA, demonstrated that the amount of cytokine
released from the scaffolds over 4 weeks was small, but the release rates were similar for VEGF
and bFGF (Figure 3-2F). A significant amount of each cytokine remained in the scaffolds after
28 days (Figure 3-2G).
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Figure 3-2 Characterization of scaffolds. (A) Stress-strain curves for cytokine-free scaffolds (solid lines) and
cytokine-enhanced scaffolds (dashed lines) showed similar tensile strength with or without cells. (B) Young’s
modulus was not significantly different between cytokine-free and –enhanced scaffolds. (C) Ultimate tensile
52
strength was significantly lower in cytokine-enhanced scaffolds in all groups but the Old group (**p < 0.01).
However, this measure was obtained outside of the physiological range of strain. Young’s modulus provides a better
estimate of the physical characteristics of the patch in vivo. (D) Scanning electron microscopy at 100× (scale bar =
500 μm) and 500× (scale bar = 100 μm) magnification revealed similar porosity in scaffolds with and without
growth factors. (E) Enzyme-linked immunosorbent assay (ELISA) demonstrated that both vascular endothelial
growth factor (VEGF) and basic fibroblast growth factor (bFGF) (150 ng of each added to the scaffolds) were
immobilized on the scaffolds, with little in the supernatant. Scaffolds without 1-ethyl-3-(3-dimethylaminopropyl)
carbodiimide HCl/N-hydroxysulfosuccinimide treatment served to show physical bonding of the cytokines to the
scaffolds (not covalent immobilization). (F) ELISA demonstrated that little VEGF and bFGF was released from the
scaffolds over 28 days. (G) After day 28, ELISA showed that substantial amounts of VEGF and bFGF remained in
the scaffolds. n = 3/group. bFGF = basic fibroblast growth factor; EDC = 1-ethyl-3-(3-dimethylaminopropyl)
carbodiimide HCl; GF = growth factor; PBS = phosphate-buffered saline; VEGF = vascular endothelial growth
factor.
The described covalent immobilization strategy using EDC chemistry is effective in
immobilizing growth factors or peptides to both porous scaffold and hydrogels. We characterized
collagen sponge modified with FITC-QHREDGS with the critical addendum that all steps in the
method were protected from light to prevent photobleaching of the fluorophore (Figure 3-3).
The presence of fluorescently labelled immobilized molecule was visible using even simple
fluorescence microscopy in comparison to the blank scaffolds and the scaffolds that relied on
physical adsorption alone (Figure 3-3A).
The immobilization efficiency was characterized using sample preparation described in 3.2.1 and
fluorescence-based assay described in 3.2.3, demonstrating up to eight-fold higher amount of the
immobilized QHREDGS peptide in the EDC treated scaffold compared to the physical
adsorption alone (Figure 3-3B). Importantly, as the scaffold degrades using cell culture or
incubation with PBS or culture media, the peptide will slowly be released into the environment.
This obviates the need for continuous application of the biomolecule with culture media changes,
thus it decreases the totally applied dose. Importantly, the pore structure and porosity was not
appreciably changed with EDC based covalent immobilization of biomolecules (Figure 3-3C).
53
Figure 3-3 Characterisation of peptide immobilization. (A) FITC-QHREDGS modified collagen sponges under
the microscope (all images acquired at same exposure time). Left: control sponge modified with EDC reaction
solution without adding FITC-QHREDGS; Middle: physical adsorption sponge modified with FTIC-QHREDGS
without adding EDC reaction solution; Right: immobilization sponge modified with EDC reaction solution with
presence of FITC-QHREDGS. Scale bar = 200 μm. (B) Quantification of the amount of FITC-QHREDGS peptide in
freshly prepared collagen sponges. The signals in the control group come from collagen auto-fluorescence. (C) SEM
images of the collagen sponges with (right) or without (left) EDC modification.
Using the immobilization protocol described in 3.2.1, the immobilization efficiency should be
above 20% as we described previously [181]. The amount of growth factors or peptides released
over 4 weeks in aqueous media (e.g. PBS) should be small with significant amount remaining in
54
the scaffolds after 4 weeks [181]. The mechanical properties of collagen sponges will change as
we showed an increased Young’s modulus due to crosslinking of collagen molecules to
themselves [425]. The collagen sponge modified with VEGF and bFGF has been shown to
promote proliferation and down-regulate aging-related gene expression of human MSCs from old
donors, which improves the cell function to restore cardiac function after SVR [181].
Using the conjugation protocol described in 2.3.1, the conjugation efficiency of the peptide to
chitosan should be above 50% as we described previously [9]. Characterized by rheology, the
modified 1:1 chitosan-collagen hydrogel should be mechanically stable at 37°C based on the
values of its storage modulus [9]. As characterized by live/dead staining and lactate
dehydrogenase (LDH) assay, the presence of QHREDGS peptide should improve cardiomyocyte
viability and metabolic activity in vitro as we showed previously [9]. Compared to unmodified
hydrogel, the peptide-conjugated chitosan-collagen hydrogel also improves the morphology of
cardiomyocytes in vitro [9]. Moreover, the modified hydrogel inhibits paclitaxel-induced
apoptosis of endothelial cells and stimulates tube-like structure formation in vitro [10].
3.4 Conclusion
Here we have described our protocols for preparation and characterization of collagen-based
biomaterials (both porous scaffolds and hydrogels) modified with growth factors or peptide by
EDC chemistry. As detailed above, quick handling during EDC crosslinking is critical for
effective immobilization. In addition, the physical and biological properties of the modified
biomaterials should be carefully characterized for different tissue engineering applications. The
bioactivity of immobilized growth factors or peptides should be tested on in vitro cell cultures
before in vivo application.
3.5 Acknowledgments
This work is funded by the Heart and Stoke Foundation GIA T6946, the Canadian Institutes of
Health Research (CIHR) Operating Grant (MOP-126027), NSERC-CIHR Collaborative Health
Research, Grant (CHRPJ 385981-10), NSERC Discovery Grant (RGPIN 326982-10), and
National Institutes of Health grant 2R01 HL076485. M.R. is supported by Canada Research
Chair (Tier 2) and Steacie Fellowship.
55
Chapter 4
4 Diabetic wound regeneration using peptide-modified hydrogel targeting the epithelium
4.1 Introduction
Chronic ulcers are considered a major healthcare challenge as they affect 6.5 million people in
the United States [426]. Non-healing wounds, including chronic ulcers, can be caused by a
number of common diseases and medications, such as vascular insufficiency, diabetes mellitus,
and local-pressure effects, which disrupt the well-orchestrated cellular and molecular interactions
during the wound healing process [235]. Specifically, diabetic foot ulcers affect 15% of people
with diabetes and are a leading cause of amputation [427]. The mechanism underlying diabetic
chronic wounds remains elusive and new interventions for diabetes-impaired wound healing are
needed. After almost two decades without new chemical entities approved by the Food and Drug
Administration (FDA) (Regranex® was approved in 1997), it has been recognized that an
optimal wound healing outcome requires a multifaceted approach that addresses different issues
(e.g. persistent inflammation, insufficient angiogenesis, and impaired re-epithelialization) at once
[235].
Keratinocytes are the major cell type in the epidermis, the outermost layer of skin. Upon injury,
keratinocytes migrate from the wound edge into the wound to re-epithelialize the damaged tissue
and restore the epidermal barrier. The hallmark of non-healing human wounds is non-migratory
and hyper-proliferative keratinocytes, resulting in epidermis thickening at the wound edge and an
absence of wound closure [428]. Scarless embryonic wound healing and complete healing in
animals with a high regenerative potential such as newts critically depend on rapid re-
epithelialization [429, 430].
Additionally, non-healing diabetic wounds are trapped in a state of prolonged inflammation,
characterized by supra-physiological oxidative stress that can induce keratinocyte injury,
dysfunction and apoptosis [431-433]. It results from the excess production of reactive oxygen
species (ROS) by macrophages and neutrophils, coupled with an impaired antioxidant defense
capability in response to hyperglycemia [253]. Moreover, altered extracellular matrix (ECM)
56
composition in non-healing wounds and an enhanced ECM degradation rate due to elevated
matrix metalloproteinase levels can impair keratinocyte attachment, leading to aberrant cell
signaling and impaired migration [250, 251, 434].
To address these challenges, we sought to develop a novel wound healing approach that could
recapitulate key aspects of scarless embryonic wound healing by 1) promoting effective
keratinocyte migration, 2) protecting the wound-bed cells against oxidative stress and 3)
providing a new matrix for cell attachment.
Our group has recently described a novel angiopoietin-1derived peptide, QHREDGS, which
interacts with integrins, receptors that function in cell-adhesion and ECM-binding. The
QHREDGS peptide was shown to enhance endothelial cell metabolism, tube formation kinetics,
and survival in response to apoptotic stimuli [10]. QHREDGS was also shown to promote
neonatal rat cardiomyocyte attachment and survival [7], to inhibit human induced pluripotent
stem cell (hiPSC) apoptosis during cells expansion [11], to induce osteoblast matrix deposition
and mineralization [12], and to have cardiac protective effects in a chitosan-collagen hydrogel
both in vitro and in vivo [9, 13].
We therefore hypothesized that the QHREDGS peptide could promote keratinocyte survival and
migration and thereby accelerate diabetic wound healing. We investigated the effect of the
QHREDGS peptide as a soluble supplement on the survival of normal human keratinocytes upon
oxidative stress. The effect on attachment, survival upon oxidative stress, and collective
migration of both normal and diabetic keratinocytes was assessed by immobilizing the
QHREDGS peptide within a chitosan-collagen film coating. We further investigated the ability
of the QHREDGS peptide to promote diabetic wound repair in vivo using a full-thickness
excision wound model in db/db diabetic mice.
4.2 Materials and methods
4.2.1 Primary human keratinocytes cell culture
Primary neonatal human epithelial keratinocytes (HEKs) were purchased (Cascade Biologics)
and cultured in EpiLife medium supplemented with EpiLife Defined Growth Supplement (EDGS)
as recommended by the manufacturer (Cascade Biologics; referred to as complete medium).
57
HEKs were cultured on surfaces coated with a coating matrix kit (Cascade Biologics) and
passaged using 0.025% trypsin/EDTA (Cascade Biologics) until 70-80% confluence was reached.
Third and fourth passage HEKs were used in our experiments.
Diabetic human adult epithelial keratinocytes (DHEKs) from a patient (72 years old female) with
type II diabetes were purchased from Lonza and cultured in KGM-GoldTM BulletKitTM
medium as recommended by the manufacturer (Lonza). DHEKs were passaged using
ReagentPackTM Subculture Reagents (CC-5034, Lonza) until 70-80% confluent. Second and
third passage DHEKs were used in our experiments.
4.2.2 Evaluation of soluble QHREDGS in vitro
The effect of the QHREDGS peptide as in the soluble form was assessed on HEKs in vitro. 24-
or 96-well plates were coated with 0.05 mg/mL type I collagen (BD Biosciences) in 0.02 N
acetic acid overnight and then rinsed once with phosphate buffered saline (PBS) (Lonza). HEKs
were seeded in complete EpiLife medium and attached for at least 2 h. 100 µM or 650 µM
QHREDGS was supplemented to the EpiLife medium as Low or High dosage, respectively.
4.2.3 Proliferation assay
HEKs were seeded at a density of 1×104 cells/cm2 in collagen-coated 96-well plates. After 4 h,
media was removed and replenished with EpiLife media supplemented with 20 µM
bromodeoxyuridine (BrdU) (Sigma Aldrich) in the presence or absence of QHREDGS peptide.
After incubating for 8 h, proliferating HEKs were identified by BrdU staining. Briefly, HEKs
were fixed with 4% paraformaldehyde (PFA), permeablized with 0.25% triton X, treated with
DNase for 30 min at 37 °C, and blocked with 5% bovine serum albumin (BSA). The HEKs were
then incubated with a rat anti-BrdU antibody (AbD Serotec) overnight at 4 °C followed by an
anti-rat TRITC-labelled secondary antibody (Jackson Immuno Research). BrdU-positive cells
were counted in 5 randomly chosen fields under 20× magnification and then normalized to the
total number of cells labelled by 4',6-diamidino-2-phenylindole (DAPI) counterstaining.
4.2.4 H2O2 treatment on HEKs with soluble QHREDGS peptide
HEKs were treated by H2O2 following the regimen shown in Fig 4-1B. HEKs were seeded at a
sub-confluent density in a 96-well plate for the cell integrity assay or a 24-well plate for Western
blot analysis, and serum starved overnight. The HEKs were then pretreated with soluble
58
QHREDGS (100 µM for Low and 650 µM for High) for 2 h. The cells were then exposed to
fresh EpiLife basal medium supplemented with 500 µM H2O2 in the presence or absence of
QHREDGS peptide. For the cell integrity assay, the cells were stained using the EarlyToxTM Cell
Integrity kit (Molecular Devices) after 2 h of H2O2 treatment. For the Western analysis, another
set of wells were seeded in parallel for the Ctrl, Low and High conditions and protein samples
were collected after 0, 15 min and 2 h of H2O2 treatment.
4.2.5 Conjugation of QHREDGS to chitosan
The QHREDGS peptide was conjugated to chitosan using 1-ethyl-3-(3-dimethylaminopropyl)
carbodiimide (EDC) chemistry as previously described [88]. Briefly, chitosan (UP G 113,
Novamatrix) was dissolved at 20 mg/ml in 0.9% normal saline and the QHREDGS peptide at 10
mg/ml in PBS. These were then mixed with EDC and N-hydroxysulfosuccinimide (S-NHS),
dissolved in PBS, to a final solution concentration of 5mg/ml chitosan and 0.5 mg/ml of
QHREDGS peptide (Low peptide group) or 3 mg/ml of QHREDGS peptide (High peptide
group). In the reaction solution, the mass ratio of [EDC]/ [peptide] and [S-NHS]/ [EDC] were
kept constant at 0.8 and 2.75, respectively. The reaction solution was left on a vortex mixer
(VWR) at 650 rpm for 3 h, diluted 4× with PBS and dialyzed against distilled water for 48 h
(Spectra/POR MWCO 3500, Spectrum Labs). The dialyzed solution was then filter sterilized,
lyophilized for 48 h and stored at -20 °C until use.
4.2.6 Solvent casting of chitosan-collagen films
The chitosan samples were reconstituted at 2 mg/ml in 0.5 N acetic acid and mixed with 2 mg/ml
type I collagen (BD Biosciences) to obtain a film coating solution composed of 1 mg/ml each of
chitosan and collagen. 24-well plates were coated with 250 µL film coating solution per well and
96 well plates with 50 µL per well. The film coating solution was fully evaporated in a biosafety
hood and the chitosan-collagen films were cast in the wells. The coated plates were rinsed three
times with ample PBS before use.
4.2.7 Coating validation
Fluorescently labeled peptide, FITC-QHREDGS (Biomatik), was used to validate peptide
concentrations in the film. FITC-QHREDGS was substituted for regular peptide in the protocol
above and all steps were protected from light. Standards of the FITC-QHREDGS in PBS were
59
made ranging from 0.025 pg/ml to 0.5 pg/ml. After rinsing with PBS three times, the coated 96-
wells were filled with 200 µl PBS and then run, with the standards in the same plate, through a
fluorometer (SpectraMax i3, Molecular Devices) at excitation and emission wavelengths of 490
nm and 520 nm, respectively. The final amounts of peptide were quantified using SoftMax Pro
6.4 software.
4.2.8 Keratinocyte attachment on chitosan-only films
Chitosan-only films were prepared by casting 1 mg/ml chitosan with the presence or absence of
conjugated QHREDGS peptide in 0.5 N acetic acid in the wells of a 96-well plate and drying in
the a biosafety hood overnight. The coated plates were rinsed three times with ample PBS before
use. HEKs or DHEKs were seeded in supplemented EpiLife or KGM medium and allowed to
attach on the chitosan-only films. After 2 h, unattached cells were carefully rinsed off using PBS
and cells were then fixed with 4% paraformaldehyde. The number of attached cells were
quantified using DAPI counterstaining and the cell numbers on chitosan-only films in the
presence or absence of conjugated QHREDGS peptide were normalized to the number of cells
attached to regular tissue culture polystyrene (TCP).
