Modular Approach to Adipose Tissue Engineering...ii Modular Approach to Adipose Tissue Engineering...
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Modular Approach to Adipose Tissue Engineering by Mark James Butler A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Graduate Department of Chemical Engineering and Applied Chemistry & Institute of Biomaterials and Biomedical Engineering University of Toronto © Copyright by Mark James Butler 2011
Transcript of Modular Approach to Adipose Tissue Engineering...ii Modular Approach to Adipose Tissue Engineering...
Modular Approach to Adipose Tissue Engineering
by
Mark James Butler
A thesis submitted in conformity with the requirements
for the degree of Doctor of Philosophy
Graduate Department of Chemical Engineering and Applied Chemistry &
Institute of Biomaterials and Biomedical Engineering
University of Toronto
© Copyright by Mark James Butler 2011
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Modular Approach to Adipose Tissue Engineering
Mark James Butler Doctor of Philosophy
Department of Chemical Engineering and Applied Chemistry University of Toronto, 2011
Abstract Despite the increasing clinical demand in reconstructive, cosmetic and correctional surgery there
remains no optimal strategy for the regeneration or replacement of adipose tissue. Previous
approaches to adipose tissue engineering have failed to create an adipose tissue depot that
maintains implant volume in vivo long-term (>3 months). This is due to inadequate mechanical
properties of the biomaterial and insufficient vascularization upon implantation. Modular tissue
engineering is a means to produce large volume functional tissues from small sub-mm sized
tissues with an intrinsic vascularization. We first explored the potential of a semi-synthetic
collagen/poloxamine hydrogel with improved mechanical properties to be used as the module
biomaterial. We found this biomaterial to not be suitable for adipose tissue engineering because
it did not support embedded adipose-derived stem cell (ASC) viability, differentiation and
human microvascular endothelial cell (HMEC) attachment. ASC-embedded collagen gel
modules coated with HMEC were then implanted subcutaneously in SCID mice to study its
revascularization potential. ASC cotransplantation was shown to drive HMEC vascularization in
vivo: HMEC were seen to detach from the surface of the modules to form vessels containing
erythrocytes as early as day 3; vessels decreased in number but increased in size over 14 days;
and persisted for up to 3 months. Early vascularization promoted fat development. Only in the
case of ASC-HMEC cotransplantation was progressive fat accumulation observed in the module
implants. Although implant volume was not maintained, likely due rapid collagen degradation,
the key result here is that ASC-HMEC cotransplantation in the modular approach was successful
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in creating vascularized adipose tissue in vivo that persisted for 3 months. The modular system
was then studied in vitro to further understand ASC-EC interaction. Coculture with ASC was
shown to promote an angiogenic phenotype (e.g. sprouting, migration) from HUVEC on
modules. RT-PCR analysis revealed that VEGF, PAI-1 and TNFα was involved in ASC-EC
paracrine signalling. In summary, ASC-HMEC cotransplantation in modules was effective in
rapidly forming a vascular network that supported fat development. Future work should focus
on further elucidating ASC-EC interactions and developing a suitable biomaterial to improve
adipose tissue development and volume maintenance of engineered constructs.
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Acknowledgements
I first would like to thank Dr. Michael Sefton for his help and guidance through the years.
I have been a student in his lab from the beginning of my research experiences in my 4th year
undergraduate project. I remember waiting for our very first meeting outside his office trying to
digest my very first journal publication, thinking it was written in a language I did not
understand. I have come a long way over the years learning new terms, concepts, techniques,
and how to develop and test hypotheses. From Dr. Sefton I have learned two things: First,
effective research must tell a story and second, learning is a life long adventure that does not end
with the awarding of a PhD.
I thank all members of the Sefton lab, past and present, that contributed to my life both
academically and personally over the years. Thanks to Omar Khan, Brendan Leung, Lindsay
Fitzpatrick, Rohini Gupta, Dean Chamberlain, Ema Ciucurel, and Derek Voice for the many
brainstorming sessions, help with experimental work, research discussions and the times outside
of the lab that made graduate school so enjoyable. A special thanks to Chuen Lo for his
expertise and help with animal work.
I would like to thank all other collaborators on this project: Kim Woodhouse for her help
and guidance as a cosupervisor; Yu-Ling Cheng for being part of the reading committee; and
Alison McGuigan for her help early on in my research career as a 4th year thesis supervisor and
her mentorship over the years. A special thanks to Dr. John Semple and his colleague Dr. Bell
for the adipose tissue samples. Without your help this work would not have been possible.
Finally, I thank my family, my parents, Jim and Barb, and sisters, Sarah and Glenda for
being there for me and encouraging me to do my best every step of the way. Thank you to
Nicole and Kostas for listening to some of my crazy ideas and providing the needed distraction
from work. If you’ve made it this far, thanks to you for reading my thesis. It was a long journey
with some ups and downs, but such a worthwhile experience.
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Table of Contents
Abstract .....................................................................................................................................................ii
Acknowledgements..............................................................................................................................iv
Table of Contents ................................................................................................................................... v
List of Figures.........................................................................................................................................ix
List of Tables ........................................................................................................................................ xii
Abbreviations...................................................................................................................................... xiii
1 Introduction, Hypothesis and Experimental Plan.............................................................1
1.1 Clinical Impetus .............................................................................................................. 1
1.2 Modular Tissue Engineering........................................................................................... 2
1.3 Objectives and Hypothesis.............................................................................................. 4
1.4 Thesis Organization ........................................................................................................ 5
2 Adipose Tissue Engineering & Adipose‐derived Stem Cell/Endothelial Cell
Cotransplantation as a Revascularization Strategy...................................................................6
2.1 Adipose-derived Stem Cells and Soft Tissue Engineering............................................. 6
2.1.1 Adipose Tissue........................................................................................................ 6
2.1.2 Adipocyte Differentiation ....................................................................................... 8
2.1.3 Adipose Tissue Extracellular Matrix .................................................................... 12
2.1.4 Stem Cells for Adipose Tissue Engineering ......................................................... 13
2.1.5 Adipose Tissue Engineering Strategies ................................................................ 14
2.1.6 Semi-Synthetic Collagen/Poloxamine Hydrogel .................................................. 16
2.2 Angiogenesis & Revascularization Strategies .............................................................. 18
2.2.1 Mechanisms of Angiogenesis ............................................................................... 18
2.2.1.1 Vascular Functions in Adipose Tissues ............................................................ 21
2.2.1.2 Angiogenic Modulators .................................................................................... 22
2.2.1.3 Relationship between ASC and EC .................................................................. 24
2.2.2 Revascularization Strategies ................................................................................. 26
2.2.2.1 Growth Factor Delivery .................................................................................... 27
2.2.2.2 Biomaterial-mediated Angiogenesis................................................................. 28
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2.2.2.3 Vascular Pedicle................................................................................................ 29
2.2.2.4 Cell-Based Revascularization Strategies .......................................................... 29
3 Semi‐Synthetic Collagen/Poloxamine Hydrogel as a Potential Biomaterial for
Adipose Tissue Engineering............................................................................................................ 33
3.1 Abstract ......................................................................................................................... 33
3.2 Introduction................................................................................................................... 33
3.3 Materials and Methods.................................................................................................. 36
3.3.1 Cells ...................................................................................................................... 36
3.3.2 Hydrogel Synthesis ............................................................................................... 37
3.3.3 Cell Viability......................................................................................................... 38
3.3.4 Hydrogel Characterization .................................................................................... 39
3.3.5 Oil Red O .............................................................................................................. 39
3.3.6 Glycerol-3-phosphate Dehydrogenase Activity.................................................... 40
3.3.7 EC Attachment to Gels and Modules.................................................................... 40
3.3.8 Implantation of Collagen/Poloxamine Modules into SCID mice ......................... 41
3.3.9 Statistical Analysis................................................................................................ 42
3.4 Results........................................................................................................................... 43
3.4.1 ASC Viability........................................................................................................ 43
3.4.1.1 Addition of Laminin to Improve ASC Viability............................................... 46
3.4.2 Hydrogel Characterization .................................................................................... 47
3.4.3 ASC Differentiation .............................................................................................. 49
3.4.3.1 Oil Red O .......................................................................................................... 49
3.4.3.2 GPDH Activity.................................................................................................. 50
3.4.4 Endothelial Cell Attachment to Gels and Modules............................................... 52
3.4.5 Surface Modifications to Improve Cell Attachment............................................. 54
3.4.6 In Vivo Implantation............................................................................................. 56
3.5 Discussion..................................................................................................................... 58
3.6 Conclusions................................................................................................................... 61
3.7 Acknowledgements....................................................................................................... 61
4 Cotransplantation of ASC and HMEC in a Modular Construct Drives Early
Angiogenesis, ASC differentiation and Long‐term Fat Regeneration in SCID Mice ..... 62
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4.1 Abstract ......................................................................................................................... 62
4.2 Introduction................................................................................................................... 63
4.3 Materials and Methods.................................................................................................. 64
4.3.1 Cells ...................................................................................................................... 64
4.3.2 Module Fabrication............................................................................................... 65
4.3.3 Module Implants ................................................................................................... 66
4.3.4 Histology Quantification....................................................................................... 67
4.3.5 Perfusion Studies .................................................................................................. 68
4.3.6 Volume Measurements ......................................................................................... 68
4.3.7 Statistical Analysis................................................................................................ 68
4.4 Results........................................................................................................................... 69
4.4.1 Angiogenesis & Remodeling Study (Day 3, 7, 14, 21) ........................................ 69
4.4.1.1 Appearance at Explant ...................................................................................... 69
4.4.1.2 Effect of ASC on HMEC Viability and Blood Vessel Formation .................... 70
4.4.1.3 Host Tissue Infiltration and Angiogenesis........................................................ 75
4.4.2 Implants with HUVEC.......................................................................................... 77
4.4.3 Fat Development and Volume Maintenance Study (Day 30, 60, 90)................... 79
4.4.3.1 Fat Development............................................................................................... 79
4.4.3.2 Angiogenesis and HMEC Viability .................................................................. 83
4.4.3.3 Volume Measurements ..................................................................................... 85
4.5 Discussion..................................................................................................................... 86
4.6 Conclusions................................................................................................................... 89
4.7 Acknowledgements....................................................................................................... 89
5 Investigating the Relationship between Adipose‐dervied Stem Cells and
Endothelial Cells In Vitro.................................................................................................................. 90
5.1 Abstract ......................................................................................................................... 90
5.2 Introduction................................................................................................................... 91
5.2.1 Cells ...................................................................................................................... 92
5.2.2 Optimized In Vitro Angiogenesis Sprouting Assay.............................................. 92
5.2.3 Sprouting Counts and Measurements ................................................................... 94
5.2.4 RNA Isolation ....................................................................................................... 95
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5.2.5 cDNA Synthesis.................................................................................................... 96
5.2.6 Real-time RT PCR ................................................................................................ 97
5.2.7 Statistical Analysis.............................................................................................. 100
5.3 Results......................................................................................................................... 101
5.3.1 Effect of ASC on HUVEC Sprouting from Modules in a Fibrin Gel................. 101
5.3.2 RT-PCR Analysis ............................................................................................... 106
5.4 Discussion................................................................................................................... 111
5.5 Conclusions................................................................................................................. 114
5.6 Acknowledgements..................................................................................................... 115
6 Synopsis.......................................................................................................................................116
6.1 Biomaterials for Adipose Tissue Engineering ............................................................ 117
6.2 Revascularization Strategies ....................................................................................... 119
6.3 Understanding the relationship between ASC and EC ............................................... 122
6.4 Future Directions ........................................................................................................ 124
7 Conclusions ................................................................................................................................128
8 Reference List............................................................................................................................130
9 Appendix 1: HMEC Harvesting............................................................................................143
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List of Figures
Figure 1-1: Modular approach to adipose tissue engineering....................................................... 3
Figure 2-1: Major phases in the development of mature adipocytes from multipotent stem cells
found in the stromal-vascular fraction of adipose tissue. ............................................................... 9
Figure 2-2: Poloxamine is modified with methacrylate groups (IEM) that can undergo free
radical polymerization. ................................................................................................................. 17
Figure 2-3: Mechanisms of angiogenesis. .................................................................................. 19
Figure 2-4: Functions of adipose tissue vasculature................................................................... 22
Figure 2-5: Modulators of angiogenesis in adipose tissue.......................................................... 23
Figure 3-1: Alamar Blue® viability of ASC embedded in collagen (control), poloxamine-only
and collagen/poloxamine hydrogels over 7 days .......................................................................... 44
Figure 3-2: Live/Dead™ confocal microscopy images of ASC embedded in collagen and
collagen/poloxamine hydrogels. ................................................................................................... 45
Figure 3-3: Live/Dead™ confocal microscopy images of ASC embedded collagen/poloxamine
hydrogels with the addition of laminin. ........................................................................................ 47
Figure 3-4: Immunostaining of Collagen and Collagen/Poloxamine/Laminin hydrogels.......... 48
Figure 3-5: High magnification images of immunostained collagen/poloxamine hydrogels..... 49
Figure 3-6: ASC differentiation in collagen, poloxamine and collagen/poloxamine hydrogels 50
Figure 3-7: GPDH activity of ASC during differentiation. ........................................................ 51
Figure 3-8: GPDH activity of ASC during differentiation normalized for GAPDH expression.52
Figure 3-9: HMEC-1 attachment to poloxamine hydrogel films................................................ 53
Figure 3-10: HMEC-1 attachment to poloxamine modules........................................................ 54
Figure 3-11: HMEC-1 attachment to modules with fibronectin treatment................................. 55
Figure 3-12: HMEC-1 attachment to modules with collagen coating. ....................................... 56
Figure 3-13: Collagen/Poloxamine modules implanted subcutaneously in SCID mice............. 57
Figure 4-1: Photographs of the modules at explant day 7 and 14............................................... 70
Figure 4-2: UEA-1 Lectin stained sections of HMEC+ASC modules and HMEC-only modules
at day 3, 7 and 14. ......................................................................................................................... 71
Figure 4-3: Magnified images of serial sections of the tissue surrounding a single module at day
3..................................................................................................................................................... 73
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Figure 4-4: Magnified images of serial sections of a single blood vessel surrounding a module
at day 3.......................................................................................................................................... 74
Figure 4-5: MVD counts from HMEC+ASC implants at day 3, 7 and 14. ................................ 75
Figure 4-6: CD31 images of host tissue infiltration into the modular implant at day 14. .......... 76
Figure 4-7: Magnified images of serial sections of tissue surrounding several modules. .......... 76
Figure 4-8: MicroCT images of HMEC+ASC (A) and HMEC-only (B) module implants under
mouse skin at day 21..................................................................................................................... 77
Figure 4-9: UEA-1 Lectin stained sections of HUVEC+ASC modules and HUVEC-only
modules at day 3, and 14............................................................................................................... 78
Figure 4-10: Oil Red O images of frozen sections from HMEC+ASC and ASC-only module
implants at 30, 60 and 90d post implantation. .............................................................................. 80
Figure 4-11: H&E and Masson’s Trichome micrographs of HMEC+ASC modules at day 60. 81
Figure 4-12: Masson’s Trichrome micrograph of HMEC+ASC modules at day 90.................. 81
Figure 4-13: Percentage of area stained red for Oil Red O on micrograph sections of
HMEC+ASC and ASC-only module implants at day 30, 60 and 90............................................ 82
Figure 4-14: H&E and Masson’s Trichrome micrographs of HMEC+ASC and ASC-only
modules 30 days after implantation. ............................................................................................. 84
Figure 4-15: UEA-1 stained sections of HMEC+ASC modules 30 days and 90 days after
implantation. ................................................................................................................................. 85
Figure 4-16: Volume measurements of HMEC+ASC and ASC-only module implants. ........... 86
Figure 5-1: Experimental set up of the in vitro angiogenesis assay. .......................................... 94
Figure 5-2: Image of capillary like-formations growing out from the surface of a HUVEC-
coated module in fibrin gel with the presence of ASC in coculture at day 3. .............................. 95
Figure 5-3: Photographs of HUVEC sprouting from modules in a fibrin gel with and without
ASC coculture............................................................................................................................. 103
Figure 5-4: Magnified image of HUVEC sprouting from modules with ASC coculture at day
14................................................................................................................................................. 104
Figure 5-5: Average number of HUVEC sprouts from the module surface in fibrin gel with and
without the presence of ASC in coculture. ................................................................................. 105
Figure 5-6: Average length of HUVEC sprouts from the module surface in fibrin gel with and
without the presence of ASC in coculture. ................................................................................. 106
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Figure 5-7: Relative fold change in mRNA expression of adipogenic genes of ASC in coculture
with HUVEC, vs. ASC cultured alone........................................................................................ 108
Figure 5-8: Relative fold change in mRNA expression of angiogenic genes of ASC in coculture
with HUVEC, vs. ASC cultured alone........................................................................................ 109
Figure 5-9: Relative fold change in mRNA expression of HUVEC in coculture with ASC, vs.
HUVEC cultured alone. .............................................................................................................. 110
Figure 9-1: Light microscopy images of cells obtained from HMEC separation trials............ 145
Figure 9-2: Cytospin slides of cells from Trial 6 and 7 immunostained with vWF ................. 146
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List of Tables
Table 1: Histology stains. ........................................................................................................... 66
Table 2: RT-PCR Primer List ..................................................................................................... 99
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Abbreviations
ASC: Adipose-derived Stem Cells
bFGF: Basic Fibroblast Growth Factor
BM-MSC: Bone Marrow-derived Mesenchymal Stem Cells
C/EBPα: CCAAT/enhancer binding protein α
CD31: Cluster of differentiation molecule 31, PECAM
DHAP: Dihydroxyacetone phosphate
EC: Endothelial Cells
ECM: Extracellular Matrix
GPDH: Glycerol-3-Phosphate Dehydrogenase
H&E: Hematoxylin and Eosin
HMEC: Human Microvascular Endothelial Cells
HMEC-1: Human Microvascular Endothelial Cell line 1
HUVEC: Human Umbilical Vein Endothelial Cells
MMP: Matrix MetalloProteinases
MSC: Mesenchymal Stem Cells
MVD: MicroVessel Density
NADH: Nicotinamide adenine dinucleotide
PAI-1: Plasminogen Activator Inhibitor 1
PDGF: Platelet Derived Growth Factor
PECAM: Platelet-Endothelial Adhesion Molecule
PPARγ: Peroxisome Proliferator-Activated Receptor γ
SCID: Severe Combined Immunodeficiency
SD: Standard Deviation
SMA: α-Smooth Muscle Actin
SMC: Smooth Muscle Cells
SVF: Stromal-Vascular Fraction
TNFα: Tumor Necrosis Factor
TUNEL: Terminal deoxynucleotidyl Transferase Biotin-dUTP Nick End Labeling
UEA-1: Ulex Europaeus Agglutinin I
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VEGF: Vascular Endothelial Growth Factor
vWf: Von Willebrand Factor
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1 Introduction, Hypothesis and Experimental Plan
1.1 Clinical Impetus
There is a critical need for soft tissue reconstruction for patients suffering from cancer,
trauma, deep burns and congenital defects. These patients suffer from contour defects due to
loss of soft tissue, mainly comprised of subcutaneous adipose tissue. In particular, the surgical
treatment of breast cancer involves lumpectomy or mastectomy which leads to abnormal
cosmesis affecting the emotional well-being of patients and may also impair function (eg. range
of motion of arms). The American Society of Plastic Surgeons cites that over 5.2 million
reconstructive procedures were performed in 2009. Yet, despite the increasing clinical demand
in reconstructive, cosmetic and correctional surgery, there remains no optimal strategy for the
regeneration and replacement of adipose tissue. The use of autologous adipose tissues for soft
tissue replacement is one clinical approach that has proven problematic due to absorption,
subsequent volume loss and fibrosis of the transplanted tissue[1,2]. Synthetic materials such as
saline and silicone implants are widely used to replace soft tissue with various levels of clinical
success. These materials have certain drawbacks such as rupture, leakage, dislocation and
biocompatibility issues. Recently, tissue engineering strategies have been applied to soft tissue
regeneration and have the potential to overcome many of the deficiencies with autologous grafts
and synthetic implants. In this approach, adipose tissue is reconstructed by combining adipose-
derived stem cells (ASC) with an appropriate substrate, typically a polymeric scaffold. Seeding
of adipose precursor cells, onto synthetic and biodegradable scaffolds has led to adipose tissue
formation in vivo. Yet, these approaches have failed to maintain a volume-stable fat pad in vivo
over the long term due to lack of rapid vascularization upon implantation leading to ischemia and
transplanted cell death. Hence, a critical challenge in engineering large portions of adipose
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tissue is early vascularization of the engineered tissue upon implantation to avoid considerable
cell death. Diffusion alone can only supply cells with oxygen and nutrients at a distance of
150μm from a blood supply[3], which would be insufficient for a three-dimensional construct of
clinical relevance. Therefore, volume loss of adipose tissue can only be avoided through early
vascularization of the engineered tissue.
1.2 Modular Tissue Engineering
Modular tissue engineering, as developed by our group, has the potential to shift the
traditional cell + scaffold paradigm because of its inherent uniformity, scaleability and most
importantly for adipose tissue engineering, vascularity. In this approach, functional cells (here,
human adipose-derived stem cells (ASC)) are embedded in small hydrogel rods (0.5mm
diameter, 1mm length) coated with a human microvascular endothelial cell layer (HMEC).
Many modules (hundreds or thousands) can be implanted subcutaneously at the site of a soft
tissue defect to restore volume loss (Figure 1-1). The interstitial spaces created from random
packing of the modules results in endothelialized channels that remodel and connect to the host
vasculature upon implantation. With rapid vascularization of the construct, the embedded ASC
have the potential to remain viable and differentiate into mature adipocytes, resulting in the
creation of a fat pad over time.
Bovine collagen type I gel used thus far as the module material is mechanically weak and
undergoes contraction with volume reductions of up to 90% over time, perhaps making it
unsuitable as a scaffold for a volume persistent culture of adipose tissue in vivo longterm. In
order to address the mechanical limitations presented with collagen gels, semi-synthetic
collagen-containing networks were produced by photopolymerization of a Tetronic- T1107
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(poloxamine) aqueous solution. These matrices demonstrated increased stiffness while
maintaining the advantageous cell-adhesive property of collagen.
Figure 1-1: Modular approach to adipose tissue engineering.
Each module contains embedded functional cells (ASC) and is surface coated with an human microvascular endothelial cell (HMEC) layer. Many modules implanted subcutaneously at the site of a soft tissue defect can restore normal cosmesis.
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1.3 Objectives and Hypothesis
Premise:
Modular tissue engineering is a viable platform for producing vascularized adipose tissue
in vivo.
Objectives:
1. Fabricate and optimize the modular platform for adipose tissue engineering to promote
embedded ASC viability, differentiation and attachment of surface coated endothelial cells
(EC).
Hypothesis: The semi-synthetic collagen/poloxamine hydrogel used as the module material
promotes high viability of embedded ASC and allows differentiation of these cells into mature
adipocytes, while enabling good adhesion and coverage of EC on the module surface.
2. Evaluate the regenerative potential of the modular construct to produce vascularized adipose
tissue in vivo.
Hypothesis: The endothelialized perfusion channels created by packing many modules into a
larger construct provide an immature capillary network that remodels and anastomoses with the
host vasculature upon implantation to promote transplanted ASC viability and facilitates a long-
term (> 3months) volume-persistent culture of adipose tissue in vivo.
3. Investigate ASC/EC interaction in an in vitro sprouting assay.
Hypothesis: Coculture of ASC and HUVEC (no direct cell-cell contact) promotes an angiogenic
phenotype (i.e. migration and sprouting) of surface covered HUVEC in a fibrin gel.
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1.4 Thesis Organization
I describe the utility of the modular approach for adipose tissue engineering. Chapter 2 is
a literature review that highlights current strategies to engineer adipose tissue, as well as
approaches used to vascularize tissue engineering constructs. In chapter 3 I examine ASC cell
interaction with the semi-synthetic collagen/poloxamine hydrogel as the module material [ie. will
ASC survive and differentiate into mature adipocytes in this extracellular environment? What
modifications are necessary to the collagen/poloxamine hydrogel to make it suitable for adipose
tissue engineering?]. In chapter 4, I investigate the regenerative potential of the modular
platform for adipose tissue in vivo. [ie. will the modular approach facilitate a long-term (>3
months) volume-persistent culture of vascularized adipose tissue in vivo?]. In chapter 5, I begin
to study the relationship between ASC and EC in in vitro coculture [ie. will the coculture of ASC
and EC promote EC migration and sprouting in vitro? What key angiogenic and adipogenic
genes are influenced by ASC/EC coculture?]. Chapter 6 summarizes the work with a synopsis
and gives recommendations for future work. Chapter 7 states the overall conclusions of this
thesis.
