Thesis_Final_w_cover.pdf - Worcester Polytechnic Institute

Acknowledgements 2

Acknowledgements

I would first like to thank my advisor, Marsha Rolle, for her endless support and guidance over

the past 5 years. I would also like to thank my committee, Kris Billiar, Glenn Gaudette, Suzanne

Scarlata, and Eben Alsberg for their feedback and guidance on this project.

I would like to thank our many collaborators over the years. I would like to thank Monica, Ben,

and Marco for designing our bioreactors and for continued assistance with setup and

troubleshooting. I would also like to thank Anna and Rui from the Alsberg lab at Case Western

Reserve University for fabricating the gelatin microspheres that are critical to this project, Yibing

and Jiesi from Yale University for providing iPSCs, and Tabby Ahsan from RoosterBio for

providing hMSCs.

I would like to thank the many graduate students who have helped me throughout my time at

WPI, especially Dalia Shendi, Jennifer Cooper, Zoe Reidinger, Beth Calamari, Joni Grosha,

Emily Caron, David Dolivo, Lindsay Lozeau, and Katrina Hansen. Your technical assistance

proved invaluable, and your support was greatly appreciated.

I would also like to thank the countless undergraduate students who have assisted with this

project, often doing endless staining, imaging, and image analysis. I would like to thank our

histology technicians Hans Snyder and Jyotsna Patel for their assistance with histology and

training undergraduate students.

Finally, I would like to thank my friends and family for their support throughout my time at

WPI, especially my husband Michael, for his patience with my many late nights and long

weekends in the lab.

Table of Contents 3

Table of Contents

Acknowledgements ...................................................................................................................... 2

Table of Contents ......................................................................................................................... 3

Abstract ...................................................................................................................................... 10

Abbreviations ............................................................................................................................. 11

Table of Figures ......................................................................................................................... 13

Table of Tables........................................................................................................................... 18

Chapter 1: Executive Summary .................................................................................................. 19

1.1. Introduction ..................................................................................................................... 19

1.2. Overview of aims ............................................................................................................. 20

Aim 1: Develop a system to locally deliver bioactive factors within tissue rings. ............... 20

Aim 2: Fuse human SMC rings into tissue tubes and evaluate the effects of dynamic culture.

............................................................................................................................................ 21

Aim 3: Create vascular tissue tubes with spatially distinct regions ..................................... 22

1.3. Summary ......................................................................................................................... 23

1.4. References ....................................................................................................................... 24

Chapter 2: Background ............................................................................................................... 26

2.1. Smooth muscle phenotype ............................................................................................... 26

2.2. Intimal hyperplasia .......................................................................................................... 27

2.3. Treatments for IH ............................................................................................................ 28

2.4. Model systems for studying IH ........................................................................................ 29

2.5. Tissue engineered blood vessels as in vitro human vascular models. ............................... 30

2.6. Modular fabrication of vascular tissue constructs from self-assembled cell ring units. .... 31

2.7. Engineering custom agarose molds for self-assembled tissue ring fabrication ................. 31

2.8. Microsphere incorporation and modular assembly to create focal regions of IH .............. 34

2.9. Bioactive molecule release from tissue engineered blood vessels .................................... 34

2.10. Microsphere-mediated growth factor delivery in engineered vascular tissue ................. 35

2.11. Platelet-derived growth factor ........................................................................................ 35

2.12. Gelatin microspheres for controlled delivery of PDGF .................................................. 36

Table of Contents 4

2.13. Summary ....................................................................................................................... 37

2.14. References ..................................................................................................................... 38

Chapter 3: Cellular self-assembly with microsphere incorporation for growth factor delivery

within engineered vascular tissue rings ...................................................................................... 50

3.1. Introduction ..................................................................................................................... 50

3.2. Materials and methods ..................................................................................................... 52

3.2.1. Gelatin microsphere preparation ................................................................................ 52

3.2.2. Human smooth muscle cell culture ............................................................................ 52

3.2.3. Smooth muscle cell ring self-assembly and unloaded microsphere incorporation ..... 52

3.2.4. TGF-β1-loaded microsphere preparation and incorporation within tissue rings ........ 53

3.2.5. Histology and immunohistochemistry ....................................................................... 53

3.2.6. SMC ring thickness and diameter measurements ...................................................... 54

3.2.7. Mechanical testing .................................................................................................... 54

3.2.8. Western blot analysis ................................................................................................ 54

3.2.9. Statistical analysis ..................................................................................................... 55

3.3. Results ............................................................................................................................. 55

3.3.1. Gelatin microsphere characterization ........................................................................ 55

3.3.2. Effects of microsphere incorporation on self-assembled SMC rings cultured in growth

medium ............................................................................................................................... 56

3.3.3. Effects of microsphere incorporation on self-assembled SMC rings cultured in

differentiation medium ........................................................................................................ 57

3.3.4. TGF-β1 delivery from incorporated microspheres within self-assembled SMC rings 57

3.4. Discussion ....................................................................................................................... 62

3.5. References ....................................................................................................................... 66

Chapter 4: Fabrication and characterization of electrospun polycaprolactone cuffs for self-

assembled vascular tissue ........................................................................................................... 72

4.1. Introduction ..................................................................................................................... 72

4.2. Methods ........................................................................................................................... 74

4.2.2. Fabrication of PCL electrospun cuffs ........................................................................ 74

4.2.3. Fiber diameter measurement ..................................................................................... 75

4.2.4. Tensile testing of electrospun cuffs ........................................................................... 75

4.2.5. Cell culture ................................................................................................................ 75

Table of Contents 5

4.2.6. TEBV fabrication from self-assembled tissue rings................................................... 75

4.2.7. Longitudinal pull to failure testing ............................................................................ 76

4.2.8. Hoechst staining ........................................................................................................ 76

4.3. Results ............................................................................................................................. 76

4.3.1. Characterization of electrospun PCL cuffs ................................................................ 76

4.4. Discussion ....................................................................................................................... 78

4.5. References ....................................................................................................................... 79

Chapter 5: Generate modular vascular tissue tubes with luminal flow ....................................... 83

5.1. Introduction ..................................................................................................................... 83

5.2 Methods ............................................................................................................................ 85

5.2.2. Tissue ring fabrication ............................................................................................... 85

5.2.3. Tissue tube fusion with varying pre-culture time....................................................... 85

5.2.4. Fusion angle, length, and thickness measurements .................................................... 86

5.2.5. CellTracker labeling .................................................................................................. 87

5.2.6. Polycaprolactone (PCL) cannulation cuff fabrication ................................................ 87

5.2.7. Bioreactor culture ...................................................................................................... 87

5.2.8 Histology and immunohistochemistry ........................................................................ 88

5.2.9. Statistics .................................................................................................................... 89

5.3. Results ............................................................................................................................. 89

5.3.1. Effect of ring pre-culture time on human SMC tube fusion rate ................................ 89

5.3.2. Structure and morphology of fused human SMC tubes ............................................. 90

5.3.3. Spatial positioning of SMCs within rings during fusion ............................................ 90

5.3.4. PCL cannulation cuffs and dynamic tube culture .................................................. 92

5.4. Discussion ....................................................................................................................... 94

5.5 References ........................................................................................................................ 98

Chapter 6: Create vascular tissue tubes with spatially distinct regions ..................................... 103

6.1. Introduction ................................................................................................................... 103

6.2. Methods ......................................................................................................................... 104

6.2.1. Cell culture .............................................................................................................. 104

6.2.2. Ring fabrication ....................................................................................................... 105

6.2.3. Tube fabrication for fusion comparison ................................................................... 105

Table of Contents 6

6.2.4. Fabricating tubes with spatially defined regions of microsphere incorporation ....... 106

6.2.5. PDGF treatment of 2D cell cultures ........................................................................ 106

6.2.6. PDGF treatment of self-assembled SMC rings ........................................................ 106

6.2.7. Histology and immunohistochemistry ..................................................................... 106

6.2.8. Statistical analysis ................................................................................................... 107

6.3. Results ........................................................................................................................... 108

6.3.1. Effect of microspheres on tube fusion ..................................................................... 108

6.3.2. Fabrication of a focal region of microsphere incorporation ..................................... 109

6.3.3. Effect of PDGF on proliferation of 2D SMC cultures ............................................. 110

6.3.4. Effect of microsphere-mediated PDGF release on SMC rings ................................. 110

6.4. Discussion ..................................................................................................................... 111

6.5. References ..................................................................................................................... 114

Chapter 7: Induced pluripotent stem cells as an alternative human smooth muscle cell source for

vascular tissue engineering ....................................................................................................... 117

7.1. Introduction ................................................................................................................... 117

7.2. Methods ......................................................................................................................... 118

7.2.1. Ring culture ............................................................................................................. 118

7.2.2. Tube culture ............................................................................................................ 118

7.2.3. iPSC-vSMC response to PDGF in 2D ..................................................................... 119

7.2.4. iPSC-vSMC response to PDGF in 3D ..................................................................... 119

7.2.5. Mechanical testing .................................................................................................. 120

7.2.6. Histological analysis and immunohistochemistry .................................................... 120

7.2.7. Western blotting ...................................................................................................... 120

7.3. Results ........................................................................................................................... 120

7.3.1. Ring formation and characterization ........................................................................ 120

7.3.2. iPSC-vSMC response to PDGF ............................................................................... 121

7.3.3. Tube fabrication ...................................................................................................... 122

7.4. Discussion ..................................................................................................................... 123

7.5. References ..................................................................................................................... 125

Chapter 8: Growth factor delivery for phenotypic modulation of human mesenchymal stem cell

rings ......................................................................................................................................... 129

Table of Contents 7

8.1. Introduction ................................................................................................................... 129

8.2. Methods ......................................................................................................................... 130

8.2.1. Cell culture .............................................................................................................. 130

8.2.2. Ring culture ............................................................................................................. 130

8.2.3. Ring thickness measurements .................................................................................. 132


8.2.5. DNA quantification ................................................................................................. 132

8.2.6. PDGF loading efficiency ......................................................................................... 133

8.2.7. Statistical analysis ................................................................................................... 133

8.3. Results ........................................................................................................................... 133

8.3.1. Effects of microsphere-mediated PDGF release on hMSC rings ............................. 133

8.3.2. Effect of microsphere-mediated FGF release on hMSC rings .................................. 137

8.3.3. Effect of microsphere-mediated TGF-β1 release on hMSC rings ............................ 141

8.4. Discussion ..................................................................................................................... 144

8.5. References ..................................................................................................................... 149

Chapter 9: Vascular tissue tubes with distinct phenotypic and structural regions ..................... 154

9.1. Introduction ................................................................................................................... 154

9.2. Methods ......................................................................................................................... 155

9.2.1. Cell culture .............................................................................................................. 155

9.2.2. Ring fabrication ....................................................................................................... 155

9.2.3. hMSC tube fabrication ............................................................................................ 155


9.3. Results ........................................................................................................................... 157

9.3.1. Fabrication of tissue tubes from hMSC rings .......................................................... 157

9.3.4. Focal region of synthetic SMCs .............................................................................. 158

9.4. Discussion ..................................................................................................................... 158

9.5. References ..................................................................................................................... 161

Chapter 10: Conclusions and future work ................................................................................ 163

10.1. Summary ..................................................................................................................... 163

10.2. Other applications of the ring system ........................................................................... 164

10.3. Limitations ................................................................................................................... 165

Table of Contents 8

10.4. Future work ................................................................................................................. 166

10.5. References ................................................................................................................... 168

Appendix A: Reprint permission for Chapter 2.7 ..................................................................... 172

Appendix B: Reprint permission for Chapter 3 ........................................................................ 173

Appendix C: Chapter 3 supplemental data ............................................................................... 174

Supplemental methods .......................................................................................................... 174

Cell culture ........................................................................................................................ 174

Supplemental figures ............................................................................................................ 174

Appendix D: Reprint permission for Chapter 4 ........................................................................ 177

Appendix E: Chapter 4 supplemental data ................................................................................ 183

Appendix F: Chapter 5 supplemental data ................................................................................ 184


Cell culture ........................................................................................................................ 184


Appendix G: Microsphere characterization .............................................................................. 187

Appendix H: Supplemental data for Chapter 8 ......................................................................... 188



Appendix I: Automation and scale-up of tissue tube production .............................................. 191

Abstract ................................................................................................................................ 191

I.1. Introduction .................................................................................................................... 192

I.2. Methods .......................................................................................................................... 195

I.2.1. Mold design ............................................................................................................. 195

I.2.2. Cell culture .............................................................................................................. 195

I.2.3. Agarose gel preparation ........................................................................................... 196

I.2.4. Ring fabrication ....................................................................................................... 196

I.2.5. Mechanical testing ................................................................................................... 196

I.2.6. Histology ................................................................................................................. 197

I.2.7. Robotic punch design ............................................................................................... 197

I.2.8. Tube fabrication ....................................................................................................... 198

I.2.9. Statistics ................................................................................................................... 199

Table of Contents 9

I.3. Results ............................................................................................................................ 199

I.3.1. Ring fabrication in PEI-MED610 plate system ........................................................ 199

I.3.2. Automation system .................................................................................................. 200

I.3.3. Tube fusion following automated ring stacking ....................................................... 201

I.4. Discussion ...................................................................................................................... 202

I.5. References ...................................................................................................................... 205

Abstract 10

Abstract

Tissue engineered blood vessels (TEBVs) have great potential as tools for disease

modeling and drug screening. However, existing methods for fabricating TEBVs create

homogenous tissue tubes, which may not be conducive to modeling focal vascular diseases such

as intimal hyperplasia or aneurysm. In contrast, our lab has a unique modular system for

fabricating TEBVs. Smooth muscle cells (SMCs) are seeded into an annular agarose mold,

where they aggregate into vascular tissue rings, which can be stacked and fused into small

diameter TEBVs. Our goal is to create a platform technology that may be used for fabricating

focal vascular disease models, such as intimal hyperplasia. Because tubes are fabricated from

individual ring units, each ring can potentially be customized, enabling the creation of focal

changes or regions of disease along the tube length. In these studies, we first demonstrated our

ability to modulate cell phenotype within individual SMC ring units using incorporated growth

factor-loaded degradable gelatin microspheres. Next, we evaluated fusion of ring subunits to

form composite tissue tubes, and demonstrated that cells retain their spatial positioning within

individual rings during fusion. By incorporating electrospun polycaprolactone cannulation cuffs

at each end, tubes were mounted on bioreactors after only 7 days of fusion to impart luminal

medium flow for 7 days at a physiological shear stress of 12 dyne/cm2. We then created focal

heterogeneities along the tube length by fusing microsphere-containing rings in the central region

of the tube between rings without microspheres. In the future, microspheres may be used to

deliver growth factors to this localized region of microsphere incorporation and induce disease

phenotypes. Due to the challenges of working with primary human SMCs, we next evaluated

human mesenchymal stem cells (hMSCs) as an alternative cell source to generate vascular

SMCs. We evaluated the effects of microsphere-mediated platelet-derived growth factor

(PDGF), fibroblast growth factor (FGF), and transforming growth factor beta-1 (TGF-β1)

delivery on ring thickness, proliferation, and contractile protein expression over a 14 day period.

Finally, we created a structurally distinct region of smooth muscle within tissue tubes by fusing

human aortic SMCs in a central region between hMSC rings. In summary, we developed a

platform technology for creating modular tubular tissues that may be further developed into an in

vitro intimal hyperplasia model. It may also be modified to model other focal vascular diseases,

such as aneurysm, or to create other types of multi-tissue tubular structures, such as trachea.

Abbreviations 11

Abbreviations

ANOVA – Analysis of variance

BCA – Bicinchoninic acid

DMEM – Dulbecco’s modified eagle medium

CALP – Calponin

CAM – Computer-aided manufacturing

CNC – Computer numerical control

ECM – Extracellular matric

ECs – Endothelial cells

EDTA – Ethylenediaminetetraacetic acid

EdU – 5-ethynyl-2’-deoxyuridine

EGF – Epidermal growth factor

FBS – Fetal bovine serum

FGF – Fibroblast growth factor

H&E – Hematoxylin and Eosin

hMSC – Human mesenchymal stem cell

HRP – Horse radish peroxidase

IGF – Insulin-like growth factor

IH – Intimal hyperplasia

IHC – Immunohistochemistry

IL-1 – Interleukin 1

IL-6 – Interleukin 6

iPSC-vSMCs – Induced pluripotent stem cell-derived vascular smooth muscle cells

ITS – Insulin transferrin selenium

MAPK – Mitogen activated protein kinase

MS – Microspheres

MTM – Maximum tangent modulus

NBF – Neutral buffered formalin

NGS – Normal goat serum

NO – Nitric oxide

NRS – Normal rabbit serum

PBS – Phosphate buffered saline

PCL – Polycaprolactone

PDGF – Platelet derived growth factor

PDMS – Polydimethylsiloxane

PGA – Poly-glycolic acid

PLA – Poly-lactic acid

PLGA – poly(lactic-co-glycolic) acid

SDS – Sodium dodecyl sulfate

Abbreviations 12

SEM – Scanning electron microscopy

SMA – Smooth muscle alpha actin

SM-22α – Smooth muscle protein 22 alpha

SMC – Smooth muscle cell

SVAS – Supravalvular aortic stenosis

TBST – Tris buffered saline plus tween

TEBV – Tissue engineered blood vessel

TFE – Tri-fluoroethanol

TGF-β1 – Transforming growth factor beta 1

UTS – Ultimate tensile strength

VEGF – Vascular endothelial growth factor

Table of Figures 13

Table of Figures

Figure 1.1: Aim 1 overview: Develop a system to locally deliver bioactive factors

within tissue rings ………………………………………………………………………... 20

Figure 1.2: Aim 2 overview: Fuse human SMC rings into tissue tubes and evaluate the

effects of dynamic culture………………………………………………………………... 21

Figure 1.3: Aim 3 overview: create vascular tubes with distinct regions………………… 22

Figure 2.1: Characteristics of IH model lesion …………………………………………... 30

Figure 2.2: Cross-sectional view of 3D printed mold ……………………………………. 33

Figure 2.3: Fabrication of self-assembled tissue rings …………………………………... 33

Figure 2.4: Fabrication of vascular tissue tubes …………………………………………. 34

Figure 2.5: Schematic of method for fabricating TEBV with intimal lesion ……………. 34

Figure 3.1: Schematic of microsphere incorporation within self-assembled tissue rings... 52

Figure 3.2: Gelatin microsphere incorporation within rings …………………………….. 56

Figure 3.3: Effects of microsphere incorporation on thickness of rings cultured in

growth medium …………………………………………………………………………... 56

Figure 3.4: Mechanical properties of 14 day-old rings cultured in growth medium……... 57

Figure 3.5: Microsphere incorporation in rings cultured in differentiation medium……... 58

Figure 3.6: Effects of microsphere incorporation on thickness of rings cultured in

differentiation medium …………………………………………………………………... 58

Figure 3.7: Mechanical properties of 14 day rings with incorporated microspheres

cultured in differentiation medium ………………………………………………………. 59

Figure 3.8: Microsphere incorporation in TGF-β1-treated rings ………………………… 59

Figure 3.9: Effect of TGF-β1 treatment on ring morphology ……………………………. 60

Figure 3.10: Smooth muscle contractile protein expression in rings treated with TGF-β1. 61

Figure 4.1: SEM image of electrospun PCL material ………………………………......... 77

Figure 4.2: Longitudinal pull to failure testing of fused tubes …………………………... 77

Table of Figures 14

Figure 4.3: Cellular infiltration within cuff materials …………………………………… 77

Figure 5.1: Schematic of tube fabrication process, and tissue tube culture experimental

groups for the ring pre-culture duration experiment ……………………………………... 86

Figure 5.2: Fusion kinetics of human SMC rings ………………………………………... 90

Figure 5.3: Histological assessment of human SMC tubes …………………………........ 91

Figure 5.4: Spatial position of rings during fusion ………………………………………. 92

Figure 5.5: Cell proliferation during fusion …………………………………………........ 92

Figure 5.6: PCL cannulation cuff incorporation for bioreactor culture ………………….. 93

Figure 5.7: Histological images of tubes cultured in a luminal flow bioreactor ………… 94

Figure 5.8: Matrix deposition in fused tissue tubes …………………………………........ 94

Figure 6.1: Fabrication of modular tissue tubes with focal heterogeneities ……………... 104

Figure 6.2: Effect of microspheres on ring fusion ……………………………………….. 108

Figure 6.3: Fusion of rings with and without microspheres ……………………………... 108

Figure 6.4: Focal region of microsphere incorporation …………………………………. 109

Figure 6.5: Coronary artery SMC tubes with a focal region of microsphere incorporation 109

Figure 6.6: Effect of PDGF on 2D cell culture proliferation ……………………………. 110

Figure 6.7: Morphology of PDGF treated rings …………………………………………. 111

Figure 6.8: Ki67 staining of rings with PDGF treatment ………………………………... 112

Figure 6.9: Contractile protein expression in SMC rings ………………………………... 112

Figure 6.10: Schematic of future IH model ……………………………………………… 113

Figure 7.1: Images of 14 day iPSC-vSMC rings ………………………………………… 121

Figure 7.2: iPSC-vSMC ring morphology and collagen deposition ……………………... 121

Figure 7.3: Effect of PDGF on 2D iPSC-vSMC cultures ……………………………....... 122

Figure 7.4: Effect of PDGF on iPSC-vSMC ring contractile protein expression ….......... 122

Figure 7.5: Effect of PDGF on ring smooth muscle alpha actin expression …………….. 123

Table of Figures 15

Figure 7.6: Fusion of iPSC-vSMC rings …………………………………………………. 123

Figure 7.7: Fusion rate of iPSC-vSMC rings ……………………………………………. 123

Figure 8.1: Schematic of growth-factor induced focal lesion……………………………. 130

Figure 8.2: Effect of PDGF treatment on ring thickness ………………………………… 134

Figure 8.3: Effect of PDGF on total DNA content ………………………………………. 134

Figure 8.4: Cellular proliferation in rings treated with PDGF ………………………….... 135

Figure 8.5: Collagen deposition in rings with PDGF treatment …………………………. 136

Figure 8.6: Morphology of rings with PDGF treatment …………………………............. 137

Figure 8.7: Contractile protein expression in PDGF treated rings ………………………. 137

Figure 8.8: Effect of FGF treatment on ring thickness …………………………………... 138

Figure 8.9: Collagen deposition in rings with FGF treatment …………………………… 139

Figure 8.10: Morphology of FGF treated rings …………………………………….......... 139

Figure 8.11: Proliferation in rings with FGF treatment …………………………………. 140

Figure 8.12: Cellular proliferation in FGF treated rings ……………………………........ 140

Figure 8.13: Effect of FGF on total DNA content ……………………………………….. 140

Figure 8.14: Contractile protein expression in FGF treated rings ……………………….. 141

Figure 8.15: Effect of TGF-β1 and BMP-4 on ring thickness ………………………........ 142

Figure 8.16: Collagen deposition in TGF-β1 treated rings …………………………......... 142

Figure 8.17: Cellular proliferation in TGF-β1 treated rings ……………………………... 143

Figure 8.18: Proliferation in hMSC rings treated with TGF-β1 …………………............. 143

Figure 8.19: Effect of TGF-β1 on total DNA content …………………………………… 144

Figure 8.20: Contractile protein expression in rings treated with TGF-β1 ………………. 145

Figure 9.1: Schematic of focal lesion experimental setup ………………………….......... 156

Figure 9.2: Tubes fabricated from hMSCs after 7 days of fusion………………………... 157

Figure 9.3: Contractile protein expression in fused hMSC tubes ………………………... 157

Table of Figures 16

Figure 9.4: Alignment of hMSCs within hMSC tubes …………………………………... 158

Figure 9.5: hMSC tube with hole ………………………………………………………... 158

Figure 9.6: Morphology and matrix deposition of vascular tissue tubes ………………… 159

Figure 9.7: Contractile protein expression in hMSC and human aortic SMC tubes……... 160

Figure 10.1: Luminal flow bioreactor in custom stand for endothelialization ……........... 166

Figure 10.2: Endothelialization of SMC tubes ………………………………………....... 166

Figure C.1: Mechanical properties of rings treated with TGF-β1 ………………….......... 174

Figure C.2. Effects of TGF-β1 treatment in smooth muscle cell rings sourced from a

different donor ………………………………………………………………………........ 175

Figure C.3: Effects of TGF-β1 treatment on smooth muscle cell protein expression in

rings self-assembled from human SMCs from a different donor ………………………... 176

Figure E.1: Assembly of custom grips for longitudinal pull to failure test ……………… 183

Figure F.1: Fusion of human SMC rings ……………………………………………....... 184

Figure F.2: Fusion of human coronary artery SMC rings ………………………….......... 185

Figure F.3: Fluorescent images of human coronary artery SMC ring fusion …………… 185

Figure F.4: Contractile protein expression in aortic SMC tubes ………………………… 186

Figure H.1: Effect of FGF treatment on ring thickness…………………………………... 188

Figure H.2: Collagen deposition in rings with FGF treatment…………………………… 189

Figure H.3: Proliferation in rings with FGF treatment…………………………………… 189

Figure H.4: Contractile protein expression in FGF treated rings………………………… 190

Figure I.1: Manual method for fabrication of self-assembled vascular tissue rings and

tubes ……………………………………………………………………………………… 193

Figure I.2: Overview of proposed method of automated assembly of TEBVs fabricated

from self-assembled vascular tissue rings ……………………………………………….. 194

Figure I.3: PEI-MED610 plate system ………………………………………………....... 195

Figure I.4: Schematic of robotic process to remove tissue rings from a PEI-MED610

plate ………………………………………………………………………………………. 197

Table of Figures 17

Figure I.5: Robotic assembly system in cell culture hood ………………………………. 198

Figure I.6: Structure and strength of tissue rings grown in PEI-MED610 plates

compared to control agarose gels ………………………………………………………... 200

Figure I.7: Morphology of self-assembled tissue rings cultured in agarose gels or PEI-

MED610 plates …………………………………………………………………………... 201

Figure I.8: Fusion of automatically or manually fabricated tissue tubes …………............ 202

Table of Tables 18

Table of Tables

Table 7.1: Mechanical characterization of iPSC-vSMC rings, compared to previously

published primary cell rings …………………………………………………………... 121

Table 8.1: Exogenous growth factor concentrations in culture medium for

microsphere (MS)-mediated growth factor delivery experiments ……………............. 131

Table G.1 Characterization of gelatin microspheres used for each experiment……….. 187

Table I.1: Failure rates of tissues rings stacked manually or automatically …………... 201

Chapter 1: Executive Summary 19

Chapter 1: Executive Summary

1.1. Introduction

Every 40 seconds an American dies from cardiovascular disease, the leading cause of

death in the US. Within 15 years, 43.9% of Americans will be living with some form of

cardiovascular disease [1]. Many of these diseases lead to blood vessel occlusion, requiring

bypass procedures utilizing autologous or synthetic grafts, balloon angioplasty, or stent

placement. A side effect of these procedures is intimal hyperplasia (IH), an over-proliferation of

vascular smooth muscle cells following vascular injury that can lead to vessel occlusion. Up to

15-50% of angioplasties, 16-30% of saphenous vein bypass grafts, and up to 90% of synthetic

coronary bypass grafts fail due to IH within 1-3 years [2-6]. IH can reduce blood flow to vital

organs such as the heart and brain, causing chest pain, shortness of breath, and dizziness, which

adversely impact patient quality of life [7, 8]. Severe occlusion may lead to ischemia (oxygen

deprivation) and permanent tissue damage. If this occurs in vessels of the heart or brain, it can be

fatal. While some preventative treatments for IH are available, there are no drugs to reverse

existing intimal growth [9]. Thus, there is a strong need to develop new treatments.

A major obstacle to the development of new, effective drugs is the lack of models that

mimic human IH initiation and progression. Vascular disease research predominantly depends on

mouse models, which do not accurately mimic the progression of IH in humans [10]. As a result,

approximately 90% of drugs that succeeded in animal studies failed in clinical trials [11-13].

Human 2D cell culture and 3D cadaveric tissues have been used as in vitro and ex vivo models to

test vascular therapies. However, cadaveric vessels are limited in supply, and 2D cultures fail to

replicate the complex 3D cell-cell, cell-ECM, and mechanical interactions in human vessels with

IH lesions.

The overall goal of this project is to engineer 3D human vascular tissue with spatially

distinct regions that may serve as a platform for creating an IH model. Existing vascular tissue

engineering approaches are designed to create homogenous tubes not conducive to inducing local

regions of cell hyper-proliferation. In contrast, our lab developed a unique system to fabricate

tissue from modular ring units of self-assembled cells. These rings can be fused to form tubes


with spatially defined regions [14, 15]. We have also shown that gelatin microspheres (MS) can

be incorporated within tissue rings during self-assembly for growth factor delivery [16] . To

create IH lesions, growth factor-loaded microspheres may be incorporated within rings to

stimulate SMC proliferation. Ring units with growth factor-loaded MS may be fused with control

SMC rings to form tubes with localized regions of growth factor delivery and thus intimal

growth.

This innovative, modular tissue fabrication approach allows spatial customization of tube

structure and function, which may ultimately lead to the creation of a 3D human IH model for

drug screening.

1.2. Overview of aims

Aim 1: Develop a system to locally deliver bioactive factors within tissue rings.

The goal of this aim was to evaluate the effects of gelatin microsphere incorporation on

ring morphology and mechanical properties, and to demonstrate the feasibility of using growth

factor-loaded microspheres to modulate SMC phenotype (Figure 1.1). Cellular self-assembly has

been used to generate living tissue constructs as an alternative to seeding cells on or within

exogenous scaffold materials. However, high cell and extracellular matrix density in self-

assembled constructs may impede diffusion of growth factors during engineered tissue culture.

We first assessed the feasibility of incorporating gelatin microspheres within vascular tissue

rings during cellular self-assembly to achieve growth factor delivery. To assess microsphere

incorporation and distribution within vascular tissue rings, gelatin microspheres were mixed with

a suspension of human smooth muscle cells at 0, 0.2 or 0.6 mg per million cells and seeded into

agarose wells to form self-assembled cell rings. Microspheres were distributed throughout the

rings, and were mostly

degraded within 14 days in

culture. Rings with

microspheres were cultured in

both smooth muscle cell

growth medium and

differentiation medium, with

Figure 1.1: Aim 1 overview: Develop a system to locally deliver

bioactive factors within tissue rings. Gelatin microspheres (purple dots)

are co-suspended with SMCs (pink dots) in agarose molds. Cells aggregate

to form rings with incorporated microspheres. Arrows point to individual

rings with incorporated microspheres. Microspheres can be pre-loaded with

growth factors.


no adverse effects on ring structure or mechanical properties. Incorporated gelatin microspheres

loaded with transforming growth factor beta 1 (TGF-β1) stimulated smooth muscle contractile

protein expression in tissue rings. These findings demonstrate that microsphere incorporation can

be used as a delivery vehicle for growth factors within self-assembled vascular tissue rings [16].