4.2.9 H2O2 treatment on keratinocytes on the chitosan-collagen films
HEKs or DHEKs were seeded in complete EpiLife or KGM medium and allowed to attach for 4
h. HEKs were then changed to basal EpiLife medium supplemented with 500 µM H2O2 and
DHEKs were changed to basal KBM medium supplemented with 2 mM H2O2. After 2 h, cells
were changed to a complete medium with EarlyTox™ Cell Integrity staining reagent and
subjected to cell integrity assay. Cells treated by H2O2 for 15 min were used for Western blotting
together with non-treated controls.
4.2.10 EarlyToxTM Cell Integrity assay
The EarlyTox™ Cell Integrity Kit (Molecular Devices, R8213) is based on two nuclear dyes: live
red dye is cell permeant and marks both live and dead cells (Excitation: 622 nm/Emission: 645
nm); dead green dye is cell impermeant and stains only cells with damaged outer membranes
(Excitation: 503 nm/Emission: 526 nm). To avoid cell detachment, half of the medium in each 96
well was removed by micropipette and the equal volume of the staining solution with double
concentration of the dyes was added into the well carefully, such that the final concentration in
60
the well was the one suggested by the manufacturer. The plate was then incubated at 37 °C for
30 min and imaged using a SpectraMax i3 plate reader (Molecular Devices). The percentage of
viable cells after H2O2 treatment on chitosan-collagen films in the presence or absence of
QHREDGS peptide was normalized to the viability percentage of non-treated controls on the
same coating film condition.
4.2.11 Western blotting
Protein was isolated from keratinocytes in the 24-well plate after H2O2 treatment using 60-80 µL
Lysis Buffer per well (10× Cell Lysis Buffer, Cell Signaling Technology; complete Mini,
EDTA-free protease inhibitor cocktail tablet, Roche; in ddH2O). Proteins were separated by
electrophoresis in Novex Tris-Glycine gels (Life technologies) and transferred using the iBlot
(Life technologies) to a PVDF iBlot Transfer Stack (Life technologies). Membranes were probed
for phospho-p44/42 MAPK (pMAPKp42/44), p44/42 MAPK (MAPKp42/44), phospho-Akt, Akt, or
GAPDH as a loading control (Millipore). All primary antibodies were purchased from Cell
Signaling unless stated otherwise. HRP conjugated goat anti—mouse or goat anti—rabbit
secondary antibodies were used (DAKO). Membranes were developed with Amersham ECL or
Amersham ECL Prime Western Blotting Detection Reagent (GE Healthcare) and exposed to the
films. The films were scanned and densitometry was performed using ImageJ or Image Studio™
Lite (LI-COR Biosciences).
4.2.12 Migration assay
Culture-inserts were purchased from ibidiⓇand carefully placed on the chitosan-collagen films in
24-wells. HEKs were seeded in complete EpiLife medium at 0.3-0.4×106 cells/mL with 70-100
µL per chamber and allowed to attach for 2 h. The culture-inserts were then carefully lifted and
the unattached HEKs were immediately removed by rinsing twice using warm PBS. Basal
EpiLife medium was then added to the wells and the wells were imaged every 2 h for 8 h and the
next day using the SpectraMax i3 plate reader. After 24 h, HEKs were fixed with 4%
paraformaldehyde.
The Ca2+ level in the EpiLife medium was increased from 0.06 mM to 0.12 mM to ensure HEKs
migrated collectively. After initiating the HEK migration by lifting the ibidiⓇ culture inserts,
HEKs were maintained in EpiLife basal medium supplemented with CaCl2 at a final Ca2+
61
concentration of 0.12 mM. HEKs were fixed with 4% paraformaldehyde after exposure to
increased Ca2+ concentration for the indicated period of time.
Similarly, DHEKs were seeded in complete KGM medium with CellMask Green (1000× dilution)
at a density of 0.15-0.2×106 cells/mL in ibidi migration chambers and allowed to attach for 2 h.
Migrations were initiated by lifting the culture-inserts as mentioned above and the wells were
imaged every 2 h for 6 h using the SpectraMax i3 plate reader. After 6 h, DHEKs were fixed
with 4% paraformaldehyde.
4.2.13 Immunostaining
At the end of migration, HEKs and DHEKs were fixed and stored in PBS at 4 °C. Cell
monolayers were permeablized with 0.25% triton X, blocked with 5% bovine serum albumin
(BSA), and then incubated with mouse anti-E-cadherin primary antibody (BD Biosciences; 1:200)
overnight at 4 °C followed by goat anti-mouse Alexa 488 secondary antibody (Jackson Immuno
Research; 1:400). Cell nuclei were counterstained with 4',6-diamidino-2-phenylindole (DAPI)
(Biotium; 1:100). The 24-well plates were imaged at 20× magnification using a fluorescence
microscope (Olympus IX81).
4.2.14 Animals, wound model, and treatment
The Animal Care Committee of the University of Toronto approved all described animal studies.
8 weeks old, genetically diabetic, maleBKS.Cg-Dock7m +/+ Leprdb/J mice (db/db) (Stock 000642)
were ordered from Jackson Laboratories (Bar Harbor, USA). Mice were acclimatized for one
week and their blood glucose levels were tested with a glucometer (Accu-ChekⓇ Aviva) to
confirm plasma glucose levels were over 300 mg/dL the day before surgery.
The chitosan-collagen hydrogel was prepared in a similar manner as previously described.[88]
The final hydrogel consisted of 2.5 mg/ml chitosan (with or without conjugated QHREDGS
peptide) and 2.5 mg/ml type I collagen neutralized by 1 N NaOH and 10× PBS. The final
hydrogel solution was mixed thoroughly and kept on ice until use. For in vivo application, only
the Low-peptide chitosan-collagen was applied and the pre-gel solution was warmed for about 10
min at 37 °C to initiate the gelling process and applied to the wound site with a 23½ G needle.
62
Mice were anaesthetized with inhaled isoflurane (5%) and the dorsal surface of the mouse was
shaved with an electric shaver, followed by treatment with a hair removal cream (VeetⓇ).
Betadine and 70% ethanol were applied in series to the surgical site. 8 mm Biopsy punches
(VWR) were used to create mid-dorsal full-thickness wounds by excising the epidermis and
dermis, including the panniculus carnosus. Either 50 µL control hydrogel without conjugated
QHREDGS peptide or 50 µL hydrogel with Low peptide (containing a total of 2.2nmol peptide)
was applied topically to the wound beds, or the wounds were left untreated as blank. The wound
beds were then covered by Tegaderm™ film (Fig. 4-1A). Buprenorphine (0.03 mg/kg) was given
subcutaneously before and right after the surgery as an analgesic. Thereafter, the mice were
housed individually and observed every other day. Digital photographs of wounds were taken at
the same distance by a camera (Canon) with a calibration scale on the side every two days. Mice
were sacrificed using CO2 asphyxiation, followed by cervical dislocation, on day 14.
4.2.15 Histology analysis
Following euthanasia, the wound tissue was excised together with surrounding tissue and fixed
in 10% formalin (Sigma). Tissue samples were embedded in paraffin blocks and then sliced into
5 µm-thick sections. Sections were processed and stained with hematoxylin and eosin (H&E),
Masson’s trichrome, or immunostained using anti-CD31 and anti-smooth muscle actin (SMA)
antibodies at the University Health Network Pathology Research Program laboratory. Stained
slides were scanned (20×) using the Aperio ScanScope XT (Aperio Technologies, USA) at the
Advanced Optical Microscopy Facility (AOMF, Toronto, Canada). The images of scanned slides
were analyzed using the Aperio ImageScope (Version 11).
In order to characterize the healing of the wounds, Masson’s trichrome stained slides were
scanned with a ScanScope XT whole slide scanner and measured using Aperio ImageScope (v11,
Aperio Technologies). The wound edge was defined as the panniculus carnosus muscle gap, the
epithelial gap as the distance between the epithelial tongues, and the re-epithelization percentage
as the ratio of epithelial gap over wound edge. The size of the granulation tissue was defined by
the highly cellular tissue between epidermis and the fat/muscle tissue. The epidermal thickness
was defined as the average thickness of the leading epithelial tongue (300 µm) from both ends.
63
In order to characterize angiogenesis, six views (300 µm × 300 µm) within the granulation tissue
were randomly selected for each scanned CD31 slide and then the number of CD31 positive
vessel-like structures was counted and normalized to the area of granulation tissue in the view to
determine the vessel density. Similarly, the SMA positive area percentage was determined from
six random views within the granulation tissue. An automated algorithm built-in with the Aperio
ImageScope (Microvessel Analysis Algorithm) was used to quantify the lumen area, vascular
area, vessel area, vessel perimeter, and vessel wall thickness of each CD31 positive vessel-like
structure over the entire granulation tissue.
4.2.16 Microvessel analysis algorithm
Scanned images of the histology slides stained with anti-CD31 antibody were analyzed using the
Microvessel Analysis Algorithm with Aperio ImageScope (Version 11). The entire granulation
tissue was selected using a drawing tool and CD31 positive staining was thresholded by Color
Deconvolution. Microvessels were thresholded using a Region Joining Parameter of 8 μm, a
Vessel Completion Parameter of 10 μm, and a Vessel Area between 1 and 20000 μm2. The
automated algorithm calculated the Lumen Area, Vascular Area, Vessel Area, Vessel Perimeter,
and Vessel Wall Thickness for each vessel within the selected granulation tissue area and output
the results in histograms.
4.2.17 Statistical analysis
All results are presented as mean ± SD. Statistical analysis was performed using SigmaPlot 11.0.
Differences between experimental groups were analyzed using one-way or two-way ANOVA
followed by a Tukey post hoc test for pairwise comparison. A value of P < 0.05 was considered
as statistically significant.
4.3 Results
4.3.1 QHREDGS peptide prevents H2O2-induced apoptosis in human primary keratinocytes and upregulates Akt and MAPKp42/44 signaling
To evaluate the effect of the QHREDGS peptide on keratinocytes, we first focused on normal
neonatal human epidermal keratinocytes (HEKs) cultured with the soluble QHREDGS peptide at
doses previously reported to be effective for endothelial cell survival (Low: 100 µM; High: 650
64
µM) [10]. The percentage of proliferating HEKs in the population was not affected by the
presence of the QHREDGS peptide at either concentration as quantified by bromodeoxyuridine
(BrdU) incorporation (Fig. 4-1A). No significant difference in the HEK migration rate was
observed with the soluble QHREDGS peptide at either concentration (Fig. 4-2).
To investigate the effect of the soluble QHREDGS peptide on keratinocyte survival under
oxidative stress, we pre-conditioned HEKs by incubating the cells with or without the
QHREDGS peptide and then exposed the HEKs to 500 µM H2O2 for 2 h (Fig. 4-1B). An
endpoint cell integrity assay showed a significant dose-dependent increase in the percentage of
viable HEKs in the presence of supplemented QHREDGS (Fig. 4-1C).
Given that the full-length protein from which the QHREDGS peptide was derived, angiopoietin-
1, is known to protect skin cells from oxidative damage and to increase the activation of the
prosurvival Akt and MAPKp42/44 pathway[435], we investigated whether improved survival upon
H2O2 stress in the presence of the QHREDGS peptide was associated with the upregulation of
Akt and MAPKp42/44 phosphorylation. HEKs treated with 500 µM H2O2 showed transient
phosphorylation of both Akt (Fig. 4-1D) and MAPKp42/44 (Fig. 4-1E) at 15 min by Western blot
analysis. Indeed, the presence of the soluble QHREDGS peptide during pre-conditioning and
H2O2 treatment increased the phosphorylation of Akt and MAPKp42/44and the increase was dose-
dependent (Fig. 4-1D and E).
65
Figure 4-1 Soluble QHREDGS peptide prevents H2O2-induced cell death in human primary keratinocytes
with up-regulation of Akt and MAPK phosphorylation. (A) Kct-positive HEKs cultured in the presence or
absence of different concentrations of soluble QHREDGS peptide did not incorporate significantly different
amounts of BrdU, indicating similar proliferation rates in all three condition (Low: 100 μM; high: 650 μM). Scale
bar = 50 μm. n=3-4. (B) Hydrogen peroxide treatment regimen for HEKs in the presence or absence of QHREDGS
peptide. HEKs were allowed to attach for 2 h and serum-starved overnight. For the peptide groups, HEKs were pre-
conditioned with QHREDGS peptide for 2 h and then treated with 500 μM hydrogen peroxide in the presence or
absence of the peptide. (C) HEK survival after hydrogen peroxide treatment was determined by the EarlyTox Cell
Integrity assay. The QHREDGS peptide protected HEKs against H2O2-induced cell death in a dose-dependent
manner (scale bar = 200 μm). One representative experiment is shown of n=3 independent experiments with 9
replicates for each condition in one experiment. (D and E) Immunoblotting with phosphorylated Akt or MAPKp42/44
66
and Akt or MAPK p42/44 antibodies showed transient activation of Akt and MAPK p42/44 pathways signaling under
H2O2 stress. GAPDH was used to ensure even loading. The presence of QHREDGS peptide in the culture medium
up-regulated Akt and MAPK p42/44 phosphorylation. n=4 independent experiments and each experiment performed in
duplicates or triplicates. Data presented as mean ± SD. * indicates P < 0.05.
Figure 4-2 The presence of soluble QHREDGS peptide does not accelerate HEKs migration on collagen
coated surfaces. (A) Representative images of HEKs on collagen-coated substrates in EpiLife basal medium in the
presence or absence of soluble QHREDGS peptide at the different times indicated. Scale bar = 200 μm. (B) Image
analysis showed no difference in HEK migration over 24 h among the three groups. One representative experiment
is shown of n=3 independent experiments with at least four replicates for each condition in one experiment. Data
presented as mean ± SD.
4.3.2 Immobilized QHREDGS peptide promotes human primary
keratinocytes attachment, survival and migration in vitro
While we observed increased survival without excessive proliferation in the presence of soluble
QHREDGS, we did not observe enhanced keratinocyte migration (Fig. 4-2). This motivated our
further optimization of the method by which we presented the peptide to the cells. Given that the
67
QHREDGS peptide is reported to primarily function through integrin interactions [10, 11] and
there is an ever-growing body of literature showing the increased efficacy of integrin ligands
when immobilized to a matrix [436, 437], we covalently immobilized the QHREDGS peptide to
a chitosan-collagen hydrogel. Chitosan and collagen interact through a combination of thermal
and ionic mechanisms, stabilized by polyanion (collagen) and polycation (chitosan) electrostatic
interactions.[438] Conjugation of the QHREDGS peptide to chitosan was achieved using
previously described methods [88] and chitosan-collagen films with or without immobilized
QHREDGS peptide were cast in the wells of 24-well or 96-well plates. Quantification using
fluorescently labelled peptide, FITC-QHREDGS, demonstrated effective immobilization in both
Low (4.7 ± 0.1 nmol/cm2) and High (13.8 ± 1.4 nmol/cm2) peptide concentrations and absence of
the peptide in the Ctrl condition (Fig. 4-3A). Normalized to the mass of chitosan in the films, the
amount of immobilized QHREDGS peptide was 14.9 ± 0.3 nmol/mg in the Low condition and
44.1 ± 4.6 nmol/mg in the High condition.
There was no significant difference in the attachment of HEKs to the various chitosan-collagen
films (Fig. 4-3B). However, in the settings of chitosan-only films wherein adhesion was poor,
the QHREDGS peptide clearly promoted HEK attachment in a dose-dependent manner (Fig. 4-4).
This indicates that while the QHREDGS peptide can promote HEK attachment, the presence of
collagen adhesion sites in the setting of the chitosan-collagen film masks this effect. Furthermore,
Western blot analysis showed the increased activation of Akt and MAPKp42/44 during 2 h
attachment on chitosan-collagen films in the presence of immobilized QHREDGS peptide (Fig.
4-3C).
We then investigated the effect of the immobilized QHREDGS peptide on HEK survival
following 500 µM H2O2 treatment. HEKs were allowed to attach to the chitosan-collagen films
for 4 h, then treated with H2O2 for 2 h. Subsequent cell integrity assessment showed an increased
percentage of viable HEKs in the presence of the immobilized QHREDGS peptide (Fig. 4-3D).