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2 Adipose Tissue Engineering & Adipose‐derived Stem Cell/Endothelial Cell Cotransplantation as a Revascularization Strategy
2.1 Adipose‐derived Stem Cells and Soft Tissue Engineering
Adipose-derived stem cells (ASC) are found in the stromal vascular fraction (SVF) of
adipose tissues. These adult stromal cells are similar in phenotype and multipotency to bone
marrow derived mesenchymal stem cells (BM-MSC) and have gained much interest in recent
years because of the near limitless supply of adipose tissue and the relative ease with which it
can be harvested via liposuction or abdominoplasty procedures. Traditional soft tissue
engineering approaches combine ASC and a suitable polymeric scaffold. Once implanted in the
body, the ASC are expected to proliferate and differentiate into adipocytes, while the scaffold
remodels and degrades resulting in a mature fat pad that retains the volume needed to correct the
soft tissue defect.
Adipose-derived stem cell (ASC) is a term that has been proposed to include the entire
population of adipose precursors, ie. both non-committed and committed (preadipocyte) stem
cells found in the stromal-vascular fraction of adipose tissue. For the purposes of this work, the
terms ASC and preadipocyte are used interchangeably.
2.1.1 Adipose Tissue
Adipose tissue is a highly specialized connective tissue that is found throughout the
human body. It can be classified into two forms: white adipose tissue (WAT) and brown adipose
tissue (BAT). While both of these forms serve to insulate and cushion the body, they have
individualized functions as well. BAT is so named because of the colour due to its highly
vascular nature and it functions primarily as a heat source for the body. Since BAT is primarily
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only found in neonates and hibernating animals, WAT is the type of fat of interest in the
development of a tissue-engineered adipose substitute for reconstructive purposes.
The primary cellular component of WAT is a collection of lipid filled cells known as
adipocytes held in place by a loose connective tissue comprising mainly collagen, laminin and
fibronectin. This extracellular matrix (ECM) interconnects adipocytes where they exists in
clusters or fat lobules in the adipose tissue. Adipocytes are rounded cells, characterized by a
single large lipid vacuole, comprising approximately 90% of the cell volume, and a flattened
eccentric nucleus[4]. Other cellular components found in adipose tissue are the stromal-vascular
cells which include endothelial cells, smooth muscle cells, fibroblasts, blood cells and adipose
progenitor cells or preadipocytes[5].
While WAT has long been recognized as an inert mass with roles in maintenance of body
contours, insulation and mechanical protection, it is now recognized as a secretory organ that
functions in energy storage and balance under tight hormonal control. WAT acts as the major
energy reserve by storing triacylglycerol in periods of energy excess and mobilization during
energy deprivation. A much more complex and dynamic role of WAT has emerged recently
with realization that adipocytes secrete factors known to play a role in immunological responses,
vascular diseases and appetite regulation[6]. Leptin, for example, is a hormone that is secreted
by mature adipocytes that interacts with the hypothalamus to control satiety. When body fat
decreases the levels of leptin also decrease which in turn leads to increase in appetite and food
intake. Conversely, when the levels of body fat increase, levels of leptin also increase which
suppresses appetite. Other hormones secreted by adipocytes include immune system related
proteins such as adipsin, acylation stimulation protein (ASP), adipocyte complement-related
protein (Acrp30/AdipoQ), macrophage inhibitory factor (MIF) and tumor necrosis factor-α
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(TNFα), and vascular function related proteins such as angiotensinogen and plasminogen
activator inhibitor type I (PAI-1). The immune system related proteins may be involved in
energy homeostasis and insulin resistance. TNFα, for example, has been shown to inhibit
differentiation of adipose precursors into adipocytes and may decrease the expression of markers
of adipogenesis[7]. TNFα may also contribute to insulin resistance by inhibition of the insulin
receptor[8]. It has been shown to be present at elevated levels in obese individuals and may act
upon adipocytes to increase secretion of the proinflammatory cytokine MIF[9]. PAI-1 is
secreted by adipose tissue and higher levels are observed in omental fat compared with
subcutaneous fat. This may be involved in vascular diseases related to an increase in abdominal
adipose tissue in obesity[10]. It is clear that adipocytes act not only as a paracrine/autocrine cell
but adipose tissue has many endocrine functions as well.
2.1.2 Adipocyte Differentiation
Adipocytes are derived from multipotent mesenchymal stem cells. Adipogenesis, the
formation of mature, terminally-differentiated adipocytes through the differentiation of precursor
cells, can occur throughout life and is likely the primary cause of increase in body fat mass[11].
The molecular mechanisms that govern the differentiation of adipose precursors to mature
adipocytes has been reviewed in the literature[6,12-15]. The differentiation cascade may be
initiated by hormones such as insulin growth factor-1 (IGF) and insulin which have been clearly
shown to be necessary for adipogenesis[16,17]. Since it is difficult to classify distinct cellular
intermediates between stem cells and mature adipocytes, adipogenesis is generally broken down
into three distinct phases: 1. Determination or commitment; 2. Growth arrest and clonal
expansion and; 3. Terminal differentiation. An overview of these events is presented in Figure
2-1.
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Figure 2-1: Major phases in the development of mature adipocytes from multipotent stem cells found in the stromal-vascular fraction of adipose tissue.
Multipotent stem cells with the capacity to differentiate into several mesenchymal lineages give rise to the preadipocyte as a result of adipogenic inducers. The cells undergo growth arrest and clonal expansion and subsequent terminal differentiation. The cells change shape from a splindle-like fibroblastic morphology to a rounded cell with increased size while accumulating lipid droplets. The mature adipocyte is a unilocular cell. Changes in gene expression are indicated on the right with arrows suggesting their approximate duration. Reprinted by permission from Macmillan Publishers Ltd: Journal of Craniofacial Surgery [12], copyright (2007).
Determination or Commitment
The first phase, known as determination or commitment, involves the commitment of the
pluipotent stem cells to the adipocyte lineage, which results in the conversion of the multi-potent
stem cell to the preadipocyte. These precursors cells are small (diameter 10-12μm), fusiform or
fibroblast-like in appearance and do not contain any intracellular lipid[4]. The factor or gene
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responsible for the commitment of the stem cell to the adipocyte lineage is unknown. One
theory is that the signal is secreted by nearby mature adipocytes, as they have been shown to
secrete factors that promote preadipocyte proliferation and differentiation[18]. Bone
morphogenic protein 4 (BMP4) has been shown to cause 10T1/2 cells (multipotential cell line) to
commit to the adipocyte lineage at high frequency in vitro, suggesting its role in preadipocyte
committal[19]. Pre-adipocyte factor-1 (Pref-1) is another factor that has been proposed to
maintain the phenotype of the preadipocytes. Commencement of differentiation of preadipocytes
has been shown to coincide with decreased expression of pref-1 and that pref-1 is highly
expressed in 3T3 preadipocytes but not mature fat cells[20].
Growth Arrest and Clonal Expansion
Growth arrest of the proliferating phenotype of preadipocytes is the next step in the
differentiation process. It appears that growth arrest and not the actual cell-cell contact is needed
for differentiation[6]. The transcription factors, CCAAT/enhancer binding protein α (C/EBPα)
and peroxisome proliferator-activated receptor γ (PPARγ) are two key transcription factors that
control adipocyte differentiation. While their expression increases dramatically during adipocyte
differentiation, low levels of these factors expressed in preadipocytes may be responsible for
initiating growth arrest in the cells. Studies have demonstrated both the antimitotic activity of
C/EBPα[21] and the ability of PPARγ to induce cell cycle withdrawl[22]. After growth arrest,
studies on preadipocytes cell lines demonstrate that these cells undergo at least one round of
DNA replication and cell division leading to the clonal expansion of the committed cells[23].
Yet, for primary human preadipocytes clonal expansion may not be an important step as they do
not appear to require cell division to begin differentiation[24].
11
One of the earliest changes in gene expression during differentiation is the increased
expression of lipoprotein lipase (LPL) which is often cited as an early marker for adipocyte
differentiation[25]. LPL is secreted by mature adipocytes and is involved in controlling lipid
accumulation[26]. LPL expression has been shown to occur spontaneously at confluence
without adipogenesis inducers and therefore may be involved in growth arrest and may not be a
part of an early differentiation step[27].
Terminal Differentiation
In the final and third phase, the preadipocytes undergo morphological and biochemical
changes where cells become rounded in shape and increase their diameter by up to 20 times by
accumulation of triacylglycerol and lipid vacuoles and become sensitive to insulin[6]. Terminal
differentiation is controlled through the expression of CCAAT/Enhancer binding proteins
(C/EBP) and peroxisome proliferator-activated receptors (PPAR). Both C/EBPα and PPARγ act
synergistically to activate the transcription of genes that produce the adipogenic phenotype[28].
The activity, protein and expression levels of mRNA of many enzymes involved in
triacylglycerol synthesis including adenosine triphosphate citrate lyase, malic enzyme, glycerol-
3-phosphate dehydrogenase (GPDH) and fatty acid synthase increase by up to 100-fold[6,29,30].
Glucose transporters are also increased[31] as well as proteins not directly associated with
triacylglycerol synthesis such as aP2, a fatty acid binding protein which is considered an
intermediate marker of adipocyte differentiation[29,32]. Other factors that are produced by the
maturing adipocytes include angiotensinogen II, adipsin, and leptin which are viewed as late
markers of differentiation[6,25,33].
12
2.1.3 Adipose Tissue Extracellular Matrix
The extracellular matrix of adipose tissue is a critical component that gives not only
mechanical support of the tissue but also provides sites for cell attachment, proliferation and
migration and signaling that controls a variety of processes including adipogenesis, angiogenesis,
and immune response. Structurally, the ECM of adipose tissue serves to connect adipocytes
where they form clusters or fat lobules in vivo. These lobules are highly vascular and therefore
also include endothelial cells and their respective ECM. Adipose tissue ECM is divided into two
different compartments, the basement membrane that surrounds individual adipocytes and the
reticular fibre network that forms a soft skeleton that supports the tissue. The basement
membrane is comprised of networks of collagen IV and laminin-8 linked together with entactin
and heparan sulfate proteoglycan. The reticular fibres are made of collagen type III connected to
fibres of collagen type I[12]. During differentiation, not only are there changes in cell
morphology where preadipocytes enlarge and acquire lipids, but also on the level of secreted
ECM as well. One of the first signs of differentiation, for example, is deposition of collagen at
the cell-ECM border and creation of the basement membrane[34]. Alterations in the adipose
tissue ECM by the differentiating preadipocytes are likely needed to change cell adhesion
properties, remodeling of cell components and cellular organization into fat lobules. Studies
with preadipocyte cell lines have shown that when undergoing differentiation these cells lose
their fibroblastic characteristic along with decreased expression of collagens type I and III[35]
and increased expression of collagen IV, laminin, entactin and glycosaminoglycans[36-38]. In
addition, differentiation of 3T3-F442A cells was shown to have decreased synthesis and
assembly of actin and fibronectin, with concurrent disappearance of fibrillar bundles of
extracellular matrix fibronectin in the pericellular space[39]. While preadipocytes have been
shown to alter secretion of ECM components during differentiation, the ECM may in turn have a
13
role in regulating the differentiation process. Inhibition of collagen synthesis, for example, was
shown to prevent preadipocyte differentiation, highlighting that collagen synthesis is a
requirement for differentiation[40]. Furthermore, studies on fibronectin have shown that it has
both pro and anti-adipogenic properties[29,41].
2.1.4 Stem Cells for Adipose Tissue Engineering
Adult stem cells, known as mesenchymal stromal cells (MSC), are mature adult cells that
are undifferentiated and found in a particular tissue or organ. These cells have the capacity for
self-renewal and are able to differentiate into the majority of specialized cell types found in the
body. In addition, they are easily isolated from bone marrow and adipose tissues and can be
expanded in vitro, making them an attractive cell source for tissue engineering. Bone marrow
derived mesenchymal stromal cells (BM-MSC) are likely the most recognized adult stem cell
used for therapeutic purposes. They are multipotent cells and can be differentiated into several
cell types from different lineages, including osteocytes, myocytes, adipocytes and neural
cells[42-44]. Similar to bone marrow, adipose tissue represents another source of mesenchymal
stromal cells. Adipose tissue is typically present in an excess and expendable quantity in the
body which makes ASC harvesting a relatively safe and easy procedure by liposuction or
abdonminoplasty. When given the proper stimuli, ASC harvested from the stromal vascular
fraction (SVF) of adipose tissue have been shown to differentiate into cells of adipogenic,
chondrogenic, myogenic and osteogenic lineages[45,46]. It is not surprising that both ASC and
BM-MSC are found to be very similar in both morphology and phenotype[47] since both adipose
tissue and bone marrow are derived from the mesenchyme. Directly comparing the phenotype
between ASC and BM-MSC shows that they are >90% identical[48]. Typical of MSC, ASC
have been shown to express high levels of stem cell surface markers CD29, CD73, CD90,
14
CD105 and low levels of CD45[49]. In contrast to BM-MSC, ASC have been shown in several
reports to be CD34+ at early passages, with transient decrease in expression level with
subsequent expansion in culture[49-51]. It is still not clear whether CD34 can be used as a stem
cell marker to identify the ASC lineage. One group working with human adipose tissue found
that the population of the SVF that was positive for CD34 contained the ASC[52]. While species
differences may exist, another group studying murine adipose tissue found that the CD34
negative SVF population was enriched for ASC[53].
2.1.5 Adipose Tissue Engineering Strategies
There have been several strategies investigated to engineer adipose tissue. Scaffold
guided tissue regeneration is the method most widely investigated. In this approach ASC are
cultured on absorbable polymeric scaffolds and implanted in vivo such that simultaneous cell
proliferation/differentiation and scaffold resorption results in a mature fat pad. An important
design criterion here is the artificial extracellular matrix (ECM) that is used as a three-
dimensional support for the transplanted cells. Not only must this ECM provide the necessary
mechanical support and desired degradation properties but must also provide the appropriate
cell-matrix interaction. ASC have been cultured successfully on a wide array of materials for
adipose tissue regeneration including synthetic materials such as biodegradable PLGA[54],
protein-coated polytetrafluroethylene[55], polyglycolic acid[56] and natural scaffolds such as
hyaluronan-based[57], collagen[58] and acellular placenta[59]. Hydrogel materials, such as
collagen[60] and alginate[61] gels may prove optimal because their mechanical properties
closely match soft tissue, providing a “feel” more like native adipose tissue. Synthetic hydrogels
have also been investigated, yet as ASC are anchorage-dependent cells[62], synthetic materials
typically need to be modified to promote cellular attachment in order to avoid cell death. A
15
poly(ethylene glycol) hydrogel system was shown to support preadipocyte viability, for example,
only upon modification with the laminin-binding peptide sequence YIGSR[63], which is
consistent with research that demonstrates preadipocyte adhesion and migration on Laminin-1
via the α1β1 integrin[62]. The optimized hydrogel system that supported both viability and
proliferation also included the peptide sequence LGPA to permit polymer degradation by cell-
secreted collagenase[63]. Remodelling of the matrix by the cells may therefore be necessary to
create space to accommodate proliferation and cellular expansion of the ASC during
differentiation.
Another approach to repair soft tissue deficits involves the use of an acellular tissue
engineering device. This strategy, known as de novo adipogenesis, uses a stimulus, such as a
growth factor, to cause migration of ASC to the implant site where they will proliferate and
differentiate into adipocytes to form a local accumulation of adipose tissue. This method was
demonstrated by injections of Matrigel (gelatinous protein mixture secreted by Engelbreth-
Holm-Swarm (EHS) mouse sarcoma cells) with basic fibroblast growth factor (bFGF)[64,65].
The authors concluded that the visible fat pad that formed was the result of ASC and EC
migration to the injection site. In another example, photocured styrenated gelatin microspheres
with bFGF and insulin and insulin-like growth factor I (IGF-1) with a view to stimulating both
angiogenesis and adipogenesis[66]. They were successful in creating adipose tissue in vivo yet
de novo adipogenesis seemed limited in the quantity of fat tissue that can be created and may not
be clinically relevant for large soft tissue voids.
While many current strategies are successful in initially generating an accumulation of
adipose tissue in vivo, they fail to engineer an adipose tissue graft that will remain viable and
retain its volume over the long-term (> 3 months). In one of the few published long-term
16
studies, the volume of adipose tissues engineered with ASC seeded onto a PLGA scaffold and
implanted subcutaneously into a rat was found to be maximal at 2 months, decreased by 3
months and completely resorbed by months 5-12[67]. In another study, ASC seeded onto
collagen scaffolds implanted subcutaneously into mice decreased in volume by approximately
70% after 3 weeks implantation[58]. The volume loss of adipose tissue may be the result of
insufficient mechanical strengths of the construct to withstand the compressive forces in vivo and
the lack of sufficient neovascularization upon implantation.
2.1.6 Semi‐Synthetic Collagen/Poloxamine Hydrogel
Poloxamine is a collagen-mimetic as developed by our group[68] to overcome some of
the limitations of collagen hydrogel scaffolds, mainly weak mechanical properties and
progressive remodeling and contraction of the matrix. Matrix contraction, which can result in
volume reductions of up to 90% over time, is undesirable when engineering a soft tissue
replacement where maintenance of implant volume is critical.
Poly(ethylene oxide) and poly(propylene oxide) block copolymers (PEO-PPO) are
commonly used as materials in biomedical engineering. They have shown good
cytocompatibility with different cell types and they are non-irritating when applied topically or
subcutaneously, and produce little in vivo response. The poloxamines, also known as
Tetronic™) are different from the more common PEO-PPO-PEO triblock coplymers in that they
contain four arms and as such their higher functionality makes a superior degree of crosslinking
achievable with lower polymer concentrations. Collagen-containing poloxamine hydrogels are
produced by functionalization of the four-arm PEO-PPO block copolymer with methacrylate
groups (Figure 2-2) and subsequent UV free radical polymerization of water solutions of the
modified polymer in the presence of collagen. This forms a network of collagen suspended in
17
the crosslinked poloxamine hydrogel, where the collagen imparts the cell adhesive properties to
the matrix while the poloxamine (non cell adhesive) provides the mechanical strength. The
crosslinked collagen/poloxamine hydrogel was found to be much stiffer than a pure collagen gel
with a storage (G’) and loss modulus (G’’) of 7400Pa and 1000Pa as compared with 70Pa and
10Pa respectively for collagen-only gel[68].
Figure 2-2: Poloxamine is modified with methacrylate groups (IEM) that can undergo free radical polymerization.
18
2.2 Angiogenesis & Revascularization Strategies
2.2.1 Mechanisms of Angiogenesis
Angiogenesis is the process of new blood vessel formation and is a critical for tissue growth,
function and repair. Figure 2-3 highlights some of the pathological steps during new blood
vessel formation. Angiogenesis can occur though three methods: sprouting, bridging and
intussusception. Spouting is the most common process and begins with vessel destabilization;
the activation of the endothelial cell layer, vasodilation of the existing vessels, an increase in
vascular permeability and degradation of the basement membrane [69,70]. Largely in reponse to
nitric oxide (NO) production, vasodilation is one of the earliest steps in angiogenesis. VEGF, in
part due to the increase in NO production [71], mediates an increase in vascular permeability due
to the redistribution of intercellular adhesion molecules, including PECAM-1, and vascular
endothelial (VE)-cadherin, and alterations in cell membrane structure via induction of a series of
kinases [69,72]. Angiopoietins (Ang-1 and Ang-2) are ligands for the EC receptor Tie-2. While
Ang-1 is a natural anti-permeability factor, Ang-2 helps detach smooth muscle cells and loosens
the underlying matrix allowing EC to migrate as the inter-endothelial cell contacts are
broken[69]. Subsequently, the extravasation of plasma proteins, similarly induced by VEGF, is
required to provide a temporary support for the migration of activated EC. The activated EC
modify proteinase expression and release proteinases causing degradation of the basement
membrane. This is an important step as it will allow the EC to invade into the stroma of
neighborouring tissue. The degradation of the basement membrane is caused by the cooperation
of urokinase-plasminogen activator (uPA) and metalloproteinase (MMP) systems [70]. The
degradation of the ECM, not only provides “room” for the EC to migrate, but results in the
liberation of growth factors, including bFGF, VEGF, and insulin-like growth factor-1 (IGF-1),
19
Figure 2-3: Mechanisms of angiogenesis.
Sprouting is the most common angiogenic process and begins with activiation of the endothelial cell layer and destablization of the basement membrane. Ang-2 helps detach smooth muscle cells and loosens the underlying matrix allowing EC to migrate by VEGF stimulation. Migrating EC assemble, form solid cords and subsequently acquire a lumen. These capillaries then synthesize a new basement membrane. Vessels are stabilized and mature via the recruitment of periendothelial support cells such as pericytes and smooth muscle cells (SMC) by PDGF. Adapted by permission from Macmillan Publishers Ltd: Nature Medicine [73], copyright (2000).
which would otherwise remain immobilized in the matrix [73]. These liberated growth factors
act on the activated EC as mediators of migration and proliferation.
Once the physical barriers in the ECM are dissolved, the next step in the process of
angiogenesis is the migration of proliferating EC into the interstitial space. At this stage there
are a myriad of factors that are responsible for mediating embryonic, neonatal and pathological
angiogenesis, including VEGF, angiopoietins, FGFs and their receptors while many other factors
20
have been implicated in the process [73]. FGFs for example, stimulate EC growth and recruit
mesenchymal and/or inflammatory cells, producing countless angiogenic factors [74]. Platelet-
derived growth factor (PDGF) encourages microvascular sprouting of EC and recruits pericytes
and smooth muscle cells around nascent vessel sprouts [75].
As EC migrate into the ECM, they assemble, form solid cords and subsequently acquire a
lumen. This is achieved through the intercalation and thinning of the EC along with fusion of the
existing vessels to create longer and greater diameter vessels. The formation of the capillary
lumen requires the polarization of the EC and is established via cell-cell and cell-matrix
interactions. These capillaries then synthesize a new basement membrane. Vessels are
stabilized and mature via the recruitment of periendothelial support cells such as pericytes and
smooth muscle cells (SMC) [70].
Finally, upon fusion of the newly formed vessels with the preexisting vasculature, blood
flow is initiated. When sufficient neovascularization has occurred there is an upregulation of
anti-angiogenic factors, such as VEGF-R1, Ang-1, angiostatin, with downregulation of many of
the pro-angiogenic stimulators discussed. Upon completion of the angiogenic cascade, the EC
return to their quiescent state with survival measured in years.
Following a similar mechanism, angiogenesis can also occur through bridging and
intussusception. In bridging, vessels can be split into individual daughter cells through
development of transendothelial cell bridges. Branching by intussusception takes place when
there is an insertion of interstitial tissue columns into the lumen of pre-existing vessels [76].
Subsequent growth and stabilization of these colums, due in part to the ingrowth of
periendothelial cells, results in partitioning of the vessel lumen and remodeling of the vascular
network.
21
2.2.1.1 Vascular Functions in Adipose Tissues
With increasing interest in the development of a therapeutic target for obesity, the
complex interplay between the different cellular components of adipose tissue is only now being
increasingly understood and has been reviewed in [77]. Figure 2-4 shows the functional
relationship of adipose tissue to its vasculature. In growing adipose tissue, angiogenic vessels
contribute to adipogenesis through several distinct mechanisms. First and most obvious, is that
the vessels bring nutrients and oxygen to nourish the main component of the fat tissue – the
adipocytes. Like any other tissue in the body, they require sustenance for growth and survival.
Secondly, the circulating plasma provides an environment rich in growth factors and cytokines.
Leptin and adiponectin, for example, are secreted adipose-derived hormones that are responsible
for maintenance of adipose tissue mass[78]. Thirdly, the angiogenic vessels permit infiltration of
inflammatory cells, such as monocytes and neutrophils, which tend to be observed in high
numbers in obese individuals[79]. The fourth purpose of the vasculature is to supply circulating
stem cells from the bone marrow. These mesenchymal cells can differentiate into preadipocytes,
adipocytes and vascular cells, including pericytes and endothelial cells[80,81]. Fifth, a recent
study has shown that adipocytes descend from a pool of proliferating progenitor cells that reside
in the mural cell compartment of adipose vasculature, but not in the vasculature of other tissues.
Mural cells including pericytes, have been shown to be capable of differentiating into both
adipocytes and preadipocytes making the interactions between the fat and vascular compartments
even more complex[81]. The sixth functional relationship of adipose tissue vasculature is the
paracrine relationship shared between activated EC and adipocytes/preadipocytes which is
discussed in more detail in section 2.2.1.3.