Aim 2: Fuse human SMC rings into tissue tubes and evaluate the effects of dynamic culture.

The goal of this aim was to evaluate ring fusion kinetics and develop a system for

dynamically culturing modular tissue tubes (Figure 1.2). This was divided into three primary

objectives. Our first objective was to evaluate ring fusion kinetics, with the goal of reducing

fusion time and culture duration to generate cohesive tissue tubes. Our lab has previously

published our ability to fuse rings into tubes, however ring boundaries were still visible. To

address this, we hypothesized that decreasing ring pre-culture time prior to fusion would

accelerate and improve fusion. It was determined that while ring pre-culture time did not affect

fusion rate, rings cultured for less time prior to fusion appeared more cohesive and had less

distinct ring boundaries [17].

Next, we aimed to

determine if cells maintained

their spatial positioning along

the tube length during fusion,

with the goal of fusing rings

into tissue tubes with distinct

tissue regions along the tube

length. This is important for

creating focal lesions within

the tissue, as diseased cells

must maintain their position in the diseased region of the tube, and not spread along the tube

length. Otherwise, the model cannot mimic the focal nature of the disease. Cells were pre-loaded

with red or green CellTracker dye and seeded into rings. Rings with alternating colors were fused

for 7 days. It was determined that rings maintain their spatial position within rings, with minimal

“mixing” of green and red cells between adjacent rings [17].

Figure 1.2: Aim 2 overview: Fuse human SMC rings into tissue tubes

and evaluate the effects of dynamic culture. After 3 days of culture, rings

are threaded onto silicone tubing and fused into a tube with PCL cuffs on

ends. After 7 days of fusion, tubes are mounted onto a luminal flow

bioreactor for dynamic culture.


The final objective was to develop a system for dynamically culturing vascular tissue

tubes with luminal fluid flow at physiological shear stresses. To do this, we first had to develop a

method to reliably handle and cannulate tissue tubes. Self-assembled tissues such as ours are

fragile at early time-points in culture, and may not be able withstand forces from forceps or

suture material necessary to handle and secure them to bioreactors. To address this, we designed

and fabricated an electrospun PCL cuff material that incorporates onto tube ends via cellular

attachment and infiltration [18]. This provides a reinforced extension of the tube to aid handling

and cannulation. We then cannulated tissue tubes into custom designed luminal flow bioreactors,

and demonstrated that they remained intact for 7 days of dynamic culture at physiologically

relevant shear stresses [17]. Overall, Aim 2 resulted in the accelerated fabrication of spatially-

controlled, fused tissue tubes that can be dynamically cultured on custom flow bioreactors and

endothelialized.

Aim 3: Create vascular tissue tubes with spatially distinct regions

The primary goal of this aim was

to create spatially distinct regions along

the length of vascular tissue tubes that may

potentially be used for modeling focal

vascular diseases (Figure 1.3). Towards

this goal, we first created a focal region of

microsphere incorporation within tissue

tubes. Degradable, cross-linked gelatin

microspheres were incorporated into select

rings and fused in a central region of a

tube, with rings without incorporated

microspheres on either side. This

demonstrated that we can create distinct

tissue regions along the length of the tissue

tube [17]. Ultimately, microspheres may

be utilized to locally deliver growth factors within these regions.

Figure 1.3: Aim 3 overview: create vascular tubes with

distinct regions. Rings with incorporated microspheres are

fused between rings without microspheres, with PCL cuffs on

either end. The resulting construct is a fused tissue tube with

focal region of microsphere incorporation [17].


The second objective of this aim was to evaluate the effects of microsphere-mediated

growth factor delivery on SMC phenotype and proliferation, with the goal of selectively de-

differentiating smooth muscle rings. However, we observed that primary human aortic SMCs in

self-assembled cell rings failed to produce contractile proteins, even with TGF-β1 treatment.

Thus, we evaluated human mesenchymal stem cells (hMSCs) as an alternative cell source of

SMCs for ring self-assembly. We observed that hMSCs successfully formed rings and expressed

smooth muscle contractile proteins. PDGF-loaded microspheres increased hMSC ring thickness

but did not appear to reduce contractile protein expression. We next evaluated the effects of FGF

treatment on hMSC rings, as FGF is another potent SMC mitogen. However, FGF-loaded

microspheres appeared to have minimal effects on ring thickness or contractile protein

expression. Following this, we incorporated TGF-β1-loaded microspheres into tissue rings as in

Aim 1, to determine if it would be more effective to selectively differentiate, rather than de-

differentiate, hMSC rings. TGF-β1-loaded microspheres caused only a small increase in

contractile protein expression. Microspheres in these experiments may have degraded too rapidly

to provide a sustained growth factor release and modulate cell phenotype. Modifications to

microspheres may be necessary for future growth factor delivery experiments.

Our final goal was to demonstrate spatial control over smooth muscle phenotype.

Because of the challenges observed with localized growth factor delivery, we instead used

human aortic SMCs to create a focal region of synthetic smooth muscle, as human aortic SMCs

do not express contractile proteins. These rings were fused between hMSC rings, which we have

observed to express smooth muscle contractile proteins in response to TGF-β1 and BMP-4.

However, contractile protein expression was limited throughout tubes in this experiment. Still,

the region of aortic SMCs remained distinctly visible due to a clear increase in collagen

deposition compared to hMSC ring regions. Overall, Aim 3 demonstrated our ability to create

distinct structural regions along the length of vascular tissue tubes.

1.3. Summary

The following chapters describe the background, rationale, methodology and results of

experiments conducted to develop a platform technology for modular construction of tubular

tissues using customized cell ring building units. Our modular system for fabricating TEBVs

enables us to introduce focal heterogeneities along the tube length that may be used to model


focal vascular diseases. This may be done by incorporating growth factor-loaded microspheres

within select rings prior to fusion, to locally control SMC phenotype and create a diseased state.

For example, microsphere-mediated delivery of growth factors to increase SMC proliferation

may result in the formation of a focal lesion resembling intimal hyperplasia. Ultimately, such

disease models may accelerate the development of new, lifesaving treatments for cardiovascular

diseases.

1.4. References

1. Go, A.S., D. Mozaffarian, V.L. Roger, E.J. Benjamin, J.D. Berry, M.J. Blaha, S. Dai, E.S.

Ford, C.S. Fox, S. Franco, et al., Heart disease and stroke statistics--2014 update: a

report from the American Heart Association. Circulation, 2014. 129(3): p. e28-e292.

2. Kennealey, P.T., N. Elias, M. Hertl, D.S. Ko, R.F. Saidi, J.F. Markmann, E.E. Smoot,

D.A. Schoenfeld, and T. Kawai, A prospective, randomized comparison of bovine carotid

artery and expanded polytetrafluoroethylene for permanent hemodialysis vascular

access. J Vasc Surg, 2011. 53(6): p. 1640-8.

3. Siracuse, J.J., K.A. Giles, F.B. Pomposelli, A.D. Hamdan, M.C. Wyers, E.L. Chaikof,

A.E. Nedeau, and M.L. Schermerhorn, Results for primary bypass versus primary

angioplasty/stent for intermittent claudication due to superficial femoral artery occlusive

disease. J Vasc Surg, 2012. 55(4): p. 1001-7.

4. Lemson, M.S., J.H. Tordoir, M.J. Daemen, and P.J. Kitslaar, Intimal hyperplasia in

vascular grafts. Eur J Vasc Endovasc Surg, 2000. 19(4): p. 336-50.

5. Marmagkiolis, K., A. Hakeem, N. Choksi, M. Al-Hawwas, M.M. Edupuganti, M.A.

Leesar, and M. Cilingiroglu, 12-month primary patency rates of contemporary

endovascular device therapy for femoro-popliteal occlusive disease in 6,024 patients:

beyond balloon angioplasty. Catheter Cardiovasc Interv, 2014. 84(4): p. 555-64.

6. Goldman, S., G.K. Sethi, W. Holman, and et al., Radial artery grafts vs saphenous vein grafts in coronary artery bypass surgery: A randomized trial. JAMA, 2011. 305(2): p.

167-174.

7. Montalescot, G., U. Sechtem, S. Achenbach, F. Andreotti, C. Arden, A. Budaj, R.

Bugiardini, F. Crea, T. Cuisset, C. Di Mario, et al., 2013 ESC guidelines on the

management of stable coronary artery disease: the Task Force on the management of

stable coronary artery disease of the European Society of Cardiology. Eur Heart J, 2013.

34(38): p. 2949-3003.

8. Tendera, M., V. Aboyans, M.L. Bartelink, I. Baumgartner, D. Clement, J.P. Collet, A.

Cremonesi, M. De Carlo, R. Erbel, F.G. Fowkes, et al., ESC Guidelines on the diagnosis

and treatment of peripheral artery diseases. Eur Heart J, 2011. 32(22): p. 2851-2906.


9. Kim, F.Y., G. Marhefka, N.J. Ruggiero, S. Adams, and D.J. Whellan, Saphenous vein

graft disease: review of pathophysiology, prevention, and treatment. Cardiol Rev, 2013.

21(2): p. 101-9.

10. Hui, D.Y., Intimal Hyperplasia in Murine Models. Curr Drug Targets, 2008. 9(3): p. 251-

260.

11. Alexander, J.H., G. Hafley, R.A. Harrington, E.D. Peterson, T.B.F. Jr, T.J. Lorenz, A.

Goyal, M. Gibson, M.J. Mack, D. Gennevois, et al., Efficacy and Safety of Edifoligide, an

E2F Transcription Factor Decoy, for Prevention of Vein Graft Failure Following Coronary Artery Bypass Graft Surgery: PREVENT IV: A Randomized Controlled Trial.

JAMA, 2005. 294: p. 2446-2454.

12. Mann, M.J., G.H. Gibbons, P.S. Tsao, H.E.v.d. Leyen, J.P. Cooke, R. Buitrago, R.

Kernoff, and V.J. Dzau, Cell Cycle Inhibition Preserves Endothelial Function in

Genetically Engineered Rabbit Vein Grafts. J. Clin. Invest., 1997. 99: p. 1295–1301.

13. Kola, I. and J. Landis, Can the pharmaceutical industry reduce attrition rates? Nature

Reviews Drug Discovery, 2004. 3: p. 711-715.

14. Gwyther, T.A., J.Z. Hu, A.G. Christakis, J.K. Skorinko, S.M. Shaw, K.L. Billiar, and

M.W. Rolle, Engineered vascular tissue fabricated from aggregated smooth muscle cells.

Cells Tissues Organs, 2011. 194(1): p. 13-24.

15. Dikina, A.D., H.A. Strobel, B.P. Lai, M.W. Rolle, and E. Alsberg, Engineered

cartilaginous tubes for tracheal tissue replacement via self-assembly and fusion of human

mesenchymal stem cell constructs. Biomaterials, 2015. 52: p. 452-62.

16. Strobel, H.A., A.D. Dikina, K. Levi, L.D. Solorio, E. Alsberg, and M.W. Rolle, Cellular

self-assembly with microsphere incorporation for growth factor delivery within

engineered vascular tissue rings. Tissue Eng Part A, 2017. 23(3-4): p. 143-155.

17. Strobel, H.A., T.A. Hookway, M. Piola, G.B. Fiore, M. Soncini, E. Alsberg, and M.W.

Rolle, Assembly of tissue engineered blood vessels with spatially-controlled

heterogeneities. Tissue Eng Part A, 2018. In Press.

18. Strobel, H.A., E.L. Calamari, A. Beliveau, A. Jain, and M.W. Rolle, Fabrication and

characterization of electrospun polycaprolactone and gelatin composite cuffs for tissue

engineered blood vessels. JBMR Part B, 2018. 106B(2): p. 817-826.

Chapter 2: Background 26

Chapter 2: Background

Section 2.7 modified from: H. A. Strobel, E. L. Calamari, B. Alphonse, T. A. Hookway, and M. W. Rolle,

“Fabrication of Custom Agarose Wells for Cell Seeding and Tissue Ring Self-assembly Using 3D-Printed

Molds” Journal of Visualized Experiments, 2018. 134: e56618. Reprinted with permission (Appendix A.)

Authorship contributions: HAS performed the experiments shown in the manuscript and video, made all figures,

wrote and revised the manuscript, prepared and edited the video shot list and script, and prepared the materials and

performed the demonstrations in the video. ELC and BA re-designed the mold system and edited the manuscript.

TAH supervised mold re-design and edited the manuscript. MWR contributed to experimental design, supervised

data collection, data analysis, and preparation of the manuscript, and edited the manuscript.

Section 2.9 and 2.10 taken from: H. A. Strobel, E. I. Qendro, E. Alsberg, M. W. Rolle, “Targeted delivery

of bioactive molecules for vascular intervention and tissue engineering.” In Review.

Authorship contributions: HAS is the primary author and wrote the manuscript. EIQ created the figures (not

included in the chapter) and assisted with literature searches. EA revised the structure and content and edited the

final manuscript. MWR advised HAS and EIQ on structure and content and edited the manuscript.

In this Chapter, we discuss the structure and function of blood vessels in both normal and

diseased arteries. Specifically, we review the prevalence of intimal hyperplasia, the mechanisms

of lesion initiation and progression, and current therapies. Finally, we discuss clinical gaps in

treatment options, and the potential of tissue engineering for fabricating intimal hyperplasia

disease models.

2.1. Smooth muscle phenotype

Arteries are comprised of three primary layers: the adventitia, media, and intima. The

outer adventitial layer contains primarily fibroblasts and collagen, and imparts tensile strength to

the vessel at high pressures [2, 3]. The medial layer consists mainly of smooth muscle cells

(SMCs) and elastin. SMCs respond to mechanical and biochemical stimuli by contracting and

dilating to regulate blood flow [2]. The intima is comprised of a layer of endothelial cells (ECs)

on the luminal surface of the vessel, which prevent platelet adhesion and thrombosis [2].

Healthy or “contractile” SMCs are less proliferative, secrete little collagen, and express

contractile proteins such as smooth muscle alpha actin, calponin, and smooth muscle myosin

heavy chain, which allow SMCs to contract or relax to regulate blood flow [4]. In contrast,

SMCs in diseased and injured vessels exhibit a “synthetic” phenotype, characteristic of SMCs


observed in IH [4]. Synthetic SMCs proliferate, synthesize collagen and other extracellular

matrix molecules, and downregulate expression of contractile proteins [4].

ECs play critical roles in maintaining blood vessel health and homeostasis. Healthy ECs

prevent platelet aggregation and activation and secrete NO, which inhibits SMC proliferation [5-

7]. In contrast to injured or “activated” ECs, healthy ECs have increased expression of the NO-

producing enzyme, nitric oxide synthase (eNOS) [8]. When ECs become activated following

injury, there is a decrease in eNOS and NO, increase in EC proliferation, and increased

expression of pro-thrombogenic cell surface proteins such as VCAM and ICAM [9, 10].

When ECs become activated due to injury or disease, the endothelium decreases NO

production, which attenuates the preventative effects on SMC proliferation and migration, and

SMCs become less contractile and more synthetic [6, 11, 12]. In addition to measuring

expression of proteins characteristic of normal and diseased phenotypes, EC and SMC function

can be assessed by measuring contraction or relaxation of vessels in response to acetylcholine

[13-15]. Acetylcholine causes different effects on SMCs depending on the presence or absence

of functional ECs. In the absence of endothelium, acetylcholine binds to muscarinic receptors on

SMCs and triggers contraction [15, 16]. In the presence of functional ECs, acetylcholine

stimulates NO production, stimulating guanylate cyclase to form cyclic guanine monophosphate

(cGMP) and triggering SMC relaxation [15, 16]. cGMP production can be measured directly to

assess EC function [17].

2.2. Intimal hyperplasia

Intimal hyperplasia (IH) typically begins with damage to the endothelial layer [18]. This

is often caused by physical injury such as vascular bypass surgery, angioplasty, or stenting.

These procedures can also indirectly damage endothelium by creating alterations in fluid flow

and thus changes in wall shear stress, which may worsen as intimal growth progresses [19-23].

High shear stress (greater than 70 dyne/cm2) can also damage the endothelial layer [24, 25],

thereby reducing secretion of molecules such as nitric oxide (NO) and prostacyclin, which inhibit

SMC proliferation [26]. Additionally, endothelial damage can activate platelets, resulting in the

release of platelet-derived growth factor (PDGF), transforming growth factor beta (TGF-β),

interleukin 1 (IL-1), interleukin 6 (IL-6) and thrombin [27]. These factors then stimulate SMC


proliferation and migration into the intimal layer [27]. Low shear (less than 6 dyne/cm2) can

reduce flow-induced EC secretion of molecules that inhibit SMC proliferation [24, 28], and can

also upregulate PDGF expression in ECs [28]. PDGF is especially known to stimulate the SMC

proliferation, migration, and collagen deposition that contribute to IH [29-34].

SMC proliferation can begin as early as 24 hours after injury, and migration can begin in as

soon as 4 days [22]. Significant reductions in lumen area can be seen within 4-6 weeks of the

initial injury and may progress for up to 1 year before growth stabilizes [20, 35]. Lesion size

varies considerably, but the average surface area is approximately 7.4 mm2 [19]. Intimal lesions

may further progress to form atherosclerotic plaques, which have potential to rupture and trigger

a life-threatening thrombosis [20].

2.3. Treatments for IH

There are a limited number of approved drugs available to prevent IH [20]. Antiplatelet

medications such as aspirin prevent platelet aggregation, thus inhibiting platelet activation and

PDGF release, which prevents SMC proliferation associated with IH [31, 36]. However, aspirin

increases the risk of bleeding and is not appropriate for all patients [37]. Statins are prescribed

for their cholesterol-lowering effects, which prevent atherosclerosis. Statins also independently

inhibit SMC proliferation by inhibiting the MAPK (mitogen activated protein kinase) signaling

pathway [38-40], and may also improve endothelial function and accelerate re-endothelialization

[38, 41-43]. However, statins can cause side effects (e.g., muscle weakness) and may not be

tolerated by all patients [37]. These treatments are typically given to patients receiving bypass

surgery, as vascular grafts have a high rate of IH.

In cases where intimal growth becomes symptomatic, invasive procedures may be

required, such as balloon angioplasty, stent placement, or bypass surgery to restore blood flow.

Intervention is recommended for stenoses greater than 50%, although the criteria vary from

patient to patient [37]. Bypass surgery generally has better long term outcomes for patients, but

is much more invasive [44-47]. These interventions do not solve the problem, as they also trigger

endothelial injury and can stimulate the re-formation of IH, thus requiring future interventions.

Stents that elute drugs such as Paclitaxel and Sirolimus can reduce the incidence of IH by


inhibiting SMC proliferation locally. However, they may impede re-endothelialization and

endothelial function of the stented area, increasing the risk of late thrombosis [48-50].

Other drugs have been tested that directly inhibit SMC proliferation. E2F transcription

factor inhibitors such as Edifoligide, for example, directly interrupt the cell cycle [51-53].

However, these drugs were shown to be ineffective in clinical trials [51, 52]. Most potential

therapies under investigation target platelet activation or SMC proliferation, including PDGF

receptor inhibitors. PDGF is a potent stimulator of SMC migration and proliferation, thus

inhibiting PDGF receptors has been shown to prevent IH in pre-clinical trials [54, 55]. These

drugs are also advantageous because they do not inhibit EC proliferation, as macrovascular ECs

do not have PDGF receptors [56, 57]. However, many of these drugs have not been tested in

clinical trials [54, 55]. Despite the continued advancement of IH treatment, existing therapies are

not ideal for all patients, and invasive procedures are still often necessary as there are no

approved drugs that reverse existing IH.

2.4. Model systems for studying IH

Mouse models are most commonly used for studying intimal hyperplasia, due to their

well-characterized genetics, and because procedures for stimulating IH in mice are well

established and reproducible. IH is initiated in animal models by inducing a significant

endothelial injury, by arterial ligation or mechanical denudation [58, 59]. However, the time

course for IH progression following injury is significantly faster in animals than humans [60,

61]. Additionally, shear stresses are much higher in small animals, and not comparable to

humans. Thus, results from animal studies do not always accurately predict how human subjects

will respond to a particular treatment. Even experiments in larger animals do not always predict

outcomes in humans; many drugs have successfully treated IH in large animals but failed in

clinical trials [20, 51]. Thus, there is a strong need for vascular disease models that provide a

more realistic pre-clinical drug screening platform that mimics human diseases.

A number of alternatives to animals have been explored as experimental models of

human IH. Ex-vivo human arteries have been used for studying IH progression and treatment,

however there is a limited supply of cadaver vessels that are available for testing [62]. Testing on

2D human cell cultures is also not a good predictor of treatment success, as a 2D culture cannot


simulate the cell-cell and cell-matrix interactions in 3D tissue [63]. For these reasons,

development of a 3D human co-culture model for studying IH is critical. Such models may also

reduce the use of animals, and could potentially reduce the time and costs associated with pre-

clinical drug screening [64, 65].

The characteristics of an ideal model system for studying IH in vitro are shown

schematically in Figure 2.1. IH is a localized disease, and so a model must demonstrate localized

intimal growth, as shown in the center region of Figure 2.1. EC-SMC interactions also play an

important role in maintaining a healthy blood vessel,

so creating a SMC-EC co-culture environment is also

important. Fluid flow must also be applied to

maintain healthy EC phenotype, as shear stresses

promote production of NO, which affects both EC

and SMC phenotype and function. Model validation

should include testing current IH preventative

therapies for prevention of intimal growth.

2.5. Tissue engineered blood vessels as in vitro human vascular models.

Tissue engineered blood vessels (TEBVs) have potential for use as disease models and

tools for drug screening [64-66]. They can be fabricated from human cells, and are more

representative of the 3D environment than 2D cell cultures [63]. TEBVs can be fabricated using

a variety of approaches, including seeding cells on polymer scaffolds, incorporating cells in

hydrogels, or using scaffold-free cellular self-assembly approaches [67-71]. Many of these

TEBVs contract when stimulated with vasoactive substances, suggesting their potential as tools

for drug screening [67, 68, 70, 71]. However, some of these TEBVs rely on cell types such as

fibroblasts instead of SMCs [71], limiting their use as IH models. Other TEBVs rely on synthetic

polymers [67], which degrade into fragments that may weaken vessels and create acidic

degradation environment, thus de-differentiating SMCs independently of IH triggers [67, 72-74].

Most importantly, all of these tubes are homogenous in nature, and are not conducive to

developing the focal changes in SMC phenotype characteristic of IH.

Figure 2.1:

Characteristics of IH

model lesion. Local SMC

proliferation and matrix

deposition, and fluid flow.


2.6. Modular fabrication of vascular tissue constructs from self-assembled cell ring units.

Cellular self-assembly approaches to fabricating tissue engineered blood vessels are an

alternative to scaffold-based approaches. Self-assembled, scaffold-free tissues may have greater

cell density, enhanced matrix deposition and strength, and improved biological function

compared to scaffold-based tissues [75-78]. However, forming 3D tissues without the use of

exogenous scaffold support with specific sizes and shapes remains a challenge. Some methods

fuse together layers of cell sheets to form thicker constructs, although this process can be time

consuming and labor intensive [79]. Alternatively, cells can be seeded into non-adhesive molds

and allowed to aggregate into spheroids, rings, and other tissue shapes [80-82].

Self-assembled tissue ring units require fewer cells, shorter culture times, and less

reagents than larger tubular engineered tissues, but can still be mechanically tested, examined

histologically, or used for contractility and other functional testing [81, 83-85]. Because they can

be rapidly fabricated and easily tested, tissue rings are ideal for screening large numbers of

culture parameters, and have potential for use as disease models [85] or tools for drug screening

[68]. Additionally, rings can be fused into more complex tissue structures such as blood vessels

or trachea [81, 86], and rings may fuse more completely than other shapes such as spheroids [87,

88].

2.7. Engineering custom agarose molds for self-assembled tissue ring fabrication [1]

We previously reported a system for fabricating custom annular agarose cell-seeding

wells from a polydimethylsiloxane (PDMS) negative cast in a milled polycarbonate mold [69,

81]. Agarose was poured into the PDMS negative and allowed to set [69, 81]. Cells were then

seeded into agarose wells, where they aggregated to form self-assembled, scaffold-free tissue

rings in less than 24 hours [69, 81]. PDMS negatives are autoclavable, can be reused many times,

and are soft and flexible, making it easy to remove the solidified agarose wells. When this

system was initially reported in Gwyther et. al. [81], PDMS negatives were cast from milled

polycarbonate molds. After agarose casting, the cell seeding wells were individually cut out and

placed into wells of a 12-well plate [69, 81]. The design was more recently modified such that a

single agarose mold produces 5 rings and fits in a well of a 6-well plate, eliminating the need to

cut out individual wells and reducing the amount of PDMS and agarose required to produce each


ring. A smaller cell seeding trough width was used to reduce the number of seeded cells required

to achieve ring formation. Despite these changes, the resolution and customization of molds were

restricted to available standard endmill dimensions, and micromilling can be prohibitively

expensive. Additionally, computer numerical control (CNC) machining can be time consuming

and cumbersome due to the need to reserve time on heavily utilized custom equipment,

additional computer-aided manufacturing (CAM) software to convert the computer-aided design

(CAD) file to a programmable tool path, and reliable fixturing of the polycarbonate part during

machining.

To address these limitations, we examined the use of 3D printing as an alternative to

CNC machining to create the ring-shaped cell seeding well templates. 3D printing is widely used

for engineering custom implants, fabricating scaffold materials, and for direct printing of cells

and tissue spheroids [88-90]. We used a high resolution 3D printer, and specialized 3D printing

material that enabled us to print a rigid mold with a smooth, glossy surface finish. Our technique

allows for fabrication of highly customizable, high resolution plastic molds that can be used for

casting PDMS negatives and agarose wells. The mold design was further modified in the 3D

printed mold version to include tapered outer walls and center hole in order to ease removal of

both PDMS negatives from 3D printed molds and agarose wells from PDMS negatives. These

tapered features cannot be achieved with standard machining processes. The distance from the

bottom of the wells to the bottom of the mold was increased in this iteration, resulting in a

thicker agarose base below the posts to reduce the risk of posts breaking during agarose well

removal. A cross-sectional view of the 3D printed mold, and dimensions of our current design

compared to previous designs, is shown in Figure 2.2.

The current, modified mold and ring fabrication procedure is shown schematically in

Figure 2.3 [1]. Human SMCs are seeded into a ring-shaped agarose mold, where the cells

aggregate together to form a self-assembled ring within 24 hours of cell seeding. Rings can then

be stacked onto a silicone tube, where they fuse together to form a tissue tube (schematic shown


in

in

in

in

in

in

in

in

in

in

in

in

in

in

in

in

in

in

Figure 2.3: Fabrication of self-assembled tissue rings. A 3D printed mold is used to cast a PDMS

negative, which is then used to cast the agarose wells (A). Cells are then seeded directly into the

agarose wells, where they aggregate in less than 24 h to form tissue rings (B). Dashed lines in (B)

show the well outline. [1]

Figure 2.2: Cross-sectional view of 3D printed mold. Dimensions for trough width (A), trough

height (B), center hole (C), total diameter (D), outer lip (E), and outer wall height (F) are shown. The

center hole and outer walls are tapered to improve ease of removal. [1]


in Figure 2.4) [69]. This modular

system is unique because it provides

spatial control over the cellular and

molecular composition of each

segment of the tissue tube, with the

ability to customize each ring segment.

2.8. Microsphere incorporation and

modular assembly to create focal

regions of IH.

Our overall goal is to use our

modular TEBV assembly system to

create focal regions within the tube that mimic human IH. To achieve this, we elected to

incorporate growth-factor loaded microspheres into select rings during self-assembly. This

would allow us to create ring segments with microsphere-mediated growth factor delivery, and

fabricate tubes with localized regions of increased SMC proliferation and ECM synthesis

characteristic of IH. A schematic of this concept is shown in Figure 2.5.

2.9. Bioactive molecule release from tissue engineered blood

vessels

TEBVs can be fabricated in a variety of ways. A

common approach is to seed cells onto natural or synthetic

polymer scaffolds and allow the construct to mature and

remodel in a bioreactor [91, 92]. Alternatively, TEBVs can

be fabricated via cellular self-assembly approaches, where

constructs are fabricated entirely from cells and their

secreted extracellular matrix [75, 81]. While these

approaches have had some success, many challenges remain,

such as establishing a healthy, contractile SMC phenotype,

optimizing graft strength and compliance, and achieving

complete endothelialization. Localized and controlled

Figure 2.4: Fabrication of vascular tissue tubes. Tissue rings

are threaded over silicone tubing, where they fuse together to

form a modular vascular tissue tube. PCL cuffs can also be

placed over tube ends to serve as reinforced material for handling

and cannulation.

PCL Tissue PCL

Figure 2.5: Schematic of method for

fabricating TEBV with intimal

lesion. Rings with growth factor-

loaded microspheres are fused

between rings containing microspheres

with no growth factors.


bioactive factor delivery may be able to address some of these problems.

2.10. Microsphere-mediated growth factor delivery in engineered vascular tissue

Microsphere (MS)-mediated growth factor delivery has been used for years to mature and

differentiate many engineered tissues, including cartilage [93], bone [94], and stem cell

aggregates [93, 95, 96]. MS incorporation alone can increase tissue strength [86, 97], oxygen

diffusion [77, 98], cell viability [98, 99], and uniformity of matrix deposition [100]. However,

their application in vascular tissue engineering has been limited. Others have incorporated

gelatin MS into cell spheroids, which were fused into vascular tissue, but the MS primarily

served to stabilize the construct and were not used for growth factor delivery [87]. Our group has

demonstrated that MS loaded with TGF-β1 can be used to increase SMC contractile protein

expression within self-assembled SMC rings [101]. This approach may be well-suited for

applications where systemic or exogenous delivery of a growth factors may be harmful or not

possible, or for thick, high cell density engineered tissues where growth factors cannot diffuse

through the entire construct.

2.11. Platelet-derived growth factor

Many growth factors are associated with IH, including PDGF, TGF-β, IL-1, IL-6, and

fibroblast growth factor (FGF). PDGF is released from platelets upon activation, and has potent

stimulatory effects on SMC proliferation, migration, and collagen production [29, 102-105].