HEKs treated with 500 µM H2O2 and non-treated controls were also compared using Western
blot analysis, wherein phosphorylation of MAPKp42/44 was increased in the presence of
immobilized QHREDGS peptide relative to the control (Fig. 4-3E).
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Figure 4-3 Immobilized QHREDGS peptide in chitosan-collagen films promotes human neonatal primary
keratinocytes survival and migration. (A) Quantification of the amount of QHREDGS peptide immobilized
within chitosan-collagen films. n=3. (B) HEK attachment on the chitosan-collagen films in the presence or absence
69
of conjugated QHREDGS peptide. Image analysis showed no difference in the number of attached HEKs (stained
with DAPI) among the three groups (scale bar = 200 μm). n=3 independent experiments and each experiment
performed in triplicates. (C) Immunoblotting with anti-phosphorylated Akt or MAPK p42/44 and anti-Akt or
MAPKp42/44 showed up-regulation of Akt and MAPK p42/44 activation during HEK attachment. GAPDH was used to
ensure even loading. n=3 independent experiments and each experiment performed in duplicates. (D) HEK survival
after hydrogen peroxide treatment was determined by the EarlyTox Cell Integrity assay. QHREDGS peptide in the
chitosan-collagen film protected HEKs against H2O2-induced cell death in a dose-dependent manner. One
representative experiment is shown of n=3 independent experiments with four replicates for each condition in one
experiment. (E) Immunoblotting with phosphorylated MAPK p42/44 and MAPK p42/44 antibodies showed up-regulation
of the MAPK p42/44 activation in HEKs under H2O2 stress at 15 min. GAPDH was used to ensure even loading. n=3
and each experiment performed in duplicates. (F) Representative examples of HEK wounding experiments on
chitosan-collagen films in the presence or absence of conjugated QHREDGS peptide (scale bar = 200 μm).
Confluent HEK monolayers were wounded (time 0) and maintained for 24 h in EpiLife basal medium with 0.12 mM
Ca2+. The wounds were outlined and the area at the various time points was normalized to the initial wound size
(time 0). HEK migration on the QHREDGS-immobilized films was accelerated compared to the control peptide-free
films. One representative experiment is shown of n=3 independent experiments with six replicates for each condition
in one experiment. Data presented in mean ± SD. * indicates P < 0.05.
Figure 4-4 The presence of the immobilized QHREDGS peptide promotes HEK attachment on chitosan-only
films. (A) Representative images of HEKs on chitosan-only films in the presence or absence of immobilized
QHREDGS peptide. Scale bar = 200 μm. Cell nuclei are shown in blue (DAPI). (B) Image analysis showed an
increased number of attached HEKs in the presence of the immobilized QHREDGS peptide in a dose-dependent
manner. The number of attached HEKs was normalized to the number that attached to tissue culture polystyrene
(TCP) in the same experiment. One representative experiment is shown of n=3 independent experiments with three
replicates for each condition in one experiment. Data presented as mean ± SD. * indicates P < 0.05.
Keratinocyte migration is essential for wound healing as a wound cannot heal in the absence of
re-epithelialization [439]. We therefore assessed the effect of the immobilized QHREDGS
peptide on HEK migration in 2D monolayers using an Ibidi migration assay system. Importantly,
the Ca2+ concentration in the culture medium was increased from 0.06 mM to 0.12 mM upon
initiation of the migration assay to ensure collective HEK migration (essential for wound
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healing), as demonstrated by the formation of E-cadherin mediated cell-cell junctions (Fig. 4-
5A). The presence of the immobilized QHREDGS peptide accelerated collective HEK migration
in a dose-dependent manner (Fig. 4-3F). The accelerated migration was not due to increased
proliferation as there was no difference in cell density among the three groups as characterized at
the migration endpoint (Fig. 4-5B).
Figure 4-5 HEKs form calcium-induced adherens junctions during migration and the accelerated migration is
not associated with a difference in cell density. (A) Representative images of HEKs on Ctrl substrates (chitosan-
collagen films without conjugated QHREDGS peptide) in EpiLife basal medium at different times as indicated,
following an increase in calcium from 0.06 mM to 0.12 mM. Adherens junctions (green = E-cadherin) were
established as early as 2 h. Scale bar = 50 μm. Cell nuclei are shown in blue (DAPI). (B) HEK cell density
characterized by DAPI counterstaining at the end of migration (24 h). There was no difference in the number of
HEKs on the chitosan-collagen films in the presence or absence of the immobilized QHREDGS peptide. n=3 and
each experiment performed in triplicates. Data presented as mean ± SD.
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4.3.3 Immobilized QHREDGS peptide promotes diabetic human primary
keratinocytes attachment, survival and migration in vitro
In diabetic chronic wounds, keratinocytes experience hyperglycemia and supra-physiological
oxidative stress, which challenges keratinocyte’s proliferation and survival [253, 440]. Therefore,
we examined the effect of the immobilized QHREDGS peptide on adult diabetic human
epidermal keratinocytes (DHEKs) by seeding DHEKs onto chitosan-collagen films in the
presence or absence of immobilized QHREDGS peptide. Similar to the results with normal HEK
cells, we found the presence of the immobilized QHREDGS peptide promoted DHEK
attachment to chitosan-only films (Fig. 4-7) but did not affect DHEK attachment to chitosan-
collagen films (Fig. 4-6A).Western blot analysis showed that the activation of Akt was increased
in DHEKs in the presence of the QHREDGS peptide during a 2 h attachment (Fig. 4-6B).
Because of prolonged inflammation, diabetic wounds experience a higher level of oxidative
stress compared to the normal wounds [440]. To mimic this scenario, we investigated the effect
of immobilized QHREDGS peptide on DHEK survival following a 2 h treatment with 2 mM
H2O2 (4-times higher exposure than used for the HEK survival assay), after allowing DHEKs to
attach to the chitosan-collagen films for 4h. Cell integrity assessment showed that DHEK
survival under H2O2 stress was improved in the presence of immobilized QHREDGS peptide
(Fig. 4-6C), despite the higher H2O2 concentration used. DHEKs treated with 2 mM H2O2 and
non-treated controls were also compared by Western blot analysis (Fig. 4-6D) and
phosphorylation of Akt and MAPKp42/44 upon H2O2 treatment was increased in the presence of
immobilized QHREDGS peptide in a dose-dependent manner (Fig. 4-6E).
We then assessed the effect of immobilized QHREDGS peptide on DHEK migration using the
Ibidi migration assay. Importantly, the Ca2+ concentration in KGM medium (0.1 mM) was
sufficient to ensure collective DHEK migration as demonstrated by the formation of E-cadherin
mediated cell-cell junctions without additionally elevating the Ca2+ concentration (Fig. 4-8A).
The presence of the immobilized QHREDGS peptide also accelerated DHEK collective
migration (Fig. 4-6F) and this was not due to cell density differences as characterized at the
migration endpoint (Fig. 4-8B).
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Figure 4-6 Immobilized QHREDGS peptide in chitosan-collagen films promotes diabetic adult human
primary keratinocyte survival and migration. (A) DHEK attachment on the chitosan-collagen films in the
presence or absence of immobilized QHREDGS peptide. Image analysis showed no difference in the number of
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attached DHEKs (stained with TO-PRO) among the three groups (scale bar = 200 μm). n=3 independent
experiments and each experiment performed in triplicate. (B) Immunoblotting with anti-phosphorylated Akt and
anti-Akt antibodies showed up-regulation of Akt activation during DHEK attachment. GAPDH was used to ensure
even loading. n=3 independent experiments and each experiment performed in duplicates. (C) DHEK survival
following hydrogen peroxide treatment was determined by the EarlyTox Cell Integrity assay. QHREDGS peptide in
the chitosan-collagen film protected DHEKs against H2O2-induced cell death in a dose-dependent manner. n=4-6
and each experiment performed with at least six replicates for each condition. (D) Representative immunoblots of
phosphorylated Akt or MAPKp42/44 and total Akt or MAPK p42/44. (E) Quantification of immunoblotting revealed up-
regulation of the Akt and MAPK p42/44 activation of DHEKs under H2O2 stress. GAPDH was used to ensure even
loading. n=3 independent experiments and each experiment performed in duplicates. (F) Representative examples of
DHEK wounding experiments on chitosan-collagen films with or without immobilized QHREDGS peptide (scale
bar = 200 μm). Confluent HEKs were stained with CellMask Green, wounded at time 0 and maintained for 6 h in
KBM basal medium. The wounds were outlined and the area at the indicated times were normalized to the initial
wound size (time 0). DHEK migration on the films with QHREDGS peptide was accelerated compared to the
peptide-free control. n=3 independent experiments and each experiment performed with at least four replicates for
each condition. Data presented as mean ± SD. * indicates P < 0.05.
Figure 4-7 The presence of immobilized QHREDGS peptide promotes DHEK attachment on chitosan-only
films. (A) Representative images of DHEKs on chitosan-collagen films in the presence or absence of immobilized
QHREDGS peptide. Scale bar = 200 μm. Cell nuclei are shown in blue (DAPI). (B) Image analysis showed an
increased number of attached DHEKs in the presence of immobilized QHREDGS peptide. The number of attached
DHEKs was normalized to the number that attached to tissue culture polystyrene (TCP) in the same experiment. One
representative experiment is shown of n=3 independent experiments with three replicates for each condition in one
experiment. Data presented as mean ± SD. * indicates P < 0.05.
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Figure 4-8 DHEKs form adherens junctions during migration and the accelerated migration is not associated
with a difference in cell density. (A) Representative images of DHEKs in KGM basal medium (0.1 mM Ca2+) on
chitosan-collagen films in the presence or absence of QHREDGS peptide at the end of migration (6h). Adherens
junctions (green = E-cadherin) were present in all three groups. Scale bar = 50 μm. Cell nuclei are shown in blue
(DAPI). (B) DHEK cell density characterized by DAPI counterstaining at the end of migration (6 h). There was no
difference in the number of DHEKs on chitosan-collagen films in the presence or absence of QHREDGS peptide.
n=3 and each experiment performed with four replicates. Data presented in mean ± SD.
4.3.4 QHREDGS-immobilized hydrogel promotes wound healing in
db/db diabetic mice
We investigated whether the QHREDGS peptide immobilized to the chitosan-collagen hydrogel
could accelerate wound healing in diabetic mice. This hydrogel system was chosen as a delivery
vehicle because of its rapid gelation under physiological conditions and its persistence for a
period of 3 weeks in vivo [13]. In vivo biocompatibility was also demonstrated in previous
myocardial infarction model studies [13]. Therefore, only one application of the hydrogel onto
the wounds was needed for the 2 week study. A full-thickness excision wound (Fig. 4-9A) was
created on eight weeks old, male BKS.Cg-Dock7m +/+ Leprdb/J mice (db/db). This model was
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selected because the animal is leptin receptor deficient and represents a type II diabetes model
characterized by hyperglycemia, obesity, hyperinsulinemia, and impaired wound healing.
Moreover, this strain heals wounds primarily by granulation tissue formation rather than by
contraction [441].
Quantification using fluorescently labelled peptide demonstrated that in reconstituted chitosan
solutions, the amount of conjugated QHREDGS peptide was 17.5 ± 2.2 nmol/mg(chitosan) in Low
conditions and 41.5 ± 1.4 nmol/mg(chitosan) in High conditions. This was converted to a peptide
concentration in the final chitosan-collagen hydrogel of 43.8 ± 4.4 μM in Low conditions and
103.8 ± 3.5 μM in High conditions. With a view to future clinical translation, we only tested the
Low condition in the in vivo studies to minimize the amount of peptide applied to the wound. A
single application of 50 μL Low chitosan-collagen hydrogel (2.2 nmol immobilized QHREDGS
peptide; Peptide) was applied to the wound. The chitosan-collagen hydrogel alone without the
peptide (Ctrl) and a no hydrogel/no peptide (Blank) were used as controls. A secondary dressing,
Tegaderm™ film, was applied on top of the wound with or without the hydrogel, to maintain a
moist environment. As shown in Fig 4-9B, the presence of immobilized QHREDGS in the
hydrogel resulted in significantly smaller wounds on day 14 compared to the controls. Image
analysis of the wound gross morphology performed by an investigator blinded to the study
groups demonstrated faster wound healing in the Peptide group starting on day 8. Administration
of the chitosan-collagen hydrogel without the immobilized QHREDGS peptide (Ctrl) had no
significant effect on the wound closure rate compared to the Blank controls.
We also examined the wound histology by Masson’s trichrome staining and confirmed the
location of the epithelial tongue using pan-keratin staining (Fig. 4-9C). The wound edge was
defined as the distance between the two boundaries of intact skin (thin musculature of the
panniculus carnosus).There was no significant difference amongst the three groups in the wound
edge distance, indicating no difference in wound contraction (Fig. 4-9D i). The epithelial gap
was defined as the distance between the two advancing epithelial tongues (Fig. 4-9C i-vi) and
the epithelial gap in the Peptide group was significantly smaller than in the Blank and Ctrl
groups (Fig. 4-9D ii, Fig. 4-11). The re-epithelialization percentage was defined as the ratio of
the distance that has been re-epithelialized over the wound edge distance, and the re-
epithelialization percentage was higher in the presence of the QHREDGS peptide compared to
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the controls (Fig. 4-9D iii, Fig. S6). The Peptide group also developed significantly more
granulation tissue (Fig. 4-9D iv) compared to the controls. Moreover, the epidermal thickness of
the advancing epithelial tongue was significantly smaller in the Peptide group than in the Blank
and Ctrl groups (Fig. 4-9D v), which indicates more effective epidermal cell migration. There
was no difference in the epithelial thickness of the skin remote from the wounds among the three
experimental groups (Fig. 4-10).
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Figure 4-9 QHREDGS-immobilized hydrogel promotes wound healing in db/db diabetic mice. (A)
Representative images of the 8-mm full-thickness dorsal wounds on db/db diabetic mice. (B) Representative gross
images of the initial wounds on day 0 (D0) and the wounds at 14 days (D14) after treatment with no hydrogel
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(blank), ctrl chitosan-collagen hydrogel (ctrl), and QHREDGS peptide conjugated chitosan-collagen hydrogel
(peptide). Quantification of the wound size as a percentage of the original wound area revealed faster wound closure
in the peptide-treated mice at day 8-14. n=4. (C) Representative images of Trichrome stained tissue sections of
wounds treated with blank, control chitosan-collagen hydrogel, and peptide conjugated chitosan-collagen hydrogel
on day 14. Black arrows indicate wound edges; red arrows indicate the tips of the healing epithelial tongue. The tips
of the healing epithelial tongue were confirmed by pan-keratin staining as shown in the insets. Inset scale bar = 50
μm. (D) Quantification of wound size from histological samples collected 14 days after treatment. (i) Image analysis
showed no significant difference among the three groups in the wound edge distance. (ii) The peptide treatment
significantly reduced the size of epithelial gap, indicating accelerated wound closure compared with the blank and
control groups. (iii) The peptide treatment significantly increased the re-epithelialization percentage at the end of
experiment compared with the blank and control groups. (iv) The peptide treatment significantly increased the size
of the granulation tissue compared to the blank and control groups. (v) Average thickness of the epidermis within
300 μm of the leading edge of the wound. The epidermal thickness in the peptide-treated group was lower than the
blank and control groups. n=4. Data presented as mean ± SD. * indicates P < 0.05.
Figure 4-10 Thickness of the unwounded epidermis. (A) Representative images of Trichrome stained tissue
sections of unwounded epidermis in Blank, Ctrl and Peptide groups. Scale bar = 50 μm. (B) Image analysis revealed
no significant difference in unwounded epidermal thickness among the three groups.
Figure 4-11 An example of wound re-epithelialized after two weeks with a single treatment of QHREDGS
peptide in the chitosan-collagen hydrogel. Black arrows indicate wound edges; red arrows indicate tips of the
healing epithelium tongue. Re-epithelialization was quantified at 92%.