22
Figure 2-4: Functions of adipose tissue vasculature.
Blood vessels in adipose tissue provide transport of oxygen, nutrients, hormones, growth factors (GFs), inflammatory cells (ICs) and bone marrow derived mesenchymal stem cells (BM-MSC) that maintains the functions of adipocytes and adipose tissue. In the circulation, BM-MSCs, ICs and endothelial progenitor cells (EPCs) participate in both angiogenesis and adipogenesis. Endothelial cells in angiogenic vessels produce growth factors and adipokines. Adipocytes also produce angiogenic factors and adipokines that induce angiogenesis. Perivascular cells such as pericytes (PC) can differentiate into pre-adipocytes that become adipocytes. Reprinted by permission from Macmillan Publishers Ltd: Nature Reviews Drug Discovery [77], copyright (2010).
2.2.1.2 Angiogenic Modulators
As seen in Figure 2-5, adipocytes and their precursors along with inflammatory and
stromal cells in adipose tissue work together as mediators in the angiogenic process. Adipocytes
secrete angiogenic adipokines and traditional angiogenic factors such as VEGF and bFGF.
Leptin, for example, has been shown to increase vascular permeability and to synergistically
stimulate angiogenesis with VEGF and bFGF[82]. Lipids secreted by fat cells may also be pro-
angiogenic. Monobutyrin, for example, has been shown to stimulate angiogenesis in vivo and
23
Figure 2-5: Modulators of angiogenesis in adipose tissue.
Adipocytes and their precursors along with inflammatory and stromal cells in adipose tissue work together as mediators in the angiogenic process though secretion of multiple angiogenic factors, adipokines and cytokines. Adapted by permission from Macmillan Publishers Ltd: Nature Reviews Drug Discovery [77], copyright (2010).
microvascular endothelial cell migration in vitro[83]. Besides pro-angiogenic molecules, fat
tissue also produces several angiogenesis inhibitors including thrombospondin I and
plasminogen activator inhibitor (PAI-1). Overexpression of PAI-1 was found to have no effect
on de novo adipogenesis in a mouse model, while the blood vessel size in the fat pad was much
smaller than controls, indicating an inhibitory effect on angiogenesis[84].
In addition to the adipocytes, inflammatory cells play a large role in angiogenesis.
Chronic inflammation is consistently seen in obese individuals, which results in the infiltration of
24
many leukocytes and macrophages. These cells produce several angiogenic growth factors,
including VEGF, tumor necrosis factor (TNFα), interleukin-6 (IL-6), IL-8 and
chemokines[79,85].
ASC play a major role in adipose tissue angiogenesis through the secretion of soluble
angiogenic molecules and possibly by differentiating into vascular cells that become part of the
endothelium. ASC produce multiple angiogenic factors including, VEGF, hepatocyte growth
factor, bFGF, transforming growth factor β, and chemokines[86]. In addition to the secretion of
bioactive molecules, recent research has shown that ASC have the ability to differentiate into
endothelial cells or perivascular pericytes so they may actively participate in the angiogenic
process. Populations of ASC that are CD34+ have been shown to preferentially differentiate into
endothelial cells, while those that have lost CD34 expression tend to express more pericyte
markers which may be related to the process of commitment or differentiation into a specific
lineage[87,88].
2.2.1.3 Relationship between ASC and EC
Adipose tissue development is closely associated with angiogenesis but the molecular
interplay between adipogenesis on angiogenesis remains unclear. Research over the past decade
has demonstrated the paracrine relationship between adipose tissue and the vasculature and has
identified some of the pro-angiogenic and anti-angiogenic growth factors expressed by adipose
tissues or adipocytes cultured in vitro. These include VEGF, bFGF, Ang-1,-2, PAI-1, TGF-β
and leptin[89]. A recent study demonstrated that human preadipocytes and endothelial cells both
express the αvβ3 integrin and express and secrete PAI-1 which regulates preadipocyte and EC
migration in vitro[90]. This suggests that during development preadipocytes migrate with the
25
developing capillary bed during angiogenesis, and that the production and secretion of PAI-1 by
the preadipocytes ensures local coordination of adipogenic/angiogenic process.
In another study that highlights the interplay between angiogenesis and adipogenesis, the
PPARγ pathway (master regulator of adipogenesis) was blocked in 3T3-F442A preadipocytes
(preadipocyte cell line) and found to not only inhibit differentiation of these cells but
angiogenesis in vivo as well[89]. They also investigated the role of vascular endothelial growth
factor receptor 2 (VEGFR2) signaling in angiogenesis, vessel remodeling and adipocyte
differentiation. VEGFR2 is expressed on vascular endothelial cells and its signaling is critical to
the angiogenic process and VEGF is highly expressed during adipocyte differentiation[89].
They found that inhibition of angiogenesis by a VEGFR2 blocking antibody not only reduced
angiogenesis and tissue growth, but also inhibited preadipocyte differentiation as well[89].
These findings demonstrate the reciprocal relationship between adipogenesis and angiogenesis
and suggest that inhibition of VEGF signaling can reduce adipose tissue formation in vivo.
In another well-defined study, EC/ASC cocultures were studied in vitro in a system
containing serum but no additional exogenous cytokines or ECM proteins to better understand
the mechanisms at work[91]. ASC were found to have a profound effect on EC to stimulate
angiogenesis and branching networks of cord structures. During coculture, there was an
observed increase in the ECM protein production by both the ASC and EC, increased α-smooth
muscle actin expression by ASC and increased CD31/platelet endothelial cell adhesion
molecule-1 (PECAM-1) on the surface of the EC. Overall, matrix metalloproteinase activity and
paracrine cell communications between ASC and EC in the VEGF, hepatocyte growth factor and
PDGF pathways were found to impact vascular network formation. As expected, ASC were
shown to exhibit a significantly higher angiogenic effect than smooth muscle cells or fibroblasts.
26
These studies are in accordance with earlier work that showed how differentiation of
3T3-F442A stimulated EC chemotaxis in vitro and neovascularization in vivo[92]. Extracellular
matrix factors secreted by microvascular endothelial cells, derived from adipose tissue, have
shown to stimulate both preadipocyte proliferation[93] and differentiation[94] into mature fat
cells in primary culture.
Despite these observations, the exact mechanisms between adipogenesis and
angiogenesis are not well defined. It appears that a paracrine relationship exists between ASC
and EC that may be exploited when creating a vascularized adipose tissue engineering construct.
2.2.2 Revascularization Strategies
Previous attempts at adipose tissue reconstruction using either autologous tissue or tissue
engineering strategies have commonly failed as a result of implant resorption due to ischemic
cell death. Native adipose tissue is highly vascular, with each mature fat cell in contact with at
least one capillary[95]. One of the challenges of tissue engineering is to provide adequate
nutrients and oxygen to all cells embedded within the tissue engineered construct, especially
those large 3D constructs of clinical relevance, as the diffusional limitation of approximately
150µm is imposed on nutrient and waste transfer[96]. As such, a successful adipose tissue
engineering strategy will promote rapid integration with the host vasculature upon implantation.
Revascularization and inosculation are two strategies by which a tissue engineered device can
become integrated with the host vasculature[97]. In a revascularization strategy, the host blood
vessels grow into and nourish the implanted device. This approach is limited, however, by the
slow kinetics of nutrient and waste diffusion and may only be suitable for small or thin
constructs. Delivery of angiogenic growth factors could be used to accelerate blood vessel
growth, however it is difficult to orchestrate the complex temporal and spatial cues required for
27
stimulation and maturation of blood vessels by the delivery of but a few growth factors. A more
promising strategy to support cellular survival in larger matrices is inosculation, whereby a
capillary network is created within the adipose substitute and is subsequently fused with the host
vascular system upon implantation[98]. Several revascularization strategies have been
investigated in the literature, these include: growth factor delivery, biomaterial-mediated
angiogenesis, transplantation of primary endothelial cells, vascular pedicle, and stem cell
transplantation.
2.2.2.1 Growth Factor Delivery
Many current studies of therapeutic angiogenesis deliver angiogenic growth factors either
directly into the circulation or at the target site through a single or a series of bolus injections.
This approach has been used successfully in cases of both peripheral[99] and myocardial[100]
ischemia, yet has the disadvantage of possibly producing unwanted blood vessel formation at
distant sites. In addition, the half life of most bolus injections of growth factors is less than one
hour, making repeated injections necessary to maintain the therapeutic effect.
In tissue engineering approaches, growth factors are combined with polymeric
biomaterials as a vehicle for localized and sustained release. Through chemical and/or physical
means, angiogenic growth factors such as VEGF, bFGF and PDGF can be incorporated into
tissue engineering scaffolds to promote angiogenesis upon implantation. Perhaps most widely
investigated in the adipose tissue engineering literature is the incorportation of bFGF into the
tissue engineering device. bFGF has been shown to enhance adipogenesis of mesenchymal stem
cells in both 2 and 3-dimensional in vitro cultures[101,102], and also reported as effective in
inducing angiogenesis, and regeneration of bone, cartilage and nerve[103-105]. Work by
28
Thompson and colleagues showed that bFGF incorporated into a collagen type I gel produced de
novo adipogenesis accompanied by angiogenesis in mice[106]. However, their engineered
adipose tissue resorbed after 6 weeks once the collagen gel scaffold had degraded. Others have
investigated bFGF release from gelatin microspheres in Matrigel™, and found significant
adipose tissue generation, yet again volume maintenance was not achieved[64,65]. It appears
that while sustained angiogenic growth factor release is successful in initially generating
adiopose tissue in vivo, once the scaffold degrades and the growth factor delivery ceases, the
regenerated tissue diminishes.
2.2.2.2 Biomaterial‐mediated Angiogenesis
An emerging interest in the field of tissue engineering is exploring the possibility that a
biomaterial itself can be used as an agonist of an angiogenic response. Here we can imagine
incorporating ASC with an appropriate biomaterial that will elicit an angiogenic response once
implanted in the body. In previous years, we have investigated a poly(butyl methacrylate-co-
methacrylic acid) tissue engineering scaffold that was shown to be pro-angiogenic in vivo
without the use of exogenous growth factors[107]. Hyaluronan is an example of a naturally
occurring polymer that has been suggested to have pro-angiogenic activity. It is an important
structural component in the ECM that is involved in numerous cell adhesion and migration
events and has been found to be naturally angiogenic when degraded into small fragments[108].
Sacchachitin, a natural polymer derived from fungal mycelia, is another biomaterial that has
shown pro-angiogenic potential[109]. The authors found that in a full-thickness skin graft model
a larger number of blood vessels were found in the sacchachitin covered wound compared to
cotton gauze control at day 3. Hisology revealed that there was an acute and rapid inflammatory
response with a larger number of neutrophils and lymphocytes in the sacchatitin treated wound
29
compared to the control. Perhaps the invading inflammatory cells released pro-angiogenic
growth factors (ie. VEGF) which would explain why granulation tissue and angiogenesis
occurred faster in the sacchachitin treated wound as compared to the control.
2.2.2.3 Vascular Pedicle
Another revascularization strategy is the use of a defined vascular pedicle to nourish the
adipose tissue engineering construct in vivo. In this approach a vascular pedicle is elevated and
inserted into a hollow plastic chamber and implanted subcutaneously. The chamber typically
contains a biomaterial and growth factors to accelerate de novo adipogenesis and to stimulate
angiogenic sprouting from the pedicle. Once the tissue has been generated, it is possible to
transfer the construct microsurgically to a remote site to repair a soft tissue defect. Morrison and
colleagues have developed an in vivo model of angiogenesis in rats, based on the microvascular
arteriovenous loop inside a polycarbonate chamber in the groin of a rat[110]. They were able to
show than when the chamber was combined with a collagen matrix, it spontaneously developed a
vascularized, three-dimensional, fibrous tissue mass. They further developed this model for
mice and showed that when Matrigel™ was used as the matrix inside the chamber, the chambers
filled with adipose tissue, creating a highly vascularized fat flap[111].
2.2.2.4 Cell‐Based Revascularization Strategies
Cell based revascularization strategies have been studied in patients with various
ischemic diseases. These studies have shown that stem cells from bone marrow[112], skeletal
muscle[113], umbilical cord blood[114] and adipose tissue[115] can be used to treat patients
with conditions such as myocardial infarction, heart disease and peripheral vascular disease. It is
30
believed that these progenitor cells have potential in promoting tissue revascularization and
restoring function through the release of many bioactive molecules.
Transplantation of EC as a revascularization strategy is a relatively unexplored approach.
Work in our laboratory has studied HUVEC transplanted subcutaneously into severely combined
immunodeficient mice (SCID). HUVEC were found to initially form vessel-like structures in
vivo but these tended to disappear after a few weeks after transplantation. We then explored the
implantation of rat aortic endothelial cells in an allogenic rat model. Stable chimeric vessels
were seen, yet only when the animals were given immunosuppressive drug treatment[116].
Others have also observed the fragile nature of EC transplantation. Human dermal endothelial
cells (HDMEC) seeded in Matrigel™ and implanted in a PLGA scaffold in SCID mice showed
initial growth and sprouting of donor derived vessels, but these regressed quickly by 14
days[117]. In another example, HUVEC in a Matrigel™ plug implanted in SCID mice showed a
decrease in HUVEC vessel density over time[118]. It appears that EC transplantation alone is
not sufficient to avoid apoptosis and drive the formation of mature, stable and functional vessels
long-term.
Several methods to achieve viability and stability of donor EC derived vessels have been
investigated and include transfecting an anti-apoptotic gene into the EC or cotransplantation with
a supporting cell. Bcl-2 is an anti-apoptotic gene that has been shown to extend EC survival in
vitro[119]. Pober and colleagues demonstrated that Bcl-2 transduced HUVEC produced an
increased density of HUVEC-lined microvessels that were perfused in vivo as compared with
untransduced or control-transduced HUVEC[120]. It appeared that Bcl-2 overexpression
recruited ingrowth of perivascular smooth muscle cells, which organized by day 60 into
HUVEC-lined multilayered stable-vessels. Similarly, another group looked at implants of
31
collagen-fibronectin gels containing Bcl-2 transduced HUVEC and found that they produce
human EC and murine vascular smooth muscle cell chimeric vessels in SCID mice[121].
Microfil casting of the vasculature revealed perfusion of the donor derived vessels with the host
at day 60, demonstrating anastomosis and vessel functionality. Cotransplantation with a
supporting cell is another successful approach to ensure viability and stability of the donor EC
vessels. In angiogenesis, other vascular cells such as fibroblasts, smooth muscle cells and
pericytes migrate to help support and stabilize the growing vasculature. As such, it is not
surprising that cocultures of pericytes and EC have stabilized vessels in 3D collagen gels[122]
and that fibroblasts were shown to stabilize tubular network formation in tissue-engineered
reconstructed skin[123]. Perivascular precursors (10T1/2 cells) and embryonic fibrobasts have
also been co-cultured with HUVEC in PLGA scaffolds[124]. The vascular network created by
the cotransplantation of HUVEC and these precursors in SCID mice was shown to be functional
and stable for 1 year in vivo. These precursor cells likely differentiated into smooth muscle cells
which acted as a support for donor vessel maturation and stability.
In adipose tissue engineering, the use of ASC as the supporting cell in an EC
transplantation revascularization strategy has only just begun to be investigated. Increasing
evidence has shown that the ASC in adipose tissue may reside in a perivascular location and that
pericytes and endothelial cells may actually be progenies of the ASC[125]. ASC may induce
angiogenesis through paracrine support[50] of EC and suppression of their apoptosis[126,127]
and may actually integrate into the host vasculature[127]. In addition, ASC have been shown to
have pericyte-like activity in vitro by stabilizing endothelial networks[87]. Stark and colleagues
were the first to show a patent connection of tissue-engineered microvessels in adipose tissue to
a host vessel system without applying exogenous angiogenic growth factors of transfection with
32
an anti-apoptotic gene[128]. They combined HDMEC spheroids and ASC in a fibrin matrix in a
chorioallantoic membrane (CAM) model. After 7 days, many vessels consisting of HDMEC
could be detected and the vessels were perfused with chick erythrocytes. In a recently published
article, ASC were combined with endothelial progenitor cells (EPC) in collagen implants and
implanted subcutaneously in SCID mice[129]. The ASC were shown to cooperate with the
EPCs to coassemble into a functional vessel network. Synergism by cotransplantation of ASC
and EPC increased the density and complexity of the vascular networks many fold greater than
in implants with ASC or EPC alone. The authors speculated that the angiogenic effect that ASC
have on EC may be the result of secretion of diffusible proangiogenic and antiapoptotic factors
(such as VEGF and Ang-1) and direct contact with forming endothelial tubes.
Since most strategies to engineer adipose tissue employ ASC, it seems that EC
cotransplantation is a promising revascularization strategy.
33
3 Semi‐Synthetic Collagen/Poloxamine Hydrogel as a Potential Biomaterial for Adipose Tissue Engineering
3.1 Abstract
With increasing clinical demand for an adipose tissue substitute, a suitable biomaterial that
possesses the appropriate mechanical properties and ‘feel’ of fat along with the necessary
biological properties for cellular transplantation and regeneration is needed. A semi-synthetic
collagen-containing poloxamine hydrogel was investigated for its feasibility as a novel
biomaterial for adipose tissue engineering. Adipose-derived stem cells (ASC) embedded in the
photocrosslinked collagen/poloxamine hydrogel displayed lower viability over 14d in culture as
compared with ASC in collagen gels. Immunostaining of the collagen/poloxamine hydrogel
revealed inhomogeneous distribution of collagen within the hydrogel, resulting in regions devoid
of any cell attachment. Adipogenic differentiation was confirmed by Oil Red O staining in the
collagen/poloxmine hydrogel, yet was observed only in regions of the hydrogel rich in collagen.
Human microvascular endothelial cells (HMEC-1) attached and proliferated onto the surface of
both collagen/poloxamine hydrogel films and modules, yet incomplete endothelial cell coverage
was achieved over 14d in culture. The immiscibility of collagen and poloxamine made it
undesirable for a biomaterial for adipose tissue engineering.
3.2 Introduction
Tissue engineering has the potential to address the clinical need for the repair and
regeneration of adipose tissue defects. Millions of surgeries are performed each year to restore
normal cosmesis resulting from soft tissue traumas and abnormalities, including mastectomies,
tumor removal, deep burns and congenital defects. Current treatment modalities use autologous
and allogenic adipose tissues with limited success as these therapies are associated with
34
operative risk, donor site morbidity and progressive resorption/fibrosis of the implanted tissue
due to insufficient neovascularization of the fat graft[54,130].
Recent approaches to adipose tissue engineering combine Adipose-derived Stem Cells
(ASC) and a suitable biomaterial that is used as a three-dimensional support for the transplanted
cells. A biomaterial for adipose tissue engineering must provide the necessary mechanical
support and “feel” but must also provide the appropriate cell-matrix interaction for ASC viability
and differentiation. ASC have been cultured successfully on a wide array of materials for
adipose tissue regeneration including synthetic materials such as biodegradable PLGA[54],
protein-coated polytetrafluroethylene[55], polyglycolic acid[56] and natural scaffolds such as
hyaluronan[57], silk[131] and collagen scaffolds[58]. Hydrogel materials, such as collagen[60]
and alginate[61] gels may prove optimal because their mechanical properties closely match soft
tissue. Collagen has the advantage of biodegradability, low antigenicity and cell-binding
properties, yet it is mechanically weak. Upon implantation it undergoes remodeling and
contraction with volume reductions of up to 90%, making it not ideal for an application where
volume maintenance is critical[132]. Synthetic hydrogels have tunable mechanical properties,
yet may need to be modified to promote cellular attachment in order to avoid cell death. A
poly(ethylene glycol) hydrogel system, for example, was shown to support ASC viability only
upon modification with the laminin-binding peptide sequence YIGSR[63], which is consistent
with research that demonstrates ASC adhesion and migration on Laminin-1 via the α1β1
integrin[62].
A semi-synthetic collagen/poloxamine hydrogel was developed by our group[68] to
combine both the biological properties of collagen and the improved mechanical stiffness of the
synthetic hydrogel. Poly(ethylene oxide) and poly(propylene oxide) block copolymers (PEO-
35
PPO) are commonly used as materials in biomedical engineering[133]. They have shown good
cytocompatibility with different cell types and they are non-irritating when applied topically or
subcutaneously, and produce little in vivo response[133]. The poloxamines, (also known as
Tetronic™) are different from the more common PEO-PPO-PEO triblock coplymers in that they
contain four arms and as such their higher functionality makes a superior degree of crosslinking
achievable with lower polymer concentrations. The crosslinked collagen/poloxamine hydrogel
was found to be much stiffer than a pure collagen gel with a storage (G’) and loss modulus (G’’)
of 7400Pa and 1000Pa as compared with 70Pa and 10Pa respectively for collagen-only gel[68].
In addition, cytocompatibility of embedded and surface-attached cells was high [134].
Mechanical strength is not the only criterion in designing a hydrogel for adipose tissue
engineering. The effective attachment of EC to the module surface and the phenotype of
embedded ASC and their differentiation into mature adipocytes must be considered. It was
thought the unique 2-phase network of collagen/poloxamine may be advantageous for ASC
differentiation. The ASC are anchorage dependent and need to attach to the matrix to proliferate
and avoid cell death by anoikis[63]. During differentiation, these cells must be able to remodel
the matrix (ie. using cell secreted collagenase) to provide room for volume expansion during
lipogenesis. Mature adipocytes, on the other hand, may prefer a non cell-adhesive matrix. It has
been suggested that a biomaterial suitable for adipose tissue engineering may need a balance
between cell adhesive qualities (attachment and spreading) and non-adhesive properties that
favour differentiation[135].
Here, we report on the feasibility of using collagen/poloxamine as a novel biomaterial for
adipose tissue engineering. Studies were performed to assess embedded ASC viability,
differentation and EC cell attachment.
36
3.3 Materials and Methods
3.3.1 Cells
A protocol for the isolation and culture of human adipose-derived stem cells (ASC) has
been adopted from Dr. Lauren Flynn, Queen’s University[136]. Fresh, sterile abdominal fat
samples were obtained from patients undergoing elective surgery, with consent. The tissue was
minced and digested in a 2mg/mL collagenase solution at 37oC, containing Kreb’s Ringer
bicarbonate buffer, supplemented with 3mM glucose, 25mM HEPES and 20mg/mL BSA (all
Sigma, Oakville, ON). Upon digestion the sample was filtered to remove undigested tissue
fragments and centrifuged to separate the desired stromal-vascular cell population from the
buoyant mature adipocytes. The cell suspension was gently agitated in erythrocyte lysing buffer
and filtered again to remove red blood cell fragments. Several centrifugation steps with washes
in complete media finished the processing. The cells were plated in a 1:1 mixture of Dulbecco’s
Modified Eagle’s Medium and Ham’s F-12 nutrient mixture (DMEM:Ham’s F-12) (Sigma,
Oakville, ON), supplemented with 10% fetal bovine serum (FBS) (Sigma, Oakville, ON), 100
U/mL penicillin and 0.1 mg/mL streptomycin (Gibco, Burlington, Canada). The growth medium
was changed every 2-3 days. To passage the cells, cultures at 90% confluence were trypsin-
released (0.25% trypsin/0.1% EDTA, Gibco, Burlington, Canada), washed, counted and re-
plated in new flasks at 30,000 cells/cm2. Cells at passage 2 were used for the differentiation
experiments, while higher passage cells (but < passage 10) were used for the viability
experiments.
To induce adipogenic differentiation, cells were cultured in serum-free DMEM:Ham’s F-
12 supplemented with 15 mM NaHCO3, 15 mM HEPES, 33 μM biotin, 17 μM pantothenate, 10
μg/mL transferrin, 100 nM cortisol, 66 nM insulin, 1 nM triiodothyronine (T3), 100 U/mL
37
penicillin and 0.1 mg/mL streptomycin. For the first 72 hours of differentiation, 0.25mM
isobutylmethylxanthine (IBMX) and 1 μg/mL of troglitazone were added to the differentiation
medium (all Sigma, Oakville, ON).
The Human Microvascular Endothelial Cell line-1 (HMEC-1) was used in the cell
attachment studies to overcome the limitations of primary cells, mainly difficulty in isolation and
limited life span. HMEC-1 is an immortalized human microvascular endothelial cell line that
has been shown to maintain the morphologic, phenotypic, and functional characteristics of
normal human microvascular endothelial cells[137].