Inhibiting PDGF receptors has been shown to prevent IH in animals, and PDGF treatment alone

has triggered IH in ex-vivo and in vivo models [54, 62, 106]. Other growth factors such as TGF-β

stimulate collagen production but may inhibit SMC proliferation [4]. FGF also stimulates SMC

proliferation and collagen deposition, but most potential therapies target PDGF and its receptors

[20]. For these reasons, PDGF was selected as the ideal growth factor to stimulate IH in our

model system.

PDGF is a dimeric protein most commonly made up of two A or B chains linked by a

disulfide bond; PDGF-BB, PDGF-AA, or PDGF-AB [107]. PDGF-BB is the isoform most

implicated in SMC proliferation and migration associated with IH [102, 103], and was therefore

used to stimulate SMC proliferation and de-differentiation in vascular tissue rings in Aim 3

(Chapter 7). PDGF-BB is primarily released from activated platelets, although it can be secreted


by other cell types, including ECs and SMCs in their diseased states [30, 108-111]. It increases

SMC proliferation by binding to PDGF receptor beta (PDGFRβ) and activating the MAPK

signaling pathway [33]. PDGF-BB is well known to de-differentiate SMCs to synthetic

phenotype by downregulating contractile protein expression [112, 113]. However, PDGF-BB is

not known to have significant effects on ECs. While activated or angiogenic ECs may express

low levels of PDGFRβ and have some mitogenic response, quiescent ECs in healthy vessels do

not express PDGFRβ [56, 57, 110]. PDGF-BB can also be incorporated within biomaterials

[114-116], which may allow for spatially controlled PDGF-BB delivery.

2.12. Gelatin microspheres for controlled delivery of PDGF

Microspheres (MS) have been used to deliver growth factors for a wide range of tissue

engineering applications, and can be made from a variety of synthetic and natural polymer

materials. Gelatin is manufactured from animal collagen, which is denatured with an acidic or

basic treatment [117, 118]. Gelatin has several key advantages over other biomaterials for MS

fabrication. Gelatin is naturally derived, has tunable degradation rates, does not produce harmful

degradation products, and is cell adhesive [118-120]. In contrast, synthetic polymer MS such as

poly(lactic-co-glycolic) acid (PLGA), must be dissolved in organic solvents which can denature

the growth factors they are designed to release, their degradation products can cause SMC de-

differentiation, and they are generally non-cell adhesive [73, 118, 120].

Gelatin MS are typically manufactured using an oil in water emulsion process [97, 121],

and are crosslinked to prevent swelling and to control degradation [122, 123]. Crosslinking can

be achieved with several different chemicals, including glutaraldehyde, carbodiimide, and

genipin [117, 124, 125]. Genipin is derived from the gardenia plant, and has low cytotoxicity

compared to other cross-linkers [117, 124]. Gelatin MS can be soaked in a solution of growth

factors, which bind electrostatically to gelatin. Growth factors are then released during the

proteolytic degradation of the MS [97, 118]. Gelatin degradation rates and growth factor release

kinetics can be controlled by altering crosslink density [97, 117, 118]. Growth factor release

from MS has been shown to improve differentiation within tissue constructs compared to

exogenous growth factor treatment [100]. MS incorporation has also been shown to improve

differentiation in some constructs, even without growth factor loading [98, 99, 126]. In addition,

MS incorporation can increase tissue strength [86, 97], oxygen diffusion [77, 98], cell viability


[98, 99], and uniformity of matrix deposition [100]. We used gelatin MS to achieve localized

delivery of PDGF to spatially control stimulation of SMC proliferation to create model IH

lesions.

Tissue maturation may further be enhanced by adjustments to growth factor release

kinetics. Release of multiple growth factors sequentially may be beneficial for vascular tissue

engineering. Gong et al. developed an optimized procedure for fabricating TEBVs by treating

them in vitro with exogenous PDGF for 4 weeks to stimulate new tissue growth, and then TGF-

β1 for 4 weeks to promote differentiation [105]. This sequential delivery could also be obtained

by designing biomaterials for controlled dual-delivery, an approach that is already being used to

create microvasculatures within other engineered tissues [127, 128].

Spatiotemporal control of the release of multiple growth factors may also be applied to

modeling focal vascular diseases. Because many vascular diseases such as atherosclerosis,

intimal hyperplasia, and aneurysm affect only one region of the vessel, delivering growth factors

or other molecules specifically within a focal region within TEBVs may enable the creation of

such models. This could potentially be accomplished with modular TEBV approaches and

controlled release systems. Spatially controlled release may also be advantageous for culturing

and maintaining distinct tissue phenotypes in multi-tissue constructs such as trachea, which have

alternating smooth muscle and cartilage regions [86, 129].

2.13. Summary

Intimal hyperplasia is an injury that frequently occurs following treatments for invasive

vascular diseases. There are few treatment options, and limited model systems for evaluating

new therapeutics. TEBVs have potential as models for screening new therapies, but most are

homogenous in nature and not conducive to modeling focal vascular diseases. As a step toward

addressing the need for an in vitro IH model, we have developed a modular system for

fabricating TEBVs from self-assembled ring units. With this method, we can incorporate

biomaterials such as gelatin microspheres into select ring sub-units. This may enable growth

factor delivery in a localized region of the TEBV, potentially creating a focal region of vascular

disease. In the following chapters, we evaluate microsphere incorporation within tissue rings,


microsphere-mediated growth factor delivery, ring fusion, and the fabrication of tubes with

distinct regions. This work is a critical step towards the fabrication of an in vitro IH model.

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Advanced Science, 2017.

Chapter 3: Cellular self-assembly with microsphere incorporation for growth factor delivery within engineered

vascular tissue rings 50

Chapter 3: Cellular self-assembly with microsphere

incorporation for growth factor delivery within engineered

vascular tissue rings

H. A. Strobel, A. D. Dikina, K. Levi., L. D. Solorio, E. Alsberg, and M. W. Rolle, “Cellular self-assembly

with microsphere incorporation for growth factor delivery within engineered vascular tissue rings.”

Tissue Engineering Part A, 2017. 23:(3-4), p. 143-155. Reprinted with permission (Appendix B).

Supplemental figures presented in Appendix C.

Authorship contributions: HAS designed and performed the experiments, collected and analyzed the data presented,

prepared the figures and wrote the manuscript; ADD prepared and characterized microspheres for the growth

factor release study, provided feedback on experimental design and data analysis, and edited the manuscript. KL

contributed to initial study design. LDS prepared and characterized microspheres used in microsphere

incorporation experiments and contributed to the initial study design. EA and MWR contributed to experimental

design, supervised data collection, data analysis, and preparation of the manuscript, and edited the manuscript.

3.1. Introduction

Vascular tissue engineering has become a viable approach to meet the growing clinical

need for blood vessel substitutes [1-4]. In addition to meeting the need for transplantable grafts,

functional vascular constructs could also serve as in vitro models to screen potential therapies [5,

6]. There are a variety of approaches currently employed for development of tissue engineered

blood vessels, including use of cell-seeded degradable synthetic polymer scaffolds [2, 3, 7] and

hydrogels [8, 9], as well as scaffold-free cellular self-assembly strategies [1, 4, 10, 11].

Our lab developed a cellular self-assembly system to fabricate living engineered human

vascular tissue constructs entirely from smooth muscle cells (SMCs) [10]. Briefly, SMCs were

seeded into annular agarose wells, where they aggregated and self-assembled to form tissue

rings. The rings were then stacked together and fused in culture to form 2mm diameter tissue

tubes [10, 12]. In addition to SMC rings and tubes, this versatile cellular self-assembly system

may enable fabrication of rings and tubes of other tissue types, including human cartilage [13].

Cellular self-assembly may have advantages over scaffold-based approaches for vascular

tissue engineering. Compared to cells seeded on scaffold materials, self-assembled cellular

constructs may have greater cell density, enhanced ECM production and tissue strength,

improved biological function, and lower susceptibility to degradation and infection [11, 14-16],

and thus may be more similar in structure and function to native tissue. However, existing



methods for fabricating self-assembled blood vessels create homogenous tubes not conducive to

creating focal heterogeneities characteristic of certain diseases such as aneurysm or intimal

hyperplasia. Our self-assembled cell rings can be used as building units to fabricate tubes by

modular assembly of ring subunits. This allows introduction of spatial heterogeneity along the

length of the tube may enable customization of distinct regions at the anastomoses, or within the

tubes to model focal changes characteristic of disease. To create these changes within rings, we

proposed the incorporation of degradable gelatin microspheres within the tissue constructs

during self-assembly. Microspheres have been used to deliver growth factors such as

transforming growth factor beta 1 (TGF-β1) within dense tissue constructs, to help overcome

diffusion limitations and permit spatiotemporal control over growth factor release [17-19].

Degradable gelatin microspheres were used as the delivery vehicle for TGF-β1, as gelatin

microspheres are naturally biocompatible and cell adhesive [18, 20], and have been well-

characterized [19, 21, 22]. Gelatin degradation, and therefore growth factor release rate, can be

controlled by modifying the polymer cross-link density [21, 23-26].

The first goal of this study was to test the feasibility of incorporating microspheres into

self-assembled human SMC rings, and evaluate the effects on ring structure and mechanical

properties. We first tested microsphere incorporation in rings cultured in a commercially

available SMC growth medium, which supports SMC proliferation and self-assembly into tissue

rings. However, growth medium contains EGF and FGF, which have been shown to interfere

with TGF-β1-mediated differentiation to a healthy “contractile” SMC phenotype [27-29]. Thus,

we also tested incorporation in a differentiation medium, which does not contain growth factors,

and supports SMC differentiation to a healthy “contractile” phenotype [30]. The second goal of

this work was to evaluate the feasibility of utilizing gelatin microspheres to deliver TGF-β1 to

3D self-assembled SMC constructs to improve ring structure and function. TGF-β1 is important

in vascular tissue engineering because it stimulates ECM synthesis (e.g., collagen and elastin

[31-36]) induces contractile protein expression in SMCs (e.g., smooth muscle alpha actin and

calponin [27, 28, 37, 38]) and enhances vascular graft contractility [39, 40]. These studies may

be essential for future work aimed at modelling focal changes in the vascular wall characteristic

of disease.



3.2. Materials and methods

3.2.1. Gelatin microsphere preparation

Microspheres were formed and characterized using methods described previously [13].

Briefly, a water-in-oil emulsion was created with 11.1 w/v % Type A gelatin (Sigma-Aldrich)

and olive oil (GiaRussa). Gelatin microspheres were cross-linked with 1 % w/v genipin (Wako)

for 3 hours at room temperature. Ninhydrin assay was used to quantify the degree of polymer

cross-linking. Images of microspheres were taken on a TMS microscope (Nikon) with Coolpix

995 camera (Nikon). Microsphere diameters were measured using ImageJ software.

3.2.2. Human smooth muscle cell culture

Human coronary artery smooth muscle cells (Lifeline) were cultured in Lifeline complete

growth medium (Lifeline Vasculife Growth Medium) supplemented with 0.2% penicillin-

streptomycin (Mediatech) and 1% amphotericin B (Corning Cellgro). Differentiation medium

(adapted from [30]) consisted of a 1:1 ratio of Dulbecco’s Modified Eagle Medium (DMEM;

Mediatech) and Ham’s F-12 (Mediatech) with 1% insulin-transferrin-selenium (ITS), 1% FBS

(PAA Laboratories), 1% L-glutamine (Mediatech, glutaGro supplement), 1% penicillin-

streptomycin (Mediatech), 1% amphotericin B (Mediatech) and 50 µg/ml ascorbic acid (Wako).

3.2.3. Smooth muscle cell ring self-assembly and unloaded microsphere incorporation

Agarose molds were prepared using methods described previously [10, 12] with some

modifications to the mold design. Briefly, a solution of 2% agarose in DMEM (w/v) was

autoclaved, pipetted into molds made from cured polydimethylsiloxane (PDMS; Sylgard 184;

Dow Corning), and cooled to room temperature to solidify. Agarose wells were transferred into a

6 well plate and equilibrated

overnight in growth medium.

Each mold consisted of 5

wells, each with a 2mm

diameter center post (Fig.

3.1D).

Figure 3.1: Schematic of microsphere incorporation within self-

assembled tissue rings. (A), Gelatin microspheres (purple circles) were

mixed in suspension with SMCs (black dots) at 0, 0.2 or 0.6mg/106 cells. (B),

Cells and microspheres were seeded into agarose molds. (C), Cells aggregate

to form self-assembled rings with incorporated microspheres. (D),

Photograph of an agarose mold with aggregated human SMC-microsphere

rings. Arrowheads point to rings on agarose posts.



Prior to ring seeding, microspheres were UV sterilized for 10 minutes. The unloaded

(growth factor free) microspheres were hydrated in phosphate buffered saline (PBS) for two

hours at 37ºC. Then, microspheres were diluted to twice the desired concentration (9.6mg

microspheres per ml for 0.6mg/106 cells, and 3.2mg microspheres per ml for 0.2mg/106 cells) in

serum free growth medium. SMCs were resuspended at a concentration of 16x106 cells/ml, and

mixed 1:1 with microspheres to achieve final concentrations of 0mg, 0.2mg, or 0.6mg

microspheres per million cells. The cell-microsphere suspension was seeded into the agarose

wells (shown schematically in Fig. 3.1) with 400,000 cells per ring. All rings were seeded in

growth medium, then cultured in growth medium or switched to differentiation medium after one

day. Rings were cultured for a total of 7 or 14 days.

3.2.4. TGF-β1-loaded microsphere preparation and incorporation within tissue rings

UV-sterilized microspheres were incubated in a solution of 80ng/µl TGF-β1 (Peprotech;

400ng/mg microspheres; 5µl/mg microspheres) in PBS for two hours at 37 ºC [21]. Rings were

seeded with 0.6mg microspheres per million SMCs (as described above) in growth medium, and

switched to differentiation medium after 24 hours. In the designated control groups, 10ng/ml

exogenous TGF-β1 was added to differentiation medium on day 1 and continued until day 14.

3.2.5. Histology and immunohistochemistry

Tissue rings were fixed for 1 hour in 10% neutral buffered formalin, embedded in

paraffin, sectioned in 5µm slices, and adhered to charged slides (Superfrost Plus; VWR).

Hematoxylin and Eosin staining was used to examine ring morphology and Picrosirius Red/Fast

Green (Sigma) was used to visualize collagen.

To examine contractile protein expression, deparaffinized slides were blocked with 1.5%

normal rabbit serum (NRS, Vector) in PBS for 45 minutes at room temperature. Antigen retrieval

was performed on samples stained for calponin by incubating slides in 10 mM Tris, 1 mM

EDTA, 0.05 % Tween-20, (pH 9.0) in a pressure cooker for 5 minutes. Samples were incubated

at 4ºC overnight with the primary antibodies calponin (Dako, monoclonal mouse anti-human

clone CALP) or smooth muscle alpha actin (Dako, monoclonal mouse anti-human clone 1A4)

diluted 1:100 in 1.5% NRS. Control slides were incubated with mouse IgG (Vector). Samples

were incubated in secondary antibody (Invitrogen, Alexa Fluor 488 rabbit anti-mouse) at a 1:400



dilution in NRS for one hour at room temperature, and stained with Hoechst 33342 (Invitrogen;

1:6000 dilution in DI water for 6 minutes) to visualize cell nuclei.

3.2.6. SMC ring thickness and diameter measurements

Rings were removed from the agarose wells and placed in a PBS filled dish under a

machine vision system (model 630; DVT Corporation). Ring thickness was averaged from

measurements in four locations around the circumference of each sample using edge detection

software as described previously (Framework 2.4.6; DVT; [10]). For microsphere incorporation

experiments, these thicknesses were used to calculate cross sectional area and ultimate tensile

stress.

For the TGF-β1 treatment experiments, rings treated with TGF-β1 contracted upon

removal from agarose posts, causing changes in thickness. To control for this, thickness was

calculated from images taken prior to removing rings from molds using ImageJ. After removal,

additional images of rings were taken using a stereoscope (Leica EZ4D). Final diameter (two

measurements per ring) and thickness (four measurements per ring) were measured using ImageJ

to determine changes after contraction.

3.2.7. Mechanical testing

After 14 days, rings were pulled to failure with a uniaxial testing system (ElectroPuls

E1000; Instron) as described previously [10, 12]. Ring cross-sectional areas were calculated from

thickness measurements, and samples were mounted over two stainless steel wires. After

applying a tare load, each ring was subjected to 8 pre-cycles and pulled to failure at 10mm/min

[12]. Data were analyzed in a custom MATLAB (The MathWorks Inc) program to calculate

ultimate tensile stress (UTS; failure load/cross-sectional area), maximum load, maximum strain,

and maximum tangent modulus (MTM; maximum slope of stress/strain curve) of each ring [10,

12].

3.2.8. Western blot analysis

Western blotting was performed with samples flash frozen in liquid nitrogen following

mechanical testing. Samples were lysed for 30 minutes in lysis buffer (diluted from 5X solution

of 200mM Tris at pH of 7.5, 750mM NaCl, 40% glycerol, 0.0635% Triton X-100, 0.025%



Tween-20, and 0.01% NP-40) containing protease inhibitors (Thermo Fisher), mechanically

homogenized, and briefly sonicated. A BCA assay (Thermo Fisher) was then used to determine

protein concentration in each sample, to allow equal amounts of protein to be loaded into each

lane. Samples were boiled for 5 minutes in sample buffer (5X solution of 60mM pH 6.8 Tris-

HCl, 25% glycerol, 2% SDS, 14.4mM β-mercaptoethanol, and 0.1% bromphenol blue) prior to

loading. 15µg of protein per sample was loaded into lanes of polyacrylamide gels with a 10%

resolving and 5% stacking gel. After transfer, PVDF membranes were blocked with 5% nonfat

dry milk powder (BioRad) in Tris Buffer Saline plus Tween 20 (TBST) for 1 hour at room

temperature. Membranes were incubated in smooth muscle alpha actin (1:1,000, Dako,

monoclonal mouse anti-human clone 1A4) or calponin (1:500, Dako, monoclonal mouse anti-

human clone CALP) antibodies diluted in 1% milk powder in TBST overnight at 4ᴼC.

Membranes were incubated for 1 hour at room temperature in secondary antibody (1:3,000 goat

anti-mouse, BioRad). Antibodies were detected using an HRP substrate kit (Thermo Fisher) and

imaged using a BioRad gel documentation system. After imaging, membranes were incubated

overnight at 4ᴼC with anti-Histone (1:250, H3, Santa Cruz) primary antibody as a loading

control, and then one hour at RT with goat anti-rabbit HRP conjugate (1:5000, BioRad) before

imaging. Blots were analyzed using ImageJ. Smooth muscle alpha actin and calponin were both

normalized to histone in each blot.

3.2.9. Statistical analysis

Mechanically tested samples that failed during loading or pre-cycling were omitted from

analysis. Statistical analysis was performed using SigmaPlot software (version 12.5, Systat

Software Inc.). One way ANOVA tests with Holm-Sidak post-hoc analysis were used to

determine statistical significance (p < 0.05) of normal datasets. For datasets that failed a

normality test, a one way ANOVA on ranks test was performed with a Dunn’s multiple

comparison test. Data is represented as mean ± SD.

3.3. Results

3.3.1. Gelatin microsphere characterization

Two batches of cross-linked gelatin microspheres were prepared for microsphere

incorporation and growth factor delivery studies, with average microsphere diameters of 47.5 ±



42.7µm and 48.4 ± 41.9µm (mean ± SD) and

cross-link densities of 32.6 ± 6.1% and 35.7 ±

15.4%, respectively. Previous reports have

characterized degradation and TGF-β1 release

profiles from similarly sized gelatin microspheres

prepared using the same protocol and materials as

this study [21, 22].

3.3.2. Effects of microsphere incorporation on

self-assembled SMC rings cultured in growth

medium

Microspheres were incorporated during ring

self-assembly as shown schematically in Fig. 3.1.

Microspheres appeared incorporated within rings,

with better distribution around the rings when

seeded with 0.6mg/106 cells compared to

0.2mg/106 cells (Fig. 3.2 A-C; G-I). Microspheres

were clearly

visible

within 7-day

rings, but were difficult to discern after 14 days (Fig. 3.2

D-F; J-L), suggesting degradation between 7 and 14 days.

Inclusion of 0.6mg/106 cells significantly

increased ring thickness at 14 days compared to rings with

0.2 or 0mg/106 cells (Fig. 3.3). Microsphere incorporation

caused a significant decrease in ring UTS (Fig. 3.4A) and

MTM (Fig. 3.4B). Significant changes in failure load (Fig.

3.4C) and strain (Fig. 3.4D) were not observed, however

there was a slight decrease in failure load in rings with

0.6mg/106 cells.

Figure 3.2: Gelatin microsphere incorporation

within rings. SMC rings were seeded with 0, 0.2,

or 0.6mg/106 cells and cultured for 7 or 14 days in

growth medium before harvesting for histological

analysis. Hematoxylin and eosin (A-F) and

Picrosirius Red/Fast Green stain (G-L,

red=collagen). Example microspheres marked with

asterisks. Scale = 100µm.

Figure 3.3: Effects of microsphere

incorporation on thickness of rings

cultured in growth medium. Images of

rings seeded with (A) 0, (B) 0.2, or (C)

0.6mg/106 cells and cultured in growth

medium for 14 days and (D) their average

wall thicknesses. Scale = 1mm, n = 6,

*p<0.05. Values are mean ± SD.



3.3.3. Effects of microsphere

incorporation on self-assembled

SMC rings cultured in

differentiation medium

Similarly, histological

analysis of rings cultured in

differentiation medium showed

that microspheres were

incorporated on day 7, with clear

evidence of degradation by day

14 (Fig. 3.5). Rings without

microspheres were significantly

thinner than with 0.6mg/106 cells,

whereas 0.2mg/106 cells did not

significantly increase ring thickness (Fig. 3.6). Overall, rings cultured in differentiation medium

were thinner than rings cultured in growth medium (0.25-0.31mm v. 0.59-0.72mm; Fig. 3.6 and

Fig. 3.3, respectively).

Uniaxial tensile testing of rings cultured in differentiation medium with 0.6mg/106 cells

showed a significant increase in failure load (Fig. 3.7C) and failure strain (Fig. 3.7D). No

significant changes in UTS (Fig. 3.7A) or MTM (Fig. 3.7B) were observed, although a slight

increase in UTS and decrease in MTM was observed in the 0.6mg/106 cells group compared to

rings without microspheres.

3.3.4. TGF-β1 delivery from incorporated microspheres within self-assembled SMC rings

To assess the effects of microsphere-mediated TGF-β1 delivery within rings,

microspheres were loaded with TGF-β1 and incorporated into rings. Unloaded gelatin

microspheres (0.6mg/106 cells) were incorporated into control rings to assess the effects of

microspheres alone on rings with or without exogenously added TGF-β1. Control rings without

microspheres were prepared with and without TGF-β1 supplementation to assess the effects of

exogenous TGF-β1 on SMC contractile protein expression.

Figure 3.4: Mechanical properties of 14 day-old rings cultured in

growth medium. Mean values for (A) UTS, (B) MTM, (C) failure load

and (D) failure strain were calculated for each ring sample. n = 6,

*p<0.05. Values are mean ± SD.



Histological analysis at day 14

showed that microspheres are well

incorporated and still visible at day 14. They

did not appear degraded (Fig. 3.8 C-E, H-J) to

the same extent as observed in initial microsphere incorporation experiments (Fig. 3.5).

Representative images of 14-day rings are shown in Fig. 3.9 A-E. Samples treated with

either exogenous TGF-β1 (Fig. 3.9B, D), or TGF-β1-loaded microspheres (Fig. 3.9E) appeared

to spontaneously contract upon release from the agarose wells to a greater extent than rings that

were not exposed to TGF-β1 (Fig. 3.9A-E). To quantify contraction, the inner diameter of each

ring was measured and the change in diameter was calculated (Fig. 3.9F). Change in ring

thickness was also calculated (Fig. 3.9G). Rings treated with TGF-β1 exhibited a greater

reduction in diameter (Fig. 3.9F) and a greater increase in thickness (Fig. 3.9G) compared to

rings that were not exposed to TGF-β1. In this experiment, before removal from agarose posts,

rings treated with TGF-β1, either exogenously or via microspheres, were significantly thicker

compared to unloaded microspheres without TGF-β1. Specifically, rings had average thicknesses

Figure 3.5: Microsphere incorporation in rings

cultured in differentiation medium. Rings were seeded

with 0, 0.2, or 0.6mg/106 cells, harvested at 7 or 14 days

and stained with (A-F) Hematoxylin and Eosin and (G-L)

Picrosirius Red/Fast Green stain (red = collagen, green =

counterstain). Example microspheres marked with

asterisks. Scale = 100µm.

Figure 3.6: Effects of microsphere

incorporation on thickness of rings cultured

in differentiation medium. Images of rings

seeded with (A) 0, (B) 0.2, or (C) 0.6mg/106

cells and cultured for 14 days and (D) their

average wall thicknesses. Scale = 1 mm; n = 8

for the 0mg group; n = 9 for the 0.2 and

0.6mg/106 cells groups, *p<0.05. Values are

mean ± SD.



(± SD) of 0.14 ± 0.02mm without

microspheres or TGF-β1, 0.12 ±

0.02mm with exogenous TGF-β1 but

no microspheres, 0.25 ± 0.03mm

with microspheres but no TGF-β1,

0.19 ± 0.03mm with microspheres

and exogenous TGF-β1, and 0.21 ±

0.05mm with TGF-β1 loaded

microspheres. A small number of

samples failed during culture or

removal from molds, resulting in the

varying sample sizes in Figure 3.9F

(8 rings per group were originally

seeded). An additional two rings

were excluded from Figure 3.9G, because there was insufficient contrast between the ring and

agarose mold to obtain an initial thickness using the DVT.

Figure 3.7: Mechanical properties of 14 day rings with

incorporated microspheres cultured in differentiation medium.

Mean values for (A) UTS, (B) MTM, (C) failure load and (D) failure

strain were calculated from stress-strain curves for each ring sample.

n = 6, *p<0.05. Values are mean ± SD.

Figure 3.8: Microsphere incorporation in TGF-β1-treated rings. (A,F) Control (untreated) rings. (B,G)

Rings without microspheres cultured with exogenous TGF-β1 (10ng/ml). Rings with unloaded

microspheres are (C,H) untreated or (D,I) treated with exogenous TGF-β1. (E,J) Rings with TGF-β1-

loaded microspheres but without exogenous TGF-β1. (A-E) Hematoxylin and Eosin stain and (F-J)

Picrosirius Red/Fast Green stain, (F-J; red = collagen, green = counterstain). Example microspheres

marked with asterisks. Scale =100µm.



Contractile protein expression was visible in rings from all three TGF-β1-treated groups

(Fig. 3.10B, D, E; G, I, J). While positive staining could be seen throughout the TGF-β1-treated

rings, the strongest signal was observed around ring edges (Fig. 3.10). Smaller amounts of

smooth muscle α-actin and calponin were also observed around the outer edges of rings cultured

without added TGF-β1 (Fig. 3.10 A, C; F, H). Similar observations of ring contraction

(Appendix C, Fig C.2) and contractile protein expression (Appendix C, Fig C.3) with TGF-β1

treatment were observed when the experiment was repeated with SMCs from a different

manufacturer.

These trends were also apparent when contractile protein expression was quantified with

western blotting (Fig 3.10K). Smooth muscle alpha actin expression was significantly higher in

rings with loaded microspheres and rings with unloaded microspheres and exogenous TGF-β1

than in rings without microspheres or TGF-β1 (Fig 3.10L). There were also increases in groups

with exogenous TGF-β1, although the difference was not significant. A small increase was also

seen in the group with microspheres but without TGF-β1. Calponin expression also increased in

Figure 3.9: Effect of TGF-β1 treatment on ring morphology. (A) Untreated control ring with

no microspheres. (B) Ring treated with 10ng/ml exogenous (exo) TGF-β1. (C, D) Ring with

unloaded gelatin microspheres either (C) untreated or (D) treated with exogenous TGF-β1. (E)

Ring with TGF-β1 loaded microspheres. Change in (F) inner diameter and (G) ring thickness

after removal from agarose posts. Scale = 1 mm, *p<0.05. Values are mean ± SD, sample size for

each group shown on bars.



groups treated with TGF-β1 either exogenously or via microspheres, although this was only

significant in the group with microspheres and exogenous TGF-β1 delivery (Fig 3.10M).

When uniaxial tensile testing was performed on rings, several rings failed during loading

or pre-cycling, resulting in low sample sizes (Appendix C, Fig C.1). There were no significant

differences between sample groups (Appendix C, Fig. C.1 A, B, D), except that rings cultured

with unloaded microspheres and no exogenous TGF-β1 had a higher failure load than rings

without microspheres but with exogenous TGF-β1 (Appendix C, Fig. C.1 C). A total of 7 rings

were tested for the group without microspheres or exogenous TGF-β1 and the group with

microspheres but without TGF-β1, 8 rings for the group with microspheres and exogenous TGF-

β1 and the group without microspheres but with exogenous TGF-β1, and 6 rings were tested for

Figure 3.10: Smooth muscle contractile protein expression in rings treated with TGF-β1. (A, F) Control

(untreated) rings. (B, G) Rings without microspheres cultured with exogenous TGF-β1 (10ng/ml). Rings

with unloaded microspheres (C, H) untreated or (D, I) treated with exogenous TGF-β1. (E, J), Rings with

TGF-β1-loaded microspheres. Rings were stained for either (A-E) smooth muscle alpha actin (green) or (F-

J) calponin (green). Nuclei are shown in blue (Hoechst). Scale = 100µm. Corresponding western blots are

shown below (K), with histone (H) loading control shown below each protein. Densitometry analysis is

shown of smooth muscle alpha actin (L) and calponin (M) normalized to histone. Lanes are marked as with

or without exogenous TGF-β1 (Tβ) and with or without microspheres (MS). Loaded MS are marked as with

Tβ. N=4, *P<0.05 (One Way ANOVA on Ranks, Dunn’s Post hoc analysis).



the loaded microsphere group. Rings that failed during pre-cycle or loading were not included in

analysis, resulting in the varying sample sizes.

3.4. Discussion

The goal of this work was to determine the feasibility of microsphere incorporation

within self-assembled SMC rings for the purpose of growth factor delivery, and evaluate effects

on ring mechanical properties and morphology. Microsphere incorporation was tested in two

medium types that have been shown to have different effects on SMC growth and differentiation,

respectively. Therefore, this medium was not used in TGF-β1 delivery experiments. Ring tissue

assembly and microsphere incorporation was successfully demonstrated independent of the

medium in which the tissue rings were cultured.