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4.3.5 Accelerated QHREDGS-induced diabetic wound healing does not involve changes in the extent of angiogenesis of the granulation tissue
Figure 4-12 The improvements in the diabetic wound healing process induced by the QHREDGS peptide are
not associated with increased angiogenesis within the granulation tissue. (A) Representative images of CD31-
stained tissue sections of wounds treated with no hydrogel (blank), control chitosan-collagen hydrogel (ctrl), and
QHREDGS peptide conjugated chitosan-collagen hydrogel (peptide) on day 14. Scale bar = 300 μm. (B)
Representative images of smooth muscle actin (SMA)-stained tissue sections from blank, control, and peptide
groups on day 14. Scale bar = 300 μm. (C) Image analysis showed no significant difference among the three groups
in (i) vessel density, (ii) CD31-positive area percentage, and (iii) SMA-positive area percentage. n=4. Data presented
as mean ± SD. * indicates P < 0.05.
To further characterize the granulation tissue, we compared the density of microvessels and
contracting myofibroblasts in the three experimental groups by immunohistochemistry with
antibodies against CD31 (Fig. 4-12A) and α-smooth muscle actin (α-SMA) (Fig. 4-12B). There
was no difference in the vessel density and CD31 positive area percentage among the three
groups based on measurements obtained from six random locations within the granulation tissue
(Fig. 4-12C i and ii). This was further confirmed by an automated algorithm analysis of the
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entire granulation tissue that showed no significant difference amongst the three groups in terms
of vessel density, CD31 positive percentage, lumen area, vascular area, vessel area, vessel
perimeter, or vessel wall thickness (Fig. 4-13). There was also no significant difference in the
density of myofibroblasts amongst the three groups as determined by α-SMA staining (a
common, albeit non-specific, marker of myofibroblasts) (Fig. 4-12C iii). Myofibroblasts are
considered to be the main contributors to wound contraction [442] and the absence of a
difference in myofibroblast cell density is consistent with the histological results showing no
difference in the wound edge size amongst the groups (Fig. 4-9D i). Taken together, the
accelerated wound healing in the presence of the conjugated QHREDGS peptide cannot be
attributed to changes in granulation tissue blood vessel density or to myofibroblasts, although
there was more granulation tissue overall in the Peptide group (Fig. 4-9D iv).
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Figure 4-13 QHREDGS peptide does not affect microvessel number and size within granulation tissue. Micro-vessel analysis of (A) CD31 positive area percentage and microvessel density, (B) lumen area, (C) vascular
area, (D) vessel area, (E) vessel perimeter, and (F) vessel wall thickness within the entire granulation tissue. Data
presented as mean ± SD. n=4.
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4.4 Discussion
Angiogenesis has been a primary target of therapeutic interventions in wound healing by local
application of growth factors (e.g. epidermal growth factor [443, 444], fibroblast growth factor
[445, 446], vascular endothelial growth factor [447, 448], and angiopoietin [449]) or angiogenic
cells [450, 451]. However, diabetic patients are often reported to have dysfunctional endothelium,
which cannot respond efficiently to growth factor stimulation [452, 453]. Diabetics also suffer
from the insufficient recruitment of circulating endothelial progenitor cells (EPCs) critical for
wound repair [240, 241]. As a result, many of these approaches failed to meet the FDA-accepted
primary efficacy endpoint standard of “complete wound closure within 12 weeks for any dose”
[454].
We chose to take a different approach here, by considering the hallmarks of true regenerative
healing as observed during the closure of embryonic wounds and scarless healing in model
organisms such as flies, zebrafish and newts. In all of these models, rapid, coordinated and
collective migration of epidermal cells was critical for regenerative healing [429, 430]. Hence,
we focused on developing a hydrogel treatment that would promote the collective migration of
keratinocytes and enhanced granulation tissue formation. Different mechanisms have been
proposed to explain the cell-cell interactions in collective cell migration, including local tractions
pulling cooperatively towards unfilled space (termed “kenotaxis”) [455], mechanical exclusion
interactions between cells [456], and intercellular adhesion and tension [457]. In our in vitro
studies, calcium concentration was increased when necessary to ensure that both normal and
diabetic keratinocytes migrated collectively rather than individually so that the treatment effect
on collective migration could be accurately described.
Following its discovery [458], there has been a growing recognition that angiopoietin-1 is an
important regulator of cellular processes including vascular protection [459], cardiac remodeling
[460-462], inflammation [463-465], and wound healing [449]. Due to the insolubility of
angiopoietin-1, its derivatives have been developed to promote wound healing by angiogenesis
[466-468]. Here, we assessed the effect of a novel water-soluble peptide, QHREDGS, derived
from the fibrinogen-like domain of angiopoietin-1 and conserved amongst species [460], on
promoting normal and diabetic keratinocyte survival and collective migration. In this study,
soluble QHREDGS peptide supplemented in the culture medium protected human keratinocytes
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against H2O2 stress and induced up-regulation of Akt and MAPKp42/44 phosphorylation. These
findings are consistent with previous reports for angiopoietin-1 in human keratinocytes and
melanocytes [435]. Our previous studies have also reported that a scrambled peptide
(DGQESHR or DQSHGER) does not exert prosurvival effects similar to the QHREDGS peptide
in cardiac cells [7, 469], endothelial cells [10], and iPSCs [11] motivating the omission of the
scrambled peptide in the studies described here.
Importantly, the QHREDGS peptide functions through interactions with 1- containing integrins
that are involved in cell-matrix interactions, rather than Tie2 receptors that reside mainly on
endothelial cells [10, 11, 13]. This enables the QHREDGS peptide to act on various cell types
including keratinocytes, which express the 31 integrin implicated in their enhanced survival
and migration [470-472]. The integrin interaction also motivated the presentation of the
QHREDGS peptide as a matrix bound ligand. Furthermore, the short peptide sequence
QHREDGS can be modified using versatile chemical methods and does not require a specific
orientation or conformation to function, as does the full length protein [473]; and peptides are
less susceptible to degradation due to proteolysis or hydrolysis during modification and after
delivery to the native environment compared to full length proteins [2].
Here, we used EDC chemistry to conjugate the QHREDGS peptide to the backbone of chitosan,
which is a zero-length cross-linker that does not add any other moieties to the final product,
features important for further clinical translation. Peptide immobilization also provides the
advantages of localized action in the target tissue and a lower total dose requirement over
application in an encapsulated or soluble form. We confirmed not only that the prosurvival effect
of the QHREDGS peptide was preserved after conjugating and immobilizing it in the chitosan-
collagen films, but that the immobilized QHREDGS peptide was able to promote keratinocyte
collective migration whereas the soluble peptide did not. Notably, in the context of the chitosan-
collagen film system, the conjugated QHREDGS peptide is presented in close proximity to
collagen, thereby creating an environment reminiscent of the collagen-glycosaminoglycan
interaction in the native ECM [474].
Using DHEKs from an adult diabetic patient we determined that the prosurvival, promigratory
effects of immobilized QHREDGS peptide could be translated to diabetic keratinocytes. Diabetic
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chronic wounds often occur to elderly population (the source of the primary DHEKs in this study)
due to age-associated comorbidities [475]; and advanced protein glycosylation has been
associated with diabetes and aging that can alter the functional properties of important ECM
components such as collagen [243]. Despite these challenges, the immobilized QHREDGS
peptide was effective in the in vitro diabetic wound healing model, which suggested the
possibility of in vivo benefits to diabetic wound healing.
Although our previous in vitro study showed that QHREDGS peptide promoted endothelial cell
survival, metabolism, and tube formation mediated by integrin interactions [10], the accelerated
diabetic wound healing shown in the animal studies here was not associated with enhanced
angiogenesis per area of the granulation tissue. Since there was more granulation tissue in the
Peptide group, consequently higher total number of blood vessels was achieved in the wound
with the Peptide hydrogel treatment, although blood vessel density was the same amongst the
groups. This is consistent with our previous study in a rat myocardial infarction (MI) model,
which showed an increase in large vessels within the MI border zone but no difference in the
total vascularization [13]. Alternatively, the diabetic endothelium is associated with enhanced
degradation of nitric oxide, an important regulator of inflammation, angiogenesis, and re-
epithelialization, due to the presence of excessive ROS [242]; and in our previous study it was
shown that the QHREDGS peptide can induce enhanced endothelial cell nitric oxide production
[10]. Hence, it is possible the improved wound healing effects induced by the QHREDGS
peptide could be attributable in part to nitric oxide production.
Our previous study demonstrated the retention of the chitosan-collagen hydrogel over two weeks
post-injection in the infarcted rat heart, an extremely dynamic, inflammatory environment [13].
In this study, we applied the QHREDGS peptide in the same chitosan-collagen hydrogel used as
a single treatment and found it to be sufficient to accelerate wound healing in a clinically
relevant, genetically modified db/db diabetic mouse model at two weeks. The ability to induce an
effective wound healing response with a single application is an important clinical consideration
as it removes the need for frequent repeated applications that can disturb the healing process, it
reduces the pain and discomfort associated with frequent dressing changes, and it requires fewer
healthcare resources [475, 476]. We also selected the use of a single mouse model for the
analysis of therapeutic efficacy, which can be seen as a potential limitation of our study.
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However, the db/db diabetic mice are used most often as a model of human diabetic chronic
wound [477] and the genetically modified db/db diabetic mouse has been used previously to
support clinical trial development with other molecules [478].
To tease out the extent to which the QHREDGS peptide could induce wound healing, we
selected a larger 8-mm diameter initial wound size rather than the commonly used 6-mm wounds
[282, 362]. We also selected the chitosan-collagen hydrogel with the low concentration of
QHREDGS peptide for our in vivo study based on the concept that pharmaceutical agents should
be applied at a minimal effective dosage to avoid potential side effects. For example, an
increased rate of mortality secondary to malignancy was reported for patients treated with > 45 g
of Regranex® (180 nmol PDGF-BB) [479]. Here a single dose of 4.4 nmol peptide/cm2 of wound
was found to be an effective dose to promote granulation tissue formation and wound closure in
vivo, whereas daily application of 0.4 nmol PDGF-BB (Regranex®)/cm2 of wound was found to
promote granulation tissue formation but did not shorten the time to wound closure in a db/db
mouse model [480].
The QHREDGS peptide is available by cost-effective synthesis with a precisely defined
composition, offering an additional advantage to potential clinical applications. The cost of the
synthetic QHREDGS peptide used in our study was $1.6CAD/mg, which is approximately 20
000-times cheaper than the commercially available rhPDGF-BB, an active component of the
FDA approved Regranex®, which costs $33,000CAD/mg (Fisher Scientific). The choice of the
hydrogel components, collagen and chitosan, has also been motivated by clinical translation
considerations. Chitosan has been approved by the FDA for use in humans in topical applications
(e.g. HemCon® bandages). Similarly, collagen is a component of numerous wound dressings
currently on the market in different formulations such as freeze-dried sheet, pastes, pads,
powders, and gels (e.g. INTEGRA™ Matrix Wound Dressing, BIOSTEP Collagen Matrix, BGC
Matrix®, Stimulen™ Collagen).
Here, we have demonstrated that keratinocyte survival and collective migration represent
promising alternative therapeutic targets for diabetic chronic wounds that support re-
epithelialization, a hallmark of wound regeneration and closure. In a genetically modified
diabetic mouse, the immobilized QHREDGS peptide accelerated the wound healing process by
promoting re-epithelialization rate and granulation tissue formation, without significantly
86
affecting angiogenesis. Moreover, the immobilized QHREDGS peptide did not increase the
number of -SMA positive cells, e.g. myofibroblasts, that can induce the antithetic wound
regeneration processes of wound contraction, collagen overproduction, and scar formation [481].
On the basis of our reported results, a number of questions arise. The efficacy of a potential
therapeutic intervention providing synergistic regulation of angiogenesis from angiogenic growth
factors (e.g. VEGF and bFGF) and re-epithelialization from QHREDGS peptide in diabetic
chronic wounds is yet unknown. Previously, we have successfully immobilized angiogenic
growth factors (VEGF and bFGF) on collagen scaffold and observed promoted angiogenesis in
vivo [88]. Notably, the chitosan-collagen hydrogel system is capable of delivering other
molecules at the same time as the chitosan backbone is amenable to various chemical
modification [482, 483].
In conclusion, the QHREDGS peptide promoted keratinocyte adhesion and collective migration
in vitro, as well as survival against H2O2 stress through Akt and MAPKp42/44 pathways. In vivo,
the QHREDGS peptide immobilized to a chitosan-collagen hydrogel accelerated diabetic wound
healing by enhanced re-epithelialization and granulation tissue formation. Together, our data on
both normal and diabetic human primary keratinocytes and in a db/db diabetic mouse model
demonstrate the translational relevance of the QHREDGS peptide in treating diabetic wounds.
We propose the QHREDGS peptide as a therapeutic candidate for promoting diabetic wound
healing.
4.5 Conclusion
In conclusion, the QHREDGS peptide promoted keratinocytes adhesion, survival against H2O2
stress, and collective migration in vitro involving up-regulation of Akt and MAPKp42/44 pathways.
When immobilized in a chitosan-collagen system, the QHREDGS peptide accelerated diabetic
wound healing in a manner not dependent on angiogenesis or wound contraction. Together, our
data on human primary keratinocytes, both normal and diabetic, and in a db/db diabetic mouse
model demonstrate clinical relevance and we propose QHREDGS peptide as a therapeutic
candidate for promoting diabetic wound healing.
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4.6 Acknowledgments
We thank L.A. Reis for training on the chitosan-collagen hydrogel preparation. We thank L.E.
Fitzpatrick and M.V. Sefton for input on animal study and useful discussion. This work is funded
by the NSERC Steacie Fellowship to M.R., Canadian Institutes of Health Research (CIHR)
Operating Grants (MOP-126027 and MOP-137107), NSERC Discovery Grant (RGPIN 326982-
10), and National Institutes of Health Grant 2R01 HL076485. The authors declare that they have
no competing interests. All data and materials are available.
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Chapter 5
5 Microfabricated perfusable cardiac biowire: a platform that mimics native cardiac bundle7
5.1 Introduction
Cardiovascular diseases are important targets for pharmacological therapy because they are
associated with high morbidity and mortality rates[484]. In vitro engineered models may serve as
cost-effective alternatives to animal models due to improved system control and higher
throughput. In recent years, tissue engineering methods have been significantly advanced to
generate functional three-dimensional (3D) cardiac tissues in vitro[485-487], which better
recapitulate the complexity and electro-mechanical function of native myocardium compared to
conventional in vitro systems of single cell suspensions or monolayers. Moreover, with the
opportunities brought about by human pluripotent stem cells (hPSC), tissue engineering methods
hold a great promise in developing patient-specific medical treatment[488].
In native myocardium, cardiomyocytes are highly anisotropic, usually in a length of 80-100 μm
and 20-30 μm in diameter[489]. Each cardiomyocyte adjoins neighboring cardiomyocytes by
specialized intracellular junctions, such as gap junctions and desmosomes, to form a complex 3D
network, or syncytium. On tissue level, native cardiomyocytes are organized into spatially well-
defined cardiac bundles with supporting vasculature. This highly organized architecture is
critical for electro-mechanical activation, propagation of electrical signals, and global cardiac
function[490]. Cardiac tissues generated by current tissue engineering methods often poorly
recapitulate this architecture.
Increased cardiovascular risk is one of the major unwanted side effects of new drug candidates,
which frequently leads to usage restriction or even withdrawal from the market[491]. Cardiac
7 Copyright © 2014 Royal Society of Chemistry. Contents of this chapter have been published in Lab Chip: Xiao Y,
Zhang B, Liu H, Miklas JW, Gagliardi M, Pahnke A, Thavandiran N, Sun Y, Simmons C, Keller G, Radisic M.
Microfabricated perfusable cardiac biowire: a platform that mimics native cardiac bundle. Lab Chip. 2014; 14:869–
82. Reuse with permission from Royal Society of Chemistry. A link to the published paper can be found at:
http://pubs.rsc.org/en/content/articlelanding/2014/lc/c3lc51123e
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toxicity was the main reason behind withdrawal of numerous drugs from the market, including
well known examples such as Vioxx or Avanida, accounting for up to 20% of all drug
withdrawals[492, 493]. Thus, it is essential to identify these risks at an early stage in drug
development process to define safety profile and avoid cost escalation. Despite the exceptional
progress in developing cardiac disease models with hPSC (Timothy[494], long QT[495],
LEOPARD syndrome[496] and dilated cardiomyopathy patients[497]), most studies still use
cardiac monolayers that do not capture architectural complexity of the native cardiac niche. After
pharmacologic agents are administrated into human body, they are circulated through the
vasculature and delivered to the myocardium by the blood in capillaries. Current in vitro drug
testing systems, however, expose the cardiac cells to the pharmacologic agents directly from the
culture media in conventional well plates[411, 498-500]. Thus, developing in vitro cardiac
systems that can recapitulate the perfusion scenario could provide improved physiological
relevance when assessing pharmacological effects on cardiac tissue in vitro.