3.3.2 Hydrogel Synthesis
Poloxamine (T1107-methacrylate) is synthesized under nitrogen by a reaction with
Tetronic 1107 (a gift from BASF, New Jersey, NJ) and isocyanato ethylmethacrylate (IEM)
using stannous (II) 2-hexyl-hexanoate (SnOct) as the catalyst (Sigma, Oakville, ON), as
described elsewhere[68].
To prepare poloxamine hydrogels, an aliquot of T1107-methacrylate solution (25 wt% in
distilled water) was mixed with a 3mg/mL collagen type I solution (PureCol™, Inamed,
Fremont, CA) or phosphate buffered saline (to produce poloxamine hydrogels without collagen)
to the desired final concentration. Then 10X MEM (10.1 g α-Minimum Essential Medium
powder (Gibco, Invitrogen, Paisley, PA), and 2.2g endotoxin free NaHCO3, in 100mL endotoxin
free water) was added and mixed thoroughly and neutralized to pH ~ 7.4 by adding the required
volume of 0.8N NaHCO3. Irgacure I2959 ((2-hydroxy-1-[4-(2-hydroxyethoxy)phenyl]-2-methyl
propanone) (Ciba, Basel, Switzerland), dissolved in ethanol, was added so that the final
concentration of photoinitiator is 0.07wt %. The poloxamine solution was then mixed with the
desired cell number (ie. 106 cells/mL) and transferred into the wells of a 96-well plate (to make
38
hydrogel films) or injected into the lumen of PE tubing (which is subsequently cut after
crosslinking to produce cylindrical modules). The poloxamine-cell solutions were incubated at
37oC for 30min (to promote collagen fibrillogenesis) and then crosslinked by exposure to UV
light for 10min (Spectroline EN-180 lamp, one 8-W 365nm tube, with intensity of
1.850mW/cm2(at source)) at 10cm (distance from source UV). Hydrogel films (or modules)
were washed repeatedly upon crosslinking with complete media and subsequently cultured at
37oC. Collagen-only gels were prepared in the same manner, only by omitting the poloxamine-
methacrylate.
In an attempt to improve cell attachment, hydrogels of collagen/poloxamine/laminin were
prepared by adding laminin (derived from Engelbreth-Holm-Swarm murine sarcoma basement
membrane in 1 mg/mL in Tris buffered NaCl, Sigma) with the collagen/poloxamine solution (5
μg/mL and 50μg/mL laminin) prior to mixing with cells and photocrosslinking
3.3.3 Cell Viability
A preliminary alamarBlue® viability assay (Invitrogen) (106 ASC/mL hydrogel, 10min
UV, 0.07wt% I2959) comparing collagen, poloxamine-only and collagen/poloxamine hydrogels
was conducted to assess the viability of the ASC in culture over 7 days. AlamarBlue® contains
a redox indicator that changes color in response to metabolic activity. Briefly, alamarBlue®
reagent was added directly into culture media of the gels in a 96-well plate, after 1, 3, and 7 days
after hydrogel fabrication, to a final concentration of 10% and the plate was returned to the
incubator. Optical density was measured at 540 and 630 nm with a standard spectrophotometer
after 4 hours of incubation at 37oC. The percent reduction was calculated and compared against
ASC embedded in collagen-only gels.
39
A Live/Dead® kit (Invitrogen) was used to stain live and dead ASC cells within the
crosslinked hydrogel. Confocal microscopy was used to visualize cells and estimate viability by
observing the presence of green (live) and red (dead) staining cells.
3.3.4 Hydrogel Characterization
Immunostaining of collagen and laminin was used to visualize the distribution of the
components in the hydrogels. Briefly, the monoclonal anti-collagen type I produced in mouse
(Sigma, Oakville, ON) and anti-laminin produced in rabbit were diluted 1:100 in PBS and 100uL
of this solution was added to each well and incubated for 10min. After several PBS washes the
secondary antibodies, Alexa Fluor® goat anti-mouse 568 (red) and Alexa Fluor® goat anti-rabbit
488 (green) (Molecular Probes) were diluted 1:1000 in PBS and added to the gels and incubated
for 10min. After several PBS washes, the hydrogels were visualized under confocal microscopy.
3.3.5 Oil Red O
Lipid accumulation within the collagen, poloxamine, collagen/poloxamine and
collagen/poloxamine/laminin hydrogels was assessed by Oil Red O (Sigma, Oakville, ON)
staining at 7 and 14 and 21 days after the induction of differentiation. At each time point, three
hydrogels from each group were fixed for 24 hours in 10% neutral buffered formalin and rinsed
thoroughly with PBS. The hydrogels were labelled prior to Oil Red O staining with the anti-
collagen type I and Alexa Fluor® 588 carboxylic acid, succinimidyl ester, as described in section
3.3.4 to visualize the distribution of collagen in the hydrogel. Hoescht (Invitrogen) was added at
a concentration of 1µg/mL and allowed to incubate at room temperature for 10min followed by
extensive PBS rinsing.
An Oil Red O stock solution (3 g/L) was prepared in isopropanol and was diluted 3:2
(v:v) with deionized water. This solution was agitated at room temperature for 10 minutes and
40
then filtered through a type 1 qualitative filter paper. Each hydrogel was incubated in 0.5 mL of
the prepared stain for 2 hours under agitation at room temperature. All samples were rinsed
thoroughly overnight in deionized water, with three solution changes to reduce non-specific
staining. The scaffolds were analyzed using confocal microscopy (Zeiss LSM510, C-APO
63x/1.2 NA water immersion (DIC) objective lens, excitation with a HeNe laser at 543 nm and
an Enterprise laser at 351 nm).
3.3.6 Glycerol‐3‐phosphate Dehydrogenase Activity
Glycerol-3-phosphate dehydrogenase activity (GPDH, a mid to late marker of ASC
differentiation) was measured for each hydrogel configuration at 72h, 7d and 14d post induction
of differentiation using a GPDH activity measurement kit (Kamiya Biomedical Co.) to assess the
impact of each type of hydrogel on the differentiation of ASC (n=4). The kit provides a substrate
reagent containing NADH and DHAP that is mixed with the prepared samples. The decrease in
the NADH concentration, as DHAP is converted into glycerol-3-phosphate by the GPDH in the
sample, can be measured by monitoring the change in the absorbance of the mixture at 340nm.
A larger change in the absorbance during the analysis period correlates with greater GPDH
activity in the sample. Using the extinction coefficient of NADH, it is possible to calculate the
GPDH (Units/mL) in the sample. To allow comparison of the different samples (varying cell
viabilities) the results were normalized for cell number as determined by semi-quantitative
Western blotting for the housekeeping gene GAPDH.
3.3.7 EC Attachment to Gels and Modules
To characterize endothelial cell attachment to the collagen/poloxamine hydrogel, both
hydrogel films (100μL in wells of a 96-well plate) and modules were prepared as outlined in
section 3.2.2. To the hydrogel films, 30,000 HMEC-1 were seeded on top of the films and
41
cultured in ‘coculture media’ (DMEM-F12 Ham’s, 10% FBS, 1% P/S, 10μg/mL hydrocortisone,
1ng/mL EGF, 10μg/mL L-glutamine) (Sigma, Oakville, ON). Modules were dynamically and
then statically seeded with HMEC-1 (2x107 HMEC-1/mL of settled modules) by first rocking the
modules in 10mL of cell suspension for 1h at 37oC and subsequently allowing the cells to settle
and attach to the modules overnight in a petri dish. At 1, 7 and 14d post-seeding, films or
modules were rinsed in PBS and stained using the Live/Dead® (Molecular Probes) viability
stain, which stains live cells green and dead cells red, and visualized with confocal microscopy
(Zeiss LSM510).
In an attempt to improve HMEC-1 attachment, several modifications including
fibronectin and collagen coatings on the surface of the modules were tested. Fibronectin (FN)
was selected as it is typically used as an adsorbed layer on the surface of tissue-culture surfaces
to promote attachment, proliferation and spreading of a variety of cell types via the Arg-Gly-Asp
(RGD) domain. Collagen/poloxamine modules were suspended in a fibronectin solution (500ug
FN per mL of settled modules, BD Biosciences) at 37oC for 1h to allow adsorption. To coate the
modules with a layer of collagen, collagen/poloxamine modules were suspended in a collagen
solution (3mg/mL, 1.5mg/mL or 0.75mg/mL bovine collagen type I (PureCol)) and neutralized
to pH 7.4 with NaHCO3. Modules were strained from excess collagen solution on a 100μm
mesh filter and the collagen coating allowed to gel at 37oC for 1h.
3.3.8 Implantation of Collagen/Poloxamine Modules into SCID mice
Adult male (6 weeks of age) male SCID mice were purchased from Charles River
Laboratories (Wilmington, MA). They were individually housed in sterile cages and fed ad
libitum under the approval by the University of Toronto animal care committee. As a pilot
study, ~ 250 modules suspended in PBS were injected subcutaneously on the dorsum of mice.
42
After 3 days, the mice were euthanized, and the implants excised and fixed in 4% neutral
buffered formalin (Sigma Aldrich, Oakville, ON) and fixed for 48h. Tissue samples were
embedded in paraffin and sections were processed and stained for hematoxylin and eosin (H&E,
Fisher) and Masson Trichrome (Fisher). Histology was performed by the Research Pathology
lab at Toronto General Hospital.
3.3.9 Statistical Analysis
All statistical analyses were performed using the software program STATISTICA Version
5.1 (Statsoft, USA). Student’s t-tests were used to compare the means between two groups
analyzed and considered significant at p<0.05.
43
3.4 Results
3.4.1 ASC Viability
The first step in demonstrating the efficacy of a poloxamine/collagen hydrogel system for
adipose tissue engineering was to investigate the viability of embedded ASC. A preliminary
alamarBlue® (AB) viability assay comparing collagen, poloxamine-only and
collagen/poloxamine hydrogels was conducted to assess the viability of the ASC in culture over
7 days (Figure 3-1). It was observed that cells in poloxamine and poloxamine/collagen
hydrogels had similar AB reductions over the first 3 days in culture as compared to the collagen
controls (with and without UV/initiator), suggesting that the fabrication procedure
(photocrosslinking) had little impact on cell viability. After 7 days, however, a 50% loss in the
AB reduction in the poloxamine-only hydrogel suggested cell death, likely the result of lack of
cell attachment to the matrix and death by anoikis. The viability of cells in the
collagen/poloxamine hydrogels was higher in comparison with the poloxamine-only hydrogel at
7d, yet lower when compared with collagen controls at day 7.
44
0
10
20
30
40
50
60
Control (collagen) Poloxamine Poloxamine + Collagen
% A
B re
duct
ion
1d
3d
7d
Figure 3-1: Alamar Blue® viability of ASC embedded in collagen (control), poloxamine-only and collagen/poloxamine hydrogels over 7 days
Low viabilities in the poloxamine-only hydrogel after 7 days was likely due to lack of cell attachment to the matrix. ASC in collagen/poloxamine hydrogels had improved viabilities compared with poloxamine-only hydrogels at day 7, but not as good as collagen-only control (n=3). Error bars represent 1SD.
A Live/Dead™ viability stain was used to visualize the viability of the embedded ASC
over 7 days in collagen and collagen/poloxamine hydrogels (Figure 3-2). High viability was
observed in both the collagen control and collagen/poloxamine hydrogels at day 1 indicating that
the fabrication and photocrosslinking process was likely non-toxic to cells. After 7 days
however, ASC in the collagen gels spread, adopted a typical fibroblast-like morphology and
appeared to have proliferated in the matrix. In the collagen/poloxamine hydrogel at day 7,
however, pockets of viable cells similar in appearance to those found in the collagen controls
were observed, but there were also regions of the hydrogel with a high number of dead cells.
This was thought to have been the result of inhomogeneity in the distribution of the collagen and
45
poloxamine phases in the hydrogel. Regions in the matrix rich in collagen would permit ASC
adherence and proliferation, while regions in the hydrogel composed of mostly poloxamine
would result in cell death over 7 days.
Figure 3-2: Live/Dead™ confocal microscopy images of ASC embedded in collagen and collagen/poloxamine hydrogels.
High viability of cells in both hydrogels at day 1 show that the fabrication process is non-toxic to cells. Two images of collagen/poloxamine at 7d show different regions of high and low cell viability in the same gel. This suggests inhomogeneity in the mixing of the collagen and poloxamine components (live cells = green, dead cells = red, scale bar = 100µm). Images are typical of 4 replicates.
Collagen
Collagen/ Poloxamine
1d 7d
46
3.4.1.1 Addition of Laminin to Improve ASC Viability
Several groups have investigated including laminin-derived peptide sequences into their
synthetic polymers to promote the attachment and proliferation of ASC[63,138]. Laminin (LN)
is a major component in adipose extracellular matrix and has been shown to promote the
adhesion, migration of ASC and may play an important role in adipogenesis[62]. It was of
interest to incorporate laminin into the collagen/poloxamine hydrogels to see if this had any
effect on ASC viability. Hydrogels of Collagen/Poloxamine/Laminin revealed similar results to
those obtained with collagen/poloxamine matrices (Figure 3-3). High cell viability throughout
the matrix was observed at day 1, with regions of viable and non-viable cells seen at day 7.
Interestingly, the addition of laminin appeared to have an effect on cell attachment. Even at day
1, cells in the hydrogel containing 50μg/mL LN appeared to be adopting a fibroblast-like
morphology and spreading onto the matrix. This was clearly seen for both the hydrogels
containing 5μg/mL and 50μg/mL laminin at day 7.
47
Figure 3-3: Live/Dead™ confocal microscopy images of ASC embedded collagen/poloxamine hydrogels with the addition of laminin.
ASC in the hydrogels with laminin appear to adopt a more elongated and fibroblast-like morphology indicative of adhesion and spreading onto the matrix (live cells = green, dead cells = red, scale bar = 100µm). Images are typical of 4 replicates.
3.4.2 Hydrogel Characterization
Immunostaining of collagen and laminin was used to visualize the distribution of the
components in the hydrogels to help explain the observed pockets of live and dead cells in the
matrix after 7 days (Figure 3-4). Collagen is stained red, laminin green and poloxamine is not
stained and appears black. Large black regions void of biological components are observed in
the collagen/poloxamine/laminin hydrogels. Laminin association with collagen is apparent at
higher magnification as both red and green colours are colocalized. Higher magnification
1d
7d
0µg/mL LN 50µg/mL LN 5µg/mL LN
48
images of collagne/poloxamine show that black regions devoid of biological components span
diameters greater than 100µm (Figure 3-5).
Figure 3-4: Immunostaining of Collagen and Collagen/Poloxamine/Laminin hydrogels.
Immunostaining enables visualization of the distribution of components in the collagen/poloxamine/laminin hydrogel. Collagen is stained red, laminin green. Poloxamine is unstained and appears black. Images are typical of 4 replicates.
Collagen
Collagen/ Poloxamine/Laminin
49
Figure 3-5: High magnification images of immunostained collagen/poloxamine hydrogels.
Higher magnification images of immunostaining of collagen/poloxamine hydrogels shows collagen (red) and large areas of poloxamine (black) greater than 100µm in diameter. Images are typical of 4 replicates.
3.4.3 ASC Differentiation
3.4.3.1 Oil Red O
The next step in demonstrating the efficacy of a poloxamine/collagen hydrogel system for
adipose tissue engineering was to investigate the differentiation capacity of the embedded ASC.
Three hydrogel configurations were tested: poloxamine, collagen/poloxamine and
collagen/poloxamine/laminin (50μg/mL LN) and compared against ASC on TCPS as a control.
As seen in Figure 3-6, in the poloxamine (POL) hydrogels lipid accumulation (red) was seen in
the majority of cells after 7 days of differentiation. However, lipid accumulation appeared to
halt after 7 days with no further increase in lipid droplet size over time and only very few nuclei
appeared to stain for Hoescht (blue) suggesting that most of these cells had died after 21 days.
Collagen/poloxamine (COL POL) and collagen/poloxamine/laminin (COL POL LN) hydrogels
diplayed intracellular lipid accumulation at day 7, with an increase in the size of the fat droplets
50
at day 21 indicating a more-differentiated phenotype. Differentiating cells were found to cluster
in regions rich in collagen within the hydrogels (green by immunostaining), and only very few
cells stained positive for hoescht (blue) outside of these regions, similar to what is seen in POL
hydrogels. No observable difference was seen in the hydrogels with laminin compared to those
without. These results indicate that poloxamine hydrogels did not support ASC differentiation,
likely because of cell death due to anoikis since this hydrogel did not support cell attachment.
Collagen/poloxamine hydrogels appeared to support ASC differentiation with levels similar to
that of collagen gels, yet only in regions of the hydrogel rich in collagen.
Figure 3-6: ASC differentiation in collagen, poloxamine and collagen/poloxamine hydrogels
Oil Red O staining (red) of ASC embedded in collagen (COL) poloxamine (POL), collagen/poloxamine (COL POL) and collagen/poloxamine/laminin (COL POL LN) hydrogels cultured with adipogenic medium at 7 and 21 days. Hoescht stains cell nuclei blue. Immunostaining of collagen is green. Scale bar = 20μm. Images are typical of 4-6 replicates.
3.4.3.2 GPDH Activity
The overall GPDH expression for each sample is shown in Figure 3-7 with the data
normalized for total number of cells in Figure 3-7. ASC in all samples appear to be expressing
COL POL COL POL COL POL LN
7d
21d
51
GPDH at all time points investigated with the ASC in collagen gel samples at 72h expressing a
statistically significant higher amount of total GPDH compared to cells on TCPS or the other
hydrogel configurations. When normalized for the total number of cells, cells in the collagen
gels at 72h and 7d expressed a statistically significant lower amount of GPDH (ie. GPDH per
cell) compared to all other samples.
Overall GPDH
0
5
10
15
20
25
TCPS COL POL COL POL COL POL LN
mU
72h7d14d
Figure 3-7: GPDH activity of ASC during differentiation. GPDH activity in mU per sample (not normalized for cell number) of each hydrogel group at 72h, 7d and 14d after the induction of differentiation (n=4). Collagen gels expressed a significantly higher total amount of GPDH than other samples at 72h (p<0.05). Error bars represent 1 SD.
*
52
Overall GPDH (normalized with GAPDH)
0
24
68
1012
1416
18
TCPS COL POL COL POL COL POL LN
mU
/GA
PDH
72h7d14d
Figure 3-8: GPDH activity of ASC during differentiation normalized for GAPDH expression.
GPDH activity in mU per sample, normalized for the number of viable cells by GAPDH expression (western blot), of each hydrogel group at 72h, 7d and 14d after the induction of differentiation (n=4). Collagen gels expressed a significantly lower amount of GPDH than other samples at 72h when normalized for total cell number (p<0.05). Error bars represent 1 standard deviation.
3.4.4 Endothelial Cell Attachment to Gels and Modules
As expected, HMEC-1 on either TCPS or collagen gel films (COL) quickly formed a
confluent layer of cells covering the surface of the films (Figure 3-9). HMEC-1 on the surface of
the poloxamine-only hydrogel (POL) appeared in rounded clusters on day 1, indicative of a non-
adhesive substrate and very few cells were found to be attached to the poloxamine-only hydrogel
film by day 14. The incorporation of collagen in the collagen/poloxamine (COL POL)
hydrogels greatly improved the attachment of the HMEC-1, yet complete coverage of the entire
surface of the film was not achieved. These holes or gaps in the surface coverage of the cells
were also observed in the collagen/poloxamine hydrogels with laminin (50μL/mL LN) (COL
POL LN). No improvement in HMEC-1 attachment to the hydrogels with laminin was observed,
likely because laminin self-associates with the collagen phase in the gels. Similar results were
found for the attachment of HMEC-1 to the surface of the collagen/poloxamine modules (Figure
* *
53
3-10). Here the gaps in the surface coverage appear larger, likely due to the curved geometry of
the cylindrical module.
The lack of complete HMEC-1 coverage on the surface of the collagen/poloxamine
hydrogel was consistent with the non-uniform distribution of collagen and poloxamine within the
hydrogel (Figure 3-5). Regions that are devoid of HMEC-1 are likely regions of the gel rich in
non-cell adhesive poloxamine, while regions that are rich in collagen promote HMEC-1
attachment.
Figure 3-9: HMEC-1 attachment to poloxamine hydrogel films.
Live/Dead™ images show attachment of HMEC-1 to collagen (COL), poloxamine-only (POL), collagen/poloxamine (COL POL) and collagen/poloxamine with laminin (COL POL LN) hydrogels. Lack of HMEC-1 surface attachment is clearly observed on the POL hydrogel. HMEC-1 attachment was improved with the COL POL hydrogel yet complete surface coverage is not achieved after 14 days as compared to HMEC-1 cultured on collagen gels. The addition of laminin did not appear to improve HMEC-1 attachment. Images are typical of 4-6 replicates.
TCPS POL COL POL COL POL LN
1d
7d
COL
14d
54
Figure 3-10: HMEC-1 attachment to poloxamine modules.
Live/Dead™ images show attachment of HMEC-1 to collagen/poloxamine (COL POL) and collagen/poloxamine with laminin (COL POL LN) modules. Holes or gaps in HMEC-1 coverage are likely the result of inhomgeneous distribution of the collagen/poloxamine components. Images are typical of 4 replicates.
3.4.5 Surface Modifications to Improve Cell Attachment
Live/Dead™ images of HMEC-1 seeded onto the FN-modified modules display a similar
pattern of attachment as unmodified collagen/poloxamine modules, with incomplete coverage of
the module surface after 14 days (Figure 3-11). Again, due to the inhomogeneous mixing of the
collagen/poloxamine components, regions of the module surface rich in poloxamine (PEG-based
material) likely resisted protein adsorption and as such did not aid in promoting universal
HMEC-1 attachment over the entire surface.
1d 7d 14d
COL POL
COLPOLLN
55
Figure 3-11: HMEC-1 attachment to modules with fibronectin treatment.
Live/Dead™ images show attachment of HMEC-1 to collagen/poloxamine modules modified with surface adsorbed FN show no improvement in HMEC-1 attachment. Regions of the module surface rich in poloxamine likely resisted FN adsorption. Images are typical of 4 replicates.
To overcome the issue with lack of protein adsorption to regions of the surface rich in
poloxamine, the next strategy employed was coating the modules in a collagen solution, and
gelling the collagen to create a layer of collagen gel surrounding individual modules.
Live/Dead® images of the HMEC-1 seeded onto collagen-coated modules demonstrated very
high initial attachment at day 1 for all the collagen concentrations tested (Figure 3-12). Yet, over
time the attached cells appeared to proliferate and contract the surrounding collagen gel, pulling
it off the surface of the modules, since the surrounding collagen gel was not attached well to the
module surface. This appeared as highly fluorescent regions on the surface of the modules that
seemed to detach from the module surface at day 7 and 14.
1d 7d 14d
56
Figure 3-12: HMEC-1 attachment to modules with collagen coating.
Live/Dead™ images show attachment of HMEC-1 to collagen/poloxamine modules coated with different concentrations of collagen gel. Initial attachment of the cells was good (day 1) but over time proliferating cells were found to detach from the modules. Images are typical of 4 replicates.
3.4.6 In Vivo Implantation
As a pilot study to test the feasibility of exploring the collagen/poloxamine hydrogel in
vivo, we implanted modules made from collagen/poloxamine into SCID mice using the same
model for the implantations as seen in Chapter 4. A large infiltration of neutrophils was
observed in collagen/poloxamine module implants at day 3 suggesting a highly inflammatory
reaction in vivo (Figure 3-13). This was the first implantation of poloxamine in vivo and a
biocompatibility study is needed. Perhaps additional washing steps are required to remove
residual chemicals from the synthesis steps or initiator from UV photocrosslinking.
0.75mg/mL 3mg/mL 1.5mg/mL
1d
7d
14d
57
Figure 3-13: Collagen/Poloxamine modules implanted subcutaneously in SCID mice.
H&E and Masson’s Trichrome photomicrographs of collagen/poloxamine modules at day 3. Many neutrophils are surrounding the modules suggesting a large inflammatory reaction (n=3). Scale bar 100µm.
H&E Trichrome
58
3.5 Discussion
There is a need for a suitable biomaterial for adipose tissue engineering that has the
appropriate mechanical properties combined with bioactive cues that will enable ASC
attachment and differentiation into mature adipocytes, with the goal of creating a volume-stable
fat pad. Hydrogels based on synthetic polymers are advantageous in that they can be tailored
based on their physicochemical properties to suit a specific application. Hydrogels made from
poly(ethylene oxide) and poly(proplylene oxide) block copolymers (PEO-PPO) have been
extensively explored as a biomaterial because of their intrinsic biocompatibility, low protein
binding and hydrophilicity[133]. Yet, these properties make cell attachment difficult. Here we
report on the feasibility of using a semi-synthetic collagen/poloxamine hydrogel for adipose
tissue engineering. This network consists of 2 components; first the poloxamine provides the
crosslinks to form the hydrogel while collagen is a non-binding element that provides improved
biological properties.