The effects of microsphere incorporation on mechanical strength were evaluated, as

polymer fragments within tissue engineered constructs can create focal weaknesses [41]. When

rings were cultured in growth medium, a decrease in UTS was measured, which may be due to

the increase in ring thickness and cross-sectional area (given that stress is calculated as force

divided by cross-sectional area). However, the load at failure was not significantly different

between groups. Interestingly, when rings were grown in differentiation medium, significant

increases in failure load were observed in rings with microspheres, although UTS only slightly

increased. This suggests microspheres will not adversely affect ring mechanical strength when

grown in differentiation medium. Others have reported increases in tissue strength and stiffness

with gelatin microsphere incorporation [13, 21], which may be due to improved oxygen and

nutrient diffusion in dense tissues [16, 42]. The decrease in ring MTM was unexpected, as

microsphere incorporation has been shown to increase tissue stiffness [21, 43]. However,

microspheres in this experiment appear to be degraded by 14 days, and may no longer be directly

contributing to stiffness.

It may be noted in this study that there was a large variation in microsphere size,

however, the size distribution between batches are relatively consistent, and there is precedent

for the use of similarly sized microspheres with large size variations [13, 16, 19, 22, 44, 45]. A

large size variation could potentially result in ring failure due to presence of some large

microspheres, as any remaining fragments may create local stress concentrations [41]. However,



standard deviations in failure load were relatively small, suggesting rings were failing

consistently despite the variation in microsphere size. Additionally, the majority of microspheres

in these studies were degraded before mechanical testing. In the few groups where microsphere

fragments were still apparent, ring failure load was not negatively impacted. In future studies, we

may utilize a sieve to obtain microspheres of a consistent size and assess the effect of

microsphere size on tissue ring structure and mechanics.

A second batch of gelatin microspheres was prepared for the TGF-β1 delivery

experiments. It was apparent from histological images that microspheres used for the TGF-β1

studies did not appear completely degraded at day 14 as in the initial microsphere incorporation

experiments. This may be due to differences in cross-link density between the two microsphere

batches, as increased cross-link density has been shown to slow degradation [21, 26].

In vivo, SMCs in healthy blood vessels exhibit a “contractile” phenotype and contract or

relax to regulate blood flow in response to stimuli [38]. Following vascular injury or disease,

SMCs shift to a “synthetic” phenotype characterized by increased proliferation and ECM

deposition, and decreased contractile protein expression [38, 46]. SMCs in culture typically

adopt this synthetic phenotype, making it necessary to differentiate cells in vascular constructs

by switching to a differentiation medium with TGF-β1 [30, 38, 46, 47]. TGF-β1 is well known to

stimulate differentiation to a contractile phenotype and increase contractile protein expression

[27, 28, 37, 38]. Our results are consistent with these observations, as rings supplemented with

TGF-β1, either exogenously or through microspheres, displayed visible increases in expression

of the contractile proteins smooth muscle alpha actin and calponin (Fig 3.10,). These results were

confirmed when contractile protein expression was quantified with western blotting. Smooth

muscle alpha actin was significantly increased compared to untreated controls without

microspheres in the TGF-β1 loaded microsphere group and unloaded microspheres with

exogenous TGF-β1 (Fig 3.10L). Calponin was significantly increased with unloaded

microspheres and TGF-β1 treatment, although trends were visible in all three TGF-β1 groups

(Fig 3.10M). This suggests that microspheres successfully delivered TGF-β1 within tissue rings,

and the bioactivity of TGF-β1 was maintained. Interestingly, there was also a notable, though not

significant, increase in smooth muscle alpha actin in rings with unloaded microspheres but

without exogenous TGF-β1, suggesting microspheres alone may stimulate contractile protein



expression. This is not entirely surprising, as microsphere incorporation alone has been shown to

increase differentiation of other cell types, such as pluripotent stem cells, chondrocytes, and

adipose derived stem cells [17, 48, 49].

Controlling SMC phenotype and ring contractility is an important step for developing in

vitro vascular disease models. In addition to increased contractile protein expression, rings

treated with TGF-β1 visibly contracted when removed from agarose posts, resulting in

significant decreases in diameter (Fig. 3.9), which is an expected outcome of the increased

contractile protein expression. Others have also reported increases in vascular graft contractility

in response to TGF-β1 treatment [39, 40]. Future work will include quantification of active

contraction in response to vasoactive substances, compared to passive tension on the ECM

released upon ring harvest from the agarose posts.

In the TGF-β1 delivery study, TGF-β1 treatment appeared to reduce ring thickness,

although this difference was only significant between unloaded microsphere groups with and

without TGF-β1 supplementation. This may be due to reduced proliferation and matrix

deposition, or increased tissue compaction [50]. While no significant differences in UTS were

observed, the group with the highest failure load contained unloaded microspheres and no TGF-

β1. However, the low sample sizes in some groups limit the conclusions that can be drawn from

this experiment. It also should be noted that fewer rings in the unloaded microsphere groups

failed prior to testing. This supports our conclusion that microspheres alone do not adversely

affect ring strength, and may in fact increase failure load, which is an important criteria for

implantation.

While TGF-β1 release from the gelatin microspheres reported herein has already been

well-characterized for cartilage differentiation [21], delivery may need to be optimized

specifically for SMC differentiation. Future studies will include testing different types and

concentrations of growth factors to increase ring strength and contractility. The effects of TGF-

β1 may be dependent on microsphere distribution within ring tissues, delivery rate, cell density

and the amount of growth factor available per cell [27, 51]. Due to the high cell density of our

constructs, a higher dosage of TGF-β1 or delayed release may be necessary to further enhance

contractility and strength, as TGF-β1 responses have been shown to be dose dependent [28, 35,

37, 52]. It may be possible to delay TGF-β1 supplementation or deliver additional growth



factors, such as platelet-derived growth factor (PDGF) and fibroblast growth factor (FGF) to

promote SMC proliferation and collagen deposition [53, 54]. Gong et al. created a culture system

for growing tissue engineered blood vessels with human mesenchymal stem cells, where grafts

were initially grown in medium supplemented with PDGF to encourage proliferation and

collagen deposition, and then switched to TGF-β1 [28]. This resulted in increased cell number

and collagen deposition, as well as increased contractile protein expression [28]. Modifications

to microspheres, such as increased cross-link density, may be used to slow degradation, resulting

in a longer growth factor delivery period [24]. Others have also demonstrated the use of

polymeric microsphere coatings to reduce burst release and delay growth factor delivery [55].

Published studies have reported that microsphere-mediated delivery of cytokines results

in more homogenous delivery throughout dense tissue constructs and improved cell

differentiation, compared to exogenous treatment [56, 57]. While these data do not include direct

measurements of TGF-β1 diffusion, contractile protein expression is an important outcome of

TGF-β1 delivery. Within rings, exogenous TGF-β1 supplementation and microsphere mediated

delivery resulted in contractile protein expression (Fig. 3.10), which were visible primarily

around ring edges. These increases were uniform around the entire circumference of the rings.

Thus, we have concluded that any slight heterogeneity of microsphere distribution and TGF-β1

delivery does not affect our control over SMC phenotype within each individual ring. This is

consistent with the observation that diffusion limits within dense tissues are typically 100-200µm

[58, 59]. Since rings cultured in differentiation medium with or without TGF-β1 were 200-

300µm thick, and microspheres were in the central portion of the ring, it is expected that TGF-β1

can diffuse throughout the ring.

We have demonstrated that degradable gelatin microspheres can be incorporated into

self-assembled vascular tissue constructs without adversely affecting ring strength, and even

increase ring failure load, supporting the use of microspheres in future studies. Since gelatin is

cell adhesive, it may provide tissue stability at early time points, which will be beneficial when

rings are harvested for tube formation. This stability was evident when commercially available

cross-linked gelatin beads were used by Twal et. al. as microcarriers for forming vascular tissue

constructs [20]. SMCs and endothelial cells were cultured on the beads, which were then seeded

into a mold and fused to form small tissue tubes. In contrast to our ring constructs, there was



lower cell density, gelatin appeared to make up the bulk of the construct, and gelatin bead

degradation was minimal over a 17 day culture period [20]. In our system, microspheres

appeared to be mostly degraded and replaced by ECM within 14 days, and successfully

demonstrated delivery of bioactive TGF-β1 as indicated by induction of SMC differentiation.

One critical advantage of microsphere-mediated delivery is the spatial control over

growth factor release. Microsphere-mediated growth factor delivery stimulated cell

differentiation within self-assembled vascular tissue rings. This will be essential in future studies

aimed at fabricating more complex tubular structures where spatial control of growth factor

delivery may be required. After rings are fused together to form tubes, we may be able to create

localized phenotypic changes by delivering TGF-β1 or other growth factors to specific ring

segments within the tube. This could be applied to modeling diseases such as intimal

hyperplasia, which is characterized by a localized de-differentiation and increased proliferation

of SMCs [60]. Using this system, we may be able to spatially control growth factor release and

differentiation of multiple tissue types within a single tissue construct in order to fabricate more

complex multicellular tissues. Modular fabrication of vascular tissue from microsphere-

incorporated cell rings may enable spatial and temporal regulation of tissue structure and

function, and has the potential to address the need for functional human vascular tissue as model

systems for screening therapies in vitro.

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56. Carpenedo, R.L., A.M. Bratt-Leal, R.A. Marklein, S.A. Seaman, N.J. Bowen, J.F.

McDonald, and T.C. McDevitt, Homogeneous and organized differentiation within

embryoid bodies induced by microsphere-mediated delivery of small molecules.

Biomaterials, 2009. 30(13): p. 2507-15.

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58. Curcio, E., S. Salerno, G. Barbieri, L. De Bartolo, E. Drioli, and A. Bader, Mass transfer

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Engineering. Tissue Engineering: Part B, 2009. 15(3).



Chapter 4: Fabrication and characterization of electrospun polycaprolactone cuffs for self-assembled vascular tissue

72

Chapter 4: Fabrication and characterization of electrospun

polycaprolactone cuffs for self-assembled vascular tissue

Taken from part of: H. A. Strobel, E. L. Calamari, A. Beliveau, A. Jain, and M. W. Rolle, “Fabrication

and characterization of electrospun polycaprolactone and gelatin composite cuffs for tissue engineered

blood vessels.” JBMR Part B, 2018. 106B(2): p. 817-826. Reprinted with permission (Appendix D)

Supplemental figures in Appendix E

Authorship contributions: HAS fabricated material, performed all experiments and data analysis except for porosity

measurements, and wrote and revised the manuscript. ELC developed and optimized all protocols and assays during

her Master’s thesis project, performed porosity measurements, and designed experiments. AB and AJ contributed to

experimental design and protocol development, particularly regarding electrospinning parameters. MWR

contributed to experimental design, supervised data collection, data analysis, and preparation of the manuscript,

and edited the manuscript.

4.1. Introduction

Tissue engineered blood vessels (TEBVs) are being evaluated in clinical trials as vascular

grafts for dialysis access [1-3] and pediatric cardiovascular surgery [4], with the potential to treat

millions of patients requiring blood vessel repair or replacement. In addition, TEBVs are being

developed to serve as in vitro tools for disease modeling and drug screening [5-7]. Several

approaches for fabricating TEBVs have been reported, including seeding cells on synthetic

polymer scaffolds [8, 9], seeding cells in natural hydrogels [10, 11], or facilitating cells to self-

assemble and produce their own extracellular matrix (ECM) without artificial scaffold materials

[12, 13]. These “scaffold-free”, self-assembled tissues are advantageous due to their natural

biocompatibility, lack of synthetic or xenogeneic components, and enhanced cell signaling and

ECM deposition compared to scaffold-based approaches [14, 15].

A disadvantage of cellular self-assembly is that scaffold-free TEBVs may be fragile in

the early stages of fabrication and culture, before deposition of sufficient cell-derived ECM to

support tissue structure and strength. Because of this, securing tissues to cannulas or other

mechanical testing apparatus may result in failures, as suturing is likely to tear the tissue.

Extensions or “cuffs” integrated into the ends of the TEBVs allow tissue to be secured and

handled, without damaging the tissue itself. Previous studies report the use of sewing cuffs

attached at each end of the TEBV to facilitate suturing in vivo [16], and for mounting samples

onto bioreactors in vitro [17, 18]. For example, Huang et al. report using Dacron cuffs fastened


73

to the ends of TEBVs comprised of poly-glycolic acid (PGA) scaffolds seeded with cells to

cannulate the vessel for dynamic culture [17]. Poly-lactic acid (PLA) cuffs were integrated into

the ends of fibrin gel-based constructs by Syedain et al. [18] to mount TEBVs in a bioreactor.

Sewing cuffs made from polyurethane sponges have also been used to provide a reinforced

interface for graft anastomosis with native vessels in vivo [16]. For each of these studies, cuffs

were sutured to the TEBV, encapsulated by the tissue during graft maturation, or embedded

within the TEBV scaffold [16-18]. Integration of cuffs with scaffold-free TEBVs has not been

reported, and may require a different approach. Scaffold-free TEBVs do not utilize hydrogel

materials that facilitate cuff embedding, and unlike polymer scaffold-based constructs, suturing

to cuffs may tear cell-derived tissues in the first several days or even weeks following self-

assembly. Although scaffold-free TEBVs created by cellular self-assembly may ultimately

develop mechanical strength equivalent to saphenous vein grafts [19], suturing or embedding the

cuff material may not be possible in the early stages of graft maturation.

To create materials that allow cell and tissue integration during scaffold-free TEBV

maturation, we fabricated custom cuff materials using electrospinning. Electrospinning enables

fabrication of porous, nanofiber materials conducive to cellular infiltration. The high surface area

to volume ratio allows improved cellular attachment and proliferation compared to polymer films

[20, 21]. Additionally, variables such as porosity, which affects cellular infiltration, can easily be

controlled by altering electrospinning parameters [22-24].

We selected poly-caprolactone (PCL) as the electrospun cuff material for use in these

studies. The use of PCL in electrospinning is well-established, including for vascular

applications [25, 26]. Additionally, PCL is biocompatible and FDA-approved as a drug delivery

and suture material [27]. Although PCL is biodegradable, degradation can take 1-2 years, and

thus will maintain its integrity during culture periods of weeks to months [26].

The goal of this study was to develop an electrospun cuff material that promotes cellular

infiltration and creates a strong interface with scaffold-free TEBVs. We incorporated cuffs into

TEBVs using a modular cellular self-assembly and fusion approach developed in our laboratory

([13]). Cuff materials were placed at each end, and tubes were cultured for 7 days to allow fusion

between cell rings and tissue integration with cuffs. We hypothesized that cell migration into

cuffs from adjacent tissue rings would result in cuff integration with the TEBVs. To evaluate the


74

strength of the tissue-cuff interface, we developed a custom system to cannulate and grip TEBVs

and longitudinally pull them to failure. To our knowledge, this is the first report describing

design and integration of electrospun cuffs in scaffold-free self-assembled TEBVs.

In addition to the work discussed here, the incorporation of gelatin within electrospun

cuffs was evaluated and compared to PCL alone. Preliminary experiments for the comparison

were discussed in the Master’s thesis of Elizabeth Mayor [28], with additional results of gelatin

incorporation experiments presented in Strobel et al. [29], which demonstrated that gelatin did

not have any significant effects on cellular attachment or strength of the cuff-tissue interface. In

this chapter, we discuss the development and characterization of electrospun PCL cuffs (without

gelatin), which were used for all subsequent experiments in Chapters 5, 6, and 7.

4.2. Methods

4.2.1. Electrospinning setup

A custom setup was fabricated with a syringe pump (SP200i, World Precision

Instruments), high voltage power supply, and a stainless steel collecting mandrel (2mm diameter)

attached to a variable speed motor (2Z846, Grainger) via a series of custom-machined couples. A

3cc syringe with 19-gauge blunt needle was used for dispensing polymer solution from the

syringe pump at a rate of 5 ml/hour for 12 minutes. The collecting mandrel was rotated at

approximately 265 rpm, with a tip-to-collector distance of 15 cm and applied voltage of 15-20

kV. To spin flat sheets used for cell attachment assays, a custom aluminum drum (5cm diameter)

was used instead of the collecting mandrel, with a spin time of 6 minutes instead of 12. Collector

distance, voltage, and flow rate were optimized in preliminary experiments, described in the

Master’s thesis of Elizabeth Mayor [28].

4.2.2. Fabrication of PCL electrospun cuffs

Poly-ε-caprolactone (PCL; 440744, Sigma Aldrich) was dissolved in 2,2,2 Tri-fluoro-

ethanol (TFE, T63002, Sigma Aldrich) at 12% (w/v) and mixed overnight on a stir plate at room

temperature. All electrospun materials were sterilized with ethylene oxide and allowed to degas

for 24 hours prior to cell culture experiments.


75

4.2.3. Fiber diameter measurement

Scanning electron microscopy (SEM) was performed at the University of Massachusetts

Medical School (UMMS) Core Electron Microscopy Facility. Three samples from each of three

batches of the three material types were sputter-coated with gold/palladium (80:20, thickness 8

nm) and imaged using an FEI Quanta 200 EFEG MKII scanning electron microscope. Fiber

diameter then was measured using a plugin for ImageJ, DiameterJ, as detailed in Hotaling et al.

[30]. The program maps fibers visible in SEM images and measures the diameter at multiple

locations along each fiber. Two SEM images taken at 2,500X were analyzed per batch of

material, with greater than 13,000 measurements per image.

4.2.4. Tensile testing of electrospun cuffs

Materials were hydrated in phosphate-buffered saline for 30 minutes prior to mechanical

testing. Thickness (average of 4 measurements) and length (average of 2 measurements) of cuffs

were measured using calipers and used to calculate cross sectional area. Samples were loaded

over the tips of two wires (bent 90º) and pulled to failure at 10 mm/min using a uniaxial testing

system (Instron, ElectroPuls E1000) with a 50N load cell. The cuff load at failure, ultimate

tensile stress (UTS, calculated from load/cross sectional area), elastic modulus, and strain at

failure were measured. Three batches per group and three samples per batch were tested.

4.2.5. Cell culture

Rat aortic smooth muscle cells (RaSMCs, WKY 3M-22) were cultured in DMEM

(VWR) supplemented with 10% fetal bovine serum (Atlanta Biologicals), 1%

penicillin/streptomycin (VWR), 1% nonessential amino acid solution (VWR), 1mM sodium

pyruvate (VWR) and 2mM L-glutamine (VWR). Cells were passaged at 70% confluence.

4.2.6. TEBV fabrication from self-assembled tissue rings

Rings and tubes were grown as described previously [13] with slight modifications to

mold design. Briefly, 500,000 RaSMCs per ring were seeded into custom agarose wells with a 2

mm central post and ring-shaped, round-bottomed seeding channel. Rings were cultured for 3

days prior to harvesting for tube formation. Three rings per tube were threaded over a beveled

piece of silicone tubing (2 mm outer diameter, Specialty Manufacturing Inc.), and electrospun


76

cuffs were stacked onto either end of the tube. The tubes were then mounted into a custom

polycarbonate holding device as described previously [13, 31], and cultured for 7 days prior to

longitudinal pull to failure testing. Culture medium was changed every 3 days.

4.2.7. Longitudinal pull to failure testing

A custom grip was designed for longitudinal pull-to-failure tests based on Berry et al.

[32] with modifications. A detailed photographic description of grip assembly is given in

Appendix E (Fig E.1). Briefly, tubes were cannulated and clamped between two pieces of 3D

printed ABS plastic (Dimension SST 1200es 3D printer), which are mounted onto a support base

until testing. PDMS spacers threaded over a screw maintained the distance between the two sides

of the support base during loading. Once the sample and grip assembly were loaded onto the

uniaxial tensile testing system (Instron, ElectroPuls E1000), the support base was removed, and

tubes were pulled to failure at 1 mm/min using a 1N load cell (Instron). Tubes remained

submerged in PBS while being secured in the grip assembly. After clamping grips onto the

Instron, samples were hydrated by applying PBS onto the tissue using a pipet until the start of the

test. The maximum load at failure and failure location for each tube was recorded. Three samples

per batch were tested, with three batches per material. Tubes that failed during loading were

excluded from analysis.

4.2.8. Hoechst staining

After mechanical testing, tubes were fixed for 1 hour in 10% neutral buffered formalin.

Tubes were incubated in 30% sucrose overnight, and embedded in OCT. Tubes were sectioned

on a cryostat (Leica CM3050) at -18ºC, 10 µm per section. Sections were stained with Hoechst

33342 (Invitrogen, 1:6000 dilution in DI water) for 6 minutes. Images were acquired using an

upright microscope (Leica DMLB2) with a digital camera (Leica DFC 480).

4.3. Results

4.3.1. Characterization of electrospun PCL cuffs

We successfully fabricated electrospun materials from PCL. Representative SEM images

of flat sheets of electrospun materials are shown in Fig. 4.1. PCL sheets had an average fiber

diameter of 0.50 ± 0.01 µm (mean ± SEM, n = 3). To determine the mechanical strength and


77

stiffness of electrospun cuffs, cuff samples were pulled

uniaxially to failure. Cuffs had an average failure load of 4.96 ±

0.18 N, UTS of 2.32 ± 0.53 MPa, failure strain of 2.29 ± 0.21

mm/mm, and elastic modulus of 1.34 ± 0.32 MPa.

The ultimate test of cuff materials is the strength of their

integration with scaffold-free TEBVs. To test this, we fabricated

TEBVs with integrated cuffs, cannulated and gripped the cuffs in

a custom grip system, and longitudinally pulled tubes to failure.

Tubes failed at 105 ± 0.47 mN, and failed an approximately even

number of times at either the tissue-cuff

interface or within the tissue. Figure 4.2

shows representative time-lapse images of

tube failure within the tissue (top) and at

the tissue-cuff interface (bottom). Several

tubes with PCL cuffs failed during the

loading procedure, which required some

refinement.

After mechanical testing, tubes

were sectioned and stained with Hoechst

dye to visualize cell migration into the

material (Fig. 4.3). Cells appeared to

migrate along the outside of the cuff

material to form a coating several cell

layers thick. A thinner cell layer was

visible along the lumen, where cuffs

were in contact with silicone tubing.

Additionally, cells appeared to migrate

into the material.

Figure 4.2: Longitudinal pull to failure testing of fused

tubes. PCL tubes failed at either the cuff-tissue interface or

within the tissue (A). Representative time-lapse images of

tubes in custom grips failing at the cuff-tissue interface (B,

top), and within the tissue (B, bottom)

A B

Figure 4.3: Cellular infiltration within cuff materials. Tubes

were stained with Hoechst dye following pull to failure testing

(A). Higher magnification image of tube (B) are shown to

visualize cellular ingrowth. Blue = nuclei. Scale = 100 µm. Images

representative of 3 samples.

Figure 4.1: SEM image of

electrospun PCL material. Scale

= 10µm. Representative of 3

samples, 3 images per sample.


78

4.4. Discussion

Many factors affect the pore size, fiber diameter, and porosity of electrospun materials,

including the viscosity of the polymer solution (affected by polymer concentration and solvent),

flow rate, applied voltage, and collector distance [22, 33-35]. We selected electrospinning

parameters based on optimization experiments to test the effects of these factors on Taylor cone

formation and fiber morphology, with the goal of obtaining a material with mean pore size to

allow cellular ingrowth and integration of the cuff material with the self-assembled vascular

tissue. Based on optimization studies, we chose a collector distance of 15cm, voltage of 15-

20kV, and flow rate of 5 ml/hr. This allowed for a pore size that allowed cellular infiltration of

rat aortic SMCs, as is visible in Hoechst staining of samples.

It is important that electrospun cuffs have a strong tissue-cuff interface to prevent damage

or leakage when handling and cannulating TEBVs. Thus, we developed a custom system for

cannulating and gripping tubes on a uniaxial tensile testing machine and pulling them

longitudinally to failure to test the strength of the cuff-tissue interface. Longitudinal pull-to-

failure testing is typically performed on rectangular or dog-bone tissue sections secured with

clamps [9, 36, 37]. Early in culture, self-assembled tissue may be too fragile to be secured using

a clamp system without tearing. Our method is advantageous because tubular tissues retain their

shape and structure during testing. This procedure can be applied broadly to test fusion of tissue

constructs, or integration of electrospun materials with other types of engineered tissues.

Vascular tissue tubes with PCL cuffs failed approximately equally at the cuff-tissue interface and

within tissue tubes. These results suggest that the cuff-tissue interface is as strong as the tissue

itself.

Cuffs may also have broader applications as a reinforced interface for suturing TEBVs in

in vivo studies. While the strength of PCL is favorable, its higher stiffness compared to native

vessels [26] could be problematic if cuffs are used as reinforced interfaces for in vivo

transplantation, as compliance mismatch between graft and native vessel can trigger intimal

hyperplasia [38].

We previously determined that there are no significant advantages to using PCL blended

or coated with gelatin compared to PCL alone [29]. Thus, we will continue to use pure PCL cuffs


79

fabricated using the above protocol due to the simplicity of fabrication and the greater stability of

PCL alone during storage. Overall, the incorporation of electrospun cuffs may be a critical step

in the fabrication and testing of scaffold-free tubular tissues, by allowing cannulation at early

time points in tissue culture and maturation. We further explore this possibility in Chapter 5,

where cannulation cuffs are used to aid in dynamic tissue tube culture.

4.5. References

1. Lawson, J.H., M.H. Glickman, M. Ilzecki, T. Jakimowicz, A. Jaroszynski, E.K. Peden,

A.J. Pilgrim, H.L. Prichard, M. Guziewicz, S. Przywara, et al., Bioengineered human

acellular vessels for dialysis access in patients with end-stage renal disease: two phase 2

single-arm trials. The Lancet, 2016. 387(10032): p. 2026-2034.















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9. Soletti, L., Y. Hong, J. Guan, J.J. Stankus, M.S. El-Kurdi, W.R. Wagner, and D.A. Vorp,

A bilayered elastomeric scaffold for tissue engineering of small diameter vascular grafts.

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10. Syedain, Z.H., L.A. Meier, M.T. Lahti, S.L. Johnson, and R.T. Tranquillo, Implantation

of completely biological engineered grafts following decellularization into the sheep

femoral artery. Tissue Eng Part A, 2014. 20(11-12): p. 1726-34.

11. Swartz, D.D., J.A. Russell, and S.T. Andreadis, Engineering of fibrin-based functional

and implantable small-diameter blood vessels. Am J Physiol Heart Circ Physiol, 2005.

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12. L'Heureux, N., N. Dusserre, G. Konig, B. Victor, P. Keire, T.N. Wight, N.A. Chronos,

A.E. Kyles, C.R. Gregory, G. Hoyt, et al., Human tissue-engineered blood vessels for

adult arterial revascularization. Nat Med, 2006. 12(3): p. 361-5.










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16. Watanabe, T., K. Kanda, H. Ishibashi-Ueda, H. Yaku, and Y. Nakayama, Development of

biotube vascular grafts incorporating cuffs for easy implantation. J Artif Organs, 2007.

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17. Huang, A.H. and L.E. Niklason, Engineering biological-based vascular grafts using a

pulsatile bioreactor. J Vis Exp, 2011(52).

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grafts from human fibroblasts and fibrin using a multi-graft pulsed flow-stretch

bioreactor with noninvasive strength monitoring. Biomaterials, 2011. 32(3): p. 714-22.

19. Konig, G., T.N. McAllister, N. Dusserre, S.A. Garrido, C. Iyican, A. Marini, A. Fiorillo,

H. Avila, W. Wystrychowski, K. Zagalski, et al., Mechanical properties of completely

autologous human tissue engineered blood vessels compared to human saphenous vein

and mammary artery. Biomaterials, 2009. 30(8): p. 1542-50.

20. Guarino, V., M. Alvarez-Perez, V. Cirillo, and L. Ambrosio, hMSC interaction with PCL

and PCL/gelatin platforms: A comparative study on films and electrospun membranes.

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21. Wu, S.-C., W.-H. Chang, G.-C. Dong, K.-Y. Chen, Y.-S. Chen, and C.-H. Yao, Cell

adhesion and proliferation enhancement by gelatin nanofiber scaffolds. Journal of

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22. Nottelet, B., E. Pektok, D. Mandracchia, J.C. Tille, B. Walpoth, R. Gurny, and M.

Moller, Factorial design optimization and in vivo feasibility of poly(epsilon-

caprolactone)-micro- and nanofiber-based smhall diameter vascular grafts. J Biomed

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23. Lee, J., G. Tae, Y.H. Kim, I.S. Park, S.H. Kim, and S.H. Kim, The effect of gelatin

incorporation into electrospun poly(L-lactide-co-epsilon-caprolactone) fibers on

mechanical properties and cytocompatibility. Biomaterials, 2008. 29(12): p. 1872-9.

24. Zhang, Y., H. Ouyang, C.T. Lim, S. Ramakrishna, and Z.M. Huang, Electrospinning of gelatin fibers and gelatin/PCL composite fibrous scaffolds. J Biomed Mater Res B Appl

Biomater, 2005. 72(1): p. 156-65.

25. Ma, Z., W. He, T. Yong, and S. Ramakrishna, Grafting of Gelatin on Electrospun

Poly(caprolactone) Nanofibers to Improve Endothelial Cell Spreading and Proliferation

and to Control Cell Orientation. Tissue Eng, 2005. 11: p. 1149-1158.

26. de Valence, S., J.C. Tille, D. Mugnai, W. Mrowczynski, R. Gurny, M. Moller, and B.H.

Walpoth, Long term performance of polycaprolactone vascular grafts in a rat abdominal

aorta replacement model. Biomaterials, 2012. 33(1): p. 38-47.

27. Cipitria, A., A. Skelton, T.R. Dargaville, P.D. Dalton, and D.W. Hutmacher, Design,

fabrication and characterization of PCL electrospun scaffolds—a review. Journal of

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28. Mayor, E., Fabrication and characterization of electrospun poly-caprolactone-gelatin

compoosite cuffs for tissue engineered blood vessels, in Biomedical Engineering. 2015,

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30. Hotaling, N.A., K. Bharti, H. Kriel, and C.G. Simon, Jr., DiameterJ: A validated open

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34. Sisson, K., C. Zhang, M.C. Farach-Carson, D.B. Chase, and J.F. Rabolt, Fiber diameters

control osteoblastic cell migration and differentiation in electrospun gelatin. J Biomed

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35. Panzavolta, S., M. Gioffre, M.L. Focarete, C. Gualandi, L. Foroni, and A. Bigi,

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Chapter 5: Generate modular vascular tissue tubes with luminal flow 83

Chapter 5: Generate modular vascular tissue tubes with luminal

flow

Modified from: H. A. Strobel, T. A. Hookway, M. Piola, M. Soncini, G. B. Fiore, E. Alsberg, and M. W.