We recently described a development of a human cardiac micro-tissue, termed biological wire,
which captures some of the architectural complexity of the native myocardium and enables
maturation of cardiomyocytes derived from human pluripotent stem cells with the application of
electrical stimulation[413]. However, these original biowires lacked perfusion, a critical aspect
for mimicking native physiology and mass transfer. We describe here technological
developments required to create a perfusable biowire conducive to electrical field stimulation
and we prove the feasibility of drug testing in this system.
Perfusable cardiac biowires were generated using a polytetrafluoroethylene (PTFE) tubing
template in microfabricated bioreactors, which provided contact guidance for cells to align and
elongate. To demonstrate the feasibility of this platform for drug testing, we supplied nitric oxide
(NO) in the cell culture channel to provide biochemical stimulation to cardiomyocytes within the
biowire. NO was released from perfused sodium nitroprusside (SNP) solution and passed
through the tubing wall to reach the tissue constructs with cardiomyocytes. This bioreactor was
also integrated with electrical stimulation to further improve phenotype of cardiomyocytes.
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5.2 Materials and methods
5.2.1 Biowire bioreactor design and fabrication
The biowire bioreactor consisted of two main units, the microfabricated platform made of
poly(dimethysiloxane) (PDMS) and the suspended template made of either silk 6-0 suture or
PTFE micro-tubing. To fabricate the PDMS platform, standard soft lithography technique was
used to make to a two-layer SU-8 (Microchem Corp., Newton, MA) master[501]. The first layer
included the template channel and the cell culture chamber, while second layer included only the
cell culture chamber. Then PDMS was cast onto the SU-8 master and baked for 2 hr at 70 °C. A
biowire template was then anchored to the two ends of the PDMS platform followed by the
bioreactor sterilization in 70% ethanol and overnight UV irradiation.
5.2.2 Perfusion system design and fabrication
In order to provide perfusion through the tubing template, two microfabricated modules, drug
reservoir and connecting channel, were added to the biowire bioreactor. Both modules were
fabricated by first molding PDMS with a single-layer SU-8 master (length × width × height =
10 × 1 × 0.3 mm). The drug reservoir was created by cutting through the PDMS using an 8 mm
biopsy punch (Sklar). The biowire bioreactor channel was connected to the drug reservoir and
connecting channel with the PTFE tubing (inner diameter (ID) = 0.002 inch, outer diameter (OD)
= 0.006 inch, Zeus). Tygon tubing (ID = 0.01 inch, OD = 0.03 inch, Thomas Scientific)
connected the perfusion system to external negative pressure generated by a peristaltic pump.
The perfusion rate was characterized by the liquid volume collected at the outlet from the
peristaltic pump (n = 3). All the connecting points were secured by epoxy glue and three
microfabricated modules were plasma bonded to a glass slide.
5.2.3 Cell culture
Neonatal rat cardiomyocytes were obtained from 2-day old neonatal Sprague-Dawley rats as
described previously[9] and according to a protocol approved by the University of Toronto
Committee on Animal Care. The culture media contained 10% (v/v) fetal bovine serum, 1% (4-
(2-hydroxyethyl)-1-piperazineethanesulfonic acid) (HEPES), 100 U/ml penicillin-streptomycin,
1% Glutamine, and the remainder Dulbecco`s modified Eagle`s medium.
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Cardiac differentiation in embryoid bodies (EBs) of HES-2 human embryonic stem cell (hESC)
line was performed as described previously[388, 502]. Briefly, EBs were first cultured in
StemPro-34 (Invitrogen) media containing BMP4 (1 ng/ml). On day 1, they were transferred to
the induction medium (StemPro-34, basic fibroblast growth factor (bFGF; 2.5 ng/ml), activin A
(6 ng/ml) and BMP4 (10 ng/ml)). On day 4, the EBs were removed from the induction medium
and re-cultured in StemPro-34 supplemented with vascular endothelial growth factor (VEGF; 10
ng/ml) and Inhibitor of Wnt production-2 (IWP2; 2 μM). On day 8, the medium was changed
again and the EBs were cultured in StemPro-34 containing VEGF (20 ng/ml) and bFGF (10
ng/ml) for the remainder of EB culture as well as for the biowire culture. EBs were maintained in
hypoxic environment (5% CO2, 5% O2) for the first 12 days and then transferred into a 5% CO2
for the remainder of the culture period. EBs were dissociated for seeding in biowires at day 23
(EBd23).
5.2.4 Generation of cardiac biowires
Cardiac cells (from neonatal rat isolation or hESC differentiation) were first suspended at 200
million /ml (unless specified otherwise) in Collagen Type I based gel (2.5 mg/ml of rat tail
collagen type I (BD Biosciences) neutralized by 1N NaOH and 10× M199 media as described by
the manufacturer) with the supplements of 4.5 μg/ml glucose, 1% (v/v) HEPES, 10% (v/v)
Matrigel (BD Biosciences), and 2 μg/ml NaHCO3. Suspended cardiac cells were then seeded
into the cell culture channel (3 μl per biowire). After 30 min incubation at 37 °C to induce the
gelation, appropriate media were added. Cardiac biowires were kept in culture for up to 14 days
with media change every 2-3 days.
Cardiac biowires starting with different cell densities (100 and 200 million/ml) were seeded to
study the effect of the cell seeding density. Collagen-based gel was seeded into the cell culture
channel without loading cardiac cells, to serve as a cell-free control. Ultra-long cardiac biowires
were generated with customized biowire bioreactor that was 5 cm long fabricated in a similar
manner as described above and seeded with neonatal rat cardiomyocytes.
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5.2.5 Quantification of compaction rate
After seeding, brightfield images of the biowires were taken every day (n = 3 per group) using an
optical microscope (Olympus CKX41) and the diameters of the biowires at five distinct locations
were averaged with image analysis.
5.2.6 Immunostaining and Fluorescent Microscopy
Biowires were fixed with 4% paraformaldehyde, permeablized by 0.25% Triton X-100, and
blocked by 10% bovine serum albumin (BSA). Immunostaining was performed using the
following antibodies: mouse anti-cardiac Troponin T (cTnT) (Abcam; 1:100), rabbit anti-
Connexin 43 (Cx-43) (Abcam; 1:200), mouse anti-α-actinin (Abcam; 1:200), goat anti-mouse-
Alexa Fluor 488 (Jackson Immuno Research; 1:400), anti-rabbit-TRITC (Invitrogen; 1:200),
anti-mouse-TRITC (Jackson Immuno Research; 1:200). Nuclei were counterstained with 4',6-
diamidino-2-phenylindole (DAPI) (Biotium; 1:100). Phalloidin-Alexa 660 (Introgen; 1:600) was
used to stain F-actin fibers. For confocal microscopy, the stained cardiac biowires were
visualized under an inverted confocal microscope (Olympus IX81) or an upright confocal
microscope (Zeiss LSM 510).
5.2.7 Quantification of nuclei elongation and alignment
Cell nuclei within the biowires were visualized by DAPI staining and z-stack images were
obtained by confocal microscopy with 3 μm interval. Each stack of the confocal images was
analyzed in ImageJ 1.45s (National Institutes of Health, USA) with an automated algorithm
described by Xu et al[503] with approximately 1000 nuclei analyzed per sample. Nuclei
elongations were characterized as nucleus aspect ratios, the ratio of long axis over short axis of
the nuclei, and nuclear alignment was characterized by orientation angles. In the control
monolayer group, orientation of the nuclei was characterized compared to an arbitrarily defined
orientation, while in the biowire group, the orientation of the suture templates was set as
reference.
5.2.8 Characterization of perfusable biowires
Neonatal rat cardiac cells were seeded into the perfusable biowire reactors with tubing template.
After cultivation for 7 days, the cardiac biowires were sectioned and visualized under
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environmental SEM (Hitachi S-3400 N). The biowires were imaged under variable pressure
mode at 70 Pa and 15 kV and the chamber temperature was −20°C.
To visualize the cross-section, perfusable cardiac biowires were stained with cTnT antibody and
then TRITC. Stained biowires were then cryo-sectioned into 500 μm thick sections using a
cryostat (Leica CM3050S) and mounted to Superfrost Plus glass slide (VWR). Images of the
cross-sectioned biowires were acquired by Olympus fluorescent microscope (Olympus IX81).
To demonstrate the feasibility of the perfusable biowire bioreactor, FITC-labeled polystyrene
beads (Spherotech Inc.) were added into the drug reservoir and perfused through the rat cardiac
biowire, while it beat spontaneously on day 8. Bright-field and fluorescent videos and images
were acquired with a fluorescence microscope (Olympus IX81).
5.2.9 Quantification of NO perfusion
Sodium nitroprusside (SNP) (Sigma) was dissolved in distilled water to make 200 mM SNP
solution and then added to the drug reservoir. Perfusion through the tubing template was driven
by the external peristaltic pump. Once the SNP solution perfused through the tubing, the
peristaltic pump was stopped and the entire perfusion system was kept in cell culture incubator.
NO amount in the cell culture channel (outside the PTFE tubing) was quantified with a
fluorometric Nitric Oxide Assay Kit (Calbiochem, 482655). In brief, samples collected from the
cell culture channels (8 μl, n = 3) at different time points (0.5 hr, 6 hr, and 24 hr) were converted
to nitrite by nitrate reductase and then developed into a fluorescent compound 1-H-
naphthotriazole. The fluorescent signals were quantified by a plate reader (Apollo LB 911,
Berthold Technologies) and compared to the nitrate standard.
5.2.10 NO treatment of human cardiac biowires
On day 7, the NO treatment of human cardiac biowire was initiated by perfusing the 200 mM
SNP solution and the peristaltic pump was stopped once the SNP solution was perfused through
the tubing. The beating activities of the human cardiac biowires were recorded at 16.67
frames/second before treatment and 24 hr post-treatment by Olympus IX81 while the biowires
were kept at 37°C. The beating activities of the human cardiac biowires were quantified by the
image analysis method described by Sage et al[504]. In brief, the movements of one spot at the
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same location on the human cardiac biowire before and after the NO treatment were
characterized.
5.2.11 Electrical stimulation
Different electrical stimulation conditions were applied to the rat cardiac biowires as described
previously [485]. The parallel stimulation chambers were fitted with two 1/4-inch-diameter
carbon rods (Ladd Research Industries) placed 2 cm apart, perpendicular to the biowires (such
that the electrical field was parallel with the biowire long axis), and connected to a stimulator
(S88X, Grass) with platinum wires (Ladd Research Industries). The perpendicular stimulation
chambers were built with two carbon rods 1 cm apart placed parallel with the biowires (i.e. the
filed was perpendicular to the long axis of the biowire). The biowires were pre-cultured for 4
days until the biowire structures were established and their spontaneous beating was
synchronized, and then subjected to the electrical field stimulation (biphasic, rectangular, 1 ms
duration, 1.2 Hz, 3.5-4 V/cm) for 4 days with 10 μM ascorbic acid supplemented in the culture
media while control biowires were cultured without electrical stimulation. At the end of
electrical stimulation, the rat cardiac biowires were double stained for cTnT with Alexa 488
conjugated antibody and Cx-43 with TRITC conjugated antibody, or their mechanical properties
were measured by atomic force microscopy (AFM). Confocal images were acquired using
identical microscope settings for all groups. Areas stained positive for cTnT staining (green
pixels) or Cx-43 staining (red pixels) were quantified by ImageJ using the identical thresholding
parameters in all groups.
For the human perfusable cardiac biowire, only parallel electrical stimulation was applied as
described above. Starting on day 4, electrical field stimulations (biphasic, rectangular, 1 ms
duration, 1 Hz, 3.5-4 V/cm) were applied for 4 days while control biowires were cultured
without electrical stimulation. Both stimulated and control biowires were perfused with culture
medium at a flow rate of 2 μl/min within the PTFE tubing driven by an external syringe pump
(PHD Ultra; Harvard Apparatus). At the end of electrical stimulation, the electrical properties of
the stimulated and control human cardiac biowires were characterized in terms of excitation
threshold (ET) and maximum capture rate (MCR) under external field pacing as previously
described [505].
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5.2.12 Atomic force microscopy (AFM)
After application of electrical stimulations for 4 days, rat cardiac biowires were tested using a
commercial AFM (Bioscope Catalyst; Bruker) mounted on an inverted optical microscope
(Nikon Eclipse-Ti). The force-indentation measurements were done with a spherical tip (radius =
5-10 μm) at nine distinct spots to evenly cover the center of the cardiac biowires with 5 nN
trigger force at 1 Hz indentation rate. The cantilever (MLCT-D, Bruker) had a nominal spring
constant of 0.03 N/m. Hertz model was applied to the force curves to estimate the Young's
modulus and detailed data analysis was described elsewhere[506]. All AFM measurements were
done in fluid environment at room temperature.
5.2.13 Statistical analysis
Statistical analysis was performed using SigmaPlot 11.0. Differences between experimental
groups were analyzed using t-test or one-way ANOVA with significant difference considered as
P < 0.05.
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5.3 Results
5.3.1 Generation and characterization of cardiac biowires
Figure 5-1 Cardiac bundles in native myocardium. (a) Schematic illustration of the structure of cardiac bundles
in native myocardium. Cardiomyocytes are elongated, aligned, and grouped into bundles around capillaries. (b)
Tangential section of adult rat myocardium with CD31 staining (brown). Nuclei were counterstained as light violet
(long arrows). The blood vessels were noted as asterisks while the capillaries were noted as short arrows. Scale bar =
200 μm. (c) Fluorescent image of tangential cryosection of neonatal rat myocardium. Cardiac troponin T (cTnT) was
stained against Alexa 488-labeled (green) antibody, showing its unique striation structure. Cell nuclei were
counterstained with DAPI (blue) and the long arrows indicate elongated nuclei. Scale bar = 20 μm.
The native myocardium has a highly anisotropic structure (Figure 5-1a) with a high density of
capillaries (Figure 5-1b) and surrounding elongated and aligned cells (Figure 5-1c). In the
native heart, extracellular matrix (ECM) serves as a template for cells to align and elongate[98].
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Figure 5-2 Generation of cardiac biowires with microfabricated bioreactor. (a) Within 7 days of cultivation,
neonatal rat cardiomyocytes (8.75 million cells/ml) remodeled the gel and compacted around the suture template to
form the biowire structure. Scale bar = 200 μm. (b) Device design of the microfabricated bioreactor with suspended
6-0 suture template. (c) A customized ultra-long cardiac biowire in the length of 5 cm (scale bar = 500 μm). (d)
Quantification of gel compaction and its dependence on initial seeding density of cardiomyocytes (mean ± SD, n =
3). With no cardiomyocytes seeded (gel only), the gel did not compact and form biowire structure. Biowires with
higher seeding density (200 million cells/ml) compacted faster than those with lower seeding density (100 million
cells/ml) during the remodelling.
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In addition, a structural correlation between directionality of capillaries and cardiomyocytes can
be readily observed[489]. We aimed to emulate this in our biowire bioreactor by introducing a
perfusable tubing template and by using a hydrogel, for cell seeding, consisting of ECM
molecules normally present in the native heart. Primary neonatal rat cardiomyocytes were used
to generate 3D, self-assembled cardiac biowires by seeding within type I collagen-based gel into
microfabricated PDMS platforms with suspended templates (Figure 5-2b). Seeded cells
remodeled and contracted the collagen gel matrix around the templates within a week (Figure 5-
2a, Figure 5-4a). The gel compaction only occurred with the presence of the seeded cells, as
cell-free gels did not compact or degrade during the culture time, and the compaction rate
positively correlated with the cell seeding density (Figure 5-2d). Cardiac biowires of different
dimensions could be generated by customizing the dimensions of the biowire bioreactor. Here,
we generated biowires as long as 5 cm (Figure 5-2c). Generation of longer biowires might be
possible; however it was not explored in this work.