We found that the collagen/poloxamine is not a suitable biomaterial for adipose tissue
engineering, at least without further modification. While embedded ASC viability was higher in
collagen/poloxamine hydrogel as compared to the poloxamine-only case, it was not as high as
the collagen-only control. Even with the addition of laminin in an attempt to improve embedded
ASC viability, Live/Dead® staining revealed pockets of both viable and dead cells throughout the
collagen/poloxamine hydrogel. With regions greater than 100µm in diameter devoid of any
biological components, the heterogeneous nature is likely the cause of the regions of non-viable
cells observed after 7 days; these cells were likely in a region rich in poloxamine and were
unable to attach to the PEO-rich matrix. The immiscibility of the collagen and poloxamine and
the resultant inhomogeneous distribution of the components within the hydrogel was likely the
59
result of entropy-driven phase separation between poloxamine and collagen. A similar problem
was encountered in a recent report of a Pluronic F-127 hydrogel for adipose tissue engineering
where collagen was added to promote cell attachment and spreading. Even the addition of a very
small amount of collagen resulted in a single cluster of collagen and cells within the matrix[139].
Oil Red O staining revealed extensive lipid accumulation in the collagen-only hydrogels,
as expected. Some lipid accumulation was observed by cells in the poloxamine-only hydrogel at
day 7, yet very few cells stained positive for Hoescht, suggesting that most of the embedded cells
had apoptosed by this time. Oil red O staining was extensively found in the collagen/poloxamine
and collagen/poloxamine/laminin hydrogels, but only in the regions that also stained positive for
collagen. No observable improvement was seen in the hydrogels with laminin, likely since the
laminin and collagen were colocalized. It appears that initial ASC attachment to the matrix is
critical for cell viability, and differentiation.
Glycerol phosphate dehydrogenase (GPDH) activity demonstrated some level of ASC
differentiation within all hydrogels tested, with collagen hydrogels giving the highest amount of
GPDH per sample, likely due to the highest number of viable cells. When the GPDH activity
was normalized for cell number, interestingly collagen gels at 72h and 7d expressed a
statistically significant lower amount of GPDH (ie. GPDH per cell) compared to all other
samples. This may be explained by the fact that ASC in the collagen gels were readily
proliferating at the onset of differentiation, while cells on TCPS had reached confluence and
cells in the poloxamine based materials are less likely to proliferate due to the non cell adhesive
nature of the gel. Cells at confluence or which are unable to proliferate due to lack of cell
attachment have reached a state of terminal differentiation which is a prerequisite for
adipogenesis. This has been seen in studies examining the effect of cell density and cell shape
60
on lineage commitment of mesenchymal stem cells (MSC). MSC were seen only to differentiate
into adipocytes if they were plated on a small island of fibronectin which forces them into an
unspread and contracted morphology[140]. While the total GPDH activity from the collagen
gels may be higher, the actual fraction of cells undergoing differentiation may be lower than that
of ASC on TCPS or in poloxamine based hydrogels since many ASC were still likely
proliferating. This again hints to the possibility that an optimized biomaterial for adipose tissue
engineering will combine both a cell adhesive nature for initial attachment, spreading and
proliferation and a non-adhesive property to enhance ASC differentiation.
In the case of modular tissue engineering, not only is the viability and differentiation of the
embedded ASC important, but the biomaterial must enable EC attachment to the surface. We
found that HMEC-1 were able to attach and proliferate onto both the surface of
collagen/poloxamine hydrogel films and modules yet complete EC coverage was not achieved
over 14 days in culture. Again, the inhomogeneity of the semi-interpenetraing network likely
resulted in regions of the hydrogel surface devoid of any biological components (ie. collagen or
laminin). Modification of the module surface with either fibronectin or collagen gel coating did
not appear to be successful in creating a confluent surface layer of EC. Complete EC surface
coverage may not be critical for in vivo applications if we think of the modules simply as a
delivery vehicle for cell transplantation. Perhaps transplanted EC will remodel into vessels in
vivo regardless of whether or not they are they are in a quiescent confluent layer on the module
surface. As long as a sufficient number of EC have been transplanted this may be enough for the
creation of a vascular network in vivo. However, the inability of the collagen/poloxamine
hydrogel to support high ASC viability and differentiation makes it unsuitable for applications in
adipose tissue engineering. Perhaps, a better approach would be to chemically modify the
61
poloxamine to contain bioactive groups (e.g. YIGSR) on the polymer backbone to improve
cellular compatibility with the synthetic hydrogel.
3.6 Conclusions
The collagen/poloxamine hydrogel is not a suitable biomaterial for adipose tissue
engineering. The inhomogeneous nature of the hydrogel resulted in low viability and
differentiation of embedded ASC and incomplete EC coverage on the surface. Furthermore an in
vivo study showed that collagen/poloxamine modules caused a large inflammatory reaction. It
appears that further chemical modification of poloxamine is needed to make the hydrogel more
cell-compatible. Since this is not within the scope of this project, collagen gel was used as the
module material when studying the effect of the ASC/HMEC interaction in vivo since it avoided
any biomaterial-related complications.
3.7 Acknowledgements
The authors acknowledge the financial support by NSERC, the gift of Tetronic 1107 from
BASF and Dr. Alejandro Sosnik for his help and guidance with polymer synthesis.
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4 Cotransplantation of ASC and HMEC in a Modular Construct Drives Early Angiogenesis, ASC differentiation and Long‐term Fat Regeneration in SCID Mice
4.1 Abstract
A modular approach to adipose tissue engineering was explored by embedding adipose-
derived stem cells (ASC) in sub-mm sized collagen rods or ‘modules’ and coating with
microvascular endothelial cells (HMEC). When hundreds of these modules were injected
subcutaneously into a SCID mouse, HMEC on modules containing embedded ASC appeared to
detach from the modules to form vessels as early as day 3 as confirmed by the human EC
specific UEA-1 lectin stain, and these vessels persisted for up to 90 days. Vessels numbers
decreased over 14 days, but vessel size increased suggesting a maturing of the vasculature.
Vessel perfusion with the host was confirmed at 21 days by microCT. HMEC on modules
without embedded ASC remained attached to the module surface at day 3 and UEA-1 staining
disappeared over 14 suggesting cell death. It appeared that cotransplantation with ASC had an
anti-apoptotic and angiogenic effect on HMEC. The early revascularization strategy was
successful in supporting ASC viability and differentiation, as progressive fat accumulation in the
HMEC+ASC implants was observed: ~60% of the implant area stained positive for Oil Red O
by day 90. ASC-embedded modules without HMEC surface coating did not show fat
accumulation within the implant. All implant volumes decreased over the time-course of the
experiment, yet HMEC+ASC module implants were larger than ASC-only implants at day 90.
Collagen gel is mechanically weak and contracts in vivo making it unsuitable as a biomaterial for
adipose tissue engineering where volume maintenance is critical. When combined with an
appropriate biomaterial, the modular approach to adipose tissue engineering may represent a
successful strategy to engineer soft tissue substitutes of clinical relevance.
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4.2 Introduction
In a typical approach, adipose tissue is reconstructed by combining adipose-derived stem
cells (ASC) with an appropriate substrate, typically a polymeric scaffold. Seeding of adipose
precursor cells, onto synthetic (ie. PLGA) and biodegradable scaffolds (ie. collagen) has led to
adipose tissue formation in vivo[54,58,67]. While many of these approaches to adipose tissue
engineering have been successful in initially generating adipose tissue in vivo, they have failed to
maintain the volume of the regenerated tissue over a long-implantation period (> 3 months)[141].
Native adipose tissue is highly vascular, with resting values of blood flow and capillary filtration
coefficients two to three times higher than those in skeletal muscle. As a result, a critical
challenge in engineering large portions of adipose tissue is early vascularization of the
engineered tissue upon implantation to avoid considerable cell death. A successful adipose
tissue engineering strategy will promote angiogenesis soon after implantation to facilitate
transplanted cell survival.
Here, functional cells (adipose-derived stem cells (ASC)) were embedded in small
collagen hydrogel rods (0.5mm diameter, 1mm length) coated with a microvascular endothelial
cell layer (HMEC). Implanting many modules together is capable of filling a soft tissue defect
of any geometry, with the idea that the endothelialized lining of the modules will quickly
remodel into vessels and anastomose with the host vasculature upon implantation to promote
embedded ASC viability and differentiation into fat.
The aim of the Angiogenesis & Remodeling study (day 3, 7, 14, 21) was to study the fate
of the endothelialized lining of the ASC-embedded modules to see if they have the potential to
remodel rapidly and to anastamose with the host vasculature upon implantation to avoid
ischemic conditions. In the Fat Development & Volume Maintenance study (day 30, 60, 90),
64
longer time points will be investigated to determine if the modular approach to adipose tissue
engineering can produce volume-stable adipose tissue in vivo.
4.3 Materials and Methods
4.3.1 Cells
Primary human adipose-derived stem cells (ASC) and human microvascular endothelial
cells (HMEC) were harvested from sterile abdominal fat samples were obtained from patients
undergoing elective surgery, with consent. This procedure is based on the protocol for the
isolation and culture of human adipose-derived stem cells (ASC) from Dr. Lauren Flynn,
Queen’s University as described in section 3.3.1 with modifications to include magnetic bead
separation for HMEC. The tissue was minced and digested in a 2mg/mL collagenase solution at
37oC, containing Kreb’s Ringer bicarbonate buffer, supplemented with 3mM glucose, 25mM
HEPES and 20mg/mL BSA (all Sigma, Oakville, ON). Upon digestion the sample was filtered
to remove undigested tissue fragments and centrifuged to separate the desired stromal-vascular
cell population from the buoyant mature adipocytes. The cell suspension was gently agitated in
erythrocyte lysing buffer and filtered again to remove red blood cell fragments. Several
centrifugation steps with washes in complete media finished the processing. Half of the resultant
cell pellet was plated for ASC culture in a 1:1 mixture of Dulbecco’s Modified Eagle’s Medium
and Ham’s F-12 nutrient mixture (DMEM:Ham’s F-12) (Sigma, Oakville, ON), supplemented
with 10% fetal bovine serum (FBS) (Sigma, Oakville, ON), 100 U/mL penicillin and 0.1 mg/mL
streptomycin (Gibco, Burlington, Canada), while the other half was plated in MCDB-131 (VEC
Technologies) for HMEC purification. HMEC were purified from contaminating ASC by CD31
positive selection using MiniMacs™ magnetic bead separation columns (Miltenyi Biotech,
65
Auburn, CA). The conditions for successful HMEC separation are outlined in Appendix 1.
Primary Human Umbilical Vein Endothelial Cells (HUVEC, Lonza, Walkersville, MD), cultured
in EGM-2 culture medium (Lonza), were also used in one set of implantations to compare
different types of EC.
The growth medium on the cells was changed every 2-3 days. To passage the cells
cultures at 90% confluence were trypsin-released (0.25% trypsin/0.1% EDTA, Gibco,
Burlington, Canada), washed, counted and re-plated in new flasks at 30,000 cells/cm2. ASC at
passage 2-3 and HMEC at passage 4-6 were used for in vivo implantations. In the one
experiment using HUVEC, these cells were used at passage 4.
4.3.2 Module Fabrication
To prepare modules, bovine Type 1 collagen (3mg/mL, PureCol™, Inamed, Fremont,
CA) was mixed with 10X MEM (10.1 g α-Minimum Essential Medium powder (Gibco,
Invitrogen, Paisley, PA), and neutralized to pH ~ 7.4 by adding the required volume of 0.8N
NaHCO3. ASC (106 cells/mL) were added to the collagen solution to prepare ASC-embedded
modules. This solution was cast into the inner diameter of sterile 0.71mm ID polyethylene
tubing (PE 60, Intramedic – BD Canada, Oakville, ON) and subsequently gelled for 60 minutes
at 37oC[142]. The tubing was cut into small pieces (~ 2mm long x 0.6mm diameter) using a
custom automatic tube cutter and collected in DMEM:F12 Ham’s media, where the modules
were separated from the tubing by vortexing and cultured for 72h at 37oC in a petri dish to allow
contraction of the modules by ASC. 1mL of settled modules (produced using 3m of tubing)
with or without embedded ASC, were seeded dynamically with HMEC or HUVEC (2 x 106) for
45min on a low speed shaker and incubated overnight in a 50/50 mixture of DMEM:F12 Ham
and EGM-2. ASC-only modules were prepared as above without endothelial cell seeding.
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4.3.3 Module Implants
Adult male (6 weeks of age) male SCID/Bg mice were purchased from Charles River
Laboratories (Wilmington, MA). They were individually housed in sterile cages and fed ad
libitum under the approval by the University of Toronto animal care committee. Three different
treatment groups were studied; 1. Modules coated with EC, without embedded ASC; 2. Modules
embedded with ASC and coated with EC; 3. Modules with embedded ASC, without EC.
Approximately 250 modules suspended in PBS were injected subcutaneously using an 18G
needle on the dorsum of mice. The mice were euthanized at various timepoints (3, 7, 14, 21, 30,
60, 90days) and the implants excised and either fixed in 4% neutral buffered formalin (Sigma
Aldrich, Oakville, ON) for 48h or placed in Tissue-Tek® O.C.T. compound and flash-frozen in
liquid nitrogen for frozen tissue processing. Sections were processed and stained for
hematoxylin and eosin (H&E, Fisher) and Masson Trichrome (Fisher) and various antibodies
outlined in Table 1. Histology was performed by the Pathology Research laboratory at Toronto
General Hospital. All sections were viewed with a Zeiss Axiovert light microscope equipped
with a CCD camera.
Table 1: Histology stains.
Stain Preparation Objectives Hematoxylin & Eosin (H&E) Masson’s Trichrome
Paraffin-embedded Implant Integration Cellular Infiltration Morphology Inflammatory Response Fibrous capsule formation Locate modules
UEA-1 Lectin Paraffin-embedded Stain transplanted HMEC CD31 Paraffin-embedded Stain host (mouse)
vasculature
Oil Red O Frozen Stain fat
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4.3.4 Histology Quantification
UEA-1 stained positive vessels at day 3, 7 and 14 were counted using a Microvessel
Density Count (MVD) method adapted from the tumor angiogenesis literature[143]. At low
power (2.5X Objective lens), the number and diameter of UEA-1 stained vessels in the implants
made of HMEC coated modules with and without embedded ASC were quantified using the
ImageJ (version 1.4.3) software program provided by the National Institutes of Health. At 2.5x
objective lens the entire section of an implant could typically be visualized and the total number
of vessels per objective field (or section) was counted (not hotspots). A total of 3 sections were
counted to give an average vessel count per implant. A total of 8 animals (8 implants) were
counted for each treatment with a total of three separate experiments conducted (n=8, N=3). The
presence of a patent lumen or presence of erythrocytes was not a prerequisite for the definition of
a microvessel, yet at this low magnification only larger vessel-like structures could be counted.
MVD counts were expressed as the number of vessels per mm2 of implant.
The level of fat development in implants made from ASC-embedded modules with and
without HMEC surface coating at 30, 60 and 90d was quantified using ImageJ. Briefly, low
magnification images (2.5X objective) of Oil Red O histological sections from 3 different
animals (3 sections per implant, n=3) at each time point and treatment group were taken. Again
at this low magnification the total area of the section containing the module implant could be
visualized. The area that was stained red by Oil Red O was calculated by the software and was
divided by the total area of the implant. The result was given as a % Area of the Implant stained
positive for Oil Red O.
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4.3.5 Perfusion Studies
The level of vascular perfusion in the module implants was studied by microCT analysis
at 21 days. Following a published protocol[144], mice were first heparinized (100 units, LEO
Pharma Inc.) by subcutaneous injection for 5 minutes before the procedure. Animals were then
anesthetized with isofluorane and the descending aorta was cannulated and heparinized PBS
(5U/mL) was perfused at a constant pressure of 100 mmHg to flush away blood from the
vascular system. The mice were then perfused with 10mL of Microfil® solution (MV-122,
Flow-tech, Carver, MA) and the solution was allowed to polymerize for 2 hours. Implants were
excised into 4% formalin (Sigma Aldrich), embedded into 1% agar and visualized with a General
Electric Medical Systems MS8 microCT.
4.3.6 Volume Measurements
To assess implant volume maintenance over time, for the 30, 60 and 90d implants
precisely 0.2mL (40mm3) of settled modules in PBS was injected into each implant site. At
explant, the dimensions of each implant was carefully measured with calipers and the volume
estimated using the volume equation of an ellipsoid.
4.3.7 Statistical Analysis
All statistical analyses were performed using the software program STATISTICA Version
5.1 (Statsoft, USA). Student’s t-tests were used to compare the means between two groups
analyzed and considered significant at p<0.05.
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4.4 Results
4.4.1 Angiogenesis & Remodeling Study (Day 3, 7, 14, 21)
In the Angiogenesis and Remodeling Study the focus was to study the effect of embedded
ASC on the viability and angiogenic potential of the surface coated HMEC when implanted
subcutaneously in SCID mice at early time points (3, 7, 14, 21d). Implants containing modules
made with both surface coated HMEC and embedded ASC (HMEC + ASC) were compared
against implants made from modules without embedded ASC but with surface HMEC (HMEC-
only) and modules with only embedded ASC (ASC-only) as the controls. The results presented
here demonstrate a strong synergistic angiogenic response between the embedded ASC and
surface seeded HMEC.
4.4.1.1 Appearance at Explant
Upon visual inspection at explant, the HMEC+ASC module clump appeared to be red
and vascularized by day 7 and 14 with what appeared to be blood vessels throughout the volume
of the implant (Figure 4-1). In contrast, the HMEC-only and ASC-only controls appeared to be
opaque and avascular.
70
Figure 4-1: Photographs of the modules at explant day 7 and 14.
The HMEC+ASC explant appears red, with blood vessels visible throughout the module clump. Both the HMEC-only and ASC-only explants appear opaque and avascular.
4.4.1.2 Effect of ASC on HMEC Viability and Blood Vessel Formation
One of the goals of this experiment was to show that the embedded ASC may improve
viability of transplanted HMEC and also to see if the embedded ASC would elicit an angiogenic
response from the HMEC. Both of these were observed. As seen in Figure 4-2 , the HMEC
surrounding the modules containing ASC were found to detach from the module surface and
organize into vessels. At day 3, the vessels appeared to be disorganized and immature, but by
day 7 and 14 they appear to be organized, mature and large (diameter 10-40µm) with many
containing erythrocytes suggesting anastomosis with the host. This is in contrast to the
HMEC+ASC HMEC-only ASC-only
7d
14d
71
Figure 4-2: UEA-1 Lectin stained sections of HMEC+ASC modules and HMEC-only modules at day 3, 7 and 14.
The presence of ASC causes the HMEC to detach from the module surface to form vessels. HMEC on modules without ASC remain on the module surface and the UEA-1 staining diminishes over time suggesting cell death. Arrows highlight some large UEA-1 stained vessels containing erythrocytes, with some modules highlighted by dashed lines (n=8). Scale bar 100µm.
HMEC+ASC HMEC-only
3d
7d
14d
72
HMEC-only control where the HMEC would remain on the module surface and staining would
diminish by day 7. By day 14 virtually no UEA-1 staining could be found in the control case
suggesting HMEC death without the presence of the ASC.
Higher magnification of serial sections of the HMEC+ASC modules revealed blood
vessels surrounding modules positive for UEA-1 containing erythrocytes as early as day 3
suggesting rapid angiogenesis and anastomosis upon implantation (Figure 4-3). These blood
vessels containing erythrocytes were positive for UEA-1 and negative for mouse CD31
confirming that they originate from the implanted HMEC (Figure 4-4).
73
Figure 4-3: Magnified images of serial sections of the tissue surrounding a single module at day 3.
UEA-1+ blood vessels containing erythrocytes are observed surrounding the module, suggesting anastomosis as early as day 3 (n=8). The location of the module is highlighted by a dashed line. Scale bar 100µm.
H&E Trichrome
UEA-1
74
Figure 4-4: Magnified images of serial sections of a single blood vessel surrounding a module at day 3.
The vessel stains positive for UEA-1 but not CD31 proving human origin. CD31 successfully stains other host derived vessels surrounding the implant (bottom right panel) (n=8). Arrow points to location of vessel. Scale bar 100µm.
MVD counts (Figure 4-5) from HMEC+ASC implants show that initially a large number
(>150 UEA-1+ vessels) are present in the area of the implant. Most of these vessels appeared
unorganized and are small (~50µm in diameter). By day 14, the total number of UEA-1+ vessels
had diminished considerably but the vessels that remained were large (~200µm in diameter).
These results suggest a typical angiogenic response that begins with a large number of small
immature vessels that organize into larger functional vessels.
H&E UEA-1
CD31 (mouse)
CD31 (mouse)
75
Figure 4-5: MVD counts from HMEC+ASC implants at day 3, 7 and 14.
A large number of small vessels are observed at early time points, while by day 14 the number of vessels has decreased the average vessel diameter is larger. Error bars represent ±1SD. (n=8, N=3).
4.4.1.3 Host Tissue Infiltration and Angiogenesis
As expected, it appeared that the ASC also had an effect on the angiogenic response of
the host tissue to the implanted modules. Host tissue found infiltrating the HMEC+ASC
modules was rich in CD31 positive blood vessels (Figure 4-6), while tissue invading the HMEC-
only module clump contained only a minimal number of host derived vessels. As can be seen in
Figure 4-7, both HMEC (UEA-1+) and host derived (CD31+) blood vessels surround the
modules by day 14. The embedded ASC appear to both stimulate the transplanted HMEC to
rapidly form vessels and to anastamose with the host at very early time points (day 3) while
promoting host vessel formation by day 14. The former will help avoid ischemic conditions
upon implantation while the latter will further nourish the engineered tissue during
differentiation and remodeling into a fat pad.
76
Figure 4-6: CD31 images of host tissue infiltration into the modular implant at day 14. Many CD31+ blood vessels are seen in the HMEC+ASC module implant by day 14 showing host tissue infiltration. Dashed lines highlight some modules (n=8). Scale bar 100µm.
Figure 4-7: Magnified images of serial sections of tissue surrounding several modules.
Both UEA-1+ (arrows) and CD31+ (stars) blood vessels are observed surrounding the modules at day 14. Dashed line highlights module location (n=8). Scale bar 100µm.
HMEC+ASC HMEC-only
UEA-1 CD31
Trichrome H&E
77
Perfusion of the module implants at day 21 was confirmed by microCT imaging (Figure
4-8). The results show an image of the implant (region marked by dashed line) under the mouse
skin. In the HMEC+ASC sample blood vessels can be seen branching off from a large vessel in
the skin infiltrating the region of the implant. Two large pools of microfil were seen within the
implant and this may be the result of leaky vessels or vessel rupture caused due to high pressure
during perfusion. The HMEC-only sample remained avascular with only blood vessels in the
skin visible. Microfil perfusion does not distinguish between host and donor derived vessels and
the perfusion seen here may be a combination of both HMEC-derived and mouse vasculature.
Figure 4-8: MicroCT images of HMEC+ASC (A) and HMEC-only (B) module implants under mouse skin at day 21. Many blood vessels can be seen infiltrating the HMEC+ASC implant (A), while the HMEC-only implant (B) remains avascular (n=2). Dashed lines show region of implant.
4.4.2 Implants with HUVEC
With the success of the implants with the primary HMEC we were curious to see if the
ASC would elicit the same angiogenic response with another EC cell type. As with the HMEC,
HUVEC were seen to quickly detach from the modules containing ASC and form vessels, while
A B
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HUVEC on modules without ASC were found to remain on the module surface with a decrease
in UEA-1 staining over time suggesting cell death (Figure 4-9). In contrast with the HMEC
experiments, some small HUVEC vessels could be observed by day 14 in the case without ASC.
It appears the ASC accelerate the angiogenic response and increase vessels by both size and
number. We conclude that the angiogenic response seen in the experiments with HMEC was not
a special case (ie. due to both ASC and HMEC being harvested from the same donor) and that
ASC have an angiogenic effect on different types of EC on modules in vivo.
Figure 4-9: UEA-1 Lectin stained sections of HUVEC+ASC modules and HUVEC-only modules at day 3, and 14. Similar to the HMEC, ASC causes HUVEC to detach from the module surface to form vessels. HUVEC on modules without ASC remain on the module surface and the UEA-1 staining diminishes over time suggesting cell death. Unlike HMEC, some small HUVEC vessels are present surrounding modules without ASC at day 14. Some modules highlighted by dashed lines. Arrows highlight some large UEA-1 stained vessels containing erythrocytes (n=4). Scale bar 100µm.