Rolle. “Assembly of tissue engineered blood vessels with spatially-controlled heterogeneities”. Tissue

Engineering Part A. In Press.

Supplemental figures presented in Appendix F.

Authorship contributions: HAS designed and performed experiments, collected and analyzed all data, made all

figures and wrote and revised the manuscript. TAH contributed to experimental design and data analysis and edited

the manuscript. MP, MS, and GBF designed and fabricated bioreactors, contributed to experimental design and

data analysis, and edited the manuscript. EA provided gelatin microspheres, contributed to experimental design and

analysis, and edited the manuscript. MWR contributed to experimental design, supervised data collection, data

analysis, and preparation of the manuscript, and edited the manuscript.

5.1. Introduction

Cardiovascular disease is the leading cause of death in the United States [1]. By 2030,

43.9% of Americans will be living with some form cardiovascular disease [1]. Prior to clinical

testing, new treatments for cardiovascular diseases are tested in 2D cell cultures and animal

models. However, 2D cultures are not representative of the 3D mechanical environment and cell-

matrix interactions found in vivo [2], and animal studies do not accurately predict the success of

drugs in humans; many drugs that are successful in animals fail in human clinical trials [3, 4].

Thus, there is a strong need for 3D human tissues to model vascular diseases and serve as tools

to screen potential therapies [5-10]. Testing drugs on functional human tissues in vitro may allow

researchers to eliminate ineffective drugs earlier in the testing process, and accelerate the

development of new, lifesaving treatments.

Several approaches have been reported for fabricating functional 3D human vascular

tissue for drug screening and disease modeling [10-14]. For example, Fernandez et al. developed

a functional smooth muscle cell (SMC)-endothelial cell co-culture tube for drug testing by

seeding cells in collagen gels, which reacted to vasoactive stimuli [11]. There are also many

existing approaches for fabricating functional human tissue engineered blood vessels (TEBVs)

for implantation, including seeding cells on polymer scaffolds [15, 16], seeding cells in

hydrogels [11, 17], and using scaffold-free cellular self-assembly approaches [10]. However,

most TEBV approaches use cells seeded on or within tubular scaffolds, or use rolled cell sheets,


to create a homogenous tissue tube. In contrast, many vascular diseases, such as aneurysm and

intimal hyperplasia, create focal changes in SMC phenotype or matrix composition in localized

regions of the blood vessel wall, not the entire length of the vessel [18, 19].

Alternatively, bioprinting can be used to create complex, modular, tubular tissue

structures, which is typically done by fusing spheroidal subunits [20-22]. However, tissue

spheroids do not fuse as effectively as other cell aggregate shapes, because even tightly packed

spheroids have limited surface area where spheres are in contact with one another, compared to

other shapes such as rings [20]. Thus, TEBVs fabricated from spheroids often have distinct

fusion boundaries or gaps where spheroids were not in close contact [20, 23, 24].

While some existing homogenous TEBVs may be effective for screening drugs on

healthy tissues, they are not conducive to creating focal regions of pathological tissue.

Regardless of the tissue engineering method used, the creation of localized heterogeneities

within human TEBVs has not been previously reported. Thus, the primary goal of this study was

to establish a model system that achieves spatial control of cell position and tissue structure

within human TEBVs in order to introduce focal heterogeneities. To achieve this goal, we

utilized a unique modular system developed in our laboratory for fabricating self-assembled

vascular tissue from individual ring units [25]. Human SMCs were seeded into ring-shaped

agarose molds, where they aggregated in less than 24 hours to form self-assembled tissue rings

[25]. Within 3 days, rings can be threaded onto silicone tubing, stacked together, and fused to

form vascular tissue tubes. Unlike spheroids, rings can be pushed into close contact and fuse

without gaps between tissue units [26].

Here, we present a novel approach for fabricating spatially controlled 3D vascular tissue

from human cells, which may ultimately serve as a platform technology to introduce focal

regions of pathological tissue within TEBVs. To aid in handling and cannulation, electrospun

polycaprolactone (PCL) cannulation cuffs can be fused onto tube ends as reinforced extensions.

We then validated that after 7 days of fusion culture, tubes with PCL cuffs could be cannulated

and dynamically cultured on a custom luminal flow bioreactor.


In summary, we demonstrated a technology for creating TEBVs that allows for

customization of tissue structure and composition along the vessel length. In future studies, this

system may be modified to model focal human vascular diseases.

5.2 Methods

The first goal of the study was to enhance ring fusion and reduce fusion time. This was

accomplished by evaluating the effects of ring pre-culture time on ring fusion. The next goal was

to evaluate if rings maintain spatial positioning during fusion, to determine the feasibility of

creating focal heterogeneities. The third goal of this study was to demonstrate that tissue tubes

can be cannulated and dynamically cultured. PCL cannulation cuffs were incorporated on tube

ends as reinforced extensions for cannulation, and the use of a custom luminal flow bioreactor

for dynamic tube culture was demonstrated. The final goal was to create focal heterogeneities

within tubes, which was accomplished by creating localized regions of microsphere

incorporation.

5.2.1. Cell culture

Human aortic SMCs (Lifeline) were cultured in Lifeline VascuLife complete growth

medium containing 10mM L-glutamine, 5% FBS, 5µg/ml insulin, 5ng/ml fibroblast growth

factor-basic, 50µg/ml ascorbic acid, 5ng/ml epidermal growth factor, 30mg/ml gentamicin, and

15µg/ml amphotericin B.

5.2.2. Tissue ring fabrication

Agarose wells (2mm post diameter) were prepared as described previously from 2%

agarose (Lonza) dissolved in DMEM and autoclaved [27]. Human aortic SMC rings were seeded

into agarose molds designed to fit 5 rings in a well of a 6-well plate [27], at a density of 400,000

cells/ring. Molds were equilibrated overnight in growth medium before use. All seeded rings

were incubated overnight to allow cell aggregation, then wells were flooded with fresh growth

medium.

5.2.3. Tissue tube fusion with varying pre-culture time


To generate tissue tubes, rings

fabricated from human aortic SMCs

were removed from agarose molds at 3,

5, or 7 days in culture and threaded onto

silicone tubing mandrels (Specialty

Manufacturing Inc., O.D. 2 mm) [26].

Three rings per tube were gently pushed

together on the mandrel to ensure rings

were in contact with each other (Fig

5.1B), and the mandrel was secured in

custom polycarbonate holders, which

were placed in a 10 cm dish with 45 ml

medium [26]. The tubes were then

allowed to fuse for an additional 7 days

of static culture on the silicone mandrels.

The experiment was duplicated once

more with the same human aortic SMCs,

and once again with human coronary

artery SMCs from a different donor

(Appendix F Figures F.1-3).

5.2.4. Fusion angle, length, and thickness measurements

A Leica inverted microscope (DMIL) with a digital camera (Leica DFC 480) was used to

take brightfield images of tubes daily for one week. Image J software (NIH) was used to measure

the angle between rings (fusion angle, ɵ), tube thickness (T), and tube length (L). Four fusion

angle measurements, six thickness measurements, and two length measurements were obtained

for each tube sample at each time point and averaged to yield a single mean for each parameter

per tube per time point. Three independent tube samples were averaged for each of the three pre-

culture conditions. After 7 days of culture, tissue samples were fixed for 1 hour in 10% neutral

buffered formalin for histological analysis. Data is represented as mean ± SD.

Figure 5.1: Schematic of tube fabrication process, and

tissue tube culture experimental groups for the ring pre-

culture duration experiment. Rings are formed by seeding

SMCs into ring-shaped agarose molds, where cells

aggregate around 2 mm diameter posts and form rings in

less than 24 hours (A). Rings are then removed from molds

and threaded onto silicone tubing, where they are pushed

together and cultured for 7 additional days to allow fusion

(B). To test the effects of varying ring culture duration, rings

were cultured for 3, 5, or 7 days (“ring culture”), followed

by 7 days of “fusion culture” for all groups (C). Groups are

labeled as: days in ring culture – days in fusion culture (ex.

Group 3-7 = 3 days in ring culture followed by 7 days in

fusion culture). Black dots = SMCs.


5.2.5. CellTracker labeling

CellTracker red and green (CMTPX and CMFDA, Invitrogen) were reconstituted to

10mM in DMSO and diluted to a final concentration of 5µM in DMEM (Corning). Plates of

human aortic SMCs were rinsed with PBS and incubated with either red or green CellTracker

solution at 37°C for 45 minutes. The plates were rinsed with PBS, and growth medium was

added for an additional 30 minutes at 37°C. Cells were then passaged and seeded into ring molds

as described in section 5.2.2. After 3 days of culture, tubes were fabricated with alternating red-

and green-labeled rings and imaged with an inverted fluorescent microscope (Leica DMIL) daily

for 7 days. The experiment was duplicated with human coronary artery SMCs from a different

donor (Appendix F, Supplemental Methods). In a separate experiment examining cell

proliferation, rings were loaded with CellTracker Red dye only, pre-cultured for 3 days, and then

fused into tubes with 3 rings per tube. Tubes were fixed after 1 or 2 days of fusion, to evaluate

proliferation at earlier time points with Ki67 staining.

5.2.6. Polycaprolactone (PCL) cannulation cuff fabrication

Electrospun PCL cuffs were prepared as described previously ([28], Chapter 4). Briefly,

PCL was dissolved in 2,2,2 tri-fluoro-ethanol (TFE, T63002, Sigma) to form a 12% solution.

The solution was then electrospun onto a 2 mm diameter mandrel using a 5 ml/hour flow rate, 15

cm collector distance, and a voltage of 15-20 kV ([28], Chapter 4). Cuffs were cut into segments

approximately 3-4 mm in length, sterilized with ethylene oxide, and allowed to de-gas for a

minimum of 48 hours before use.

5.2.7. Bioreactor culture

Rings were fabricated with human aortic SMCs (Lifeline) as described above in section

5.2.2, and threaded onto silicone tubing after 3 days of ring pre-culture, with PCL cuffs adjacent

to rings on either end. Tubes were allowed to fuse for 7 days on silicone mandrels in static

culture prior to removal from the silicone tubing mandrel and cannulation onto a custom

bioreactor modified from Piola et al. [29]. Each bioreactor fits in its own individual 15 ml

conical tube, which allows for multiple units to be cultured independently, with minimal culture

medium (approximately 19 ml medium to fill each bioreactor unit and its tubing). The inner

cannulas are adjustable, to accommodate tubes with lengths ranging from a few millimeters up to


3 cm. Medium flows from a peristaltic pump (Watson Marlow, Model 323Du), equipped with a

multichannel pump head (Watson Marlow, Model 318MC). A syringe with 2 ml of medium and

3 ml of air is positioned between the pump and vessel, to dampen oscillations in medium flow

between the pump and the tissue tube. The medium then flows back into the culture chamber,

before returning to the pump. For these experiments, the bioreactor was set up to apply luminal

flow, but not pressure. We verified that pressures between cannulas are approximately zero in

benchtop experiments (not shown) prior to beginning these studies. Cannulated tubes (n=5) were

cultured with 35 ml/min applied luminal flow (corresponding to an estimated 12 dyne/cm2 wall

shear stress) for 7 days prior to fixing for histology. Two control tubes were left on silicone

mandrels in static conditions for a total of 14 days (same total culture time as tube exposed to 7

days of flow).

5.2.8 Histology and immunohistochemistry

After fixing for 1 hour in 10% neutral buffered formalin, samples were processed and

embedded in paraffin. Longitudinal sections 5 µm thick were adhered to positively-charged

slides. Hematoxylin and Eosin staining was used to examine tube morphology. Picrosirius

red/fast green and orcein stains were used examine collagen and elastin deposition, respectively.

Antigen retrieval was performed on samples to be stained for Ki67, smooth muscle alpha

actin (SMA), smooth muscle protein 22 alpha (SM22-α), and calponin by incubating slides in

10mM Tris, 1mM EDTA, and 0.05% Tween-20 (pH 9.0) in a pressure cooker for 5 minutes.

Slides were blocked in 5% normal goat serum (Ki67) or 1.5% normal rabbit serum (SMA,

SM22-α, and calponin) for 30 minutes, and were incubated overnight at 4ºC in anti-Ki67 (Abcam

Ab16667; 1:100), SMA (Dako, clone 1A4, 1:100), SM22-α (BioRad VPA00048, 1:100), or

calponin (Dako, CALP, 1:100) antibodies. Negative control samples were incubated with rabbit,

mouse, or goat immunoglobulin G (Vector). Samples were incubated in a secondary antibody

(Invitrogen; Alexa Fluor 488 goat anti-rabbit, rabbit anti-mouse, or mouse anti-goat) at a 1:400

dilution for 1 hour at room temperature. Samples with CellTracker labeling or antibody stains

were stained with Hoescht dye to visualize nuclei (Invitrogen, 1:6000 in DI water for 6 minutes).

Images were acquired using an epifluorescent microscope (Leica DMLB2) with a digital camera

(Leica DFC 480).


5.2.9. Statistics

Statistical tests were performed using SigmaPlot software (Version 11.0 Systat Software,

Inc.). A two-way analysis of variance (ANOVA) with Holm-Sidak post hoc analysis was used to

compare fusion angles, thicknesses, and lengths of tissue tubes. A p-value of less than 0.05 was

considered significant. A sample size of n=3 was used in statistically analyzed ring pre-culture

experiments.

5.3. Results

5.3.1. Effect of ring pre-culture time on human SMC tube fusion rate

The first goal of this study was to accelerate production of tissue tubes and enhance ring

fusion by examining how ring “pre-culture” time prior to tube fabrication affects fusion. In

previous studies, we observed cohesive tubes after fusion, but ring boundaries remained visible

after a total 14 day culture period (7 days as rings, 7 as tubes) [25]. By decreasing ring pre-

culture duration prior to fusion, we aimed to decrease the length of time required to generate

tissue tubes, and generate a more seamless ring fusion. Other published studies also suggest that

less mature cell aggregates fuse together more rapidly than more mature tissues [30, 31].

Therefore, we hypothesized that decreasing the ring pre-culture duration prior to fusion would

decrease the length of time required to generate tissue tubes, and lead to more seamless ring

fusion.

Human aortic SMC rings were removed from agarose molds after 3, 5, or 7 days of ring

pre-culture and cultured as tubes for 7 days, resulting in groups 3-7, 5-7, and 7-7, respectively

(shown schematically in Figure 5.1). Fusion was measured daily as the angle between adjacent

rings, to determine the time course for tissue fusion [30, 32] (Fig 5.2A). When human aortic

SMC tubes were fused, there was a significant difference in fusion angle only at day 2 between

the 7-7 group vs 5-7 group, and day 3 between the 5-7 group vs 7-7 and 3-7 groups (Fig 5.2B).

In all groups, the fusion angle appeared to plateau by day 3, with only slight increases after this

point. Tube length (Fig 5.2C) remained relatively constant over time, although tubes in the 3-7

group were significantly longer overall, and the 5-7 group was significantly shorter than the

other 2 groups. Significant differences were not observed in tube thickness (Fig 5.2D), although

tubes appear to thin slightly over time. These results are consistent with duplicate studies


performed both with human aortic

SMCs and human coronary artery

SMCs (Appendix F Figures F.1-

2).

5.3.2. Structure and morphology

of fused human SMC tubes

To evaluate fusion of

SMC ring units, tissue tube

sections were stained with

Hematoxylin and Eosin to

compare morphology of the 3-7,

5-7, and 7-7 tubes. Tubes

appeared well fused after a 7 day

fusion period, although ring

boundaries remained detectable in

all groups (Fig 5.3). Ring

boundaries are most distinct in the 7-7 group. Nearly seamless fusion was observed in the 3-7

group, although ridges at ring boundaries were still slightly visible on the tube exterior (Fig 5.3).

This suggests that rings pre-cultured for a shorter duration prior to tube fabrication may allow for

more complete tissue fusion.

5.3.3. Spatial positioning of SMCs within rings during fusion

To assess the feasibility of creating tubes with distinct tissue regions along the tube

length, we next evaluated whether cells within ring units maintain their spatial position along

tissue tubes after ring fusion. Three-day-old human aortic SMC rings were created from green or

red CellTracker-labeled cells. Alternating red and green fluorescently-labeled rings were fused in

culture for 7 days, and images were acquired daily. We did not observe “mixing” of cells at the

ring borders over the culture period (Fig 5.4A). This observation was confirmed when tubes

fused for 7 days were examined histologically and stained with Hoechst (Fig 5.4B-E). Similar

results were observed when the experiment was repeated with coronary artery SMCs (Appendix

F Fig F.3). Although some tissue compaction was visible, Hoechst-stained sections clearly show

Figure 5.2: Fusion kinetics of human SMC rings. Three human SMC

rings were threaded onto silicone tubing mandrels (A). The angle

between rings (ө), tube length (L), and thickness (T) were measured for

each sample on each day of culture (A). Fusion angles (B), tube length

(C) and thickness (D) as a function of time for tubes fabricated from

rings cultured for 3 (3-7), 5 (5-7) or 7 (7-7) days prior to 7 days of fusion

culture. N = 3 tubes per group. Data points are mean ± SD. # p<0.05 for

5-7 vs 3-7 and 7-7, ** p<0.05 for 5-7 vs 7-7, * p<0.05. Scale = 0.5mm.


that cells within rings maintain their original spatial position after tube fusion. Some decrease in

CellTracker signal was distinguishable at ring edges after fusion (Fig 5.4E). We hypothesized

that this was due to cellular proliferation at ring edges during fusion, which may dilute

CellTracker signal. To test this, rings loaded with only CellTracker red dye were fused, and were

fixed after either 1 or 2 days of fusion. Rings fused for only 1 day partially separated during

processing, indicating they were not fully fused. Ki67 staining of these sections shows

proliferating cells around the edges of individual rings (Fig 5.5A-B). After 2 days of fusion,

fewer proliferating cells are visible, and are predominately at the tube surfaces and not between

Figure 5.3: Histological assessment of human SMC tubes. H&E stained tissue tubes

comprised of rings pre-cultured for 3 (A-C), 5 (D-F), or 7 (G-I) days prior to fusion.

Low magnification longitudinal sections shown in (A, D, G). Higher magnification

views show one fusion point at the outer surfaces (solid box; B, E, H) and at the inner

surfaces (dashed box; C, F, I) of the tissue tubes. Lumen on bottom, scale bars = 250µm

(low magnification) or 100µm (high magnification). Images representative from n=3

samples/group. Sectioning schematic shown in lower right.


individual rings (Fig 5.5 C-D). This suggests that cell proliferation at the edges of the rings may

cause the decrease in CellTracker signal, and may play a role in initial ring fusion.

5.3.4. PCL cannulation cuffs and dynamic tube culture

The next step in generating an in vitro TEBV model is applying luminal flow, which is

critical for maintaining

blood vessel function

[33-35]. However, self-

assembled tissues can

be fragile at early time

points in culture, and

may not withstand

handling or suturing

forces necessary to load

the tissue into a flow

bioreactor. Thus, our

modular system for

vascular tissue

fabrication includes

electrospun PCL

Figure 5.5: Cell proliferation during fusion. Human aortic SMCs were pre-loaded

with red CellTracker dye prior to ring seeding. Rings were allowed to fuse for 1 (A,

B) or 2 (C, D) days. Tubes were then sectioned and stained for Ki67 to examine

proliferation. Green = Ki67, red = CellTracker Red, blue = nuclei. Scale = 100µm.

Images representative of n = 2 samples.

Figure 5.4: Spatial position of rings during fusion. Human aortic SMCs were pre-loaded with red or green

CellTracker dye prior to ring seeding. Rings with alternating dyes were then stacked and allowed to fuse for 7 days

(A). Tubes were then sectioned and stained with Hoechst dye. Red = CellTracker Red (B), green = CellTracker

Green (C), and blue = nuclei (D). Merged image shown in (E). Scale = 1mm (A) or 100µm (B-E). Lumen on bottom

(B-E). Images representative from n = 3 samples.


cannulation cuffs incorporated onto each end of the tube by cellular attachment and infiltration

from adjacent cell rings [28]. Previously, we incorporated PCL cuffs into tubes made from rat

aortic SMCs in static experiments [28]. Here, we assessed the feasibility of incorporating PCL

cannulation cuffs into human aortic SMC tubes to serve as reinforced extensions to aid in

cannulation and dynamic culture (Fig 5.6A). After 7 days of fusion (3 days of ring pre-culture),

we removed fused human tissue tubes from silicone tubing and mounted them onto a custom

bioreactor to demonstrate that tubes are strong enough to withstand luminal flow. Cannulation

cuffs fit snugly over bioreactor cannulas, and did not require additional suturing, as shown in

Figure 5.6B.

The bioreactor used in these studies was modified from Piola et al. [29]. An image of the

bioreactor with a cannulated SMC tube inside is shown in Figure 5.6C, and a schematic of the

bioreactor flow loop is shown in Figure 5.6D. Five tubes were successfully mounted onto

bioreactors and cultured for an additional 7 days (17 days total culture) under luminal flow (12

dyne/cm2). Tubes fixed for histology are shown in Figure 5.7, which demonstrates that tubes

remained intact and rings are fully fused. Ring boundaries are almost indistinguishable in both

static (on silicone mandrels) and dynamically cultured tubes. Tubes exposed to flow appeared to

have fewer cell nuclei on the luminal surface of the tube than static controls.

When this experiment was repeated, one tube out of six tore during loading, but the

remainder of the tubes were cultured successfully for 7 days. Picrosirius red/fast green and

Figure 5.6: PCL cannulation cuff incorporation for bioreactor culture. Electrospun PCL cuffs were

threaded onto silicone tubing and pushed into contact with cell rings at each end of the tube. Tubes were

cultured for 7 days on silicone mandrels (A) to achieve ring fusion, then mounted onto the cannulas in the

chamber of a custom luminal flow bioreactor (B). Image of bioreactor with SMC tube is shown in (C), and

a schematic of the medium flow loop is shown in (D). Scale = 1 cm.


orcein stains were used to

examine extracellular matrix

deposition of fused tubes

following static (Fig 5.8 A, B)

or dynamic (Fig 5.8 C, D)

culture. Collagen deposition is

visible throughout tubes (Fig

5.8 A, C). Elastic fibers were

not visible (Fig 5.8 B, D).

5.4. Discussion

The long-term goal of

these studies is to develop a

platform for fabricating 3D

human vascular tissue that may

potentially be used for in vitro

modeling of focal vascular

diseases such as intimal

hyperplasia or aneurysm. We

previously described a method

to rapidly generate vascular

tissue tubes from individual

self-assembled SMC ring units

[25, 26]. Here, we reduced the

total time needed to create

cohesive tissue tubes from self-

assembled rings, and utilized

the modular nature of this

system to create focal

heterogeneities within the tube

wall. Further, we incorporated

Figure 5.7: Histological images of tubes cultured in a luminal flow

bioreactor. Hematoxylin and Eosin stain of longitudinal section of tissue

tubes cultured as rings for 3 days, fused as tubes for 7 days, and then

cultured on silicone mandrels in static conditions (A, B) or with

approximately 12 dyne/cm2 shear stress (C, D) for an additional 7 days.

Lumen at bottom of image. Scale = 100µm.

Figure 5.8: Matrix deposition in fused tissue tubes. Longitudinal

sections of tubes cultured in static conditions for 14 days (A, B), or in

static conditions for 7 days followed by 7 days of dynamic culture with

approximately 12 dyne/cm2 of applied shear (C, D). Picrosirius red fast

green stain shown in (A, C; red = collagen, green = counterstain), orcein

stain shown in (C, D; dark pink = elastic fibers, purple = nuclei). Lumen

on bottom of image. Scale = 100 µm.


PCL cannulation cuffs [28] on each end, making the tubes amenable for cannulation and

dynamic culture within a custom designed luminal flow bioreactor. In future studies, this unique

modular platform may be modified to model a variety of vascular diseases.

The first goal of this study was to improve and accelerate tissue fusion. In our studies,

fusion is defined as an increase in fusion angle (angle between adjacent rings) to approximately

180ᴼ [30, 32]. While we did not observe significant differences in fusion rate with varying pre-

culture time, histological images suggest more complete fusion in the 3-7 group. This is

consistent with other reports, which suggest tissues fuse more rapidly and more completely when

pre-cultured for less time prior to fusion [30]. Others have suggested that tissues with increased

extracellular matrix (such as collagen) are generally more cohesive and difficult to remodel [36].

It is possible that increased matrix deposition at later time-points may be a reason why more

mature tissues fuse less completely than less mature tissues.

In addition to fusion angle, parameters such as thickness and length of constructs can be

used to assess fusion [30]. We did not observe significant changes in length over time, which is

contrary to previous reports of spheroid fusion [30, 37]. This may be due to differences in tissue

size or geometry, which are known to affect fusion [20, 30, 32], or due to differences in cell type.

Overall, 3-7 tubes were significantly longer than the 5-7 and 7-7 groups, and the 5-7 tubes were

significantly shorter. This may due to ring remodeling, compaction, and thinning over time

during ring-culture, resulting in the 5-7 and 7-7 groups being constructed from thinner rings.

A primary goal of varying pre-culture time was to determine a time course for ring

fusion, and develop cohesive tissue tubes in a minimal amount of time. In all experiments, fusion

angles plateaued after 3-4 days, which is consistent with other reports [20, 23, 32]. In

preliminary studies, we aimed to begin dynamic culture at this point, based on the observation

that tubes are fully fused. However, when tubes were fused for only 4 days, 66% of the tubes

tore during the cannulation procedure (data not shown). Thus, we increased fusion time to 7 days

to allow for increased ECM deposition, which improved our ability to cannulate tubes and

substantially reduced tube failure rates to an average of 8%. Importantly, this is still less time

than described in most other published reports, where engineered vascular tissue is typically

matured 2 weeks to several months in static culture prior to mounting on bioreactors for dynamic

culture [12, 38, 39], compared to the 3 days of ring pre-culture and 7 days of fusion culture used


in this study. This may be because cell-derived tissues can have enhanced ECM production and

tissue strength compared to tissue fabricated using degradable scaffold materials [40-44].

Additionally, PCL cannulation cuffs aided in handling and cannulation of tissue tubes at early

time points.

To further evaluate fusion, we examined if cells maintain spatial positioning within rings

during fusion. We observed that rings with red and green CellTracker dye are still spatially

distinct after 7 days of fusion. This result is consistent with previous reports examining fusion of

tissue sheets and spheroids, which showed that limited cellular migration or “mixing” is evident

between most fusing tissues, despite some tissue remodeling and compaction [20, 32, 45-48].

Because cells within ring units maintained their spatial positioning along tubes, we can

customize individual rings and place them in distinct regions of the tube prior to fusion. This

feature may enable us to model focal disease pathologies in future studies, by engineering

regions of tissue that contain a diseased cell phenotype in the middle of an otherwise healthy

vascular tissue tube. CellTracker dye signal was less visible around individual ring edges,

possibly due to the dye diluting as cells proliferate. We verified this by staining tubes for Ki67

after 1 or 2 days of fusion. Ki67 staining was visible predominantly around individual ring edges

after 1 day of fusion. This was also evident at day 2, although fewer cells were Ki67 positive,

likely due to contact inhibition of SMC proliferation. This suggests that proliferation may play a

role in tissue fusion.

Shear forces created from fluid flow are important for the progression of many vascular

diseases [49, 50]. Thus, it is critical to incorporate luminal flow during early culture of

engineered vascular disease models. However, self-assembled, scaffold-free tissues may be too

fragile at early time points in culture to be sutured onto cannulas for dynamic culture. The

modular nature of our system is conducive to adding biomaterial units on either end of the tissue

tube to serve as reinforced extensions, without affecting tissue structure. Previously, we

evaluated the incorporation of PCL cannulation cuffs with tubes fabricated from rat aortic SMCs

in static culture ([28], Chapter 4). Here, we applied this technology to human TEBV constructs,

enabling the successful cannulation of vascular tissue tubes in a custom bioreactor for dynamic

culture within 10 days of cell seeding. In these studies, human aortic SMC tubes remained intact

in a proof-of-concept experiment for 7 days of dynamic culture under physiologically relevant


wall shear stress. It was not surprising that fewer nuclei were observed on the luminal surface of

the dynamically cultured tube, as the endothelium typically prevents SMCs from being exposed

directly to shear forces. Others have reported that direct exposure to shear stress can trigger SMC

apoptosis [51]. Future studies will focus on establishing a functional endothelial layer by

developing a luminal cell seeding system for our custom bioreactor.

Additionally, the bioreactor chamber can also be easily modified to separate the luminal

vessel compartment from the external medium compartment [52]. In future studies, this will

allow endothelialization of cannulated tissue tubes, and enable us to flow vasoactive substances

through the tube lumen for endothelial and SMC functional testing. Alternatively, this bioreactor

can also be modified to apply cyclic stretch to tissue tubes [29] to enable mechanical

conditioning during tissue tube culture and maturation.

In studies with primary aortic SMCs, we did not observe contractile smooth muscle

markers such as smooth muscle alpha actin (Appendix F Fig F.4), or elastin deposition (Fig 5.8).

This is not surprising, since cells were cultured in a growth medium designed to support SMC

proliferation and ECM deposition consistent with a synthetic SMC phenotype. Creating a

healthy, contractile SMC phenotype will be critical for fabricating functional vascular tissue in

the future.

In future studies, we will use growth factor-loaded microspheres, different cell types, or

genetically-modified cells, in order to generate vessels with regions of cells that are

compositionally distinct from adjacent regions. For example, growth factor-loaded microspheres

may be able to create a hyper-proliferative region for modeling intimal hyperplasia. Our ability

to fabricate rings and tubes from a variety of SMC sources demonstrates the potential of this

system to model focal diseases that manifest in various types of vessels. We have shown that we

can produce rings using induced-pluripotent stem cell derived vascular SMCs (iPSC-VSMCs)

both from healthy patients (that produce elastin) and from patients with genetic disorders leading

to elastin deficiencies [13]. Using iPSC-VSMC rings to create a localized region of elastin

deficiency may allow us to model aneurysm. We have also applied this modular tube fabrication

system to create engineered cartilage rings and connective tissue rings with immature vascular

structures, and fused these rings to engineer tracheal tissue that anastomosed with host

vasculature upon subcutaneous implantation in a mouse model [47]. The controlled release of


growth factors via microsphere incorporation within individual rings may be ideal for such

multi-tissue tubular structures, where different tissue regions may require different biochemical

stimuli to maintain their differentiation and function. The ability to create high fidelity in vitro

human tissue models may provide an invaluable tool for high-throughput drug screening, and

potentially accelerate the development of new therapeutics.