Image analysis of the cell nuclei that counterstained with DAPI (Figure 5-3a, left) revealed
nuclei elongation and alignment along the axis of suture template. There was a significant
difference (p < 0.001) of nuclei aspect ratio between biowires and monolayer group (Figure 5-
3b). Compared to neonatal rat cardiomyocytes cultured in monolayer controls, biowires had a
lower frequency of cells in the smaller aspect ratio range and a higher frequency of cells in the
larger aspect ratio range (Figure 5-3c). Image analysis also revealed that cell nuclei in biowires
were oriented along with the axis of the suture template, while those in monolayers were
randomly distributed (Figure 5-3d).
Neonatal rat cardiac biowires started to beat spontaneously between 3 and 4 days post-seeding
and kept beating during gel compaction, demonstrating that the biowire bioreactor allowed for
electromechanical coupling of the cells within the hydrogel matrix. The spontaneous beating of
biowires with higher seeding density (200 million/ml) started earlier and was more prominent
than in those with lower seeding density (100 million/ml), which is thought to be a result of the
presence of more cardiomyocytes and better coupling. Immunohistochemistry staining showed
that the rat cardiac biowires expressed the sarcomeric protein, cTnT (Figure 5-3a, right).
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Figure 5-3 The suture template provides topographical cues in the biowires for the cardiomyocytes to
elongate and align. (a) Confocal images of the biowire with nuclei counterstained with DAPI and cardiac
Troponin-T (cTnT) stained with Alexa 488 (green). Scale bar = 100 μm. (b) Nuclei aspect ratios (~1000 nuclei
characterized per sample) of cardiac cells cultured as monolayer vs. seeded in the biowires plotted in box plot
showing the 1st quartile, median, and 3rd quartile with a significant difference between two groups (***, p < 0.001).
(c) Histogram showing the distribution of nuclei aspect ratios of biowire group and monolayer group (n = 3 per
group). There were significantly higher frequencies in the lower aspect ratio range in monolayer group (*, p < 0.05)
and higher frequencies in higher aspect ratio range in biowire group (#, p < 0.05). (d) Characterization of nucleus
orientation reveals random distribution of nuclei in the monolayer group (random direction as 0 ̊) and oriented
distribution of nuclei along with the suture template in the biowire group (orientation of suture template as 0 ̊).
Dashed lines indicate orientation angle, solid hemi-circular lines indicate the percentage gridlines, and grey
(monolayer) or blue (biowire) lines indicate the actual percentage of the nuclei in the specific orientation angle.
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5.3.2 Generation and characterization of perfusable cardiac biowires
Figure 5-4 Generation of perfusable cardiac biowires. (a) Neonatal rat cardiomyocytes (200 million cells/ml)
remodeled the gel and compacted around the tubing template (ID = 50.8 μm, OD = 152.4 μm). Scale bar = 200 μm.
A close-up view showing the tubing lumen at the end of the biowire is given at top-right. Scale bar = 100 μm. (b)
SEM images demonstrate that the cardiac tissue attached to the tubing surface and formed a uniform-thick layer
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after remodelling. (c) Representative phase contrast image (left) and confocal image (right) showing the circular
morphology of the cross section of the perfusable cardiac biowire with the expression of cardiac Troponin-T (cTnT).
Scale bar = 150 μm. (d) Set-up of the perfusion device. Two microfabricated modules were added to the bioreactor
system: a drug reservoir at one end of the biowire and a channel at the other end for connection to an external
negative pressure source. The entire biowire perfusion system was bonded on a glass slide (real-life image shown at
the top-left corner). (e) The tubing-templated biowire was perfused with FITC-labeled polystyrene beads (1 μm in
diameter). Dash lines illustrate the wall of the cell culture channel. FITC-labeled beads were indicated by arrows.
Asterisks indicate the auto fluorescence from the cardiomyocytes within the cardiac biowire. This image was over-
exposed to better visualize the fluorescent beads. Scale bar = 100 μm.
Primary neonatal rat and hESC-derived cardiomyocytes were used to generate perfusable cardiac
biowires with PTFE tubing template Figure 5-4d). Both cell types were able to form the cardiac
biowires and beat spontaneously (Figure 4a). As shown in SEM images, cells attached to the
smooth surface of the PTFE tubing after self-remodeling (Figure 5-4b). Cross sections of these
perfusable biowires showed that self-remodeled cells encircled the tubing template and expressed
cTnT (Figure 5-4c).
The feasibility of perfusable biowire bioreactor was demonstrated by perfusion with FITC-
labeled fluorescent beads. Perfusion rate driven by the peristaltic pump was quantified to be 2 ±
0.16 μL/min (n = 3). Bright field video showed both spontaneous beating activity of the rat
cardiac biowire and the perfusion of the fluorescent beads. The movement of the beads was
better visualized under fluorescent view. A snapshot of the video (Figure 5-4e) was overexposed
to provide better visualization of the fluorescent beads. The cardiac biowire was also visible in
this image due to the auto-fluorescence of cardiomyocytes.
5.3.3 NO treatment of human cardiac biowires by perfusion
To demonstrate feasibility of drug testing in the perfusable cardiac biowire, a pharmacological
agent, NO donor SNP, was applied to the culture media that was perfused through the tubing
lumen. As NO was generated in the tubing lumen, it diffused through the tubing wall reaching
the cell culture outer channel where the total amount of NO was quantified. The amount of NO
released from 200 mM SNP was quantified by a fluorometric assay which validated the
persistence of the NO release from SNP solution over several hours (Figure 5-5a). The
cumulative NO amount in the cell culture channel was 100 μM (800 pmol in 8μl), which
exceeded the physiological levels of NO in vivo[507].
Upon gel compaction, the hESC-derived cardiomyocytes within the biowires started spontaneous
beating. After NO treatment for 24 hr, performed by perfusion of NO-donor SNP through the
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tubing lumen, the spontaneous beating of human cardiac biowires slowed down and this was
further characterized by image analysis (Figure 5-5b, c). In order to compare beating frequency
changes between different biowires, the frequencies after 24 hr NO treatment were normalized to
the basal level (before treatment). The beating frequencies after NO treatment were significantly
lower than the basal level (74±3%, n = 3) while the control biowires remained the same (100±9%,
n = 3).
Figure 5-5 Nitric oxide (NO) treatment on human tubing-templated biowires. (a) Quantification of NO amount
passing through the tubing wall after perfusing SNP (200 mM) for 0.5 hr, 6 hr, and 24 hr. (b) 24 hr NO treatment
significantly slowed down the beating of biowires compared to the basal levels while there was no significant
change in the non-treated biowires (n = 3 per group, p < 0.01). (c) Quantified by image analysis, the beating rate of a
biowire after 24 hr NO treatment was less frequent compared to the basal level. (d) Confocal images showing the
disrupted α-actinin (green) structure within the NO-treated biowire (right) compared to the control (left) (top: lower
magnification, scale bar = 50 μm; bottom: higher magnification, scale bar = 10 μm).
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The degradation of cytoskeleton of cardiomyocytes within the biowires based on hESC derived
cardiomyocytes caused by NO treatment through perfusion was characterized using confocal
microscopy for α-actinin labeled with Alexa 488 conjugated antibody (green) and actin labeled
with Alexa 660 conjugated antibody (far red) (Figure 5-5d). It was possible to clearly discern
the striated pattern of sarcomeric Z-discs labeled with α-actinin in the control biowires, while the
NO treated biowires showed an overall punctate pattern.
5.3.4 Electrical stimulation of cardiac biowires
To demonstrate the versatility of the cardiac biowire bioreactor, electrical stimulation was
applied to further improve the phenotype of cardiomyocytes (Figure 5-6a).
Immunohistochemical staining showed that the rat cardiomyocytes in the biowires that
underwent electrical stimulation with the field parallel to the biowire long axis had more cTnT
positive structures oriented along with the axis of the suture template (indicated by the dashed
line), while those in non-stimulated biowires were randomly distributed and those in
perpendicular-stimulated biowires were found to be perpendicular to the suture template (Figure
5-6b). Moreover, the cardiomyocytes in the parallel- and perpendicular- stimulated biowires
showed stronger expression of Cx-43, a marker for the gap junctions between adjacent
cardiomyocytes, compared to the control biowires, indicating better coupling between the
cardiomyocytes in the stimulated groups (Figure 5-6b). This was also confirmed by comparing
the ratio of Cx-43 positive area over cTnT positive area under same magnification with identical
microscope settings (Figure 5-6c).
When handling the rat cardiac biowires outside the bioreactor, it was noticed that the parallel-
stimulated biowires were stiffer than the non-stimulated control. This was further assessed by
AFM analysis (n = 3 per group), which revealed significantly (p = 0.009) higher apparent
Young’s modulus of parallel-stimulated biowires compared to non-stimulated controls (Figure
5-6d).
The perfusable human cardiac biowires that underwent medium perfusion through the tubing and
parallel electrical stimulation at the same time showed improved electrical properties compared
to the non-stimulated controls as assessed by ET and MCR under electrical field stimulation. The
ET is the minimum electrical field voltage required for inducing synchronous contractions and
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the decreased ET of the stimulated biowires (Figure 5-6e) indicated better electrical excitability.
The MCR is the maximum beating frequency attainable while maintaining synchronous
contractions and the increased MCR (Figure 5-6f) of the stimulated biowires indicated improved
cell alignment and interconnectivity.
Figure 5-6 Electrical stimulation and perfusion of cardiac biowires. (a) Experimental set-up of biowires under
different electrical stimulation conditions. Carbon rods (in black) connected to an external stimulator provided either
parallel or perpendicular electrical field stimulation on cardiac biowires for 4 days starting on Day 4. (b)
Representative confocal images of biowires after application of different electrical stimulation conditions (left:
lower magnification, scale bar = 50 μm; right: higher magnification, scale bar = 10 μm). Parallel-stimulated biowires
showed more cTnT positive (green) structures oriented along with the suture template (indicated by dashed lines). (c)
Higher ratio of Cx-43 positive area over cTnT positive area indicated stronger expression of Cx-43 in both parallel-
and perpendicular-stimulated biowires compared to the non-stimulated controls (n = 4 per group, p < 0.05). (d)
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Young’s modulus of cardiac biowires characterized by AFM reveals that the parallel-stimulated biowires had higher
apparent Young’s modulus compared with the control biowires (**, p < 0.01). (e) Electrically stimulated perfused
biowires based on hESC derived cardiomyocytes had lower excitation threshold compared to the non-stimulated
controls (***, p < 0.001). (f) The electrically stimulated perfused biowires based on hESC derived cardiomyocytes
had higher maximum capture rate compared to the non-stimulated controls (*, p < 0.05).
5.4 Discussion
The native myocardium consists of spatially well-defined cardiac bundles with supporting
vasculature (Figure 1a) and the cardiomyocytes within the cardiac bundles are highly anisotropic
(Figure 1b). In this study, we have developed a microfabricated bioreactor to generate cardiac
biowires in vitro recapitulating the structure and function of native cardiac bundles. To the best
of our knowledge, this is the first study to examine the drug effects on cardiomyocytes by
perfusion within cardiac bundle model, which better mimics native myocardium mass transfer
properties compared to other engineered heart tissues. This bioreactor provided topographical
cues for the cardiac cells to elongate and align, and was also integrated with other cues, e.g.
electrical stimulation.
Gel compaction has been widely applied in tissue engineering to create 3D microtissue
constructs for in vivo implantation[508] and in vitro models[411, 509]. Compared to scaffold-
based constructs, the self-assembled constructs from gel compaction produce increased force of
contraction due to the higher cell density after the compaction[409]. Moreover, there is
increasing interest in microtissue constructs made by gel compaction as microarrays for drug
testing because they provide much higher throughput than conventional models[411, 412, 509,
510]. In this study, type I collagen was chosen as the main gel matrix as it is one of the main
ECM components of native myocardium. We noted that previous in vitro collagen-based models
only stayed intact for several days due to their poor mechanical properties[509]. In our
microfabricated system, with the mechanical support provided by the suspended templates, the
cardiac biowires remained stable in the bioreactor for weeks. We were able to generate cardiac
tissues in larger scale (up to 5 cm long) compared to other in vitro models and the dimensions of
the cell culture channel could be easily customized, which could render additional control over
the morphology of the cardiac biowires. The cell culture channels were initially designed to be
300 μm in height considering the limitations for oxygen and nutrient supply[511]. Moreover, the
presence of the templates enabled easy disassembly of the biowire from the bioreactor and facile
handling of the cardiac biowires at the end of cultivation for further characterization.
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Our microfabricated bioreactor was also able to generate cardiac biowires that are 5 cm long,
which is comparable to the height of the human heart. The feasibility of handling individual
cardiac biowire together with the ability to create macro-scale biowires raise up the prospect of
investigating the alignment of multiple cardiac biowires by bundling or weaving them together to
generate thicker structures, using similar methods as described by Onoe et al [512]. To
characterize the force generated by the cardiac biowires or cardiac biowire bundles, degradable
sutures could be used to generate template-free cardiac biowires which will be a topic of our
future studies.
To validate our microfabricated bioreactor, neonatal rat cardiomyocytes were used in preliminary
studies. Only when seeded at higher cell density (> 5×107 cells/ml), which is comparable to the
cell density in native rat myocardium (~108 cells/ml)[180]. the cardiac biowires started
spontaneous beating on day 3-4. The template provided contact guidance for the cells to elongate
and align along with, recapitulating the anisotropic properties of cardiomyocytes in the native
myocardium. The image analysis was done on cell nuclei due to the difficulty of defining cell
membranes within 3D tissue. However, nuclear alignment is a sufficient indication of cell
alignment and also one of the hallmarks of native myocardium (Figure 5-1c).
To further develop the biowire system, we used PTFE tubing as the template instead of the 6-0
silk suture. The commercially available PTFE tubing was chosen because it is biocompatible
(USP Class VI), extremely non-absorbent (ideal for drug testing), and micro-scale in dimension
(ID = 50 μm, OD = 150 μm), on the order of post-capillary venules in size[513]. Due to the small
size of the inner lumen, we used negative pressure to drive the perfusion instead of positive
pressure. Two microfabricated modules were added to the system to enable long-term perfusion
and incubation of the biowire system. Indicated by the shortening of biowire during self-
remodeling, the cell attachment on PTFE tubing was not as strong as that on silk suture, mainly
because of the smoothness of the PTFE tubing surface (Figure 5-4b). However, the cell-gel
composite was still able to assemble itself around the tubing with a circular cross-section.
In this study, NO was chosen as a model drug because of following reasons: (1) NO is produced
by endothelial cells in native myocardium, and then transported in the radial direction to
cardiomyocytes [514], the scenario we were trying to recapitulate in our biowire bioreactor; (2)
NO plays a critical role in regulating myocardial function, through both vascular-dependent and -
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independent effects[514]; (3) there is increasing evidence showing that NO is directly implicated
in cardiomyocyte disease development and prevention, such as in ischemia-reperfusion
injury[515]; (4) NO is a small gas molecular, which can readily pass the tubing wall. SNP was
chosen as the NO donor because it is a common NO donor used as vasodilator to treat pulmonary
hypertension and low cardiac output [516, 517]. Moreover, SNP aqueous solution was reported
to release NO at a constant rate over several hours in vitro[518].
For the NO treatment testing, we generated human cardiac biowires from hESC-derived
cardiomyocytes. The human cardiac biowire started spontaneous beating as early as day 1 and
the beating was synchronized within 7 days. After 24 hr of NO treatment, the beating frequencies
of the human cardiac biowires significantly slowed down compared to their basal level. This
result corresponds with the vasodilator effect of NO in vivo[519] and might be caused by
degradation of myofibrillar cytoskeleton, which has been seen by Chiusa et al[520]. However,
NO shows bi-polar inotropic effect at lower concentrations with diverse intracellular mechanisms
and there were discrepancies between studies due to the lack of standardization for in vitro
models[514]. Therefore our microfabricated bioreactor could serve as a novel platform to
uncover the effects of NO on cardiomyocytes at the tissue level.
To demonstrate the versatility of our biowire bioreactors, electrical stimulation was integrated
with the system as it has been reported to improve the phenotypes of cardiomyocytes[485, 501].