ASC + HUVEC HUVEC-only
3d
HUVEC+ASC
14d
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4.4.3 Fat Development and Volume Maintenance Study (Day 30, 60, 90)
Many approaches to adipose tissue engineering in the literature have been successful in
initially generating adipose tissue in vivo, but have failed to maintain the volume of the
regenerated tissue over a long-implantation period (> 3 months). We hypothesized that the early
angiogenesis from the cotransplantation of ASC and HMEC in the modules will nourish the
transplanted tissue to promote ASC differentiation into mature fat and limit resorption over time
to maintain implant volume. In this in vivo study we focused on:
1) Characterizing fat development in HMEC+ASC and ASC-only modules over longer
time periods (30, 60 and 90 days).
2) Characterizing the persistence of HMEC-derived blood vessels over time.
3) Comparing the volume measurements of the HMEC+ASC and ASC-only modules at
explant to monitor volume maintenance.
4.4.3.1 Fat Development
Oil Red O staining on frozen sections of the modules at explant revealed progressive fat
accumulation in the HMEC+ASC modules over the 30, 60 and 90 days (Figure 4-10). By 30
days, no fat was stained in either the HMEC+ASC or ASC-only control case. By 60 days, much
red staining was seen in the region surrounding the HMEC+ASC module implant with pockets
of red staining within the module clump suggesting fat accumulation. This is confirmed with
higher magnification images of H&E and Masson’s Trichrome slides which show clusters of fat
cells within individual modules (Figure 4-11). This suggests that at least some of the fat
accumulation seen here is attributed to the transplanted ASC differentiating into mature fat cells.
In the ASC-only case a few small pockets of fat could be observed by day 60, but much less than
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Figure 4-10: Oil Red O images of frozen sections from HMEC+ASC and ASC-only module implants at 30, 60 and 90d post implantation.
Progressive fat accumulation can be observed by the increase in red staining in the HMEC+ASC module implants over time. Little to no Oil Red O staining was seen in the ASC-only case (n=3). Scale bar = 100μm.
30d
90d
60d
HMEC+ASC ASC-only
81
what was observed in the HMEC+ASC case. By the 90 day time point, extensive Oil Red O
staining was seen throughout the region of the HMEC+ASC module clump indicating high level
of fat differentiation and/or accumulation. Again, Masson’s Trichrome images confirm this fat
accumulation where extensive clustering of adipose cells is seen within the implant (Figure
4-12).
Figure 4-11: H&E and Masson’s Trichome micrographs of HMEC+ASC modules at day 60.
Arrows highlight clusters of adipose cells within the individual HMEC+ASC modules (n=5). Scale bar = 100μm.
Figure 4-12: Masson’s Trichrome micrograph of HMEC+ASC modules at day 90.
The left panel shows a low magnification image of the implant (highlighted by a dashed line). Scale bar = 250μm. The right panel represents the boxed area and shows extensive fat accumulation within the implant reagion (n=3). Scale bar = 100μm.
H&E Trichrome
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The level of fat accumulation within the implants was quantified by analyzing the Oil
Red O stained sections with ImageJ to give a percentage of the total area of the implant stained
positive by Oil Red O (Figure 4-13). HMEC+ASC implants were shown to have progressive fat
accumulation in the implant region, growing to an average of 60% of the implant area stained
positive for Oil Red O by day 90. Implants made of modules with embedded ASC, but without
surface HMEC showed little Oil Red O staining at all time points investigated. It appears that
early vascularization, possibly driven by HMEC cotransplantation, is necessary for fat
development at later time points.
0102030405060708090
100
30d 60d 90d
% Im
plan
t Are
a O
il Re
d O
HMEC+ASCASC-only
Figure 4-13: Percentage of area stained red for Oil Red O on micrograph sections of HMEC+ASC and ASC-only module implants at day 30, 60 and 90.
Progressive fat accumulation is seen in the module implants containing both ASC and HMEC, while little fat accumulation is observed in ASC-embedded modules without HMEC (n=3). Error bars represent ± 1 SD.
83
4.4.3.2 Angiogenesis and HMEC Viability
A clear difference in the level of vascularization between the HMEC+ASC and ASC-only
modules was observed. Similar to the earlier time points, HMEC+ASC explants appeared to be
red and vascularized, while the ASC-only explants appeared to be opaque and avascular.
Histology confirmed these observations with the HMEC+ASC explants rich in large blood
vessels filled with erythrocytes while the ASC-only explants remained largely avascular (Figure
4-14). UEA-1 staining revealed that many of these large blood vessels were derived from the
implanted HMEC (Figure 4-15). HMEC-dervied vessels were shown to persist in the
HMEC+ASC modules as long as 90 days post-implantation (Figure 4-15).
84
Figure 4-14: H&E and Masson’s Trichrome micrographs of HMEC+ASC and ASC-only modules 30 days after implantation.
Many large erythrocyte-filled blood vessels are clearly visible in the HMEC+ASC case where the ASC-only case appears avascular (n=5). Scale bar = 100μm.
HMEC+ASC ASC-only
85
Figure 4-15: UEA-1 stained sections of HMEC+ASC modules 30 days and 90 days after implantation.
Large HMEC-derived vessels (arrows) are seen to persist up to 90 days in HMEC+ASC module implants (n=3-5). Scale bar = 100μm.
4.4.3.3 Volume Measurements
Volume measurements of the explants revealed progressive shrinkage of both the
HMEC+ASC and ASC-only module implants over 30, 60 and 90 days (initial volume at
implantation = 40mm3) (Figure 4-16). This is not surprising as collagen gel used as the module
material is known to contract and degrade over time in vivo. However, surprisingly at day 90,
HMEC+ASC implants were shown to be larger on average than the ASC-only implants (p<0.05).
Perhaps, fat accumulation in HMEC+ASC helped to maintain implant volume.
30 d 90 d
86
Figure 4-16: Volume measurements of HMEC+ASC and ASC-only module implants.
At explant, both HMEC+ASC and ASC-only module implants show progressive shrinkage over 90 days. At day 90, HMEC+ASC implants are statistically larger than ASC-only implants which may be attributed to the observed fat accumulation in HMEC+ASC implants. Initial volume at implantation = 40mm3. (n=3) Error bars represent 1SD.
4.5 Discussion
The successful correction of soft tissue defects represents a challenge in plastic surgery.
Autologous tissue transfers and tissue engineering approaches to make adipose tissue substitutes
have largely been unsuccessful in maintaining a stable volume of adipose tissue in vivo long-
term (>3 months). Necrosis due to insufficient blood supply after cell or tissue transplantation
results in considerable volume loss. This is because in the first few days after transplantation,
transplanted cells are only nourished by diffusion from the surrounding tissue, which is only
sufficient at distances less than 150µm. Host derived vessels may take over 7 days to invade the
implant and may only then reach the periphery of a large 3-D construct of clinical relevance.
Volume loss of the adipose tissue construct is only avoided through early vascularization of the
transplanted tissue. The goal of the cotransplantation of HMEC and ASC in the modular
approach was to rapidly create a vascular network upon implantation, that would circumvent
0
5
10
15
20
25
30
30D 60D 90D
Volu
me
mm
3
HMEC+ASC
ASC-only*
*p<0.05
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ischemic conditions to promote ASC viability and differentiation and hence a long-term volume-
stable culture of adipose tissue in vivo.
HMEC transplantation as a revascularization strategy was successful in the implants with
modules containing ASC as a supporting cell. ASC are known to secrete many bioactive
molecules, such as vascular endothelial growth factor (VEGF), hepatocyte growth factor (HGF)
and granulocyte macrophage colony-stimulating factor (GM-CSF) which have been shown to
promote EC survival and proliferation[91]. It is not surprising that the presence of the ASC
appeared to have both a protective (promoted HMEC viability) and angiogenic effect on the
transplanted HMEC. As early as day 3, HMEC were found to detach from the surface of the
modules and organize themselves into primitive vessels with lumens and many containing
erythrocytes (Figure 4-2, Figure 4-3), suggesting rapid anastomosis with the host vasculature.
By day 7 and 14, the number of vessels had reduced but the vessels that remained were large
(~200µm diameter) suggesting that the initial leaky vessels had matured and organized during
the angiogenic process. This may be the result of secreted angiogenic factors in ASC/EC
paracrine signaling events, or perhaps some ASC differentiated into pericyte-like cells that
stabilized the maturing vasculature. This will be further discussed in Chapter 5. To confirm that
HMEC were not unique in their revascularization potential, modules made with HUVEC
displayed similar results. Successful inosculation, at least by day 21, of the HMEC+ASC
implants with the host vasculature was confirmed by MicroCT (Figure 4-8). Furthermore,
vessels derived from the transplanted HMEC (with embedded ASC) were shown to persist in
vivo for up to 90 days without drugs or the addition of anti-apoptotic genes. HMEC on modules
without embedded ASC, on the other hand, were found to remain on the surface of the modules
at day 3 and UEA-1 lectin staining was shown to disappear over 14 days suggesting cell death
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without the presence of a supporting cell (Figure 4-2). A TUNEL stain would have been useful
here to confirm apoptosis of HMEC in the absence of ASC cotransplantation.
We hypothesized that a successful revascularization strategy would promote embedded
ASC viability, differentiation and long-term persistent culture of adipose tissue in vivo. We
found that implants of modules containing both embedded ASC and surface coated HMEC
started displaying Oil Red O staining indicative of fat accumulation at day 30 post implantation
(Figure 4-10). The fat accumulation progressed until the end point of the study (90 days), where
the average Oil Red O micrograph was stained ~60% of total area for Oil Red O (Figure 4-13).
While we cannot be certain that all of this fat accumulation is derived from the transplanted
ASC, higher magnification images of H&E and Masson’s Trichrome stained sections show the
unilocular appearance of fat cells within individual modules (Figure 4-14), suggesting that at
least some of the transplanted ASC had differentiated into mature adipocytes. Interestingly,
ASC-embedded modules without the surface coating of HMEC were not shown to accumulate
fat by Oil Red O staining (Figure 4-10). Perhaps, these cells had died early on in the course of
the study due to lack of early vascularization as seen in the case with HMEC+ASC
cotransplantation. Or perhaps, EC/ASC paracrine signaling as discussed further in Chapter 5
promoted ASC differentiation.
While we were successful in demonstrating the ability of cotransplantation of HMEC and
ASC in a modular platform to create adipose tissue in vivo, we were not able to show that these
were volume-stable tissues. Volume measurements of the explants showed progressive
shrinkage of both HMEC+ASC and ASC-only implants over the time-course of the experiment.
The initial volume at implantation was 40mm3. The decrease in implant size at 30 days by more
than 50% the original volume is likely attributed to contraction of the collagen gel modules by
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both the embedded ASC and surface coated HMEC as these modules were not fully pre-
contracted in vitro prior to implantation. Although we did find that the HMEC+ASC implants to
be statistically larger than the ASC-only implants at day 90, yet these still had reduced in size by
approximately 50% from day 30 likely due to collagen degradation. A 50% reduction in the size
of an adipose tissue repair strategy is not clinically acceptable. This is the result of using
collagen gel as the module material which is known to shrink and degrade in vivo. The
development of a suitable biomaterial is still needed for adipose tissue engineering.
4.6 Conclusions
Cotransplantation of ASC and HMEC in collagen gel modules implanted subcutaneously
into SCID mice was successful in rapidly creating a vascular network upon implantation. With
the presence of ASC, HMEC detached from the surface of the modules to form vessels that
contained erythrocytes as early as day 3. HMEC-derived vessels were shown to decrease in
number but increase in size over the first 14 days suggesting vessel maturation and persisted for
up to 90 days in vivo. With successful early vascularization, HMEC+ASC modules supported
ASC viability and differentiation to create a fat pad that persisted up to 90 days.
4.7 Acknowledgements
The authors acknowledge the financial support by NSERC. Special thanks to Chuen Lo and
his expertise in animal surgeries. Also, thanks to Lisa Yu (Dr. R. M. Henkleman) for microCT
imaging.
90
5 Investigating the Relationship between Adipose‐dervied Stem Cells and Endothelial Cells In Vitro
5.1 Abstract
There exists a paracrine relationship between ASC and EC, but the exact mechanisms
behind this interaction are unknown. To better understand the mechanisms of ASC-EC
interaction in the context of collagen modules we modified an in vitro angiogenesis assay to
study the sprouting of HUVEC from the surface of modules in a fibrin gel, with and without the
presence of an ASC-embedded collagen gel in a transwell insert. Although without direct cell-
cell contact, the presence of ASC in the wells promoted an angiogenic phenotype in the HUVEC.
HUVEC were seen to sprout from the modules and branch to form a dense vascular network that
intersected with vessels from neighboring modules. In the case without ASC coculture, HUVEC
were initially seen to sprout from the surface of the modules at day 3 and 7, however the average
number and length of sprouts were much less than those observed in the coculture case. By day
14, the vascular network originally created had degenerated into individual cells. It is clear that a
support cell, such as ASC, is necessary for HUVEC vascular network formation in the in vitro
conditions investigated here. The preliminary RT-PCR results were quite variable and the
presence of HUVEC did not greatly affect the mRNA expression level of adipogenic genes in the
ASC. The mRNA level of several angiogenic genes, including VEGF, PAI-1 and TNFα, were
shown to be increased in ASC with HUVEC coculture at early time points. These factors may be
involved in ASC-EC paracrine signaling.
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5.2 Introduction
Creation of a functional vascular network soon after implantation is critical when
engineering an adipose tissue substitute. In Chapter 4, we were successful in rapidly creating a
vascular network by cotransplantation of ASC-embedded modules with a HMEC surface cell
layer. Yet, without the presence of the embedded ASC, HMEC on the surface of the modules
displayed no angiogenic effect. In addition, ASC-embedded modules without HMEC
cotransplantation yielded virtually no fat development. In this chapter we wish to explore the
reciprocal relationship between ASC and EC using a modified in vitro angiogenesis assay and
RT-PCR analysis of several angiogenic and adipogenic genes.
There are numerous in vivo models for studying angiogenesis, which include the chick
CAM assay or injection of Matrigel into mice[128,145]. Yet with these models it is difficult to
purify the EC for biochemical or gene expression analyses. Nehls and Drenckhahn developed a
novel microcarrier-based assay for the study of 3-dimensional cell migration and angiogenesis in
vitro[146]. In their system, EC are cultured as a monolayer on Cytodex beads and then
embedded and cultured in a fibrin gel. This system is well characterized and used extensively in
literature to study EC sprouting and migration. We chose to modify this system, so that instead
of EC covered beads, we used EC covered modules to better mimic the modular constructs that
were implanted in the in vivo studies. Commercially available HUVEC were chosen instead of
HMEC as the EC cell source to minimize the donor-to-donor variability which may affect RT-
PCR results. In Chapter 4, we found that HUVEC-coated modules had a similar angiogenic
response to ASC as did HMEC.
The cross-talk between ASC and EC has been explored in literature, but a complete
understanding of the mechanisms of action are not fully understood. It is believed that a
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paracrine relationship exists between ASC and EC. Research over the past decade has identified
some of the pro-angiogenic and anti-angiogenic growth factors expressed by adipose tissues or
adipocytes cultured in vitro. These include VEGF, bFGF, Ang-1,-2, PAI-1, TGF-β and
leptin[89]. We explored the paracrine relationship between HUVEC-coated modules embedded
in the fibrin gel in an in vitro angiogenesis assay and ASC embedded in collagen gel in a hanging
transwell insert. Real time RT-PCR was used to study the effect of coculture on the expression
levels of both adipogenic and angiogenic genes.
5.2.1 Cells
Adipose-derived stem cells (ASC) were obtained from the stromal vascular fraction of
fresh human abdominal adipose tissue as described in Section 3.2.2. Primary Human Umbilical
Vein Endothelial Cells (HUVEC, Lonza, Walkersville, MD) were cultured in EGM-2 culture
medium (Lonza). Cells were maintained at 37ºC and 5% CO2 in a humidified incubator, medium
was changed every 2-3 days and cells were passaged when ~90% confluent. ASC were used at
passage 3. HUVEC were used between passage 4 and 6.
5.2.2 Optimized In Vitro Angiogenesis Sprouting Assay
A previously described in vitro microcarrier angiogenesis assay[146,147] was adapted to
study the spouting and migration of HUVEC from the surface of the modules in a fibrin gel, with
and without the presence of ASC in coculture. The experimental set up for the coculture assay is
shown in Figure 5-1. Briefly, a 2mg/mL fibrinogen solution (Sigma, Fibrinogen, Fraction I, type
1-S, from bovine plasma; 85% clottable) was prepared and aprotinin (Sigma, from bovine lung)
was added at 0.15 units/mL. 0.25mL of the fibrinogen solution was added to the bottom of each
well of a 24-well plate containing 0.625U/mL thrombin (Sigma) and pipetted gently to mix and
93
allowed to form a clot for 20 minutes at 37oC. This was to prepare a bottom layer of fibrin gel so
that modules would not settle to the bottom of the plate. HUVEC-coated collagen modules
(prepared as described in Section 3.2.2) were then gently mixed with the fibrinogen solution (1
package of cut modules/6mL fibrinogen; this is sufficiently dilute to avoid the modules clumping
and forming clusters once in the fibrin gel). 0.5mL of the fibrinogen/module solution was added
on top of the bottom layer of fibrin gel in each well of 24-well plate containing 0.625U/mL of
thrombin (Sigma) and pipetted gently to mix. The plate was placed in a 37oC incubator for 20
minutes to generate the clot. 1mL of EGM-2 (Lonza) was added drop-wise on top of each well
and incubated for at least 30 minutes. The medium was then aspirated and 1-1.25mL of fresh
medium was added to each well. Without the addition of ASC, this set-up was used as the
control.
For the coculture experiments, ASC-embedded collagen gels in hanging transwell inserts
were added to each well containing the HUVEC-coated modules in the fibrin gel. Briefly, ASC
(106cells/mL) were mixed with a 3mg/mL collagen type I solution (PureCol™, Inamed, Fremont,
CA) containing 10X MEM (10.1 g α-Minimum Essential Medium powder (Gibco, Invitrogen,
Paisley, PA), and 2.2g endotoxin free NaHCO3, in 100mL endotoxin free water) and neutralized
to pH ~ 7.4 with 0.8N NaHCO3. 200uL of the collagen-cell suspension was added to the bottom
of cell culture inserts (BD Bioscience, 0.4µm pore size, high density PET membrane) and
allowed to gel at 37oC for 1 hour. 0.3mL of EGM-2 was added to the hanging well insert and
0.7mL of the media was added to the bottom well containing the fibrin gel and cultured in an
incubator at 37oC. The media were changed every 48 hours.
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Figure 5-1: Experimental set up of the in vitro angiogenesis assay.
For the coculture experiments, HUVEC-coated modules were embedded in a fibrin gel layer with ASC embedded in a collagen gel in a transwell insert. Control wells consisted of collagen gel in the hanging transwell insert with an empty fibrin gel layer, and HUVEC modules in the fibrin gel layer without ASC in the transwell insert.
5.2.3 Sprouting Counts and Measurements
The level of sprouting from the HUVEC-covered modules in the coculture set-up was
investigated at 3, 7 and 14 days and compared with HUVEC-covered modules in fibrin gel
without the presence of ASC in the hanging transwell insert as the control. At each time point
investigated, a Zeiss Axiovert 135 (Zeiss, Germany) light microscope equipped with a
CoolSNAP-Pro camera (Media Cybernetics, San Diego CA) using ImpagePro software (Media
Cybernetics) was used at 2.5X objective lens to randomly locate and photograph at least 8
individual modules from both the coculture and control wells. The entire experiment was
repeated 3 times (n=8, N=3). Modules which formed clusters were not used for the analysis due
to difficulty in separating the origin of the sprouts and that the outer surface was blocked by
other modules. Mostly individual modules or modules only slightly merged (ie. 2 modules with
less than ~15% of the surfaces touching) were considered for the analysis. The images were
analysed using the ImageJ (version 1.4.3) software program. A sprout was counted and defined
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as a vessel only if it was equal to or greater than 150µm in length. The length of each sprout was
measured as the linear distance from the beginning of the sprout at the module surface to its tip
(Figure 5-2).
Figure 5-2: Image of capillary like-formations growing out from the surface of a HUVEC-coated module in fibrin gel with the presence of ASC in coculture at day 3.
Arrows highlight some counted sprouts, and show the measured linear distance of individual sprouts from the surface of the modules. Scale bar = 250µm.
5.2.4 RNA Isolation
After photographs were taken for the sprouting measurements, RNA was isolated from
both the HUVEC in the fibrin gel and ASC from the collagen gel in the transwell for RT-PCR
analysis of key angiogenic and adipogenic genes. Experimental set-ups without coculture (e.g,
HUVEC-coated modules in fibrin gel, but without ASC gel in the transwell, and ASC gel in the
transwell but without HUVEC-coated modules in the fibrin gel layer) were used as controls.
RNA was isolated using the RNeasy® Fibrous Tissue Mini Kit (Qiagen). Briefly, gel layers
96
(either fibrin containing HUVEC or collagen containing ASC) were carefully separated from
individual wells to avoid cross-contamination with the other cell type. A total of 6 gels from
each condition were carefully removed and placed in 1.5mL RNAse free centrifuge tubes. The
samples were centrifuged at 500 x g for 30 seconds to separate the cell and hydrogel from excess
medium. Following the manufacturers instructions, the samples were then homogenized in
300µL buffer RLT (extraction buffer) containing β-mercaptoethanol (Sigma) and treated with a
proteinase K solution at 55oC for 10 minutes to help digest proteins contained in either the fibrin
or collagen gels that may interfere with the RNA purification process. Following centrifugation
at 10,000 x g for 3 minutes the lysate was removed and purified on the RNeasy mini spin
column. The concentration and purity of the RNA was measured using a NanoDrop (ND1000)
Spectrophotometer with a 1µL sample volume. As a measure of purity, the 260/280 ratio was
generally between 1.9 and 2.0, within the desired 1.9 – 2.1 range for RNA.
5.2.5 cDNA Synthesis
First-strand cDNA was synthesized from 1µg of total RNA using random primers
(Invitrogen; Cat. # 48190-011) and SuperScript™ II Reverse Transcriptase (RT) (Invitrogen;
Cat. # 18064-014). Briefly, from the nanodrop concentration readings, the exact volume of stock
RNA and water required to make up 1µg RNA in 6µL water in 0.2mL reaction tubes was
determined. To each sample, 0.5µL dT primer, 0.5µL random primer and 1.0µL annealing
primer were added and the tubes were heated at 65oC for 5 minutes. After the annealing, 10µL
2x First strand reaction mix and 2µL SuperscriptIII/RNAse out enzyme mix were added to each
tube and the samples were vortex briefly and then centrifuged at 1200 x g for a few seconds.
After 5 minutes of incubation at room temperature, the tubes were put in the PCR thermocycler
97
(Applied Biosystems 2720), with a cycle set to 37oC (5 min), 50oC (50 min) and then 85oC (5
minutes). cDNA samples were then stored at -20oC until RT-PCR analysis.
5.2.6 Real‐time RT PCR
The relative expression level of several key adipogenic and angiogenic genes (see tables
for a list of gene specific primers) were used to study the effect of coculture on both ASC and
HUVEC. This was investigated at 72 h, 7 days and 14 days in culture (n=3, N=3). GAPDH
was used as the housekeeping gene for the relative quantification.
The cDNA was diluted in RNAse-free water (Gibco) so that each sample theoretically
contained 10ng of input RNA. The actual amount of cDNA in the sample was dependent on the
efficiency of the cDNA synthesis reactions. Power SYBR® Green Mastermix (Applied
Biosystems, cat# 4367569) was used for the real time analysis. To minimize pipetting errors,
mastermixes were prepared for (i) each of the samples in RNAse-treated water and (ii) the Power
SYBR® Green Mastermix with each of the gene-specific primer sets. Each reaction was
conducted in a well of a 384-well plate with a 10μL reaction volume containing the cDNA
template, 250nM forward primer, 250nM reverse primer and 5µL SYBR® Green Mastermix.
Each gene was analyzed in duplicate for each sample in separate wells.
An Applied Biosystems 7900HT Real-Time PCR system with the SDS 2.3 software
detection program was used for fluorescence detection. The PCR conditions were 95°C for 5
minutes followed by 40 cycles of 30 s at 95°C (denaturation), 30s at 58°C (annealing) and 30s at
72°C (elongation). The fluorescence was measured after each elongation period. A final
extension was conducted for 5 min at 72°C. Following this, melting curve analysis was
conducted to characterize the formed products and confirm that only one amplicon was detected.