In this study, we developed a modular system for fabricating vascular tissue tubes, which

may be modified in the future to model focal pathologies. Rings maintained their spatial position

within tubes, which may allow us to generate localized heterogeneities within tubes. After only 7

days of fusion, tubes were cohesive enough to be cannulated and cultured on a luminal flow

bioreactor. Overall, this work will serve as a platform technology for fabricating engineered

blood vessels with localized vascular diseases such as intimal hyperplasia, aneurysm, and

atherosclerosis. Such in vitro disease models may serve as tools for high throughput drug

screening, and accelerate the development of new treatments for vascular diseases.

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Chapter 6: Create vascular tissue tubes with spatially distinct regions 103

Chapter 6: Create vascular tissue tubes with spatially distinct

regions

Focal lesion experiment from: H. A. Strobel, T. A. Hookway, M. Piola, M. Soncini, G. B. Fiore,

E. Alsberg, and M. W. Rolle, “Assembly of tissue engineered blood vessels with spatially-

controlled heterogeneities.” Tissue Engineering Part A. In Press.







6.1. Introduction

While tissue engineered blood vessels (TEBVs) have enormous potential as tools for

disease modeling and drug screening, most existing approaches for TEBV fabrication create

homogenous tissue tubes [1-3]. These homogenous tubes may not be conducive to modeling

focal vascular diseases, such as intimal hyperplasia and aneurysm. To address this need, our lab

developed a modular system for fabricating tubes from individual ring subunits, which is more

conducive to introducing focal heterogeneities along the length of the tube. Our overall goal is to

develop this system into a platform technology for modeling focal vascular diseases, particularly

intimal hyperplasia (IH). In Chapter 3, we demonstrated our ability to incorporate degradable

cross-linked gelatin microspheres within tissue rings, and that these microspheres can be used to

customize cell phenotype [4]. In Chapter 5, we fused vascular tissue rings into cohesive tissue

tubes that were strong enough for dynamic culture after only 7 days of fusion with the aid of

electrospun cannulation cuffs. Here, we combine microsphere incorporation and tube fusion to

demonstrate that we can create TEBVs with focal heterogeneities along the length of our tissue

tubes (shown conceptually in Figure 6.1).

The first goal of this chapter was to create a focal region of microsphere incorporation, to

demonstrate our ability to create focal heterogeneities. To do this, we first evaluated the effect of

gelatin microspheres on ring fusion. We then tested whether modular building units comprised of

self-assembled primary human SMC rings (with or without incorporated gelatin microspheres)


can be fused together into a

contiguous and heterogeneous

tissue tube with distinct structural

regions.

Toward our goal of

creating an IH model, we next

performed preliminary

experiments evaluating the effect

of PDGF on human aortic smooth

muscle cells (SMCs). PDGF is

well established to increase SMC

proliferation and matrix

deposition and decrease

contractile protein expression, and

strongly associated with IH

initiation and progression [5-13].

Thus, we hypothesized that PDGF

would increase SMC proliferation and matrix deposition within self-assembled cell rings, and

consequently increase ring thickness. Ultimately, we aim to use these rings to create a focal

region of IH within tissue tubes. This Chapter describes the fabrication of tubes with spatially

controlled regions of microsphere incorporation, and the potential of gelatin microspheres for

PDGF delivery. Both of these are keys steps towards our ultimate goal of fabricating an in vitro

IH model.

6.2. Methods

6.2.1. Cell culture

Human aortic SMCs (Lifeline) were cultured according to manufacturer instructions, in

commercially available growth medium (Lifeline VascuLife SMC medium). Human coronary

artery SMCs (Lifeline) were used to repeat the focal region of microsphere incorporation

experiment, and were cultured according to the manufacturer’s instructions (Lifeline).

Figure 6.1: Fabrication of modular tissue tubes with focal

heterogeneities. Rings with incorporated microspheres are fused

between rings without microspheres, with PCL cuffs on either end.

The resulting construct is a fused tissue tube with focal region of

microsphere incorporation.


6.2.2. Ring fabrication

Rings were fabricated as previously described ([14], Chapter 3), with 400,000 human

aortic SMCs cells per well. After seeding, cells were allowed to aggregate 24 hours before

flooding wells with culture medium. Rings were cultured in Lifeline VascuLife growth medium

for the duration of culture unless otherwise specified.

Microspheres were incorporated within tissue rings as described in Chapter 3.2.3 and

3.2.4 [4]. Briefly, microspheres for non-growth factor experiments and unloaded control groups

were hydrated with 25µl per ml of PBS for 2 hours at 37ºC. Microspheres were then resuspended

thoroughly, combined with the cell suspension, and seeded with cells into agarose molds.

Microspheres to be loaded with PDGF were soaked in a solution of PBS containing 400

or 800 ng PDGF-BB (Peprotech) per mg microspheres (total volume of 25µl per mg), for 2 hours

at 37ºC prior to combining with cells. For fabrication of tubes with a focal lesion, and for the

PDGF release study, 0.6 mg microspheres per million cells were used (Appendix G, Batch 3).

When comparing fusion of rings with and without microsphere, 0.3 mg microspheres per million

cells was used (Appendix G, Batch 4). All microspheres were fabricated by the Alsberg Lab at

Case Western Reserve University. Diameters and cross-link densities for all batches used are

described in Appendix G.

When repeating the focal lesion experiment, coronary artery SMC rings were fabricated

as described above, with unloaded microspheres incorporated at 0.6mg per million cells. Cells

were loaded with CellTracker Red of CellTracker Green prior to ring seeding, as described in

Chapter 5.2.5. Rings with microspheres were seeded with red labelled cells, and rings without

microspheres were seeded with green labelled cells.

6.2.3. Tube fabrication for fusion comparison

Tubes were fabricated after 3 days of ring culture as described in Chapter 5.2.3. For

fusion studies, 3 rings with or without microspheres were used per tube. Brightfield images were

acquired daily, and fusion angles were calculated as described in Chapter 5.2.4.


6.2.4. Fabricating tubes with spatially defined regions of microsphere incorporation

Rings were fabricated from human aortic SMCs (Lifeline), with or without incorporated

microspheres ([4], Batch 3, Appendix G). Cells in rings with microspheres were pre-labelled

with CellTracker Red dye (Chapter 5.2.5). After 3 days of pre-culture, rings were threaded onto

silicone tubing mandrels with three microsphere-incorporated rings in the center of the tube, and

two outer regions with eight rings without microspheres per side (Fig 6.1). Cannulation cuffs

were placed adjacent to rings at each end of the tube. Tubes were cultured for 4 days on silicone

mandrels in static conditions prior to fixation and paraffin embedding. When the experiment was

repeated with coronary artery SMCs, the same procedure was followed, except rings without

microspheres were pre-labelled with CellTracker Green dye.

6.2.5. PDGF treatment of 2D cell cultures

Human aortic SMCs were seeded in wells of a 24 well plate at a density of 10,000 cells

per well in growth medium. After 24 hours, cells were switched to differentiation medium

containing a 1:1 ratio of DMEM and HAM’s F12 medium with 1% FBS, 1% L-glutamine, 1%

ITS, 1% penicillin-streptomycin, and 50µg/ml ascorbate. Four wells per group were

supplemented with PDGF-BB (Peprotech) at concentrations of 0, 1, 10, or 100 ng/ml. Cells were

fixed after 2 days of treatment for 15 minutes in 10% neutral buffered formalin (NBF) and stored

in PBS at 4ºC until staining.

6.2.6. PDGF treatment of self-assembled SMC rings

To evaluate the effects of PDGF on tissue rings, rings were seeded either with unloaded

microspheres, or with microspheres loaded with PDGF as described in section 6.2.2, with 400 or

800 ng PDGF/mg microspheres. Rings were seeded in growth medium. After 24 hours, medium

was switched to differentiation medium (described in section 6.2.4). Control rings with unloaded

microspheres were either cultured with or without the addition of 10 ng/ml exogenous PDGF.

Medium was changed daily. Rings were fixed at 3, 7, or 14 days.


Tissues were fixed for 1 hour in 10% NBF, processed, paraffin embedded, and cut into

5µm sections adhered to charged glass slides. A Hematoxylin and Eosin stain was used visualize


ring and tube morphology, and a Picrosirius Red/Fast Green stain was used to visualize collagen

deposition and gelatin microsphere degradation.

Sections to be stained for the contractile protein smooth muscle alpha actin were blocked

in 1.5% normal rabbit serum (NRS) for 45 minutes. Then, slides were incubated at 4ºC overnight

in primary anti-smooth muscle alpha actin antibody (Dako, 1:100 in 1.5% NRS). Samples were

then incubated at RT with rabbit anti-mouse AlexaFluor 488 secondary antibody (Invitrogen)

prior to counterstaining with Hoechst dye (1:6000 in DI water, 6 min). Sections of tubes

containing CellTracker were counterstained with Hoechst.

Sections stained for Ki67 were first subjected to antigen retrieval by boiling for 5 minutes

in Tris-EDTA buffer (see section 3.2.5) in a pressure cooker. Samples were blocked in 5%

normal goat serum (NGS) for 45 minutes, and incubated overnight in anti-Ki67 antibody

(Abcam, 1:100 in 3% NGS) at 4ºC. Negative controls were instead incubated with a rabbit IgG

protein (Vector). Slides were then incubated in secondary antibody (Invitrogen, AlexaFluor 488

goat anti-rabbit, 1:400 in 3% NGS) for 1 hour at RT prior to counterstaining with Hoechst.

Coverslips were adhered to slides with aqueous mounting medium, and images were taken using

an upright microscope (Leica DMLB2).

For 2D cell cultures, the same Ki67 staining procedure was followed, but cells were

permeabilized with 0.025% TritonX-100 instead of undergoing antigen retrieval. Samples were

imaged in wells using an inverted fluorescent microscope (Leica DMIL).


For fusion angle measurements, four measurements were obtained per tube, and averaged

to calculate one fusion angle per sample per time point. N = 3 samples were used for each group.

A Two-Way ANOVA test with Holm-Sidak post hoc analysis was used to determine statistically

significant differences between groups. When comparing Ki67 positive cells in response to

PDGF, the percentage of positive cells were averaged from two images per well to obtain one

measurement. Measurements from three wells per group were imaged to obtain N = 3 per group.

A One Way ANOVA test was used to calculate significance, with Holm-Sidak post-hoc analysis.

For all tests, P < 0.05 was considered significant.


6.3. Results

6.3.1. Effect of microspheres on tube fusion

To determine if microspheres would impact ring fusion, tubes were fabricated from rings

with or without microspheres and allowed to fuse for 7 days. Fusion angles (Fig 6.2 A) were

measured daily from phase contrast images (Fig 6.2 B). As shown in Figure 6.2 A, tubes with

microspheres have significantly lower fusion angles compared with tubes without microspheres.

This can also be seen in

Figure 6.2 B, where tubes

with microspheres still have

distinguishable ridges after

7 days, and rings without

microspheres do not. This is

likely due to the gelatin

microsphere material in the

ring composition

maintaining the original ring

shape.

Longitudinal sections were

stained for H&E to examine tube

morphology (Fig 6.3 A-B), and

Picrosirius Red/Fast Green to

examine collagen deposition and

presence of gelatin microspheres (Fig

6.3 C-D). Rings appear well fused in

both groups, and rings boundaries are

barely distinguishable. Collagen

Figure 6.2: Effect of microspheres on ring fusion. Fusion angles between

rings were measured daily for 7 days (A). Phase contrast images at day 0

compared to day 7 shown in (B). Scale = 1mm. Two-way ANOVA with Holm-

Sidak post hoc test (A). *P < 0.05. N = 3. Data is shown as mean ± SEM.

Figure 6.3: Fusion of rings with and without microspheres.

Hematoxylin and Eosin (A, B; H&E) and Picrosirius Red/Fast

Green (C, D; PRFG, red = collagen or gelatin, green =

counterstain) stains of tubes without (A, C) or with (B, D)

incorporated microspheres. Lumen on bottom. Scale = 100µm.

Asterisks (*) mark sample microspheres.


deposition is visible in both groups. Microsphere

degradation appears limited, as large amounts of

gelatin are still visible.

6.3.2. Fabrication of a focal region of microsphere

incorporation

An important step towards modeling focal

vascular diseases is the ability to create spatially

controlled heterogeneities within engineered vessel

walls. To do this, we incorporated degradable

gelatin microspheres within three rings (with

CellTracker Red dye) and positioned them in a

central region of the tube, between rings without

microspheres (8 per side, Fig 6.4 A). The region

with incorporated microspheres is clearly visible

due to

CellTracker Red dye and genipin cross-linked

microspheres, which both impart a purple hue to the

tissue in these regions (Fig 6.4 A). Histological

analysis demonstrated fully fused tubes with regions

of microsphere incorporation within a localized

region of the tube (Fig 6.4 B, C). This experiment

was repeated with human coronary artery SMCs.

With these cells, CellTracker Green dye was

incorporated into rings without microspheres, and

CellTracker Red dye was incorporated into rings

with microspheres, to demonstrate that cells within

each region maintain their spatial positioning along

the length of the tube during fusion (Fig 6.5)

Figure 6.5: Coronary artery SMC tubes with a

focal region of microsphere incorporation. A

central region of microsphere incorporation is

clearly visible after 7 days of fusion (A).

Fluorescent images of tubes with red and green

CellTracker show that cells within rings maintain

their spatial positioning during 7 days of fusion

(B, C), which can also be seen in Hoechst stained

sections (D). Green = CellTracker Green, red =

CellTracker Red, blue = Hoechst. Scale in mm

(A), bar = 1mm (B, C) or 100 µm (D).

Figure 6.4: Focal region of microsphere

incorporation. Rings with microspheres were

fused between rings without microspheres. Tube

after 4 days of fusion shown in (A). Hematoxylin

and Eosin stain of interface between rings with

and without microspheres shown in (B, C).

Sectioning schematic shown below figure. Lumen

on bottom. Scale in mm (A) or scale = 100µm (B,

C).


6.3.3. Effect of PDGF on proliferation of 2D SMC cultures

To create a focal disease model, we

aimed to use PDGF to locally stimulate

SMC proliferation. Prior to seeding rings

with PDGF-loaded microspheres, we first

needed to verify that human aortic SMCs

respond to PDGF in 2D cell cultures. After

2 days of PDGF treatment with varying

concentrations, 2D SMC cultures were

stained for Ki67 to evaluate the effect on

proliferation. We observed that both 10 and

100 ng/ml of exogenous PDGF significantly

increased the percentage of Ki67-positive

cells (Fig 6.6).

6.3.4. Effect of microsphere-mediated PDGF release on SMC rings

Next, we incorporated PDGF-loaded microspheres into SMC rings, to evaluate the effects

on proliferation. However, SMCs aggregated very loosely around gelatin microspheres, possibly

due to the larger size of microspheres in this batch (Batch 3, Appendix G). Microspheres in this

batch were also especially “clumpy” and difficult to break apart during resuspension. H&E

images are shown in Figure 6.7. Because there appeared to be such a large amount of

microspheres, ring edges were uneven, and ring thickness could not be reliably measured.

Despite this, rings could still be stained for Ki67 to examine cellular proliferation in response to

PDGF (Fig 6.8). It appeared at days 3 and 7 that groups treated with PDGF, either exogenously

or via microspheres, had a larger quantity of Ki67 positive cells. However, the number of Ki67

positive cells varied within and between samples of each group, making it challenging

toquantitatively assess trends in proliferating cells. Since most rings were loose in structure and

broken apart, it was not possible to quantify the total number of positive Ki67 cells per cross

sectional area. No clear differences were visible between groups at the 14 day time point. No

positive staining for the contractile protein smooth muscle alpha actin was visible in any groups

(Fig 6.9).

Figure 6.6: Effect of PDGF on 2D cell culture

proliferation. Cultures were treated with 0, 1, 10, or 100

ng/ml PDGF for 2 days. One Way ANOVA with Holm-

Sidak post hoc test. * P < 0.05 compared to 0 and 1 ng/ml.

Bars are mean ± SEM.


6.4. Discussion

This study demonstrates our ability to customize individual regions of vascular tissue

tubes through microsphere incorporation, which may ultimately enable us to create focal

vascular disease models. To do this, we first evaluated the effects of microsphere incorporation

on ring fusion. Fusion angles between stacked rings were measured daily for 7 days. When using

this method, complete fusion is defined as a fusion angle of 180º [15, 16]. We observed

significantly lower fusion angles in tubes with incorporated microspheres than in tubes without

microspheres. The reduced fusion angles are likely due to the non-degraded gelatin microspheres

maintaining the original ridges in the tissue, which are clearly visible in brightfield images (Fig

6.2 B). Despite this, histological sections showed nearly seamless fusion in both groups,

indicating that fusion angle measurements may not be the most effective means for measuring

fusion in this case. While microspheres did reduce fusion angles, they do not appear to have

affected overall tube fusion and cohesivity.

Figure 6.7: Morphology of PDGF treated rings. Rings were fixed at 3 (A-D), 7 (E-H), or 14 (I-L) days. Rings

contained unloaded microspheres were treated with no PDGF (A, E, I) or 10 ng/ml exogenous PDGF (B, F, J), or

contained microspheres loaded with 400 (C, G, K) or 800 (D, H, L) ng of PDGF per mg of microspheres.

Hematoxylin and eosin stain. Scale = 100µm


We reported previously that incorporated gelatin microspheres can be used to locally

deliver growth factors within SMC rings, for the purpose of controlling SMC phenotype ([4],

Chapter 3). The gelatin microspheres degrade within approximately 2 weeks, and do not

adversely affect ring mechanical strength ([4], Chapter 3). Here, we demonstrated that rings

containing microspheres can be localized to a central region of the tissue tubes, and successfully

Figure 6.9: Contractile protein expression in SMC rings. Rings either contained unloaded microspheres and were

treated with no PDGF (A) or 10 ng/ml exogenous PDGF (B), or contained microspheres loaded with 400 (C) or 800

(D) ng/ml PDGF. Rings were cultured for 14 days. Green = smooth muscle alpha actin, blue = Hoechst. Scale =

100µm.

Figure 6.8: Ki67 staining of rings with PDGF treatment. Rings were fixed at 3 (A-D), 7 (E-H) or 14 (I-L)

days. Rings were treated with unloaded microspheres and no PDGF (A, E, I) or exogenous 10 ng/ml PDGF (B,

F, J), or contained microspheres loaded with 400 or 800 ng PDGF per mg microsphere. Green = Ki67, blue =

nuclei. Scale = 100µm.


fuse with unmodified rings, to create a focal heterogeneity (Fig 6.4). This ability to create focal

changes is a unique attribute of our system, as other methods for fabricating self-assembled

TEBVs only create homogenous tubes.

Our overall goal is to use these microspheres to deliver growth factors to a central region

of the tube, as shown schematically in Figure 6.10. We predict that growth factors delivered from

microspheres will stimulate the formation of an intimal lesion by locally stimulating SMC

proliferation and matrix deposition. Towards our goal

of creating an IH model, we performed preliminary

experiments examining the effect of PDGF on human

aortic SMCs. PDGF is a potent SMC mitogen, that is

known to strongly contribute to IH initiation and

progression [5-13]. We observed that in 2D culture,

SMC proliferation significantly increased in response

to PDGF. PDGF also appeared to increase the number of Ki67-positive cells in 3D tissue rings.

However, some variation was visible within groups, making it challenging to quantitatively

assess trends in number of positive cells. Additionally, rings were very loosely formed in this

experiment and broke apart during histological analysis, preventing quantification of cross

sectional area to calculate the number of positive cells per unit area. For this experiment, we

switched to Batch 3 of microspheres (Appendix G), which are larger in size and tended to clump

more than previous batches. Additionally, human aortic SMCs are smaller in size than the

coronary artery SMCs used in Chapter 3. Thus, the concentration of 0.6 mg microspheres per

million cells used in earlier experiments (Chapter 3) may be too high. A lower concentration of

microspheres was used in subsequent experiments (Chapter 8). Though PDGF appeared to have

some effect on proliferation in 3D, we also observed an overall lack of contractile proteins such

as smooth muscle alpha actin, even in control groups with no PDGF. IH lesions are characterized

by a de-differentiation of SMCs. Without contractile protein expression in control, “normal”

rings without PDGF that will model healthy regions of the blood vessel, it will not be possible to

discern PDGF-mediated effects on SMC phenotype in 3D. Based on the lack of contractile

protein expression in primary human SMC lots we used to fabricate SMC rings, we decided to

re-evaluate our SMC cell source in subsequent experiments (discussed further in Chapters 7 and

8).

Figure 6.10: Schematic of future IH model.

Growth factor loaded-microspheres (purple

dots with yellow) are fused between regions

with unloaded microspheres (purple dots) to

create a focal region of growth factor delivery

and stenosis.


Future work may also include optimization of PDGF release from microspheres. When

IH occurs in vivo, SMC proliferation can begin as early as 24 hours after injury, and migration

can begin in as soon as 4 days [17]. Significant reductions in lumen area can be seen within 4-6

weeks of the initial injury [18, 19]. Thus, PDGF should be delivered from microspheres for a

minimum of 4 weeks to create a physiologically relevant IH model. The microspheres used in

this study are clearly visible at day 14, which is longer than we previously reported ([4], Chapter

3). This suggests we may have a more sustained growth factor release, but this must be verified

by determining the PDGF release curve over a minimum of 4 weeks. If needed, microsphere

cross-link density may be increased to further prolong growth factor delivery [20].

Overall, we demonstrated that we can create a focal region of microsphere incorporation

within vascular tissue tubes, that may be further developed to create an intimal lesion. This

platform technology could be modified to model other vascular diseases as well, such as

aneurysm. The controlled release of growth factors via microsphere incorporation within

individual rings may also be conducive to fabricating multi-tissue tubular structures, such as

trachea [21], where different tissue regions may require different biochemical stimuli to maintain

their differentiation and function.

6.5. References

1. Atchison, L., H. Zhang, K. Cao, and G.A. Truskey, A Tissue Engineered Blood Vessel

Model of Hutchinson-Gilford Progeria Syndrome Using Human iPSC-derived Smooth

Muscle Cells. Sci Rep, 2017. 7(1): p. 8168.














1223-1230.





growth factor-BB transactivates the fibroblast growth factor receptor to induce

proliferation in human smooth muscle cells. Trends Cardiovasc Med, 2006. 16(1): p. 25-

8.









D. Bergqvist, A. Ostman, and C.-H. Heldin, Intimal Hyperplasia Recurs After Removal of

PDGF-AB and -BB Inhibition in the Rat Carotid Artery Injury Model. Arterioscler


















41.






21(2): p. 101-9.

19. Chesebro, J. and V. Fuster, Platelet-inhibitor drugs before and after coronary artery

bypass surgery and coronary angioplasty: the basis of their use, data from animal

studies, clinical trial data, and current recommendations. Cardiology, 1986. 73: p. 292-

305.




158(2): p. 224-32.




Chapter 7: Induced pluripotent stem cells as an alternative human smooth muscle cell source for vascular tissue

engineering 117

Chapter 7: Induced pluripotent stem cells as an alternative

human smooth muscle cell source for vascular tissue engineering

7.1. Introduction

Cell sourcing is a critical issue in vascular tissue engineering, as there are several

challenges to working with primary human smooth muscle cells (SMCs). Primary SMCs are

difficult to obtain and are limited in supply due to the need to source them from human arteries

and veins. Primary SMCs also have a limited proliferative capacity, and do not readily

differentiate or may lose contractile protein expression in culture. In our experience, donor-to-

donor variation, the limited number of vials from each donor lot, and the low availability of

SMCs from human subjects younger than 30 are also problematic. A unique challenge for tissue

rings studies is that SMC differentiation and proliferation in 2D culture does not predict

successful self-assembly and ring formation, or SMC differentiation within tissue rings in 3D.

We found in Chapters 5 and 6 that certain lots of human SMCs do not express contractile

proteins in 3D culture. This observation motivated our lab to explore alternative sources of

human vascular SMCs for vascular tissue engineering.

Induced pluripotent stem cells (iPSCs) have been used as a cell source in vascular tissue

engineering [1-5]. iPSCs are reprogrammed from fibroblasts or other cell types into a pluripotent

stem cell state. Then, they are differentiated into iPSC-derived vascular SMCs (iPSC-vSMCs)

[6-9]. This method is advantageous because it may allow for the development of patient-specific

TEBVs, which may be used as implantable grafts [2], or as disease models [1, 10-12]. Protocols

for culturing and differentiating iPSCs have progressed in recent years, allowing for the

production of large numbers of iPSC-vSMCs with high purities [10]. Their high proliferative

capacity makes them ideal for vascular tissue engineering, which often requires large cell

numbers.

In collaboration with the Qyang lab at Yale University, the Rolle lab published a study in

which tissue rings were fabricated from iPSC-vSMCs using the same mold system described in

this dissertation [1]. We successfully fabricated functional tissue rings that express contractile

proteins, including the late-stage differentiation marker myosin heavy chain, and can contract in


engineering 118

response to vasoactive substances. Rings were also fabricated from iPSC-vSMCs obtained from

a patient with supravalvular aortic stenosis (SVAS). SMCs from these patients are known to be

less contractile than cells from healthy patients [10]. In a direct comparison, rings fabricated

from SVAS patients contracted significantly less when stimulated with carbachol and had a

higher percentage of proliferating cells than rings fabricated with cells from healthy patients [1].

This suggests that iPSC-vSMCs have great potential as tools for creating patient-specific disease

models. Additionally, iPSC-vSMCs successfully formed vascular tissue rings, which is

promising for our work fabricating ring-based tissue tubes.

Based on these promising results, we decided to evaluate the potential of iPSC-vSMCs to

create tissue tubes for focal disease modeling using our ring-tube system. Here, we describe

preliminary studies examining ring mechanical properties, fusion kinetics, and microsphere-

mediated growth factor delivery.

7.2. Methods

7.2.1. Ring culture

iPSC-vSMCs were provided by the Qyang lab at Yale University, where they were

differentiated according to their established protocol [1]. Cells were then expanded using

commercially available complete growth medium (Lonza) on Matrigel- (Corning) coated plates.

Rings were seeded as described previously, with 600,000 cells per ring [1]. After aggregating

overnight, rings were switched to a custom “Ring Medium” developed by the Qyang lab for ring

culture, containing DMEM (Corning), 20% FBS (Thermo Fisher), 20 µg/ml L-Alanine (Sigma),

50 µg/ml L-Proline (Sigma), 50 µg/ml Glycine (Sigma), 50 µg/ml Ascorbate (Wako), 3 ng/ml

CuSO4 (Sigma), 1 ng/ml TGF-β1 (Peprotech), 10 ng/ml PDGF-BB (Peprotech), and 1% Pen-

strep (Corning) [1]. Medium was changed daily for the duration of culture. After 7 or 14 days,

rings were fixed for histological analysis. At 14 days, additional rings were used for mechanical

testing.

7.2.2. Tube culture

Cells were labelled with CellTracker Red or Green dye (Life Technologies) as described

in Chapter 5.2.5. Rings were threaded over silicone tubing with alternating red and green colors.


engineering 119

Both fluorescent and brightfield images were taken daily for 7 days. Fusion angles were

measured from phase contrast images as described in Chapter 5.2.4.

7.2.3. iPSC-vSMC response to PDGF in 2D

iPSC-vSMCs were plated in a Matrigel-coated 24 well plate in growth medium (Lonza)

and allowed to attach for 24 hours. Wells were then switched to differentiation medium

containing a 1:1 ratio of DMEM:HAM’s F12 with 1% FBS, 1% l-glutamine, 1% ITS, 1%

penicillin-streptomycin, and 50 µg/ml ascorbate. Wells were supplemented with 0, 1, 10, or 100

ng/ml PDGF-BB (Peprotech).

After 2 days, a Click-iT EdU AlexaFluor 488 Kit (Invitrogen) was used to evaluate

cellular proliferation. EdU reagent was diluted in complete culture medium (with supplemented

PDGF) to a concentration of 20µM. Half of the culture medium was removed from each well and

replaced with the fresh medium containing EdU reagent. Cells were incubated for 6 hours, then

fixed for 15 minutes in 10% NBF at RT. Samples were permeabilized in 0.5% Triton X-100 for

20 minutes, and then incubated for 30 minutes in the EdU Click-iT reaction buffer, which was

prepared according to manufacturer instructions. A Hoechst 3342 (Invitrogen) counterstain was

used to visualize cell nuclei (1:6000 in DI water for 6 minutes). Images were taken with an

inverted microscope (Leica DMIL) and total number of nuclei and EdU positive nuclei were

counted per image. Percentage of EdU positive cells was counted from two images per well, to

get a value for each well. Averages were then taken from three wells per group.

7.2.4. iPSC-vSMC response to PDGF in 3D

UV-sterilized microspheres were soaked in a solution of 400 ng PDGF-BB (Peprotech)

per mg microspheres (25 µl growth factor solution per mg microsphere) for 2 hours at 37 ºC.

Control unloaded microspheres were soaked in phosphate-buffered saline (PBS) at a

concentration of 25 µl per mg microspheres. Rings were seeded with 600,000 iPSC-vSMCs per

ring and 0.6 mg microspheres per million cells. Rings were seeded in growth medium, and then

switched to the differentiation medium described in section 7.2.3 after 24 hours. Medium was

then changed daily. After 14 days, rings were fixed for histological analysis or flash frozen for

biochemical analysis.


engineering 120

7.2.5. Mechanical testing

Rings were imaged and mechanically tested at day 14 as described in sections 3.2.6 and

3.2.7. Briefly, thicknesses were calculated from images taken with a machine vision system

(model 630; DVT Corporation). Rings were then mounted onto a tensile testing apparatus

(ElectroPuls E1000; Instron), subjected to 8 pre-cycles and then pulled to failure at 10 mm/min.

7.2.6. Histological analysis and immunohistochemistry

At 7 and 14 days rings were fixed for 1 hour in 10% neutral buffered formalin (NBF),

processed, and paraffin embedded. Sections with a thickness of 5µm were adhered to charged

slides. H&E and Picrosirius red/fast green staining were used to visualize ring morphology and

collagen deposition. IHC for smooth muscle alpha actin and calponin was performed as

described in section 3.2.5.

7.2.7. Western blotting

Western blotting was performed on frozen 14 day-old samples as described in section

3.2.8, to quantify smooth muscle alpha actin expression. Three samples per group were tested,

except in the group without microspheres or PDGF, which had only one sample due to several

ring failures during culture.