Because the cells in the cardiac biowires were anisotropic, we studied both parallel- and
perpendicular- field electrical stimulations on the rat biowires. The higher tissue stiffness under
parallel electrical stimulation, which was closer to isolated neonatal rat cardiac tissue (6.8 ± 2.8
kPa)[521], were attributed to more organized cellular contractile apparatus as characterized by
immunohistochemical staining. The perfusable human cardiac biowires were electrically
stimulated and perfused at the same time and this brings the prospect to study the interaction
between electrical stimulation and pharmaceutical agents delivered in a physiological manner. A
more detailed study on electrical stimulation alone of biowires based on human pluripotent
cardiomyocytes has been done in our group and indicated that electrical stimulation of
progressive frequency increase markedly improved the maturation of hPSC-derived
cardiomyocytes in terms of myofibril structure and electrical properties[413].
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Medium perfusion has been recognized to improve the viability and functionality of
cardiomyocytes within cardiac constructs in vitro since perfusion significantly improves oxygen
and nutrient supply[522]. In most of previous studies, bioreactors provided medium perfusion by
sandwiching cell-laden porous scaffold, while exposing the cardiomyocytes directly to the
flow[522-524]. This does not exactly recapitulate the native myocardium where the blood supply
flows through a dense vascular network that minimizes transport distances but also protects
cardiomyocytes from shear[525]. More recently, bioreactors were developed to provide the
electrical stimulation and medium perfusion simultaneously and it was shown that perfusion and
stimulation had a synergistic effect on improving the contractile functionality of the cardiac
constructs[525, 526]. However, the cardiac constructs in these systems were based on porous
scaffolds and therefore unable to provide the information about the effect of electrical
stimulation on anisotropic cardiac tissue.
Previous studies describe the design of perfusion bioreactors that enable high-throughput in vitro
drug testing on cardiac constructs[524, 527]. Kaneko et al designed a microchamber array chip to
evaluate single cell level interactions for drug testing[527]. Agarwal et al designed a bioreactor
composed of a microarray of cantilevers that was able to characterize diastolic and systolic
stresses generated by anisotropic cardiac microtissue in real-time and the bioreactor could
provide electrical stimulation on these cardiac microtissues[524]. These two studies
characterized cardiac function on either single cell or monolayer level, which might be
insufficient to provide accurate information of cardiac disease as in our complex native system.
Moreover, the drugs investigated in these studies were directly applied to the cells, instead to the
blood compartment, and the presence of flow generated shear stress on cardiomyocytes, both of
which contributed to the generation of an unphysiological environment compared to that
cardiomyocytes experience in the native heart.
There are several advantages of our microfabricated cardiac biowire bioreactors: (1) they are
better mimic of the native cardiac bundle structure with anisotropic alignment; (2) the presence
of the template enables easier handling for later characterization and keeps the entire structure
stable for weeks; (3) the device could be easily customized and applicable for high-throughput
drug screening; (4) the platform provides topographical stimulation by itself; (5) the platform is
versatile and could be integrated with other stimuli as well (e.g. mechanical stimulation); (6) the
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perfusable cardiac biowire system is the first platform to study pharmacological agents applied to
cardiomyocytes by perfusion through cardiac bundle mimic and could provide valuable
knowledge on cardiac disease development and therapeutics.
While this perfusable cardiac biowire platform provides us many opportunities, there are still
some limitations of our current platform. The permeability of the commercially available PTFE
tubing renders limitation on the drug candidates that can be tested, as only small molecules can
diffuse appreciably through the tubing wall and proteins cannot. Ideal tubing materials should be
microporous for better permeability. Further studies are required to investigate other relevant
pharmacological agents and seeding endothelial cells in the tubing lumen to study the interaction
between endothelial cells and cardiomyocytes could also be pursued.
5.5 Conclusion
In conclusion, we have developed a microfabricated cardiac biowire bioreactor that is capable of
testing pharmacological agents applied by perfusion through the lumen. The bioreactor provides
topographical cues for the cardiomyocytes to assemble and align and it could be integrated with
other stimuli to further improve the phenotypes of the cardiomyocytes. The engineered cardiac
biowires could serve as in vitro models that recapitulate the structure and function of the in vivo
cardiac bundles for studies of cardiac development and disease.
5.6 Acknowledgments
The authors would like to thank Zheng Gong for his kind help with preparing Figure 3d. This
work was funded by grants from Ontario Research Fund–Global Leadership Round 2 (ORF-
GL2), Natural Sciences and Engineering Research Council of Canada (NSERC) Strategic Grant
(STPGP 381002-09), Canadian Institutes of Health Research (CIHR) Operating Grant (MOP-
126027), NSERC-CIHR Collaborative Health Research, Grant (CHRPJ 385981-10), NSERC
Discovery Grant (RGPIN 326982-10), NSERC Discovery Accelerator Supplement (RGPAS
396125-10), National Institutes of Health grant 2R01 HL076485 and Heart and Stroke
Foundation grant (T6946).
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Chapter 6
6 Discussion and conclusions
6.1 Discussion
The work reported in this thesis was motivated by the need for an instructive matrix designed
with biochemical and biophysical cues to promote and regulate tissue morphogenesis in situ. The
focus of tissue engineering has evolved from replacing damaged tissues or organs with functional
tissues substitutes generated in vitro into creating an instructive microenvironment to regenerate
the impaired tissue in situ. Compared with generating tissues in vitro, where biochemical cues
can be precisely controlled in term of concentration and treatment time by changing culture
media, instructive matrix design for tissue generation in vivo is challenged by difficult control
over the presentation of biochemical and biophysical cues. To facilitate optimal tissue
morphogenesis, the instructive matrix design heavily relies on our understanding of cell-ECM
interactions and the ability to properly present different cues, both biochemical and biophysical,
to regulate cellular functions. This thesis aims to contribute to this scientific endeavor with a
particular focus on designing instructive matrix with immobilized biochemical cues and
microfabricated topographical guidance.
With ever-growing evidence showing the importance of growth factor regulation in tissue
regeneration, effective delivery of growth factors and growth-factor-derived peptides from
matrix represents an important approach to promote tissue regeneration. Conventional growth
factor delivery methods as soluble supplements are usually poorly controlled with a burst release
that may lead to severe systematic side effects [528]. Moreover, soluble growth factor delivery
often requires repetitive applications, which is not desirable in clinical applications. Covalent
immobilization brings the possibility of controlled release of the biochemical cues and their
prolonged presentation in the matrix. In this thesis, we presented evidence that single application
of collagen-based matrix with covalently immobilized growth factors or short sequence peptides
promoted tissue regeneration, with specific emphases on cardiac tissue engineering and wound
healing applications.
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In Chapter 3, angiogenic growth factors, VEGF and bFGF, and ang-1-derived peptide,
QHREDGS, were covalently immobilized onto collagen scaffolds using EDC chemistry. Our
results demonstrated efficient immobilization and prolonged release of both growth factors and
the peptide. Of note, the porous structure and the tensile strength of the collagen scaffolds were
not altered by the chemical modification. The porous structure of the collagen scaffold is
important for tissue regeneration in situ as it facilitates native cell infiltration and new blood
vessel formation. The mechanical properties, such as the tensile strength, are critical for tissue
engineering applications in a dynamic microenvironment, such as in the heart.
The next study [181] used the collagen scaffold immobilized with angiogenic growth factors to
rejuvenate MSCs from aged human patients. The modified collagen scaffold enhanced MSCs
proliferation in vitro and prolonged cell survival and improved angiogenesis to restore
ventricular morphology and function in vivo. Of note, the improvement was most prominent with
patches seeded with cells from old donors. This novel cytokine-conjugated, sustained-release
system provides a practical and promising platform for cardiac repair in elderly survivors of an
extensive myocardial infarction, an important advance in an increasingly aging society.
In Chapter 4, the pro-survival effect of QHREDGS peptide was demonstrated in human primary
keratinocytes for the first time. The QHREDGS peptide has been shown to promote cell survival
through integrin interaction and there is growing body of literature demonstrating improved
efficacy of integrin ligands presented as immobilized form rather than soluble supplements. This
motivated us to conjugate the peptide onto the backbone of chitosan and then form stable
systems (e.g. films and hydrogels) with collagen. When conjugated within chitosan-only coating,
the immobilized QHREDGS peptide promoted keratinocytes attachment, while the peptide
within chitosan-collagen films promoted cell survival against H2O2 stress, and collective
migration in vitro. We presented results from both normal and diabetic human keratinocytes
showing up-regulation of Akt and MAPKp42/44 phosphorylations.
We further assessed the efficacy of QHREDGS peptide in promoting wound healing in db/db
diabetic mice model that is clinically relevant. When immobilized in a chitosan-collagen
hydrogel system, the QHREDGS peptide accelerated diabetic wound healing in a manner that
was not directly associated with improved local angiogenesis. This study indicated that
QHREDGS peptide holds promise to augment the therapeutic outcomes of current diabetic
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chronic wound therapies, the majority of which focus on addressing impaired angiogenesis.
Together, our data on human primary keratinocytes, both normal and diabetic, and in a db/db
diabetic mouse model demonstrated clinical relevance and we propose QHREDGS peptide as a
therapeutic candidate for promoting diabetic wound healing.
Importantly, both the collagen patch immobilized growth factors and the chitosan-collagen
hydrogel immobilized QHREDGS peptide were applied only once at the beginning of both in
vivo studies. This highlights the contribution of covalent immobilization of biochemical cues in
instructive matrix design for tissue engineering applications where repetitive therapeutic
interventions are sometimes not accessible, such as in the heart. In general, single application is
preferable compared with repetitive applications because of less possibilities to disrupt the
regenerating microenvironment.
Another potential contribution of covalently immobilizing growth factors or peptides is to enable
dynamic presentation of the biochemical cues. The spatial and temporal presentation of different
stimuli is crucial during embryo development and has been extensively investigated and applied
in stem cell differentiations, which indicates their importance in tissue regeneration as well.
However, this was not explored in this thesis. Other studies presenting different biochemical cues
discriminately with spatiotemporal control have shown great promise in various tissue
engineering applications such as bone regeneration [529], new vessel formation [530, 531],
osteochondral tissue engineering [532, 533], muscle regeneration [534], and brain tissue
regeneration [535, 536].
The importance of biophysical cues in stem cell differentiation and tissue morphogenesis has
been increasingly recognized and different biophysical cues including matrix mechanical
properties and topographical guidance have been applied in instructive matrix design [89-91,
537-539]. These challenges in mechanisms and applications of topographical cues became
addressable with emerging material and analytical technologies that bring the possibility to
fabricate and characterize topographical cues with precise control up to nano-scale and increased
rigor and reproducibility [94]. These studies significantly contribute to our understanding of how
cells sense, adapt, and respond to their surrounding microenvironment, which in turn is central to
designing instructive matrix for tissue engineering.
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Moreover, empowered by the advances of microfabrication, a variety of functional micro-tissue
constructs were generated in vitro to recapitulate native tissues including lung [540], liver [541,
542], heart [410, 413, 487, 543], blood vessel [544], kidney [545, 546], intestine [547, 548], and
skin [549, 550]. These engineered micro-tissues and systems composed of multiple tissues [551,
552] have been proposed to be cogent platforms for pre-clinical drug screening studies with
better physiological relevance compared with animal models. The development of these systems
marked the emergence of the field of “Organ-on-a-Chip” or “Human-on-a-Chip” [553-555].
In Chapter 5, we developed a microfabricated cardiac biowire bioreactor that was capable of
testing pharmacological agents applied by perfusion through the lumen in the center of the
engineered cardiac micro-tissues. The bioreactor provided topographical cues for the cardiac
cells to assemble and align and it was integrated with electrical stimulation to further improve the
phenotypes of the cardiomyocytes. The engineered cardiac biowires could serve as in vitro
models that recapitulate the structure and function of the in vivo cardiac bundles for studies of
cardiac development and disease. The next study [413] utilized the original bioreactor with
suture template as platform to investigate the effect of electrical stimulation on maturation of
cardiomyocytes derived from human pluripotent stem cells.
Currently, topographical cues are mainly integrated in 2D substrates in vitro with few studies
demonstrating their promise in tissue regeneration in vivo [556]. Compared with biochemical
cues applied in vitro, the importance of biophysical cues is still underestimated due to the limited
feasibility to present different biophysical cues with temporal control. For example, different
protocols have been intensively investigated for stem cell differentiation induced by biochemical
cues as a result of effortless switching of biochemical cues by changing culture media during the
treatment regimen. Current technologies have enable presenting topographical cues with precise
control up to nano-scale [44], but not the flexibility of switching between different topographical
cues over time without disrupting the cell-matrix interaction yet. Therefore, the dynamic
presentation of topographical cues is a leading challenge yet to be overcome for instructive
matrix designs.
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6.2 Significant contributions
The body of work presented in this thesis addresses different strategies to regulate tissue
remodeling with biochemical or topographical cues. The first two studies were motivated by the
limited therapeutic outcome of existing treatments and the last study was motivated by the
limitation of existing drug-screening platforms at the time I started my study.
The QHREDGS peptide is a novel peptide derived from ang1 and has demonstrated pro-survival
effect on various cell types including cardiac cells, endothelial cells, iPSCs and osteoblasts in
different studies [7-13]. Here we showed covalent immobilization of QHREDGS peptide onto
collagen scaffold by EDC chemistry. Covalent immobilization significantly prolongs the
presence of the QHREDGS peptide in the matrix compared with delivery as a soluble
supplement. Moreover, the chemical modification did not alter the porous structure and
mechanical properties of the collagen scaffolds, which are important for their integration with the
native tissue. This is the first study to immobilize the QHREDGS peptide on collagen scaffold
and it might serve as good candidate for tissue engineering applications including cardiac
regeneration and wound healing.
MSCs derived from bone marrow serve as a good candidate for cell therapy because they can be
easily obtained from autologous tissue and be expanded in vitro rapidly. However, the use of
autologous MSCs is limited in the patients most in need, the elderly, because of their age-related
dysfunctions and impaired regeneration potential [4, 557]. Here, we designed a bioactive
collagen patch with covalently immobilized angiogenic growth factors (VEGF and bFGF) and
demonstrated its ability to rejuvenate MSCs from aged donors for the first time [181]. Seeded on
the collagen patches immobilized with angiogenic growth factors, the MSCs from aged donors
showed increased proliferation and decreased differentiation in vitro and exhibited improved cell
survival and cardiac functional regeneration in vivo after implantation. This study could
significantly contribute to current cell therapies for different tissue engineering applications and
implement their applications to the patients need them the most.
Current approaches to treat diabetic wound ulcers mainly focus on angiogenesis and render
limited therapeutic outcome. Here we presented a novel approach that will provide means to
recapitulate key aspects of scarless embryonic wound healing by 1) promoting effective
115
keratinocyte migration, 2) protecting the wound-bed cells against oxidative stress and 3)
providing a new matrix for cell attachment. We used a chitosan-collagen hydrogel modified with
the QHREDGS peptide and demonstrated that the immobilized QHREDGS peptide promoted
cell survival under hydrogen peroxide stress, collective cell migration of both normal and
diabetic human primary keratinocytes. Importantly, application of a single treatment with a low
dose of QHREDGS-immobilized chitosan-collagen hydrogel accelerated wound closure and
granulation tissue formation in a type 2 diabetic mice model. Importantly, the QHREDGS
peptide can be synthesized at a cost that is 20000 times cheaper than human recombinant growth
factors that have been investigated in a number of previous studies. Together the findings in this
study propose the QHREDGS peptide as a therapeutic candidate for promoting diabetic wound
healing and augmenting current therapies.
Cardiovascular complications are the leading cause of drug withdrawal from the market and
there is a growing interest in developing physiologically relevant models with human cells to be
implemented in drug development [53]. Here we presented perfusable cardiac microtissues
generated in microfabricated bioreactors with suspended template providing topographical
guidance. This is the first platform that is able to test drug candidates by perfusion through the
center of cardiac microtissue and we validated its application by perfusing NO-releasing solution.
This work laid the foundation of the “bio-wire” system [413] and together with following
technologies our lab started TARA Biosystems Inc. to produce physiologically relevant mature
heart tissue that can be interrogated to measure physical and biological factors capable of
accurately predicting cardiotoxicity for pre-clinical applications.