For the melting curves, the temperature was increased from 72 °C to 95 °C (held 1 second), with
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fluorescence readings every 0.5 °C. Using the software program, the CT (threshold cycle) values
were obtained by analyzing the amplification curves on a logarithmic scale and setting the
threshold within the linear range for all of the genes.
The PFFAFL method[148] for RT-PCR quantification was used to determine the relative
fold-change comparing different samples. The equation for the relative expression ratio is
presented below and is calculated only from the real-time PCR efficiencies and the crossing
point deviation of an unknown sample versus a control. The PCR efficiencies were determined
using the software program LinRegPCR (version 7.2). The program determines a baseline
fluorescence and does a baseline subtraction. Then a Window-of-Linearity is set and PCR
efficiencies per sample are calculated. Unlike the comparative CT method, the PFAFFL model
needs no calibration curve. For HUVEC samples, the treated group is the HUVEC in the
coculture arrangement (ie. HUVEC-coated modules in fibrin gel in the presence of ASC-
embedded collagen gel in the transwell system) while the control is HUVEC-coated modules
cultured without the presence of ASC in the well. This will show the effect of the presence of
ASC on HUVEC gene expression in coculture. For the samples with ASC, the treated group is
ASC in the coculture arrangement (ie. ASC-embedded collagen gel in the transwell with
HUVEC-coated modules in a fibrin gel layer), while the control is ASC-embedded collagen gel
without coculture with HUVEC. This will show the effect of the presence of HUVEC on ASC
gene expression in coculture. Both GAPDH and 18S were used as the housekeeping genes, but
real time results indicated that 18S ribosomal RNA was far more abundant in the samples than
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any of the genes investigated. The 18S amplification curves generally crossed the selected
threshold at a very early cycle number, with CT values generally in the range of 5 to 7 cycles.
Since CT values below 10 may be unreliable, GAPDH was used as the housekeeping gene for all
calculations.
Table 2: RT-PCR Primer List
Gene Accession # Description Primers Length (bp)
LPL
M15856 Human lipoprotein lipase • Early marker of adipogenic differentiation. An enzyme secreted by adipocytes into the adjacent capillary endothelium. Hydrolyzes triglycerides into glycerol and free fatty acids. The adipocytes uptake the free fatty acids via a transmembrane protein, where they are processed and stored as triacylglycerols.
F: gtccgtggctacctgtcatt R: tggcacccaactctcataca
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PPARγ
NM_138712 Human peroxisome proliferator-activated receptor-γ • Early-mid marker of adipogenic differentiation. A nuclear hormone receptor involved in the differentiation of fat. Highly expressed in adipose tissue. Involved in the regulation of adipocyte-specific genes that function in lipid metabolism, such as aP2 and adiponectin. Exists in two isoforms, PPARγ1 and PPARγ2, encoded by a single gene.
F: ttcagaaatgccttgcagtg R: ccaacagcttctccttctcg
88
aP2
J02874 Human fatty acid binding protein • Mid-late marker of adipogenic differentiation. Cytoplasmic protein involved in the intracellular transport and metabolism of fatty acids in adipocytes. Binds fatty acids and transports them within the cell to facilitate triacylglycerol formation and degradation. Upregulated during adipogenic differentiation.
F: tactgggccaggaatttgac R: ggacacccccatctaaggtt
78
GPDH
NM_005276 Human glycerol-3-phosphate dehydrogenase 1 • Late marker of adipogenic differentiation. A cytosolic protein involved in the biosynthesis of fat in adipocytes. Catalyzes the conversion of dihydroxyacetone phosphate (DHAP) (an intermediate in the glycolytic pathway) into glycerol-3-phosphate using NADH as a co-enzyme. Long chain fatty acids form esters with the glycerol-3-phosphate to produce triacylglycerols.
F: gagctgatgcagacaccaaa R: ccacggccactacattcttt
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Glut4
M20747 Human insulin-responsive glucose transporter • Late marker of adipogenic differentiation. The main glucose transport protein in adipocytes. The presence of insulin activates intracellular signalling cascades that result in the transport of Glut4 transporters from specialized intracellular storage vesicles to the cell membrane, thereby facilitating the transport of glucose into the cell. When insulin levels are reduced, the Glut4 transporters return to the intracellular storage vesicles via endocytosis.
F: agcagctctctggcatcaat R: ctacccctgctgtctcgaag
66
Leptin
NM_000230 Human leptin • Late marker of adipogenic differentiation. A non-glycosylated polypeptide that is secreted by adipocytes as they accumulate lipid. Believed to function in the regulation of body fat. Leptin binding to specialized receptors in the hypothalamus is believed to influence appetite and metabolic rate. Mutations in the gene encoding leptin have been shown to result in obesity in mice.
F: ggctttggccctatcttttc R: accggtgactttctgtttgg
146
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Pecam NM_000442
Human platelet/endothelial cell adhesion molecule 1 • EC surface marker Known to be expressed on the surface of endothelial cells, circulating platelets, monocytes, granulocytes and some subsets of T-lymphocytes. PECAM-1 has been proposed as one of the main players in transendothelial migration of neutrophils, monocytes, and natural killer cells in both in vivo and in vitro models
F:aacagtgttgacatgaagagcc R:tgtaaaacagcacgtcatcctt
148
Tie-1 NM_005424
Human Tyrosine kinase with immunoglobulin-like and EGF-like domains 1 • EC surface marker Cell surface protein expressed exclusively in endothelial cells.
F:cacgaccatgacggcgaat R:cggcagcctgatatgcctg
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PAI-1 M16006
Human Plasminogen Activator Inhibitor 1 A serine protease inhibitor that inhibits fibrinolysis by inactivating urokinase-type and tissue-type plasminogen activator. PAI-1 is also involved in angiogenesis and atherogenesis. PAI-1 is normally secreted by endothelial cells, vascular smooth muscle cells, hepatocytes platelets and adipocytes. Majority of the circulating PAI-1 is contributed to by adipose tissue
F:catcccccatcctacgtgg R:ccccatagggtgagaaaacca
109
bFGF NM_002006
Human basic fibroblast growth factor • Angiogenic growth factor Basic fibroblast growth factor is present in basement membranes and in the subendothelial extracellular matrix of blood vessels
F:atcaaaggagtgtgtgctaacc R:actgcccagttcgtttcagtg
178
PDGFA NM_002607
Human Platelet derived growth factor, subunit A • Angiogenic growth factor Potent angiogenic growth factor involved in the stabilization and maturation of vessels.
F:ccagcgactcctggagataga R:cttctcgggcacatgcttagt
100
VEGF NM_113376
Human Vascular Endothelial growth factor • Angiogenic growth factor Potent angiogenic growth factor known to initiate angiogenesis events.
F:caacatcaccatgcagattatgc R: gctttcgtttttgcccctttc
164
TNFα NM_000594
Human Tumor necrosis factor alpha • Angiogenic growth factor Cytokine involved in systemic inflammation
F:atgagcactgaaagcatgatcc R:gagggctgattagagagaggtc
217
GAPDH NM_002046 Human glyceraldehyde-3-phosphate dehydrogenase • Housekeeping gene.
F: acagtcagccgcatcttctt R: acgaccaaatccgttgactc
94
18S X03205 Human 18S ribosomal RNA • Housekeeping gene.
F: accgcggttctattttgttg R: ccctcttaatcatggcctca
51
5.2.7 Statistical Analysis
All statistical analyses were performed using the software program STATISTICA Version
5.1 (Statsoft, USA). Student’s t-tests were used to compare the means between two groups
analyzed and considered significant at p<0.05.
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5.3 Results
5.3.1 Effect of ASC on HUVEC Sprouting from Modules in a Fibrin Gel
The results from the in vitro angiogenesis assay showed a clear relationship between the
presence of ASC in coculture and HUVEC sprouting from the surface of the modules into the
fibrin gel. Figure 5-3 shows representative images of HUVEC sprouting from the surface of the
modules in a fibrin gel, with and without the presence of ASC-embedded gels in a transwell
insert. In the control wells, without ASC, HUVEC could be seen to begin to migrate from the
surface of the modules by day 3 to form short and narrow cord-like structures. These structures
appeared to elongate by day 7 and begin to branch, yet some of the cords appeared to disintegrate
into disorganized groups of individual cells or cell clusters. By day 14, the entire vascular
network that had been previously established had completely degenerated into individual
rounded cells surrounding the modules. In contrast, when an ASC-embedded gel was placed in a
transwell system above the fibrin gel containing the HUVEC-coated modules, a branched
network of HUVEC cords was seen projecting from the surface of the modules as early as day 3.
Branching was thought of as a recapitulation of the sprouting process from the surface of the
module, and was seen extensively in the coculture system. HUVEC cords grew until they
reached another cord and merged. This was seen as a mature and dense network of capillaries by
day 7 with nearby vessels merging to form a vascular network surrounding individual modules.
The diameter of the capillaries at day 7 appeared to be larger than at day 3, suggesting that they
contained lumens surrounded by multiple cells. By day 14, the entire fibrin gel layer contained
an extensive web of vasculature where vessels from one module had reached the vessels of a
nearby module and connected. Again the vessels appeared to be thick with diameters up to
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50µm, suggesting that they were multicellular and contained patent lumens (Figure 5-4). No
single cells were seen migrating away from the modules that were not in contact with other cells.
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Figure 5-3: Photographs of HUVEC sprouting from modules in a fibrin gel with and without ASC coculture.
With coculture with ASC, HUVEC were seen to sprout from the modules at day 3, the sprouts elongated by day 7 and formed branches and by day 14 a dense vascular network that connected with sprouts from neighbouring was observed. Without ASC coculture, HUVEC were initially seen to form sprouts at day 3 and 7 with minimal branching but these degenerated into a cluster of individual cells surrounding the modules by day 14.
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Figure 5-4: Magnified image of HUVEC sprouting from modules with ASC coculture at day 14.
HUVEC sprouts from modules with ASC coculture appeared thick and may be multicellular and contain patent lumens. The arrow points to a sprout that is ~50µm in diameter. Scale bar = 250µm.
Measuring the number and length of HUVEC sprouts growing from the surface of the
modules, coculture with ASC gave a statistically significant improvement in both sprout number
and length as compared to the control case without ASC. As seen in Figure 5-5, there were 15 ±
3 sprouts/module at day 3 which increased to 38 ± 5 sprouts/module for the case with ASC
coculture. Comparing this to the case without ASC, the number of sprouts/module was 7 ± 3 and
13 ± 3 for day 3 and 7 respectfully. The effect of ASC coculture on HUVEC sprouting is clearly
demonstrated by an increase of 25 sprouts/module at day 7 as compared to the control wells
without ASC. Futhermore, by day 14 there were no sprouts to count in the control case and the
branching was too extensive in the coculture case to allow for accurate quantification. As seen
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in Figure 5-6 the average length of the sprouts in the coculture case was seen to increase from
227 ± 14µm at day 3 to 491 ± 43µm by day 7, demonstrating that the average sprout length had
almost doubled over this time period. In comparison, without the presence of ASC, the sprout
length did not increase much from 216 ± 22µm at day 3 to 250 ± 19µm by day 7. The average
sprout length in coculture with ASC was on average 240µm longer than the control case without
ASC at day 7.
Figure 5-5: Average number of HUVEC sprouts from the module surface in fibrin gel with and without the presence of ASC in coculture.
A greater number of HUVEC sprouts (>150µm) were observed sprouting from the surface of the modules in the fibrin gel with ASC coculture as compared to control case without ASC (n=8, N=3). Error bars represent ± 1SD. * p < 0.05.
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Figure 5-6: Average length of HUVEC sprouts from the module surface in fibrin gel with and without the presence of ASC in coculture.
At day 3 the average length of HUVEC sprouts (>150µm) was nearly identical between the coculture and the control case without ASC. By day 7, the effect of ASC coculture was seen with a near doubling in sprout length as compared to the control without ASC (n=8, N=3). Error bars represent ± 1SD. * p < 0.05.
5.3.2 RT‐PCR Analysis
Real time RT-PCR analysis of key adipogenic and angiogenic genes was conducted to
help better elucidate the reciprocal relationship between ASC and EC. The transwell system was
used so that the ASC in the collagen gel could be easily separated from the fibrin gel layer
containing HUVEC without chance of cross-contamination between the cell types.
The data presented here in Figure 5-7, Figure 5-8 and Figure 5-9 represents the relative
fold change in mRNA expression of either 1) ASC in coculture vs. ASC alone, or 2) HUVEC in
coculture vs. HUVEC alone. Due to the variable nature of the obtained results, each
experimental value from each of the three trials is plotted. The variable nature of the results may
be due, in part, to donor cell variability. Two of the experiments were conducted with ASC
derived from the same patient, while the other was from another donor. The data points derived
from the same patient are visualized on the graphs as a solid colour point, while the other data
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point is not filled with colour. A significant fold change in mRNA expression level is considered
to be anything greater than a 1.5 or less than 0.5.
From the ASC results (Figure 5-7), it appears that these cells were not being induced to
differentiate by the HUVEC under the experimental conditions examined here. If so we would
have expected at least some of these adipogenesis markers to be clearly expressed more. While
there was some evidence that LPL (an early marker of adipogenesis) was expressed more than
without HUVEC coculture at day 3, this is back down to similar levels by day 7 and 14.
Interestingly, both PPARγ and aP2 appear to be expressed less at day 3 and then return to
unchanged levels by day 7 and 14. The only clear example of a change in mRNA expression for
all 3 experimental trials is Glut4 which demonstrated fold-changes greater than 2 at day 3, but
then again it is back to unchanged levels by day 7 and 14.
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ASC (Adipogenic Genes)
LeptinGlut 4GPDHaP2PPARLPL0
0.5
1
1.5
2
2.5
3
3.5
4
Rela
tive
Fold
Cha
nge
in m
RNA
Day 3Day 7Day 14
Figure 5-7: Relative fold change in mRNA expression of adipogenic genes of ASC in coculture with HUVEC, vs. ASC cultured alone.
HUVEC coculture does not appear to induce an adipogenic phenotype of the ASC in the in vitro conditions investigated here. Only aP2 expression appears to be decreased and Glut4 expression increased for all 3 trials at day 3. Solid colour data points represent 2 trials with the same donor ASC. The unfilled data point represents ASC from a different donor.
Exploring the angiogenic genes investigated (Figure 5-8), there was a significant increase
in PAI-1 expression at day 3, with unchanged levels at day 7 and 14. Interestingly, HUVEC did
not seem to change the expression levels of bFGF or PDGF in ASC at any of the time points
investigated. VEGF expression was clearly increased at day 3, with fold change values greater
than 3.5, but returned to unchanged levels by day 7 and 14. ASC from two of the trials (same
donor, solid colour data points) appeared to be have increased expression in TNFα with fold
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changes greater than 1.5 at days 3 and 7, while fold change in ASC from the other donor
(outlined data points) were unchanged.
ASC (Angiogenic Genes)
PAI 1 bFGF PDGF VEGF TNFa0
0.5
1
1.5
2
2.5
3
3.5
4
4.5
5
5.5
Rela
tive
Fold
Cha
nge
in m
RNA
Day 3Day 7Day 14
Figure 5-8: Relative fold change in mRNA expression of angiogenic genes of ASC in coculture with HUVEC, vs. ASC cultured alone.
Coculture with HUVEC increased the expression of PAI-1, VEGF and possibly TNFα (one donor) at early timepoints. These factors may be involved in ASC-EC paracrine signalling. Expression of all other genes remained unchanged. Solid colour data points represent 2 trials with the same donor ASC. The unfilled data points represents ASC from a different donor
Examining the results of the HUVEC cells and the impact that ASC coculture had on the
relative gene expression (Figure 5-9), we can see that ASC had little impact on the EC markers
PECAM and Tie-1, with only PECAM at day 7 showing fold-changes greater than 1.5 for all
three experimental trials. Surprisingly, PAI-1, bFGF and PDGF all showed little change in
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mRNA expression with possibly the exception of bFGF which had quite variable results. It
appeared that VEGF at day 3 and 14 may be upregulated in HUVEC in the coculture system.
HUVEC
VEGFPDGFbFGFPAI 1Tie1PECAM0
0.5
1
1.5
2
2.5
3
3.5
4
4.5
Rela
tive
Fold
Cha
nge
in m
RNA
Day 3Day 7Day 14
Figure 5-9: Relative fold change in mRNA expression of HUVEC in coculture with ASC, vs. HUVEC cultured alone.
Coculture with ASC increased the expression of bFGF (one donor) at day 3 and VEGF at day 3 and 7 (one donor). Expression of all other genes was variable and unchanged. Solid colour data points represent 2 trials with the same donor ASC. The unfilled data point represents ASC from a different donor.
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5.4 Discussion
The results from Chapter 4 of this thesis showed a clear relationship between ASC and
EC in vivo. Only with ASC cotransplantation inside the modules, were the HMEC able to detach
from the surface of the modules, organize into vessels, anastomose with the host vasculature and
remain viable for up to 90 days. It appeared that the ASC promoted EC survival and had an
angiogenic effect on EC in vivo. The objective of this work was to conduct a preliminary study
to help elucidate the crosstalk between these two cell types in vitro.
The results from the in vitro angiogenesis assay recapitulated what was seen in vivo. The
presence of ASC in the transwell led to extensive sprouts and tube formation from the HUVEC
on the surface of the modules, leading to a branched and interconnected network inside the fibrin
gel layer by day 14. In contrast, without the presence of ASC, HUVEC were found to initially
form some sprouts at early time points, but by day 14 complete degeneration of the vascular
network was observed and only individual cells were seen surrounding the modules.
The in vitro sprouting assay as developed by Nehls and Drenckhahn[146], typically
involves culturing EC on Cytodex beads and then embedding these beads in a fibrin gel. They
showed that this technique works well for bovine aortic EC (BAEC), which were shown to bud
and form capillary-like structures in response to angiogenic growth factors such as bFGF and
VEGF. However, there may be phenotypic differences in the type of EC used as their sprouting
assay did not show sprouting from HUVEC. One group has modified this protocol to include
skin fibroblasts cultured on top of the gels containing the HUVEC covered beads[149]. They
found that only in the presence of these fibroblasts were the HUVEC able to sprout, divide,
migrate, form lumens and connect with neighbouring vessels, similar to what is observed in
angiogenesis in vivo. ASC share several phenotypic similarities to fibroblasts so that it is not
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surprising that we see with ASC what was seen with fibroblasts. In a recent article, coculture
of ASC and HUVEC was studied in monolayer coculture on tissue culture plastic. They found
that while both fibroblasts and smooth muscle cells promoted HUVEC tube formation in
coculture, ASC promoted vascular network formation much more strongly based on total tube
length and network branching[91]. It appeared that soluble factors secreted by a supporting cell
are needed to encourage HUVEC sprouting and maturation into a stable vascular structure in
vitro and that the paracrine relationship between ASC and EC has a stronger angiogenic effect.
But the question remains what is this paracrine relationship? Are the paracrine effects the only
contribution to the observed angiogenic effect on EC both in vivo and in the in vitro angiogenesis
assay?
The paracrine effects between adipogenesis and angiogenesis have been studied in the
literature, typically by blockade of an angiogenic growth factor or knock-out of a specific
angiogenic gene but there is no clear consensus in the literature as to what is really happening.
VEGF and hepatocyte growth factor secreted by ASC, and PDGF of endothelial origin, have
been shown to be essential for ASC-EC vascular network formation[91]. Knock-out of PPARγ
(master regulator of adipogenesis) has been shown to not only abrogate fat development in vivo,
but angiogenesis as well[150]. The RT-PCR results obtained here do not support the notion that
HUVEC have an effect on the mRNA expression levels in the ASC. This may not be surprising
as ASC differentiation in vitro is conducted in serum-free medium containing a defined cocktail
of adipogenic factors. Perhaps the EGM-2 media used in the coculture experiments contained
growth factors, such as bFGF, at concentrations that promote a proliferative phenotype of the
ASC. This was observed qualitatively, as ASC cultured on TCPS in EGM-2 appeared to grow
much more rapidly and reach confluence sooner than ASC cultured in the standard DMEM/F12-
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Ham’s media. For the angiogenic genes, expression of PAI-1, TNFα, and VEGF all appeared to
be increased at early timepoints in the ASC in the coculture system. These three growth factors
are hypoxia-inducible angiogenic factors that are known to be increased with obesity[151,152].
Interestingly expression of these three growth factors were reported to increased at early
timepoints in an in vivo model of adipose tissue development[150]. Yet without further
evidence, it is impossible to say if the increase in mRNA expression of these three growth factors
at early time points is responsible for downstream angiogenic events.
The gene expression from HUVEC, revealed no clear impact of ASC coculture besides
increase in VEGF expression at day 3 and 14. Increased expression of VEGF is not surprising as
it is a potent angiogenic growth factor responsible for the initiation of the angiogenic cascade
and the most studied growth factor in the paracrine effect between adipogenesis and
angiogenesis. It appeared that the RT-PCR data does not agree with the observed increase in
HUVEC sprout number and length with ASC coculture. Perhaps, other genes than those
investigated are involved in the observed effects including genes involved in cell apoptosis (e.g.
caspase-3, p53).
In the in vitro angiogenesis assay we studied only the paracrine effects of ASC-EC
interaction since there was no contact between the two cell types. Perhaps this is not the only
mechanism behind the observed ASC-EC interaction. Merfeld-Clauss et al. have shown that
ASC are localized in the peri-endothleial layer in native fat tissue, and can acquire properties of
pericytes both in vitro and in vivo[87,129]. In their in vitro studies they showed that vascular
network density was 60% lower for EC in direct contact with human dermal fibroblasts than that
observed for EC in direct contact with ASC in the very same well. They went on to show that
ASC were localized in proximity to the EC cords and that the areas distant from EC cords were
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clearly depleted with ASC[91]. The ASC that were adjacent to EC showed an increase in α-
smooth muscle actin expression and organization into fibres. Perhaps, the EC were
differentiating the ASC into pericyte-like cells that are able to stablize the growing vasculature.
While these are only recently published observations, it appears that both paracrine
communication and physical contact between ASC and EC may be necessary for the sequence of
events leading to the initiation, growth and stabilization of vascular structures both in vitro and
likely in native adipose tissue in vivo. We observed an increase in both HUVEC sprout number
and length with ASC cocultre, even without direct ASC-EC contact. Perhaps, the paracrine
effects are sufficient to establish a vascular network while pericytes further strengthen the
developing vasculature into stable vessels that can support blood flow.
5.5 Conclusions
Using an in vitro angiogenesis assay we have shown that the presence of ASC in
coculture promotes an angiogenic phenotype (sprouting and migration) of HUVEC. HUVEC in
the presence of ASC, were seen to sprout from the surface of the modules by day 3, elongate and
branch by day 7 and form a dense network of vessels by day 14 that merged with sprouts from
nearby modules. In contrast, HUVEC without ASC coculture initially formed sprouts, but these
degenerated into a mass of invidual cells surrounding the modules by day 14. RT-PCR analysis
of key adipogenic and angiogenic genes was variable, possibly due to different ASC donors. No
clear difference in mRNA expression of adipogenic genes was observed for ASC in HUVEC
coculture. Increased expression of PAI-1, VEGF and TNFα was observed in ASC with coculture
with HUVEC at day 3, and these may be important players in the paracrine signaling between
ASC and EC. The gene expression from HUVEC revealed no clear impact of ASC coculture
besides upregulation of VEGF at day 3 and 14. Recent reports in the literature highlight that not
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only paracrine signaling, but direct ASC-EC contact may be important in creating a vascular
network as ASC possess characteristics of pericytes.
5.6 Acknowledgements
The authors acknowledge the financial support by NSERC. Special thanks to Lindsay
Fitzpatrick for her help with RT-PCR.
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6 Synopsis
Modular tissue engineering represents a shift in the traditional cell + scaffold paradigm.
Producing small sub-mm sized hydrogel modules embedded with functional cells and coated
with a layer of EC is a means to produce scalable, vascularized tissue in vivo. For adipose tissue
engineering, the modules are embedded with ASC and surface coated with HMEC. We imagine
thousands of these modules implanted at the site of a soft tissue defect to restore normal
cosmesis. Once the modules are implanted, the transplanted HMEC are expected to quickly
form a vascular network that anastomoses with the host, while the ASC differentiate into mature
adipocytes resulting in a fat pad. This project has addressed the two main reasons where other
approaches to adipose tissue engineering have failed, mainly; 1) insufficient mechanical strength
of the cellular support structure and 2) insufficient vascularization upon implantation to support
transplanted cells. We explored the use of a semi-synthetic collagen/poloxamine hydrogel to use
as a biomaterial for adipose tissue engineering by testing its ability to support ASC viability and
differentiation capacity and EC attachment to the surface. Next we investigated the ability of the
modular approach to produce vascularized adipose tissue in vivo that persisted for the long-term
(3 months). Then to better understand the angiogenic remodeling of the transplanted HMEC in
vivo, we investigated the paracrine relationship between ASC and EC in an in vitro model of
angiogenesis with RT-PCR analysis of key angiogenic and adipogenic genes. This chapter
highlights some of the lessons learned and gives suggestions of future experiments and
strategies.