7.2.8. Statistics

Statistics were performed on the number of proliferating cells in the 2D EdU

quantification experiment. A One Way ANOVA with Dunn’s post hoc analysis was used to

determine statistical differences between groups. Measurements were taken from 2 images per

well, which were averaged into one measurement per well. Three wells per group were used to

calculate the group average. A P value less than 0.05 was considered significant.

7.3. Results

7.3.1. Ring formation and characterization

iPSC-vSMCs successfully self-assembled into vascular tissue rings. By day 14,

approximately half of rings were uniform in thickness, while half had experienced thinning on

one side of the ring (Fig 7.1). Despite this, rings fabricated from iPSC-vSMCs had an average


engineering 121

ultimate tensile stress of 1823.3 ± 359.0 kPa, which is

higher than we have previously reported for human

coronary artery SMCs (871.0 ± 251.9 kPa) [13]. iPSC-

vSMC rings also had a higher failure load, failure strain,

and maximum tangent modulus, and a lower thickness than

previously tested SMC rings (Table 7.1). Histological

analysis of fixed rings at 7 and 14 days showed large

amounts of collagen visible at both 7 and 14 days (Fig 7.2).

7.3.2. iPSC-vSMC

response to PDGF

To determine if

PDGF will increase

iPSC-vSMC

proliferation, 2D

cultures were treated with different

concentrations of PDGF, and an EdU

incorporation assay was performed to

measure the percentage of cells in S-

phase (as an indicator of proliferation).

Cells treated with PDGF had visibly

more EdU-positive nuclei (Fig 7.3),

indicating increased proliferation. Cells

treated with 10 or 100 ng/ml PDGF had

significantly higher percentages of EdU

positive cells than cells without PDGF

treatment, and the 10 ng/ml group was

also significantly higher than cells

treated with 1 ng/ml PDGF (Fig 7.3E).

We evaluated the effect of PDGF

on contractile protein expression of

14 days

H&

E

PR

FG

7 days

Figure 7.2: iPSC-vSMC ring morphology and collagen

deposition. Hematoxylin and Eosin stain (A, B; purple =

nuclei, pink = counterstain) and Picrosirius Red/Fast Green

stain (C, D; red = collagen, green = counterstain) of 7 (A, C)

and 14 (B, D) day rings. Lumen on bottom/left. Scale =

100µm

A B

C D

Figure 7.1: Images of 14 day iPSC-

vSMC rings. Approximately half of rings

were uniform in thickness (A), while half

of rings displayed visible necking (B).

Scale = 1mm.

Table 7.1: Mechanical characterization of iPSC-vSMC rings, compared to

previously published primary cell rings.


engineering 122

iPSC-vSMC in 3D by

immunohistochemical

analysis of tissue rings.

IHC suggests decreased

expression of smooth

muscle alpha actin in

rings treated with 10ng/ml

exogenous PDGF, or

rings with incorporated

PDGF loaded MS (Fig

7.4). Smooth muscle α-

actin expression in iPSC-

vSMC rings was also

measured with western blotting (Fig 7.5), although sample sizes were not high enough to

perform statistical analysis.

7.3.3. Tube fabrication

Next, we tested the potential of iPSC-vSMC rings to form modular tissue tubes. To do

this, we stacked rings together on silicone tubing mandrels, measured fusion angles over time,

and used CellTracker dye to evaluate spatial positioning of cells within rings along the length of

the tube (using methods described in Chapter 5, sections 5.2.4 – 5.2.5). Red- and green-labelled

No MS

No PDGF

No MS

+ PDGF

+ MS

No PDGF

+ MS

+ PDGF

+ PDGF

Loaded MS

SM

A

Ca

lp

Figure 7.4: Effect of PDGF on iPSC-vSMC ring contractile protein expression. Rings were stained for either

smooth muscle alpha actin (SMA; A-E) or calponin (Calp; F-J). Green = SMA or calp, blue = nuclei. Scale =

100µm. Lumen on left.

A B C D

F H G I

E

J

Figure 7.3: Effect of PDGF on 2D

iPSC-vSMC cultures. iPSC-vSMCs

treated with 0 (A), 1 (B), 10 (C), or

100 (D) ng/ml PDGF-BB. Total

percent of proliferating cells shown in

(D). Red = proliferating cell, Blue =

Hoechst, scale = 100µm. Percent

positive cells shown in (E). *P<0.05,

One Way ANOVA on Ranks with

Dunn’s Post Hoc test. Bars are mean ±

SD. N=3.


engineering 123

cells appeared to largely maintain their spatial

positioning along the length of the tube over

the 7 day fusion period (Fig 7.6 A). This is

also visible with Hoechst staining of a

longitudinal tube section (Fig 7.6 B). Fusion

angle measurements greatly increase between

day 0 and day 1, with smaller increases

observed between days 2 and 7 of fusion

culture (Fig 7.7).

7.4. Discussion

iPSC-vSMCs have potential for use in

vascular tissue engineering, due to their high

proliferative capacity and because cells can be

obtained from living patients, enabling the

fabrication of patient-specific TEBVs. Here, we presented preliminary data demonstrating the

ability of iPSC-vSMCs to be used for vascular tissue ring and tube formation. In our initial

testing, we observed that approximately 50% of rings were uniform in thickness, although on

average, iPSC-vSMCs still had a higher mechanical

strength than previously tested human coronary artery

smooth muscle rings. This may be due to their dense

collagen deposition, which is visible in Figure 7.3.

Mechanical strength is critical, as it enables us to

Figure 7.7: Fusion rate of iPSC-vSMC rings. Fusion angles

are measured daily. Schematic of fusion angle measurement

shown on left. N = 2 tubes (5-8 measurements per tube).

Figure 7.6: Fusion of iPSC-vSMC rings. Ring

fusion between day 0 and 7 (A). Longitudinal

cross-section shown in (B). Green =

CellTracker Green, red = CellTracker Red, blue

= nuclei. Scale = 1mm (A) or 100µm (B).

Lumen on bottom in (B).

Figure 7.5: Effect of PDGF on ring smooth muscle

alpha actin expression. Smooth muscle alpha actin was

normalized to histone protein (as a loading control).

Lanes labeled “L” indicates microspheres loaded with

PDGF. N = 1-3

+


engineering 124

handle rings at early time points, and potentially subject tubes to dynamic culture. Still, having

so many rings with non-uniform thickness may make tube fabrication challenging in future

experiments.

Our overall goal is to fabricate an in vitro intimal hyperplasia (IH) model, using

customized “diseased” rings, fused between healthy contractile rings. Focal regions of growth

factor-loaded microsphere incorporation may allow us to locally stimulate SMC hyper-

proliferation and de-differentiation within a tissue tube. PDGF is strongly associated with IH

initiation and progression, and is known to increase SMC proliferation and decrease contractile

protein expression [14-22]. Because of this, PDGF was proposed as the model growth factor for

stimulating SMC growth within vascular tissue tubes. Here, we first evaluated if iPSC-vSMCs

proliferate in response to PDGF in 2D cell cultures. We observed significant increases in

proliferating cells at concentrations of 10 ng/ml and 100 ng/ml, suggesting the potential of PDGF

for stimulating IH formation in vitro.

Next, we evaluated the effects of PDGF on 3D vascular tissue rings, both by exogenous

and microsphere-mediated delivery. Staining for smooth muscle alpha actin and calponin

suggested PDGF caused decreases in contractile protein expression, regardless of delivery

mechanism (Fig 7.4, 7.5). This also indicates that gelatin microspheres are an effective means to

deliver bioactive PDGF within tissue rings. Decreased contractile protein is a key indicator of

diseased SMCs. Thus, it is promising for our IH model that microsphere-mediated PDGF

delivery successfully decreased contractile protein expression in iPSC-vSMC rings.

Microsphere-mediated delivery may enable us to spatially control PDGF delivery long the length

of fused tissue tubes, in order to create a focal lesion.

Our next goal was to evaluate iPSC-vSMC ring fusion. To create an intimal lesion, we

must be able to place microsphere-loaded or disease phenotype rings within a focal region of the

tissue tube, to ensure that diseased cells stay in the diseased region of the tube. To verify that

cells retain their spatial positioning, we fused rings fabricated from red and green-labelled cells.

Although tubes did appear to contract and thin as they remodeled, fluorescently-labelled cells

largely maintained their positioning within original rings. This is consistent with our evaluation

of other cell types (Chapter 5), and work published by others on tissue fusion [23-28]. We also

evaluated fusion rate by measuring fusion angle between iPSC-vSMC rings. Fusion angles


engineering 125

showed the largest increase between day 0 and 1, followed by a slow incline to approximately

180º. This is consistent with our studies on primary SMCs, which indicated fusion was largely

complete within the first few days of fusion culture, although the initial increase in fusion angles

was more rapid with iPSC-vSMC rings. It is possible that this is due to increased proliferation

early in fusion culture compared to primary cells, as iPSC-vSMCs are highly proliferative. We

have observed that proliferation plays a role in tissue fusion (Chapter 5), as have others [23].

In summary, iPSC-vSMCs are a promising cell type for modular tissue tube assembly

from cell ring subunits, and creation of model intimal hyperplastic regions within human

TEBVs. iPSC-vSMCs self-assembled to form rings, expressed contractile proteins in 3D ring

culture, and proliferated and de-differentiated in response to PDGF delivered from incorporated

gelatin microspheres. The ability to create patient-specific disease models from fibroblast cells,

which are highly accessible compared to primary vascular SMCs, may be highly beneficial.

However, we observed significant batch-to-batch variation in the cells’ ability to form rings and

differentiate into SMCs that express contractile proteins in subsequent batches of iPSC-vSMCs

we received. This highlights the challenges associated with differentiating iPSCs reliably and

repeatedly in 3D cultures for fabrication of engineered tissues. In the future, more optimization

of iPSC differentiation protocols, and development of additional resources in the Rolle lab to

support iPSC culture and differentiation may allow us to continue utilizing iPSC-vSMCs to build

TEBVs in future studies.

Due to these challenges, we elected to explore an additional alternative source of human

SMCs, mesenchymal stem cells, which is discussed in Chapter 8.

7.5. References




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Dardik, G. Tellides, et al., Implantable tissue-engineered blood vessels from human

induced pluripotent stem cells. Biomaterials, 2016. 102: p. 120-129.

3. Eoh, J.H., N. Shen, J.A. Burke, S. Hinderer, Z. Xia, K. Schenke-Layland, and S. Gerecht,

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engineering 126

4. Wang, Y., J. Hu, J. Jiao, Z. Liu, Z. Zhou, C. Zhao, L.J. Chang, Y.E. Chen, P.X. Ma, and

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5. Sundaram, S., J. One, J. Siewert, S. Teodosescu, L. Zhao, S. Dimitrievska, H. Qian, A.H.

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6. Lee, T.H., S.H. Song, K.L. Kim, J.Y. Yi, G.H. Shin, J.Y. Kim, J. Kim, Y.M. Han, S.H.

Lee, S.H. Lee, et al., Functional recapitulation of smooth muscle cells via induced pluripotent stem cells from human aortic smooth muscle cells. Circ Res, 2010. 106(1): p.

120-8.

7. Takahashi, K. and S. Yamanaka, Induction of pluripotent stem cells from mouse

embryonic and adult fibroblast cultures by defined factors. Cell, 2006. 126(4): p. 663-76.

8. Bajpai, V.K., P. Mistriotis, Y.H. Loh, G.Q. Daley, and S.T. Andreadis, Functional

vascular smooth muscle cells derived from human induced pluripotent stem cells via

mesenchymal stem cell intermediates. Cardiovasc Res, 2012. 96(3): p. 391-400.

9. Patsch, C., L. Challet-Meylan, E.C. Thoma, E. Urich, T. Heckel, J.F. O'Sullivan, S.J.

Grainger, F.G. Kapp, L. Sun, K. Christensen, et al., Generation of vascular endothelial

and smooth muscle cells from human pluripotent stem cells. Nat Cell Biol, 2015. 17(8): p.

994-1003.

10. Ge, X., Y. Ren, O. Bartulos, M.Y. Lee, Z. Yue, K.Y. Kim, W. Li, P.J. Amos, E.C.

Bozkulak, A. Iyer, et al., Modeling supravalvular aortic stenosis syndrome with human

induced pluripotent stem cells. Circulation, 2012. 126(14): p. 1695-704.

11. Atchison, L., H. Zhang, K. Cao, and G.A. Truskey, A Tissue Engineered Blood Vessel

Model of Hutchinson-Gilford Progeria Syndrome Using Human iPSC-derived Smooth

Muscle Cells. Sci Rep, 2017. 7(1): p. 8168.

12. Granata, A., F. Serrano, W.G. Bernard, M. McNamara, L. Low, P. Sastry, and S. Sinha,

An iPSC-derived vascular model of Marfan syndrome identifies key mediators of smooth

muscle cell death. Nat Genet, 2017. 49(1): p. 97-109.





positively and negatively regulate interstitial collagen gene expression in human vascular smooth muscle cells. Arteriosclerosis, Thrombosis, and Vascular Biology, 1991. 11(5): p.

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8.

17. Jiang, B., S. Yamamura, P.R. Nelson, L. Mureebe, and K.C. Kent, Differential effects of platelet-derived growth factor isotypes on human smooth muscle cell proliferation and





















24. Sasagawa, T., T. Shimizu, S. Sekiya, Y. Haraguchi, M. Yamato, Y. Sawa, and T. Okano,

Design of prevascularized three-dimensional cell-dense tissues using a cell sheet stacking

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26. Shimizu, T., M. Yamato, T. Akutsu, T. Shibata, Y. Isoi, A. Kikuchi, M. Umezu, and T.

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engineering 128




28. Solorio, L.D., L.M. Phillips, A. McMillan, C.W. Cheng, P.N. Dang, J.E. Samorezov, X.

Yu, W.L. Murphy, and E. Alsberg, Spatially organized differentiation of mesenchymal

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Chapter 8: Growth factor delivery for phenotypic modulation of human mesenchymal stem cell rings 129

Chapter 8: Growth factor delivery for phenotypic modulation of

human mesenchymal stem cell rings

8.1. Introduction

The overall goal of this thesis was to develop a platform technology that could be used

for modeling focal vascular diseases such as intimal hyperplasia (IH). Before we can stimulate

the formation of an IH lesion, we must also have healthy, functional SMC rings. As discussed in

Chapter 7, primary SMCs are well-known to be challenging to differentiate, especially in 3D

culture. We observed previously that even with transforming growth factor-beta one (TGF-β1)

treatment, which is well-established to stimulate SMC differentiation to a contractile phenotype

[1, 2], our human aortic SMCs did not differentiate in 3D ring culture (not shown). Additionally,

primary SMCs have a limited proliferative capacity, and substantial lot-to-lot variability. Induced

pluripotent stem cells (iPSCs) have potential as source of differentiated SMCs, but as discussed

in Chapter 7, lot-to-lot variation is still poses a substantial challenge. For these reasons, we

decided to further evaluate alternative cell sources for fabrication of an in vitro IH model.

Human mesenchymal stem cells (hMSCs) are highly proliferative and can be

differentiated into contractile SMCs in the presence of stimuli such as TGF-β1 and BMP-4 [3-5].

hMSCs can also be collected from living patients, from either bone marrow or adipose tissue.

This may ultimately allow for the fabrication of patient-specific tissues for disease modeling and

drug testing. Others have also utilized hMSCs for TEBV fabrication, as an alternative to SMCs

[3, 6-8]. Because of these advantages, we decided to use hMSCs instead of primary SMCs for

the next experiments towards our goal of fabricating an in vitro IH model.

IH is a complex disease triggered by arterial injury. This injury causes numerous

cytokines to be released, including platelet derived growth factor (PDGF), fibroblast growth

factor (FGF), interleukins 1 and 6 (IL-1, IL-6), TGF-β1, and many other signaling factors [9, 10].

Upon arterial injury, the inflammatory response is activated, further contributing to the release of

cytokines. These growth factors interact in complex ways to cause SMC proliferation, migration

to the lumen, and deposition of extracellular matrix proteins. Our goal is to use degradable

gelatin microspheres to deliver growth factors to regulate SMC phenotype, thus creating regions


of healthy and diseased tissue. By fusing rings with growth factor-loaded microspheres between

rings with unloaded microspheres, we aimed to focally stimulate SMC proliferation and de-

differentiation.

PDGF-BB (PDGF) is released from activated platelets following injury, and is a potent

SMC mitogen that is well-established to trigger SMC migration, proliferation, collagen

deposition, and decrease contractile protein expression [11-

19]. Thus, our original hypothesis was that PDGF would

increase ring thickness and decrease ring contractile protein

expression in SMC ring units (shown schematically in Figure

8.1). Additionally, we evaluated the effect of FGF delivery,

another potent SMC mitogen [13, 17, 20]. Then, we evaluated

TGF-β1 delivery to selectively differentiate rings, rather than

selectively de-differentiate them.

8.2. Methods

8.2.1. Cell culture

Bone marrow-derived hMSCs were provided by RoosterBio. Cells were cultured

according to manufacturer instructions, in a proprietary growth medium (RoosterBio). hMSCs

were used for growth factor release studies and for fabricating the growth factor-induced focal

lesion.

8.2.2. Ring culture

Rings fabricated from hMSCs were seeded as described previously [21], but with

600,000 cells per ring. Culture conditions varied between experiments as medium conditions

were optimized. Microspheres were incorporated at a concentration of 0.3 mg microspheres per

106 cells. In the PDGF release experiment, microspheres had an average diameter of 51 ± 16 µm,

and 15% cross-link density. Following this experiment, it was decided that microspheres with a

higher cross-link density were needed, and Batch 4 was fabricated with an average size of 50 ±

36 µm and cross-link density of 60 ± 7% (microsphere batch characterizations listed in Appendix

G). These microspheres were used in both FGF-2 and TGF-β1 experiments. For growth factor-

Figure 8.1: Schematic of growth-

factor induced focal lesion.


loaded microspheres, a solution of PDGF-BB (Peprotech), FGF-2 (Cell Signaling Technologies),

or TGF-β1 (Peprotech), was made with 400 ng growth factor per mg microspheres, in a volume

of 25 µl solution per mg microsphere. Microspheres were hydrated for 2 hours in growth factor

solution, or in PBS for unloaded control groups. For all experiments, one group was seeded with

growth factor-loaded microspheres, and two groups were seeded with unloaded microspheres.

One of the unloaded control groups served as a control with exogenous treatment of the growth

factor being evaluated, and one was not treated with the growth factor being evaluated.

Agarose wells were flooded with growth medium 2 hours after seeding. Medium was

then switched to the designated differentiation medium after 24 hours. All differentiation

medium contained DMEM (Corning), 5% FBS (Thermo Fisher) 1% L-glutamine (Corning), 1%

ITS (Corning), 1% penicillin-streptomycin (Corning), and 50µg/ml ascorbate (Wako). Growth

factors added to the medium varied depending on the experiment. A complete list of

concentrations of growth factors for each group is shown in Table 8.1 below. Rings were

cultured for 3, 7, or 14 days for PDGF and FGF experiments, or 10 days for TGF-β1 delivery

experiments, prior to fixing for histology or flash freezing for biochemical analysis.

Unloaded MS Unloaded MS-

Exogenous control

Growth factor-

loaded MS

PDGF Experiment 5 ng/ml TGF-β1

2.5 ng/ml BMP-4

5 ng/ml TGF-β1

2.5 ng/ml BMP-4

10 ng/ml PDGF

5 ng/ml TGF-β1

2.5 ng/ml BMP-4

FGF Experiment 5 ng/ml TGF-β1

2.5 ng/ml BMP-4

10 ng/ml PDGF

5 ng/ml TGF-β1

2.5 ng/ml BMP-4

10 ng/ml PDGF

10 ng/ml FGF

5 ng/ml TGF-β1

2.5 ng/ml BMP-4

10 ng/ml PDGF

TGF-β1 Experiment 2.5 ng/ml BMP-4

10 ng/ml PDGF

10 ng/ml FGF

2.5 ng/ml BMP-4

10 ng/ml PDGF

10 ng/ml FGF

5 ng/ml TGF-β1

2.5 ng/ml BMP-4

10 ng/ml PDGF

10 ng/ml FGF

10 ng/ml PDGF

10 ng/ml FGF

10 ng/ml PDGF

10 ng/ml FGF

5 ng/ml TGF-β1

10 ng/ml PDGF

10 ng/ml FGF

Table 8.1: Exogenous growth factor concentrations in culture medium for microsphere (MS)-mediated

growth factor delivery experiments


8.2.3. Ring thickness measurements

Images of rings in agarose molds were taken using a DVT imaging system (Framework).

Ring thickness was measured from DVT images using ImageJ (NIH).


Tissues were fixed for 4-6 hours in 10% neutral buffered formalin, processed, paraffin

embedded, and cut into 5µm sections adhered to charged glass slides. A Hematoxylin and Eosin

stain was used visualize ring and tube morphology, and a Picrosirius Red/Fast Green stain was

used to visualized collagen deposition.

All slides to be used for immunohistochemistry were subjected to antigen retrieval as

described in section 3.2.5. Briefly, slides were left for 5 minutes in boiling Tris-EDTA solution

in a pressure cooker. Slides to be stained for smooth muscle alpha actin (SMA; Dako), calponin

(calp; Dako), and smooth muscle protein 22 alpha (SM22-α; Bio-Rad) were blocked for 45

minutes in 1.5% normal rabbit serum (NRS, Vector) in PBS, then incubated overnight at 4ºC in

primary antibody (1:100 in 1.5% NRS). Negative controls were incubated in either mouse (SMA,

calp) or goat (SM22-α) immunoglobulin protein (Vector) instead of primary antibody. Slides

were then incubated in the appropriate secondary antibody (AlexaFluor 488 rabbit anti-mouse for

SMA and calp, and AlexaFluor 488 mouse anti-goat for SM22-α) at a 1:400 ratio with 1.5%

NRS for 1 hour. Samples were then counterstained with Hoechst dye (1:6000 in DI water, 6

minutes) prior to mounting in aqueous mounting medium.

Following antigen retrieval, slides to be stained for Ki67 were quenched for 30 minutes

in 0.3% hydrogen peroxide, prior to blocking in 5% normal goat serum (NGS, Vector). Slides

were incubated in anti-Ki67 antibody (Abcam, 1:100 in 3% NGS) overnight at 4ºC. Negative

controls were instead incubated in a rabbit immunoglobulin protein (Vector). Samples were then

incubated in rabbit ImmPress reagent (Vector), prior to developing with a DAB kit (Vector)

according to the manufacturer instructions. Meyer’s Hematoxylin was used as a counterstain to

visualize nuclei prior to mounting coverslips on slides with aqueous mounting medium.

8.2.5. DNA quantification


Total DNA in tissue rings was quantified using a CYQUANT assay kit (Thermo Fisher).

Frozen tissue rings were lysed and homogenized as described in Chapter 3.2.8. After

homogenization, samples were diluted 1:64 or 1:128 in HBSS. A known number of hMSCs were

also lysed and used to create a standard curve. CYQUANT reagent was prepared according to

manufacturer instructions. 50µl of either sample or standard was pipetted into each well of a 96

well plate with 50µl of the CYQUANT reagent. Two replicates were used per sample. After a

one-hour incubation at 37ºC, the plate was read on a Victor3 Plate Reader, and total number of

cells was calculated from the standard curve.

8.2.6. PDGF loading efficiency

The loading efficiency of PDGF in gelatin microspheres was determined using an

enzyme-linked immunosorbent assay (ELISA; Thermo Fisher). Gelatin microspheres were

soaked in a solution of 400 ng per mg microspheres (total volume of 25 µl per mg microsphere)

for two hours at 37ºC. The microspheres were then gently rinsed in an ELISA buffer prepared

according to the manufacturer’s instructions. Buffer from three separate samples (with three

replicates per sample) was read using a Victor3 plate reader. A standard curve of known PDGF

concentrations was used to calculate the concentration and total amount of PDGF in the

supernatant from each sample.


Statistical analysis was performed for all quantifications, with a One or Two-Way

ANOVA test as appropriate, and a Holm-Sidak post-hoc analysis where applicable. A P value

less than 0.05 was considered significant for all tests. Sample sizes are specified in captions,

ranging from 3-8 samples per group.

8.3. Results

8.3.1. Effects of microsphere-mediated PDGF release on hMSC rings

Our first goal was to evaluate the effects of microsphere-mediated PDGF delivery on

hMSC rings, in order to create hyper-proliferative rings with a synthetic phenotype that may be

used for creating an intimal lesion. First, we tested the loading efficiency of PDGF in gelatin

microspheres. Using an ELISA kit, we determined that microspheres had a 51.4 ± 7.9% loading


efficiency, which is comparable to previous studies evaluating TGF-β1 release from gelatin

microspheres [22].

To evaluate the effects of PDGF

release on tissue rings, three groups were

tested. Rings had either unloaded

microspheres and no PDGF, unloaded

microspheres with exogenous PDGF, or

PDGF-loaded microspheres. Rings without

any PDGF and rings with PDGF-loaded

microspheres had comparable thicknesses at

both 3, 7, and 14 days (Fig 8.2 A). At days 7

and 14, rings with exogenous PDGF had

significantly greater thicknesses than the

other two groups (Fig 8.2 A). At 14 days, it is

clear the PDGF-treated rings are constricting the agarose posts, resulting in a smaller lumen

Figure 8.2: Effect of PDGF treatment on ring thickness. Thickness measurements of rings at 3, 7, and 14 days

with unloaded microspheres and no PDGF treatment or 10 ng/ml exogenous PDGF, or containing microspheres

loaded with 400 ng PDGF per mg microspheres (A). Representative DVT images of rings in each group shown in

(B). * P<0.05 compared to no PDGF and PDGF MS groups within time point. Two-way ANOVA with Holm-

Sidak Post Hoc test. N = 5. Bars are mean ± SD. Scale bar = 1 mm.

Figure 8.3: Effect of PDGF on total DNA content.

Rings containing unloaded microspheres were treated

with no PDGF (No PDGF) or 10ng/ml exogenous PDGF

(Ex PDGF), or contained microspheres loaded with 400

ng PDGF per mg microspheres (PDGF MS). Bars are

mean ± SD. One Way ANOVA with Holm-Sidak post

hoc test. * P<0.05 compared to Ex PDGF and PDGF MS.

N = 4. Rings are 14 days old.


diameter (Fig 8.2 B). It is unclear if the increased thickness is a result of increased ring diameter

Figure 8.4: Cellular proliferation in rings treated with PDGF. Rings were stained for Ki67 to

examine proliferation at day 3 (A-C), 7 (D-F), or 14 (G-I) days. Rings containing unloaded

microspheres were treated with no PDGF (A, D, G), 10ng/ml exogenous PDGF (B, E, H), or

contained microspheres loaded with 400 ng PDGF per mg microspheres (G-I). Rings were

stained at day 3 (C, F, I). Green = Ki67, blue = Hoechst. Top panels are Ki67, bottom panels are

Ki67 merged with Hoechst. Lumen on bottom/right. Scale = 100µm.


diameter (Fig 8.2 B). It is unclear if the increased thickness is a result of increased ring

proliferation and matrix deposition, or if the reduced inner diameter of the ring is causing only an

apparent increase in ring volume. It was also noted that tissue rings in all groups had some

degree of outgrowth up the side of the agarose posts, although this effect was slightly less in the

exogenous PDGF group.

To determine if this increase in thickness was due to an increase in total cell number, a

CYQUANT assay was used to quantify total DNA. This showed that rings treated with PDGF

both exogenously and via microspheres had significantly higher DNA content than control rings

(Fig 8.3), suggesting that the increased thickness could be due to increased cell number. A Ki67

stain was used to further evaluate if rings treated with PDGF had increased proliferation.

However, no clear differences between groups were observed, and there was considerable

variation both between and within samples of each group (Figure 8.4).

A Picrosirius Red/Fast Green stain was used to examine collagen deposition and the

presence of gelatin microspheres (Fig 8.5). Microspheres are clearly visible at day 3, but not at

days 7 or 14, suggesting that they had been degraded by this time. Large amounts of collagen are

visible in all groups at all

time points, with increasing

density over time. At 14

days, it appeared that

collagen in the exogenous

PDGF group was more

densely concentrated near the

ring lumen compared to the

other groups. Hematoxylin

and Eosin (H&E) staining is

shown in Figure 8.6.

Samples stained for

the contractile proteins SMA,

SM22-α, and calponin are

shown in Figure 8.7. While

Figure 8.5: Collagen deposition in rings with PDGF treatment. Rings

containing unloaded microspheres were treated with no PDGF (No PDGF, A-

C), 10ng/ml exogenous PDGF (Ex PDGF, D-F), or contained microspheres

loaded with 400 ng PDGF per mg microspheres (PDGF MS, G-I). Rings were

stained at day 3 (A, D, G), 7 (B, E, H), or 14 (C, F, I). Picrosirius Red/Fast

Green stain (red = collagen, green = counterstain). Lumen on bottom/right.

Scale = 100µm.


no clear differences

are observed between

groups, the increased

constriction shown in

Figure 8.2 B suggests

that rings with

exogenous PDGF

may be more

contractile, although

more analysis is

needed to verify this.

More SMA is

observed than SM22-

α, and only small

amounts of calponin

are visible. It was

also noted in all groups that

particularly thick regions of

rings expressed contractile

proteins on the inner and

outer edges, but not in the

middle region of the tissue.

8.3.2. Effect of microsphere-

mediated FGF release on

hMSC rings

Because PDGF did

not substantially decrease

contractile protein

expression, we next

evaluated the effects of FGF

Figure 8.7: Contractile protein expression in PDGF treated rings. Rings

containing unloaded microspheres were treated with no PDGF (No PDGF, A-

C) or 10ng/ml exogenous PDGF (Ex PDGF, D-F), or contained microspheres

loaded with 400 ng PDGF per mg microspheres (PDGF MS, G-I). Rings were

stained at day 14 for SMA (A, D, G), SM22-α (B, E, H), or calponin (C, F, I).

Green = contractile protein, blue = nuclei. Lumen on bottom/right. Scale =

100µm. Rings are 14 days old.