Together, the novel matrix systems designed in this thesis regulate cell-matrix interactions by
providing different biochemical cues or topographical guidance. These matrix systems regulate
tissue regeneration and remodeling in applications including cardiac remodeling, wound healing,
and generating cardiac micro-tissues, and resulted in improved therapeutic outcomes in vivo or
physiological relevance in vitro. The findings from these studies would significantly contribute
to our knowledge of how to design and present biochemical and topographical cues to regulate
cell-matrix interaction and tissue remodeling both in vitro and in vivo.
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6.3 Conclusion
In conclusion, the studies in this thesis demonstrated that cell-matrix interactions can be
facilitated by topographical and biochemical cues provided by surrounding matrix, which
regulate cell assembly, cell functions, and tissue morphogenesis. Specifically, we demonstrated
the feasibility of these instructive cues in both tissue regeneration in vivo and creating complex
micro-tissues in vitro. With immobilized biochemical cues, cardiac and cutaneous tissue
regenerations were promoted by single application of the designed instructive matrix. With
microfabricated topographical guidance, perfusable cardiac micro-tissues were generated with
improved physiological relevance. These findings, in turn, contribute to our knowledge of cell-
matrix interaction, which is critical for instructive biomaterials design. Designing instructive
matrix that promotes tissue morphogenesis in situ would significantly contribute to the transition
of tissue engineering from the laboratory bench to the patient’s bedside.
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Chapter 7
7 Recommendations for future work
7.1 Investigate cardiac regeneration by collagen patch immobilized with QHREDGS peptide
In Chapter 3, we immobilized angiogenic growth factors, VEGF and bFGF, on collagen
scaffolds and applied them in rejuvenating human MSCs from aged patients for cardiac
regeneration after SVR. The growth-factor-modified collagen patches demonstrated clear
efficacy in improving the phenotype of aged MSCs both in vitro and in vivo. We also
immobilized QHREDGS peptide on collagen scaffolds and the peptide has been reported to
promote cardiac cell survival in vitro [8, 9] and cardiac regeneration in vivo [13]. Therefore, it
would be interesting to investigate the QHREDGS-modified collagen patch and compare its
efficacy in promoting cardiac regeneration with QHREDGS-conjugated hydrogel system.
7.2 Determine the mechanism of accelerated keratinocyte collective migration promoted by QHREDGS peptide
In Chapter 4, we presented the results showing that QHREDGS peptide immobilized in
chitosan-collagen substrates promoted keratinocyte collective migration in vitro while the
underlying mechanism remains unknown. With emerging experimental techniques (e.g. laser
ablation) and analytical tools (e.g. individual cell tracking in time-lapse imaging), different
mechanisms have been proposed for collective epithelial cell migration, including local tractions
pulling cooperatively towards unfilled space (termed “kenotaxis”) [455], mechanical exclusion
interactions between cells [456], and intercellular adhesion and tension [457]. We propose using
a high throughput imaging system to acquire time-lapse images of the human primary
keratinocyte collective migration and applying individual cell tracking to look at cell speed (total
travel distance per time), cell velocity (net displacement per time), persistence (velocity per
displacement), and angle of migration of each cell. These parameters will provide insight into the
effects of QHREDGS peptide on cell locomotion, particularly whether the accelerated collective
migration is caused by directed migration (chemotactic) or increased motility in random
directions (chemokinetic).
118
Future studies can compare keratinocyte migration with QHREDGS supplemented in medium or
immobilized in substrate as our results showed no significant acceleration of migration when the
peptide was supplemented as soluble form. Understanding the mechanism on promoting
keratinocyte collective migration would be important for the clinical transition of QHREDGS
peptide.
7.3 Improve the perfusable biowire for drug candidates with high molecular weight
In Chapter 5, we presented the design of perfusable biowire and its capability to deliver
perfused nitric oxide to the surrounding cardiac tissue. However, poor permeability of the PTFE
micro-tubing significantly limited its application in testing other drug candidates with higher
molecular weight, which is particularly relevant for pharmaceutical agents screening. Therefore,
it is necessary to replace the PTFE micro-tubing with other tubular template made of
biocompatible material with better permeability. Ideally, the template material should have nano-
or micro-scale porous structure that enables efficient mass transfer and cell migration that
mimics the native blood vessel walls. Poly(octamethylene maleate (anhydride) citrate) (POMaC)
is a biocompatible material and has been microfabricated into micro-channels in our lab with
interconnected nano-pores and microfabricated micro-pores [558]. Moreover, POMaC material is
well suited for cardiac tissue engineering because it is an elastomer that can be dynamically
stretched with tunable elasticity in the range of adult human myocardium (200-500 kPa).
Therefore, microfabricated POMaC tubing would be a good candidate to replace the PTFE
tubing template in our current design. Drug candidates with high molecular weight and complex
structure, such as growth factors, can then be investigated.
7.4 Investigate the synergy between biochemical cues and topographical cues
In this thesis, our results demonstrated that topographical and biochemical cues, applied
independently in three studies, can regulate cell-matrix interaction to promote tissue regeneration
in vivo or create micro-tissue with increased complexity in vitro. It is therefore interesting to
investigate their synergistic effects and potentially provide topographical and biochemical cues
simultaneously to facilitate cell assembly, cell function and tissue morphogenesis in tissue
119
engineering applications. Specifically, keratinocyte collective migration may be accelerated by
topographical guidance as well. By microfabrication, we can create micro-grooves on the culture
substrate and immobilize QHREDGS peptide afterwards, and then investigate the keratinocyte
collective migration with topographical and biochemical cues together.
Moreover, the cardiac micro-tissues usually risk poor cell survival in the center due to limited
mass transfer, which could be potentially improved by supplementing QHREDGS peptide in the
culture medium or immobilizing QHREDGS peptide in the collagen-based hydrogel. It would be
interesting to investigate the effect of QHREDGS peptide on self-remodeling during the micro-
tissue formation in vitro as well.
120
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Appendices List of publications and contributions
Journal articles:
1. Xiao Y, Radisic M. Instructive matrix design for wound healing applications. In preparation.
2. Xiao Y, Feric N, Knee EJ, Gu J, Cao S, Laschinger CA, Londono C, McGuigan AP, Radisic
M. Diabetic wound regeneration using peptide-modified hydrogels targeting the epithelium.
Submitted to Proceedings of the National Academy of Sciences.
3. Xiao Y, Reis L, Zhao Y, Radisic M. Modifications of biomaterials with immobilized growth
factors or peptides for tissue engineering applications. Methods 2015;84:44-52. DOI:
10.1016/j.ymeth.2015.04.025
4. Davenport Huyer L, Montgomery M, Zhao Y, Xiao Y, Conant G, Korolj A, Radisic M.
Biomaterial based cardiac tissue engineering and its applications. Biomedical Materials
2015;10:034004. DOI: 10.1088/1748-6041/10/3/034004
5. Miklas JW, Nunes SS, Sofla A, Reis L a, Pahnke A, Xiao Y, Laschinger C, Radisic M.
Bioreactor for modulation of cardiac microtissue phenotype by combined static stretch and
electrical stimulation. Biofabrication 2014;6:024113. DOI: 10.1088/1758-5082/6/2/024113
6. Liu H, Wen J, Xiao Y, Liu J, Hopyan S, Radisic M, Simmons C, Sun Y. In situ mechanical
characterization of the cell nucleus by atomic force microscopy. ACS Nano 2014;8:3821–8.
DOI: 10.1021/nn500553z
7. Xiao Y, Zhang B, Liu H, Miklas JW, Gagliardi M, Pahnke AQ, Thavandiran N, Sun Y,
Simmons C, Keller G, Radisic M. Microfabricated perfusable cardiac biowire: a platform that
mimics native cardiac bundle. Lab Chip 2014;14:869–82. (Front cover; Top 10%; Hot
Article) DOI: 10.1039/c3lc51123e
8. Thavandiran N, Nunes SS, Xiao Y, Radisic M. Topological and electrical control of cardiac
differentiation and assembly. Stem Cell Res Ther 2013;4:14. DOI: 10.1186/scrt162
9. Nunes SS, Miklas JW, Liu J, Aschar-Sobbi R, Xiao Y, Zhang B, Jiang J, Masse S, Gagliardi
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M, Hsieh A, Thavandiran N, Laflamme MA, Nanthakumar K, Gross GJ, Backx P, Keller G,
Radisic M. Biowire: a platform for maturation of human pluripotent stem cell–derived
cardiomyocytes. Nat Methods 2013;10:781–7. DOI: 10.1038/nmeth.2524
10. Kang K, Sun L, Xiao Y, Li SH, Wu J, Guo J, Jiang S, Yang L, Yao TM, Weisel RD, Radisic
M, Li RK. Aged human cells rejuvenated by cytokine enhancement of biomaterials for
surgical ventricular restoration. J Am Coll Cardiol 2012;60:2237–49. DOI:
10.1016/j.jacc.2012.08.985
11. Zhang B, Xiao Y, Hsieh A, Thavandiran N, Radisic M. Micro- and nanotechnology in
cardiovascular tissue engineering. Nanotechnology 2011;22:494003. DOI: 10.1088/0957-
4484/22/49/494003
Book chapters:
1. Chiu LLY, Zhang B, Xiao Y, Radisic M. Cardiac Tissue Regeneration in Bioreactors.
Biomaterials and Regenerative Medicine, Cambridge University Press; 2014, p. 640-668.
DOI: http://dx.doi.org/10.1017/CBO9780511997839.042
2. Xiao Y, Zhang B, Hsieh A, Thavandiran N, Martin C, Radisic M. Microfluidic Cell Culture
Techniques. Microfluidic Cell Culture Systems, Elsevier; 2013, p. 303–21. DOI:
10.1016/B978-1-4377-3459-1.00012-0
Oral presentations:
1. Xiao Y, Zhang B, Liu H, Sun Y, Simmons C, Radisic M. Microfabricated Perfusable
Cardiac Biowire: A Platform That Mimics Native Cardiac Bundle. Tissue Engineering and
Regenerative Medicine International Society (TERMIS) AM Annual Conference,
Washington, D.C. Dec. 13-16, 2014.
2. Xiao Y, Zhang B, Liu H, Sun Y, Simmons C, Radisic M. Developing cardiac biofibers with
microfabricated devices. MATCH/Ontario-On-A-Chip Symposium 2013, Toronto, Canada,
May 23-24, 2013
163
Poster presentations:
1. Xiao Y, Radisic M. Chitosan-Collagen Hydrogel Modified with QHREDGS Peptide for
Wound Healing. 32nd Annual Meeting of the Canadian Biomaterials Society, Toronto,
Canada, May 28 - 30, 2015
2. Xiao Y, Radisic M. Chitosan-Collagen Hydrogel Modified with QHREDGS Peptide for
Wound Healing. Tissue Engineering and Regenerative Medicine International Society
(TERMIS) AM Annual Conference, Washington, D.C. Dec. 13-16, 2014.
3. Xiao Y, Zhang B, Liu H, Sun Y, Simmons C, Radisic M. Microfabricated Perfusable
Cardiac Biowire: A Platform That Mimics Native Cardiac Bundle. MATCH/Ontario-On-A-
Chip Symposium 2014, Toronto, Canada, May 29-30, 2014.
4. Xiao Y, Zhang B, Liu H, Sun Y, Simmons C, Radisic M. Microfabricated Perfusable
Cardiac Biowire: A Platform That Mimics Native Cardiac Bundle. 30th Annual Meeting of
the Canadian Biomaterials Society, Ottawa, Canada, May 29 - Jun 1, 2013.
5. Xiao Y, Zhang B, Liu H, Sun Y, Simmons C, Radisic M. Developing cardiac biofibers with
microfabricated devices. Launch of Boundless Campaign for Faculty of Applied Science &
Engineering, University of Toronto, Sept 18, 2012.
6. Xiao Y, Zhang B, Liu H, Sun Y, Simmons C, Radisic M. Developing cardiac biofibers with
microfabricated devices. 9th World Biomaterials Congress, Chengdu, China, Jun 1-5, 2012.
7. Xiao Y, Zhang B, Liu H, Sun Y, Simmons C, Radisic M. Developing cardiac biofibers with
microfabricated devices. MATCH/Ontario-On-A-Chip Symposium 2012, Toronto, Canada,
May 17-18, 2012.
8. Xiao Y, Radisic M. Engineering Cardiac Purkinje Fibers with Microfabricated Devices.
2011 Annual University of Toronto Institute of Biomaterials & Biomedical Engineering
Scientific Day, Toronto, Canada, May 19, 2011.
164
Other conference proceedings:
1. Nunes SS, Miklas JW, Xiao Y, Zhang B, Radisic M. Stem cell-derived cardiomyocyte
maturation by biomimetic topographical and electrical cues. North American Vascular
Biology Organization (NAVBO), Hyannis, Massachusetts, Oct. 20-24, 2013.
2. Sun L, Kang K, Xiao Y, Li SH, Wu J, Guo J, Jiang S, Yang L, Yao TM, Weisel RD, Radisic
M, Li RK. Aged human cells rejuvenated by cytokine enhancement of biomaterials for
surgical ventricular restoration. American Heart Association 2012 Scientific Sessions, Los
Angeles, California, Nov. 3-7, 2012.
3. Xiao Y, Thavandiran N, Au H, Radisic M. Microbioreactors for cardiac tissue engineering.
Tissue Engineering and Regenerative Medicine International Society (TERMIS) EU Annual
Conference, Granada, Spain, Jun. 7-10, 2011.
165
Copyright Acknowledgements
Copyright © 2012 Elsevier. Contents of this chapter have been published in J Am Coll
Cardiol: Kang K, Sun L, Xiao Y, Li SH, Wu J, Yao TM, Weisel RD, Radisic M, Li RK. Aged
human cells rejuvenated by cytokine enhancement of biomaterials for surgical ventricular
restoration. J Am Coll Cardiol. 2012;60:2237–2249. Reuse with permission from Elsevier. A
link to the published paper can be found at:
http://www.sciencedirect.com/science/article/pii/S0735109712043677
Copyright © 2013 BioMed Central. Contents of this thesis have been published in Stem Cell
Res Ther: Thavandiran N, Nunes SS, Xiao Y, Radisic M. Topological and electrical control of
cardiac differentiation and assembly. Stem Cell Res Ther. 2013;4:14. Reuse with permission
from BioMed Central. A link to the published paper can be found at:
http://www.stemcellres.com/content/4/1/14
Copyright © 2014 Cambridge University Press. Contents of this thesis have been published in:
Chiu LLY, Zhang B, Xiao Y, Radisic M. Cardiac tissue regeneration in bioreactors. Biomaterials
and Regenerative Medicine, Cambridge University Press. 2014;640-668. Reuse with permission
from Cambridge University Press. A link to the published chapter can be found at:
http://ebooks.cambridge.org/chapter.jsf?bid=CBO9780511997839&cid=CBO9780511997839A0
46
Copyright © 2014 Royal Society of Chemistry. Contents of this chapter have been published
in Lab Chip: Xiao Y, Zhang B, Liu H, Miklas JW, Gagliardi M, Pahnke A, Thavandiran N, Sun
Y, Simmons C, Keller G, Radisic M. Microfabricated perfusable cardiac biowire: a platform that
mimics native cardiac bundle. Lab Chip. 2014; 14:869–82. Reuse with permission from Royal
Society of Chemistry. A link to the published paper can be found at:
http://pubs.rsc.org/en/content/articlelanding/2014/lc/c3lc51123e
Copyright © 2015 Elsevier. Contents of this thesis have been published in Methods: Xiao Y,
Reis LA, Zhao Y, Radisic M. Modifications of collagen-based biomaterials with immobilized
growth factors or peptides. Methods. 2015;84:44–52. Reuse with permission from Elsevier. A
166
link to the published paper can be found at:
http://www.sciencedirect.com/science/article/pii/S1046202315001723
Copyright © 2015 IOP Publishing. Contents of this chapter have been published in
Biomedical Materials: Davenport Huyer L, Montgomery M, Zhao Y, Xiao Y, Conant G, Korolj
A, Radisic M. Biomedical Materials. 2015;10:034004. Reuse with permission from IOP
Publishing. A link to the published chapter can be found at:
http://iopscience.iop.org/article/10.1088/1748-6041/10/3/034004