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6.1 Biomaterials for Adipose Tissue Engineering
To successfully create a tissue engineered construct, a support structure is needed to serve
as a site for cellular attachment and guidance in three-dimensions. Not only must this support
structure possess the needed biological properties to promote the desired function of the
transplanted cell, but in adipose tissue engineering it must also demonstrate favorable
mechanical properties as well. Rigid or stiff materials (ie. PLGA) that are beneficial for load
bearing applications are not suitable for soft tissues where the esthetic shape and feel is most
important. Degradation of the scaffold material is also important. If the scaffold does not
degrade quickly enough, host fibrous tissue may impair differentiation of transplanted ASC and
vascular ingrowth. On the other hand, rapid degradation of the scaffold may result in the loss of
the desired three-dimensional shape before the maturation of the fat pad has occurred. An
appropriate balance between scaffold degradation and adipose tissue accumulation must be
achieved.
Hydrogels are considered optimal for adipose tissue engineering because they mimic the
feel of native adipose tissue most closely. Often they can be injected at the site of the tissue
defect in a minimally invasive procedure. In the module approach, while the polymer was not
injected and polymerized in situ, the modules were small enough to be injected through an 18-
gauge needle which is advantageous in a clinical setting. Yet, synthetic polymers are
disadvantageous in that they typically are not cell adhesive and must be modified to contain
bioactive moeities. The collagen/poloxamine hydrogel, for example, was unsuccessful as a
biomaterial for adipose tissue engineering (Chapter 3). Simple mixing of the collagen and
poloxamine before photopolymerization resulted in a network of the two components where
entropy-driven phase separation resulted in regions in the hydrogel devoid of collagen that
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spanned distances greater than 100µm. ASC that were in these regions were not viable and did
not differentiate into adipocytes. We originally hypothesized that the 2-phase network of
collagen and poloxamine was advantageous for ASC differentiation. While ASC are anchorage-
dependent cells, adipocytes may prefer a less cell adhesive environment as they are known to
detach from TCPS during lipogenesis in in vitro culture. We thought that the ASC would attach
to regions in the hydrogel rich in collagen and during adipogenesis cell-secreted collagenase
would degrade the collagen so that mature adipocytes would be in the less cell adhesive
poloxamine phase. Yet, because of extent of phase separation between the collagen and
poloxamine components, many cells were only in the poloxamine phase and died. It appears that
at least some initial attachment to the matrix is needed to support ASC viability before
differentiation commences. We could explore the nature of ASC differentiation in a non-
adhesive substrate by using collagen and poloxamine in a slightly different arrangement. ASC
could be embedded in small amounts of collagen gel (ie. collagen microparticles) which are
subsequently photopolymerized within a poloxamine matrix. The ASC would initially be in
collagen to enable attachment and proliferation, yet during lipogenesis the cell secreted
collagenase would dissolve the collagen support and the resulting adipocytes would be in the
non-adhesive poloxamine. Oil Red O staining and RT-PCR analysis of adipogenic genes could
be used to determine the effect of the substrate on differentiation as compared to a collagen
control.
For a synthetic hydrogel to work as a biomaterial for adipose tissue engineering it must
be tailored to support both ASC attachment and proliferation onto the matrix and subsequent
differentiation into adipocytes as well. Poloxamine could be chemically modified to contain cell
adhesive molecules such as RGD or YIGSR that would enable ASC attachment and spreading
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within the matrix as well as EC surface attachment. In addition, a degradable version of
poloxamine must be investigated as well. Optimized degradation of the matrix would occur if
degradation was controlled by the differentiation of the ASC. This would ensure that the matrix
does not degrade too quickly before maturation of the fat pad has been achieved. The
collagenase-sensitive peptide sequence Leucine-Glycine-Proline-Alanine (LGPA) as investigated
by Jennifer West and colleagues[63] could be incorporated into the poloxamine to allow cell-
mediated degradation of the matrix during adipogenesis. Poloxamine, incorporating both
YIGSR and LGPA, appears to be a better approach to enabling ASC attachment, proliferation
and differentiation as well as degradation of biomaterial and may have future applications in
adipose tissue engineering.
6.2 Revascularization Strategies
Rapid vascularization of tissue engineered constructs upon implantation is critical to
support transplanted cells and integration of the device with the host. The lack of vascularization
is the main reason why both autologous fat transfers and previous approaches to tissue
engineered adipose constructs have been unsuccessful in repairing soft tissue deficits. We
demonstrated that the modular approach to adipose tissue engineering is successful in rapidly
creating a vascular network and promoting long-term fat development (3 months) in a construct
of size greater than the diffusional limitation of oxygen and nutrient transfer. Our approach has
the advantage that both ASC and HMEC are derived from the same patient, meaning that the
adipose tissue engineered construct can be an autologous transplant and hence may avoid
immunogenic rejection.
Other reports have demonstrated that EC transplantation as a revascularization strategy
can only be successful with transfection of an anti-apoptotic gene or with the presence of a
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supporting cell[119,122]. We found that without ASC cotransplantation, HMEC on the surface
of the modules did not result in a new vasculature and died over 14 days in vivo, even in the
immunodeficient SCID mouse used as the animal model here. Yet, with ASC embedded inside
the modules, the HMEC on the surface quickly detached and formed primitive vessel-like
structures that contained erythrocytes as early as day 3, suggesting inosculation with the host.
As time progressed, the number of vessels decreased but the size of the vessels increased
indicating vessel maturation and connection with the host was proved by Microfil perfusion at
day 21. It appeared that the ASC had an angiogenic effect on the transplanted HMEC and
promoted HMEC viability. This was not unique to HMEC as implants with HUVEC revealed a
similar angiogenic response.
The angiogeneic effects may be explained by the paracrine relationship between ASC
and EC. As discussed in Chapter 5, ASC are known to produce EC-specific mitogens and
angiogenic factors, while EC produce factors that stimulate ASC proliferation and
differentiation[89]. Yet, recent reports have suggested that ASC are localized in the peri-
endothelial layer of native adipose tissue and may be able to aquire properties of pericytes both
in vitro and in vivo following harvesting and expansion[87,129]. The hypothesis here is that cell
contact between the ASC ‘pericytes’ and the EC may help to stabilize vascular network
formation. While EC vessels may initially form, without this mural layer (ie. pericytes or
smooth muscle cells) these small vessels may degrade followed by EC death, which is what was
observed in the case without ASC cotransplantation.
It would be interesting to study and track the fate of the ASC in vivo to see if
ASC/pericyte vessel stabilization is responsible for the formation of the vascular network. ASC
could be harvested from a green fluorescent protein (GFP) expressing mouse so that we could
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see if the ASC were migrating from the inside of the modules and then localizing at the vessel
walls of the HMEC-derived vessels.
While there is a lack of known ASC specific stem cell markers that could be used to track
the fate of these cells in vivo, there is some evidence that subpopulations in the stromal vascular
fraction of adipose tissues may give rise to different cell types. CD34- populations have been
shown to express more pericyte markers and may be the subpopulation of ASC that differentiate
into pericytes[87,88]. We could use flow cytometry to separate different subpopulations of ASC
and use these cells embedded into the modules. We would then compare the effect of CD34-
enriched populations of ASC on vessel formation in vivo. If ASC-EC cell contact proves
important in the vascular remodeling process, we could design modules that contain embedded
ASC with a mixture of HMEC and ASC attached on the surface. Perhaps this would accelerate
the remodeling of the HMEC-derived vascular network because supporting ASC would not have
to migrate from inside the collagen gel to the module surface.
While the understanding of why ASC promote and support HMEC vessel formation is
unclear, the formation of a rapid vascular network is needed to support transplanted cells. Only
in the case of cotransplantation did we see fat accumulation inside the modules. ASC-only
modules without HMEC on the surface showed little to no visible signs of fat accumulation by
histological analysis. This is likely the result of a lack of sufficient blood vessel network to
nourish the implant, but may also be contributed to by the paracrine effects of EC/ASC
interaction. With the lack of an appropriate stem cell marker for ASC, we cannot confirm that
the fat accumulation is due to transplanted ASC and not host cells, but the location of the fat
deposits seen inside individual modules is compelling. In the case of cotransplantation, fat
accumulation was seen to begin at day 30 and increase until day 90, yet the volume of the
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implants decreased over time. This is not surprising as collagen gel was used as the module
material and degrades too quickly to be used by itself as a biomaterial for adipose tissue
engineering. In an in vivo experiment, it would be interesting to strengthen the collagen gels,
eg., by crosslinking the gels by transglutaminase[153], to see if stiffer collagen modules when
implanted would degrade more slowly so that maturation of the fat pad happens before the
support structure has degraded to promote a long-term volume stable mass of adipose tissue.
6.3 Understanding the relationship between ASC and EC
While the in vivo experiments showed qualitatively that ASC had an angiogenic effect on
transplanted EC, we further explored the relationship between ASC and EC in vitro to help
elucidate the paracrine relationship between the two cell types. Adapting an in vitro
angiogenesis model as developed by Nehls and Drenckhahn[146], we embedded HUVEC coated
modules inside a fibrin gel with ASC-embedded in a collagen gel separated in a transwell so that
no ASC-EC direct contact occurred. The primary reason for this was to ensure easy separation
of the two cell types for RT-PCR analysis. In accordance with the in vivo results, HUVEC were
seen to sprout and form a dense vascular network over 14 days in the fibrin gel with the presence
of ASC. In the wells without the ASC, HUVEC initially formed some sprouts but these began to
degenerate by day 7 and by day 14 only an unorganized cluster of individual cells was seen
surrounding the modules. Without any direct ASC-EC contact this is an example of paracrine
signaling between the two cell types that helped establish and mature the HUVEC-derived
vascular network.
As discussed in the previous section, it would be interesting to explore the role of ASC as
pericytes to support the growing vascular network. As evidenced here, ASC as pericytes (ie.
with direct contact) were not needed to successfully form a dense sprouting network of HUVEC
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in fibrin gels, yet ASC as pericytes may (further) aid the vascular network formation in vivo. As
a next experiment, we would use ASC-embedded modules coated with HUVEC, or a mixture of
ASC and HUVEC on the module surface in the in vitro angiogenesis assay to explore the effect
of ASC-EC contact on vascular network formation. We would label the ASC with a fluorescent
dye (ie. DiI) and the EC could be tracked with CD31 antibody staining. Using confocal
microscopy we could locate the presence of ASC surrounding the HUVEC sprouts to see if both
cell types were colocalized suggesting that ASC is supporting the developing vasculature.
Others have also found that α-smooth muscle actin (αSMA) expression is upregulated in EC-
ASC cocultures[91]. αSMA antibody staining could be used to see if it is upregulated in ASC
with close proximity to the HUVEC and its organization into fibres.
The sprouting results showed the clear impact that the ASC had on the vascular network
formation of HUVEC from modules, so it was expected that RT-PCR analysis of key adipogenic
and angiogenic genes from these experiments would further confirm these observations. The
coculture with HUVEC resulted in little change in mRNA expression of early, mid and late
markers of adipogenesis in the ASC. Perhaps, the medium used in this experiment (EGM-2)
masked any possible impact because it contained numerous growth factors that promote a more
proliferative phenotype of ASC. Growth arrest (ie. serum free medium) is normally a
prerequisite for ASC differentiation in vitro. For the angiogenic factors investigated, VEGF,
PAI-1 and TNFα expression were increased in ASC at early timepoints with HUVEC coculture.
These three factors are likely players in the paracrine relationship between ASC-EC but without
further investigation (e.g. inhibition of adipogenesis or angiogenesis with a blocking antibody, or
gene transfection) this cannot be concluded here. The data tended to be quite variable, which
may be the result of ASC donor variability. Two of the experiments were conducted with ASC
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from the same donor and the results of those two experiments appeared to be less variable for
many of the genes investigated.
For HUVEC, it appeared that ASC coculture resulted in increased expression of VEGF at
early time points which is not surprising as VEGF is a potent initiator in the angiogenic cascade.
Again the results tended to be variable making it difficult to draw conclusions. Perhaps, instead
of looking at a small subset of genes a gene array should be used to look at multiple genes at
once. The TaqMan “Human Angiogenesis Array” contains 94 angiogenic targets, including
angiogenic growth factors, inhibitors, markers and targets and may provide a better insight into
the angiogenic remodeling of HUVEC by ASC. It would be interesting to also test genes
involved in cell apoptosis since the ASC were shown to promote HMEC viability in vivo.
6.4 Future Directions
The future of adipose tissue engineering is a bright one. There are two current challenges
that need to be addressed before an adipose tissue engineered construct can move from the
bench-top to the clinic:
1) A suitable biomaterial that possesses the necessary cell adhesive properties to allow
ASC attachment, proliferation and differentiation as well as the required mechanical
properties that give an appropriate ‘feel’ and degradation in the body and;
2) A successful revascularization strategy to nourish transplanted cells to encourage ASC
survival and differentiation into a mature fat pad that will retain its volume and shape.
In future work, biomaterial design will likely focus on cell-ECM interactions to provide
biological cues that will guide and direct ASC growth and differentiation. The interactions with
ASC and either synthetic or natural polymers, with and without soluble factors, could be used to
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create a biological niche that will direct cell fate. Cell shape, for example, has been shown to
drive MSC commitment. MSC cultured on fibronectin islands of restricted size (1,024µm2) were
shown to remain rounded and differentiated into adipocytes, while those cultured on larger
islands (10,000µm2) were able to spread and differentiated into osteoblasts[140]. Restricting the
cells from spreading abrogates the ability to form geometrically separated adhesions, where
when the cells are unrestricted they can form many focal adhesions that act on tensioning of the
cytoskeleton. For a hydrogel, the effect of tensioning of the cytoskeleton is not controlled by the
available surface area for attachment, since the cells are freely embedded or coated on the
surface, but rather the substrate elasticity. MSC cultured on soft substrates (Elastic modulus
(E)~0.1-1kPa) that closely mimic the mechanical properties of soft tissue were shown to highly
express neurogenic markers, while stiffer substrates with E~8-17kPa and E~25-40kPa were
shown to preferentially express myogenic and osteogenic markers respectively[154]. From this
study it appears that the elasticity of the microenvironment that closely matches that of the native
tissue helps drive differentiation into that particular lineage, so therefore a soft hydrogel is likely
optimal for ASC differentiation. Yet, substrate elasticity is not the only factor that can control
stem cell fate. While increases in gel stiffness from 0.7kPa to 80kPa increased MSC spreading
areas from 5,000 to 45,000µm2, those coated with laminin had only half of the spreading area
compared to those coated with fibronectin of the same stiffness[155]. This study shows that
incorporation of bioactive ligands into the biomaterial is another approach to direct stem cell
fate. In a recent publication, differentiation of MSC embedded in a PEG hydrogel was
controlled by incorporation of small-molecule functional groups. MSC encapsulated in a PEG
hydrogel containing charged phosphate groups were shown to undergo osteogenesis while those
cultured in hydrogels containing t-butyl groups differentiated into adipocytes[156]. These
126
studies highlight that both mechanical properties and the incorporation of bioactive
ligands/molecules are key design considerations when developing a biomaterial for adipose
tissue engineering. To help with biomaterial design, improved characterization of ASC and
understanding of the immunophenotype of the different populations of adipose progenitor cells is
needed. Furthermore, understanding the impact of in vitro cell conditioning of ASC prior to
implantation (ie. predifferentiation) may also affect the in vivo outcome.
Cotransplantation of ASC and EC in the modular approach appeared to be a successful
revascularization strategy in the work presented here. Yet further understanding of the ASC-EC
interaction on the molecular level is needed to fully exploit its potential. The capability of ASC
to not only differentiate into adipocytes but also endothelial cells and pericytes is not well
understood but is likely key to understanding ASC-EC vascular network formation in vivo. We
must look to the biology of the complex interplay between adipogenesis and angiogenesis for
answers. In addition to better understanding of the cells, is to better understand the implant
model to test the outcome of an engineered adipose construct. The majority of the adipose tissue
engineering studies to date have been subcutaneous implantation into rodent models. The next
step is to investigate other implant sites in larger animal models to study the remodeling of an
adipose tissue engineering construct of clinical relevance. All in all, to move adipose tissue
engineered constructs into clinical practice standardization of cell sourcing, biomaterial design,
and implant procedures is needed.
Looking at the bigger picture, an ASC-EC revascularization strategy may have widespread
application not only for adipose tissue engineering, but tissue engineering as a whole. Current
interest in the field of islet transplantation, for example, is the use of adipose tissue and ASC to
create a vascular network that will support islet engraftment. A recent publication demonstrated
127
the use of ASC combined with minced adipose tissue to create vascular-rich beds that were
suitable to support islet transplantation subcutaneously in diabetic mice. They were able to
correct blood glucose level within a week after islet transplantation and maintain normoglycemia
for over 8 weeks[157]. We can imagine combining ASC-EC modules with modules containing
islets where once implanted the EC rapidly create a functional vascular network that will support
the viability of the transplanted islets as a treatment for diabetes.
128
7 Conclusions
The overall objective of this project was to build a modular approach for adipose tissue
engineering to create vascularized adipose tissue in vivo that persisted long-term (3 months). In
addition we wanted to better understand the mechanism behind the vascularization seen with
transplantation of ASC in EC covered modules. We explored and tested three different
hypotheses:
1. Fabricate and optimize the modular platform for adipose tissue engineering to promote
embedded ASC viability, differentiation and attachment of surface coated endothelial cells
(EC).
Hypothesis: The semi-synthetic collagen/poloxamine hydrogel used as the module material
promotes high viability of embedded ASC and allows differentiation of these cells into mature
adipocytes, while enabling good adhesion and coverage of EC on the module surface.
Rejected: Entropic phase separation between collagen and poloxamine resulted in a 2-phase
biomaterial. ASC were not able to attach, proliferate and differentiate into regions of
the matrix rich in poloxamine. EC were unable to form a confluent layer on the
surface of the biomaterial.
2. Evaluate the regenerative potential of the modular construct to produce vascularized adipose
tissue in vivo.
Hypothesis: The endothelialized perfusion channels created by packing many modules into a
larger construct provide an immature capillary network that remodels and anastomoses with the
129
host vasculature upon implantation to promote transplanted ASC viability and facilitates a long-
term (3 months) volume-persistent culture of adipose tissue in vivo.
Accepted: Cotransplantation of ASC and HMEC in the modular approach resulted in rapid
vascular network formation that appeared to connect with the host vasculature. An
accumulation of adipose tissue was created that persisted for up to 3 months in vivo.
3. Investigate ASC/EC interaction in an in vitro sprouting assay.
Hypothesis: Coculture of ASC and HUVEC (no direct cell-cell contact) promotes an angiogenic
phenotype (i.e. migration and sprouting) of surface covered HUVEC in a fibrin gel.
Accepted: Coculture of ASC and HUVEC in an in vitro sprouting assay resulted in a dense
vascular network formation of HUVEC surrounding the modules within a fibrin gel.
HUVEC without ASC coculture showed initial sprouting and migration but this
network disintegrated over 14 days in culture. RT-PCR analysis demonstrated that
VEGF, PAI-1 and TNFα expression increased in coculture and may be involved in
paracrine signaling between ASC and EC.
The modular approach to tissue engineering was successfully applied to create
vascularized adipose tissue in vivo that persisted for at least 3 months. The success of the EC
transplantation revascularization strategy is attributed to, at least in part, to paracrine interactions
between ASC and EC. This work has potential in creating a clinically relevant adipose tissue
substitute. ASC-EC revascularization strategy may have widespread use in tissue engineering
applications.
130
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9 Appendix 1: HMEC Harvesting
The human dermal microvascular endothelial cell line (HMEC-1) was used as the EC
source in Chapter 3 because they show the characteristic morphology and protein expression of
primary human HMEC while requiring less stringent growth medium and can be passaged
indefinitely. While cell lines may be useful for preliminary experiments, they have been altered
and as such are not phenotypically equivalent to primary cells. The use of primary cells was
necessary for the experiments in Chapter 4 and 5 of this thesis. HMEC dervived from the dermis
is available commercially (Cambrex, VEC) but we feel that EC derived from the same fat source
as the ASC may have an advantage. Harvesting both the HMEC and ASC from a single tissue
sample, we can envision an engineered fat pad created entirely from the patient’s own cells. This
would overcome many of the immune/rejection issues associated with cell transplantation.
Review of the literature revealed that magnetic separation of the cells using anti-body
binding to paramagnetic beads was required to obtain a pure population of endothelial
cells[158,159]. Even with magnetic separation, obtaining a pure population of endothelial cells
is a challenge because any contaminating adipose precursor cells will quickly overgrow any
existing endothelial cell population. We employed the MiniMACS™ cell separation columns
using degradable magnetic beads bound with platelet-endothelial cell adhesion molecule 1
(PECAM-1) (Miltenyi Biotech.)
Cells were harvested from freshly excised adipose tissue as per ASC harvest protocol in
section 3.3.1. which contains a mixture of both ASC and HMEC. Table 3 highlights the
experiments, conditions and results of the cell separation. Cells obtained from the harvest were
either pre-plated in T-150 flasks or directly separated at Passage 0 (P0) using either 1 column or
2 columns in series (to improve separation efficiency). Cells were then cultured in T-75 flasks
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and were separated again at Passage 1 (P1). All trials that cultured the cells in ‘coculture media’
were found to have complete overgrowth of ASC at Passage 2. The ‘coculture media’ was
developed by a previous student in the lab and contains the ASC base media (DMEM-F12
Ham’s, 10% FBS) with additional factors EGF, hydrocortisone and L-glutamine to support
HMEC-1 proliferation. This media may not support the attachment/proliferation of primary
HMEC or promote rapid proliferation of contaminating ASC, making separation difficult. For
trail 6 and 7, MCDB-131 (VEC Technologies) complete media was used which is recommended
for primary HMEC culture and gave positive results. It was decided to separate the cells directly
from harvest as pre-plating was found to only give the ASC cells time to overgrow the HMEC
population. Trial 7, using 2 columns in series during each separation, was successful and yielded
a nearly pure population of HMEC. Light microscopy images revealed cells with a typical
cobblestone morphology similar to the HMEC-1 and characteristic of endothelial cells.
(contaminating ASC in Trial 6 can be observed as long, spindle-shaped cells which were not
apparent in Trial 7) (Figure 9-1). Immunostaining for vonWillebrand’s factor (vWF) (green) on
cells prepared by cytospin from Trial 6 and 7 confirmed the presence of endothelial cells (Figure
9-2). Many non-stained cells are observed in Trial 6, while using 2 columns in series at each
separation was key to obtaining a nearly pure culture of HMEC in Trial 7. Several successful
separations using the conditions of Trial 7 have been conducted with reproducible results.
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Table 3: HMEC Separation Trials
Trial Method Media P0 (# columns) P1 (# columns)
P2 (result)
1 Harvest coculture 1 1 ASC overgrowth
2 Preplating (1.5h) coculture 1 1 ASC overgrowth
3 Preplating (72h) coculture 1 1 ASC overgrowth
4 Harvest coculture 2 2 ASC overgrowth
5 Preplating (72h) coculture 2 2 ASC overgrowth
6 Harvest MCDB 131 1 1 ASC contamination
7 Harvest MCDB 131 2 2 EC cobblestone morphology
Figure 9-1: Light microscopy images of cells obtained from HMEC separation trials.
Light microscopy images of cells obtained from Trial 6, and 7 and HMEC-1. Cobblestone morphology of cells in Trial 7, similar to HMEC-1, is indicative of endothelial cells. ASC contamination is observed in Trial 6 with long, spindle-like cells present. Scale bar = 100µm.
HMEC-1 Trial 7 – 2 Columns Trial 6 – 1 Column
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Figure 9-2: Cytospin slides of cells from Trial 6 and 7 immunostained with vWF
Most cells in Trial 7 stain positive for vWF (green) indicating a homgeneous population of HMEC. Trial 6 cells are mostly unstained contaminating ASC. Hoescht stains cell nuclei blue.
Trial 6 – 1 column
Trail
Trial 7 – 2 columns