Figure 8.6: Morphology of rings with PDGF treatment. Rings containing unloaded

microspheres were treated with no PDGF (A-C), 10ng/ml exogenous PDGF (D-F), or

contained microspheres loaded with 400 ng PDGF per mg microspheres (G-I). Rings

were stained at day 3 (A, D, G), 7 (B, E, H), or 14 (C, F, I). Hematoxylin and Eosin stain

(purple = nuclei, pink = counterstain). Lumen on bottom/right. Scale = 100µm.


treatment on hMSC rings. FGF is a potent TGF-β antagonist and SMC mitogen. Rings were

cultured with unloaded microspheres, without or with exogenous 10 ng/ml FGF treatment, or

with microspheres loaded with 400 ng FGF per mg microsphere. Previously, we tested 5 ng/ml

exogenous FGF compared to 200 ng per mg of microsphere-mediated FGF delivery, as FGF is

typically used at lower concentrations than PDGF and TGF-β1. This is still higher than the

concentration of FGF2 in some commercial culture mediums (Lonza, 2 ng/ml) designed to

stimulate proliferation. However, we observed limited effects with 5 ng/mL FGF treatment

(Appendix H), and decided to test 10 ng/ml instead, as the effects of FGF increase with

increasing concentration [23].

At day 3, we observed significant increases in ring thickness with FGF loaded

microspheres compared to rings with exogenous FGF treatment (Fig 8.8 A). This may be

partially due to uneven thicknesses observed in rings with exogenous FGF treatment at this time

point (Fig 8.8 B). Over time, however, rings remodeled and thicknesses later became uniform in

all groups. No significant differences were observed in ring thickness at 7 days, although at day

14 rings without FGF treatment were significantly thinner (Fig 8.8 A). Additionally, some small

bumps were observed around ring edges in both FGF-treated groups (Fig 8.8 B)

Figure 8.8: Effect of FGF treatment on ring thickness. Thickness measurements of rings at 3, 7, and 14 days

with unloaded microspheres and no FGF treatment (No FGF) or 10 ng/ml exogenous FGF (Ex FGF), or containing

microspheres loaded with 400 ng FGF per mg microspheres (FGF MS, A). Representative DVT images of rings in

each group shown in (B). * P<0.05 compared to no FGF and FGF MS groups within time point. Two-way ANOVA

with Holm-Sidak Post Hoc test. N = 3-4 for days 3 and 7, n = 7-8 for day 14. Bars are mean ± SEM. Scale bar = 1

mm


Collagen deposition

was clearly visible in all groups

(Fig 8.9). However, in rings

treated with FGF more

collagen was observed around

ring edges, with less in the

middle region. H&E staining is

shown in Figure 8.10. Notably,

microspheres were not visible

in any of the stained samples,

suggesting complete

degradation by day 3.

Samples were immuno-

stained for Ki67 to evaluate

changes in cellular proliferation

(Fig 8.11). No clear

differences in number of

positive cells were visible

between groups. When

quantified, the number of

positive cells per cross-

sectional area showed no

significant differences,

except for 7-day rings with

FGF loaded microspheres

compared to 14 day rings

treated with exogenous FGF

(Fig 8.12). This may be due

to the large variation in the

number of positive cells

among samples within each

Figure 8.9: Collagen deposition in rings with FGF treatment. Rings

containing unloaded microspheres were treated with no FGF (No FGF, A-

C), 10ng/ml exogenous FGF (Ex PDGF, D-F), or contained microspheres

loaded with 400 ng FGF per mg microspheres (FGF MS, G-I). Rings were

stained at day 3 (A, D, G), 7 (B, E, H), or 14 (C, F, I). Picrosirius Red/Fast

Green stain (red = collagen, green = counterstain). Lumen on bottom/right.

Scale = 100µm.

Figure 8.10: Morphology of FGF treated rings. Rings containing unloaded

microspheres were treated with no PDGF (A-C), 10ng/ml exogenous PDGF

(D-F), or contained microspheres loaded with 400 ng PDGF per mg

microspheres (G-I). Rings were stained at day 3 (A, D, G), 7 (B, E, H), or 14

(C, F, I). Hematoxylin and Eosin stain (purple = nuclei, pink = counterstain).

Lumen on bottom/right. Scale = 100µm.


group. No significant

differences were found in total

DNA content. However, there

was a trend of decreased total

DNA in the exogenous FGF

group, which is consistent with

the decreased ring thickness

(Fig 8.13).

Samples were then

stained for the contractile

proteins SMA, SM22-α, and

calponin (Fig 8.14). It was

clear that rings with exogenous

FGF only had contractile

protein expression around ring

edges, with dense nuclei and no contractile proteins in the middle regions. These regions with

dense nuclei and minimal contractile proteins were also visible in the other two groups, but only

in particularly thick regions of the ring, whereas the effect was uniform in rings treated with

exogenous FGF. It did not appear that

Figure 8.13: Effect of FGF on total DNA content.

Rings containing unloaded microspheres were

treated with no FGF (No FGF), 10ng/ml exogenous

FGF (Ex FGF), or contained microspheres loaded

with 400 ng FGF per mg microspheres (FGF MS).

Bars are mean ± SD. One Way ANOVA, P > 0.05,

N = 4. Rings are 14 days old.

Figure 8.12: Cellular proliferation in FGF treated rings.

The number of Ki67 positive cells per image was counted,

and normalized to ring cross-sectional area for each group at

each time point. One way ANOVA with Tukey’s post-hoc

analysis. *P <0.05. N = 4. Bars are mean ± SD

Figure 8.11: Proliferation in rings with FGF treatment. Rings containing

unloaded microspheres were treated with no FGF (No FGF, A-C), 10ng/ml

exogenous FGF (Ex FGF, D-F), or contained microspheres loaded with 400

ng FGF per mg microspheres (FGF MS, G-I). Rings were stained at day 3

(A, D, G), 7 (B, E, H), or 14 (C, F, I). Ki67 immunostain stain (brown =

Ki67, purple = nuclei). Lumen on bottom/right. Scale = 100µm.


FGF-loaded microspheres

visibly reduced

contractile protein

expression compared to

rings without FGF

treatment.

8.3.3. Effect of

microsphere-mediated

TGF-β1 release on hMSC

rings

Since FGF-loaded

microspheres had a

limited ability to decrease

ring contractile protein

expression, we instead

decided to add PDGF and

FGF to the base medium

and remove TGF-β1, to prevent differentiation. We then evaluated the effect of TGF-β1 loaded

microspheres on hMSC rings, as in Chapter 3. This would allow us to selectively differentiate,

rather than de-differentiate, hMSC rings, towards the same goal of controlling ring phenotype

with microsphere-mediated growth factor delivery. Within each group, half of the rings were

treated with BMP-4 and half were not, to evaluate the necessity of BMP-4 for MSC

differentiation into SMCs. Rings treated with exogenous TGF-β1 demonstrated slight increases

in ring thickness (Fig 8.15 A) compared to rings with no TGF-β1 or with TGF-β1-loaded

microspheres, regardless of BMP-4 treatment, although this was not significant.

Picrosirius Red/Fast Green staining showed large amounts of collagen in all groups (Fig

8.16). Collagen appeared slightly more dense with exogenous TGF-β1 treatment compared to

groups without TGF-β1 or with TGF-β1-loaded microspheres, especially on the inner luminal

side. Additionally, groups without BMP-4 appeared to have slightly looser and less organized

collagen. Microspheres were not visible in any stained sections at any time points.

Figure 8.14: Contractile protein expression in FGF treated rings. Rings

containing unloaded microspheres were treated with no FGF (No FGF, A-C),

10ng/ml exogenous FGF (Ex FGF, D-F), or contained microspheres loaded with

400 ng FGF per mg microspheres (FGF MS, G-I). Rings were stained at day 14

for SMA (A, D, G), SM22-α (B, E, H), or calponin (C, F, I). Green = contractile

protein, blue = nuclei. Lumen on bottom/right. Scale = 100µm. Rings are 14 days

old.


Next, we evaluated cellular proliferation using Ki67 staining (Fig 8.17). There appeared

to be almost no Ki67 positive cells in groups without TGF-β1, or with TGF-β1 loaded

microspheres (Fig 8.17 A, D, C, F). However, groups treated with exogenous TGF-β1 appeared

to have more Ki67 positive cells (Fig 8.17 B, E). More Ki67 positive cells were visible with both

TGF-β1 and BMP-4 than with TGF-β1 alone. When quantified, there are clearly more Ki67

positive cells in

rings with

exogenous TGF-

β1. However,

there was

significant

variation within

each sample,

leading to very

high standard

deviations and

consequently no

significant

Figure 8.16: Collagen deposition in TGF-β1 treated rings. Rings in all groups were

treated with (A-C) or without (D-F) exogenous BMP-4. Within those groups, rings

contained unloaded microspheres and TGF-β1 (No TGF, A, D) or with exogenous TGF-β1

(Ex TGF, B, E), or contained microspheres loaded with 400 ng TGF-β1 per mg

microspheres (TGF MS, C, F). Picrosirius Red/Fast Green stain (red = collagen, green =

counterstain). Lumen on bottom/right of image. Scale = 100µm. Rings are 10 days old.

Figure 8.15: Effect of TGF-β1 and BMP-4 on ring thickness. Rings were seeded either with unloaded

microspheres with (Ex TGF) or without (No TGF) 5 ng/ml TGF-β1, or with microspheres loaded with 400 ng TGF-

β1 per mg microspheres (TGF MS). Within each group, rings were treated with or without exogenous BMP-4.

Average thickness measurements of 10 day old rings shown in (A), with representative DVT images shown in (B).

Bars are mean ± SEM. One Way ANOVA, P > 0.05, n = 6-8. Scale bar = 1mm. Rings are 10 days old.


differences

between groups

(Fig 8.18). When

the number of

positive cells was

normalized to

ring cross-

sectional area, it

appeared that

rings with

exogenous TGF-

β1 had

comparable

amounts of Ki67

positive cells to rings with microsphere-mediated TGF-β1 delivery. This may be because rings

with TGF-β1-loaded microspheres were thinner (Fig 8.15). However, there is still too much

variation to draw meaningful conclusions from this analysis. We then quantified total DNA using

a CYQUANT assay, to further evaluate if TGF-β1 increases total cell number (Fig 8.19). No

significant differences were observed, although there was a trend of increased total DNA with

TGF-β1 treatment, which was increased in rings also treated with BMP-4. Rings with TGF-β1-

Figure 8.17: Cellular proliferation in TGF-β1 treated rings. Rings in all groups were

treated with (A-C) or without (D-F) exogenous BMP-4. Within those groups, rings

contained unloaded microspheres and TGF-β1 (No TGF, A, D) or with exogenous TGF-β1

(Ex TGF, B, E), or contained microspheres loaded with 400 ng TGF-β1 per mg

microspheres (TGF MS, C, F). Ki67 immunostain (brown = Ki67, purple = nuclei). Lumen

on bottom/right of image. Scale = 100µm. Rings are 10 days old.

Figure 8.18: Proliferation in hMSC rings treated with TGF-β1. The average number of Ki67 positive cells per

group (A), and positive cells normalized to cross sectional area (B). One Way ANOVA on Ranks test. P < 0.05, but

Dunn’s post hoc analysis found no differences between groups. N = 4. Bars are mean ± SD.


loaded microspheres appeared to have

more total DNA than rings treated with

exogenous TGF-β1. It is possible that the

high burst release of TGF-β1 when

microspheres degraded sharply increased

proliferation early in culture, resulting in a

higher total cell number, despite the

apparent fewer actively proliferating cells

by day 10.

Rings were then stained for SMA,

SM22, and calponin to assess the effect of

TGF-β1 on hMSC ring differentiation to a

contractile phenotype (Fig 8.20). Very

limited contractile protein expression was

observed in rings with no TGF-β1, and almost none was visible in rings without TGF-β1 or

BMP-4. Rings treated with exogenous TGF-β1 exhibited higher levels of SMA and SM22

expression, except for in the middle region of the tissue, regardless of BMP-4 treatment. This is

consistent in the previous experiment, where the same combination of TGF-β1, FGF, BMP-4,

and PDGF were added exogenously. Limited amounts of calponin were visible. In the TGF-β1-

loaded microsphere group, SMA and SM22 expression was observed around ring edges, which

was more than rings without no TGF-β1, but substantially less than rings with exogenous TGF-

β1. The presence of exogenous BMP-4 seemed to slightly increase SMA and SM22 expression

within the TGF-β1-loaded microsphere group.

8.4. Discussion

IH is characterized by a focal region of SMC proliferation, migration to the vessel lumen,

and increased matrix deposition. To develop an in vitro IH disease model, it is important to have

spatial control over SMC phenotype within an engineered vessel. Such models will require SMC

rings that are of a healthy, contractile phenotype, to simulate healthy regions of the tissue, in

addition to SMC rings of a synthetic phenotype, in order to simulate diseased regions of the

Figure 8.19 Effect of TGF-β1 on total DNA content.

Rings containing unloaded microspheres were treated with

no TGF-β1 (No TGF-β1) or 5 ng/ml exogenous TGF-β1

(Ex TGF-β1), or contained microspheres loaded with 400

ng TGF-β1 per mg microspheres (TGF-β1 MS). All groups

were further divided into rings with or without 2.5 ng/ml

exogenous BMP-4 treatment. Bars are mean ± SD. One

Way ANOVA, P > 0.05, N = 3-4. Rings are 10 days old.


tissue. Due to the challenges of differentiating primary human aortic SMCs to a contractile

Figure 8.20: Contractile protein expression in rings treated with TGF-β1. Ring sections were stained

for SMA (A-F), SM22 (G-L), or calp (M-R). Rings in all groups were treated with (A-C, G-I, M-O) or

without (D-F, J-L, P-R) exogenous BMP-4. Within those groups, rings contained unloaded microspheres

and TGF-β1 (No TGF, left) or with exogenous TGF-β1 (Ex TGF, middle), or contained microspheres

loaded with 400 ng TGF-β1 per mg microspheres (TGF MS, right). Green = contractile protein, blue =

nuclei. Lumen on bottom/right of image. Scale = 100µm.


tissue. Due to the challenges of differentiating primary human aortic SMCs to a contractile

phenotype, we decided to perform these experiments with hMSCs, which more readily express

contractile proteins than primary human SMCs.

To achieve this, we aimed to use growth factor-loaded microspheres to prevent

differentiation of select rings, and maintain a hyper-proliferative state with limited contractile

protein expression. PDGF is well established to de-differentiate SMCs into a synthetic phenotype

by reducing contractile protein expression and increasing proliferation and matrix deposition,

and is strongly associated with the initiation and progression of intimal hyperplasia [6, 11, 16-

19]. Thus, PDGF seemed like the optimal growth factor to create “diseased” rings with a

synthetic phenotype. However, we observed no clear differences in contractile protein expression

with exogenous or microsphere-mediated PDGF delivery. We even observed that rings treated

with exogenous PDGF constrict more tightly around agarose posts than rings without PDGF,

suggesting that PDGF may have enhanced ring contractility.

While the effects of PDGF on primary SMCs are well established, there have been mixed

reports of its effects on hMSCs. Some reports show that PDGF has similar effects on hMSCs as

it does on SMCs, decreasing contractile protein expression and increasing proliferation [5, 24,

25]. Others reports suggest that PDGF may instead increase contractile protein expression, rather

than decrease it [26, 27]. Other stem cell types, such as embryonic stem cells, require PDGF to

differentiate into vascular precursor cells [28]. It is possible that the effect of PDGF on hMSCs

may depend on how differentiated they are. Prior to switching our cell source from primary

SMCs to hMSCs, our proposed culture medium did not contain TGF-β1. TGF-β1 is well-

established to promote SMC [1, 2] and hMSC [4, 5, 24] differentiation to a contractile SMC

phenotype, and its effects may not be strongly inhibited by PDGF [6, 29, 30]. PDGF is

sometimes used in combination with TGF-β1 to promote both proliferation and matrix

deposition and hMSC differentiation of vascular constructs [6].

For these reasons, it was unsurprising that we did not observe clear decreases in

contractile protein expression with PDGF treatment. Some rings in this experiment did thin

unevenly and neck over time, although this was observed in fewer rings and to a lesser extent in

the group treated with exogenous PDGF. Additionally, all rings treated with PDGF had a

significantly higher DNA content, suggesting increased proliferation. For these reasons, we


decided to add PDGF to our base hMSC differentiation medium in all experiments moving

forward.

As an alternative to PDGF to induce a diseased state, we decided to evaluate the effects

of FGF-2 on ring differentiation and proliferation. FGF is a potent antagonist of TGF-β1 and is

well-established to reduce contractile protein expression and increase proliferation in SMCs [13,

17, 20]. In hMSCs, FGF is known to maintain an undifferentiated state [31, 32], prevent cellular

senescence [31, 32], and reduce contractile protein expression [6]. When used together, FGF and

PDGF have been shown to more strongly stimulate SMC proliferation than either growth factor

alone [17]. Thus, we anticipated that with PDGF in our base differentiation medium, FGF

treatment would significantly increase cellular proliferation and prevent contractile protein

expression.

While rings treated with FGF still clearly exhibited SMA and SM22-α expression, it was

notably less than rings that were not treated with exogenous FGF. This decrease was not visible

in rings with FGF-loaded microspheres. It was also noted that microspheres were not visible in

histological sections at 3, 7, or 14 days. While cross-linking of gelatin microspheres does help

slow degradation, it does not prevent it. If MSCs are secreting high amounts of proteinases, it is

possible that they had degraded in this short period of time. This would have limited FGF

treatment to a short burst release early in culture, which may have hindered its ability to prevent

TGF-β1-induced increases in contractile protein expression. In the future, additional

modifications to microspheres may be needed to prolong FGF release. It is clear though that this

burst release still had some effect on hMSC rings, as evidenced by the significantly lower ring

thicknesses at day 14, which were comparable to the exogenous FGF group. Rings with

exogenous FGF treatment also had slightly lower total DNA content. This is contrary to our

hypothesis that FGF would increase proliferation and consequently increase ring thickness.

As an alternative to FGF delivery, we next decided to evaluate the effect of microsphere-

mediated TGF-β1 delivery on hMSC rings. The release of TGF-β1 from the gelatin microspheres

used in this study is well-characterized [22, 33], and we already demonstrated in Chapter 3 that

TGF-β1 loaded microspheres can increase contractile protein expression in human coronary

artery SMCs comparably to exogenous TGF-β1 [34]. Our goal would then be to fuse rings with

unloaded microspheres between rings with TGF-β1-loaded microspheres, to selectively


differentiate hMSC rings in the outer regions of the tube, leaving a focal region in an

undifferentiated state. In this case, the base differentiation medium contained 10 ng/ml PDGF

and 10 ng/ml FGF to prevent differentiation in rings without TGF-β1-loaded microspheres. All

groups were also tested with and without 2.5 ng/ml BMP-4, to determine if BMP-4 alone would

push hMSCs to a contractile vascular phenotype.

With exogenous TGF-β1 treatment, we observed clear increases in contractile protein

expression, regardless of the presence of BMP-4. Rings with incorporated TGF-β1-loaded

microspheres appeared to have a slight increase in contractile protein expression around ring

edges. This effect was slightly increased with BMP-4 treatment. While this small increase in

contractile protein expression is encouraging, it is not enough to create a fully differentiated,

contractile vascular tissue. It was noted that microspheres were completely degraded by the 10

day time point used in this study. Based on our previous experiment with FGF incorporation,

they likely degraded within the first few days of culture. Thus, it may not be possible to control

hMSC differentiation without addressing the microsphere degradation rate. The short burst

release of FGF and TGF-β1 in the first few days of culture appears to have had some effect, but

in order to fabricate a focal disease model with both differentiated and undifferentiated regions, a

longer-term release will be critical. The cross-link density of microspheres used in these

experiments was already fairly high (66%), indicating that other modifications may be needed.

Because gelatin is proteolytically degraded, it may be possible to treat rings with protease

inhibitors to delay gelatin degradation and growth factor release. Alternatively, microsphere

coatings may be needed to further delay growth factor delivery [35].

We observed slight increases in ring thickness, total DNA, and number of proliferating

cells in response to exogenous TGF-β1. The number of proliferating cells was further increased

with exogenous BMP-4 treatment. This is surprising, as TGF-β1 is well known for its inhibitory

effects on SMC proliferation [29, 30]. However, TGF-β1 is also known to contribute to IH, and

there have been a handful of reports where TGF-β1 has increased SMC proliferation [36, 37]. It

is possible that certain conditions that follow an arterial injury, such as elevated levels of

SMAD3, may cause TGF-β1 to stimulate, instead of inhibit, proliferation and intimal growth

[36]. It is also possible that our partially differentiated hMSCs may not respond in the same way

to growth factor treatment as primary SMCs. TGF-β1 is able to induce a wide range of effects on


hMSCs, including differentiation to a chondrogenic [38] or a vascular [4] phenotype, suppression

of differentiation to an osteogenic phenotype [39], or an increase in proliferation [39]. It has also

been shown that the combination of PDGF, FGF, and TGF-β1 can support undifferentiated

hMSC growth comparably to FBS, while maintaining them in an undifferentiated state [40].

With our MSC rings, cells have partially differentiated into SMCs, suggesting they may have

either SMC-like and MSC-like responses to different growth factors.

Our goal with these studies was to focally control SMC phenotype, so that certain regions

of the tube could be differentiated into a contractile phenotype, and some regions would maintain

a diseased state. However, because FGF is known to maintain MSCs in an undifferentiated state

[31, 32], even in combination with TGF-β1 and PDGF [40], we may not truly be achieving a

diseased SMC state with FGF treatment. Thus, it may be necessary to remove FGF from culture

medium. Undifferentiated hMSCs may not respond to potential therapeutics in the same way as

synthetic SMCs, just as they do not always respond to growth factors in the same way as SMCs.

It may be necessary to fully differentiate SMC rings prior to releasing growth factors such as

PDGF or FGF, and then to de-differentiate cells from a contractile to a synthetic SMC

phenotype. However, we have already observed rapid microsphere degradation in hMSC rings,

even with high cross-link densities. Thus, even further delaying growth factor release will likely

require significant modifications to microspheres.

In this chapter, we demonstrated that microsphere-mediated growth factor delivery had

small but measurable effects on hMSC rings. PDGF caused increases in ring thickness and total

DNA, FGF caused slight decreases in ring thickness, and TGF-β1 slightly increased contractile

protein expression. While this demonstrates the potential of microsphere-mediated growth factor

delivery for controlling ring phenotype, more modifications to microspheres may needed to

achieve larger changes in contractile protein expression.

8.5. References

1. Owens, G.K., M.S. Kumar, and B.R. Wamhoff, Molecular Regulation of Vascular

Smooth Muscle Cell Differentiation in Development and Disease. Physiological Reviews,

2004. 84: p. 767-801.

2. Tang, Y., S. Urs, J. Boucher, T. Bernaiche, D. Venkatesh, D.B. Spicer, C.P. Vary, and L.

Liaw, Notch and transforming growth factor-beta (TGFbeta) signaling pathways


cooperatively regulate vascular smooth muscle cell differentiation. J Biol Chem, 2010.

285(23): p. 17556-63.

3. Liu, J.Y., D.D. Swartz, H.F. Peng, S.F. Gugino, J.A. Russell, and S.T. Andreadis,

Functional tissue-engineered blood vessels from bone marrow progenitor cells.

Cardiovasc Res, 2007. 75(3): p. 618-28.

4. Wang, C., S. Yin, L. Cen, Q. Liu, W. Liu, Y. Cao, and L. Cui, Differentiation of Adipose-

Derived Stem Cells into Contractile Smooth Muscle Cells Induced by Transforming

Growth Factor-β1 and Bone Morphogenetic Protein-4. Tissue Eng Part A, 2010. 16(4):

p. 1202-1213.

5. Narita, Y., A. Yamawaki, H. Kagami, M. Ueda, and Y. Ueda, Effects of transforming

growth factor-beta 1 and ascorbic acid on differentiation of human bone-marrow-derived

mesenchymal stem cells into smooth muscle cell lineage. Cell Tissue Res, 2008. 333(3):

p. 449-59.



7. Liang, M.S. and S.T. Andreadis, Engineering fibrin-binding TGF-beta1 for sustained

signaling and contractile function of MSC based vascular constructs. Biomaterials, 2011.

32(33): p. 8684-93.

8. Cho, S.-W., S.H. Lim, I.-K. Kim, Y.S. Hong, S.-S. Kim, K.J. Yoo, H.-Y. Park, Y. Jang,

B.C. Chang, C.Y. Choi, et al., Small-Diameter Blood Vessels Engineered With Bone

Marrow Derived Cells. Annals of Surgery, 2005. 241(3): p. 506-515.





2000. 190: p. 300-309.




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22. Dang, P.N., N. Dwivedi, L.M. Phillips, X. Yu, S. Herberg, C. Bowerman, L.D. Solorio,

W.L. Murphy, and E. Alsberg, Controlled Dual Growth Factor Delivery From

Microparticles Incorporated Within Human Bone Marrow-Derived Mesenchymal Stem

Cell Aggregates for Enhanced Bone Tissue Engineering via Endochondral Ossification.


23. Horton Jr., W.E., J.D. Higginbotham, and S. Chandrasekhar, Transforming Growth

Factor-Beta and Fibroblast Growth Factor Act Synergistically To Inhibit Collagen II

Synthesis Through a Mechanism Involving Regulatory DNA Sequences. Journal of

Cellular Physiology, 1989. 141: p. 8-15.

24. Kinner, B., J.M. Zaleskas, and M. Spector, Regulation of Smooth Muscle Actin

Expression and Contraction in Adult Human Mesenchymal Stem Cells. Experimental Cell

Research, 2002. 278(1): p. 72-83.


25. Tamama, K., C.K. Sen, and A. Wells, Differentiation of bone marrow mesenchymal stem

cells into the smooth muscle lineage by blocking ERK/MAPK signaling pathway. Stem

Cells Dev, 2008. 17(5): p. 897-908.

26. Ball, S.G., C.A. Shuttleworth, and C.M. Kielty, Platelet-derived growth factor receptor-

alpha is a key determinant of smooth muscle alpha-actin filaments in bone marrow-

derived mesenchymal stem cells. Int J Biochem Cell Biol, 2007. 39(2): p. 379-91.

27. Simper, D., Smooth Muscle Progenitor Cells in Human Blood. Circulation, 2002.

106(10): p. 1199-1204.

28. Yamashita, J., H. Itoh, M. Hirashima, M. Ogawa, S. Nishikawa, T. Yurugi, M. Naito, K.

Nakao, and S.-I. Nishikawa, Flk1-positive cells derived from embryonic stem cells serve

as vascular progenitors. Nature, 2000. 408: p. 92-95.

29. Bjorkerud, S., Effects of transforming growth factor-beta 1 on human arterial smooth muscle cells in vitro. Arteriosclerosis, Thrombosis, and Vascular Biology, 1991. 11(4): p.

892-902.

30. Engel, L. and U. Ryan, TGF-beta1 reverses PDGF-stimulated migration of human aortic

smooth muscle cells in vitro. In Vitro Cell. Dev. Biol., 1997. 33: p. 443-451.

31. Bianchi, G., A. Banfi, M. Mastrogiacomo, R. Notaro, L. Luzzatto, R. Cancedda, and R.

Quarto, Ex vivo enrichment of mesenchymal cell progenitors by fibroblast growth factor

2. Experimental Cell Research, 2003. 287(1): p. 98-105.

32. Tsutsumi, S., A. Shimazu, K. Miyazaki, H. Pan, C. Koike, E. Yoshida, K. Takagishi, and

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35. Bian, L., D.Y. Zhai, E. Tous, R. Rai, R.L. Mauck, and J.A. Burdick, Enhanced MSC

chondrogenesis following delivery of TGF-beta3 from alginate microspheres within

hyaluronic acid hydrogels in vitro and in vivo. Biomaterials, 2011. 32(27): p. 6425-34.

36. Suwanabol, P.A., S.M. Seedial, X. Shi, F. Zhang, D. Yamanouchi, D. Roenneburg, B.

Liu, and K.C. Kent, Transforming growth factor-beta increases vascular smooth muscle

cell proliferation through the Smad3 and extracellular signal-regulated kinase mitogen-

activated protein kinases pathways. J Vasc Surg, 2012. 56(2): p. 446-54.


37. Chen, G. and N. Khalil, TGF-beta1 increases proliferation of airway smooth muscle cells

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39. Jian, H., X. Shen, I. Liu, M. Semenov, X. He, and X.-F. Wang, Smad3-dependent nuclear

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Yang, M.C. Vemuri, M.S. Rao, et al., PDGF, TGF-beta, and FGF signaling is important

for differentiation and growth of mesenchymal stem cells (MSCs): transcriptional

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295-307.

Chapter 9: Vascular tissue tubes with distinct phenotypic and structural regions 154

Chapter 9: Vascular tissue tubes with distinct phenotypic and

structural regions

Figure 9.3 from: H.A. Strobel, T.A. Hookway, M. Piola, M. Soncini, G.B. Fiore, E. Alsberg, and M.W.

Rolle. “Assembly of tissue engineered blood vessels with spatially-controlled heterogeneities”. Tissue

Engineering Part A. In Press.







9.1. Introduction

Human mesenchymal stem cells (MSCs) may be an alternative source of vascular smooth

muscle cells (SMCs). They are highly proliferative, easier to obtain from adult allogenetic and

autologous sources than primary human vascular SMCs, express contractile proteins in 3D

culture, and have been used in vascular tissue engineering [1-4]. In Chapter 8, we demonstrated

our ability to culture self-assembled cell rings fabricated from hMSCs. Rings expressed the

contractile proteins smooth muscle alpha actin (SMA), smooth muscle protein 22 alpha (SM22-

α), and calponin (Calp). Here, we evaluated the potential of hMSC rings for modular vascular

tissue tube fabrication.

Our ultimate goal is to create a focal region of smooth muscle de-differentiation and

increased proliferation and thickness characteristic of intimal hyperplasia (IH). We originally

intended to achieve this by delivering growth factors to a focal region of tissue tubes via

microspheres. While this approach is promising, further optimization of medium conditions and

microsphere design are needed (Chapter 8). As an alternative, we evaluated here if different cell

types can be fused to create a focal region of SMCs in a synthetic state. Human aortic SMCs do

not express contractile proteins in 3D culture (Chapter 6), which may allow us to create a focal

region of synthetic SMCs within an otherwise contractile tube. In contrast, hMSCs readily

expressed contractile proteins when treated with TGF-β1 and BMP-4 (Chapter 8). Thus, we


evaluated if a region of synthetic SMCs could be created by fusing rings fabricated from human

aortic SMCs in a central region of the tube, with differentiated hMSCs on either side.

9.2. Methods

9.2.1. Cell culture

Bone marrow-derived human mesenchymal stem cells (hMSCs) were purchased from

RoosterBio, Inc. and expanded according to the manufacturer’s instructions in a propriety growth

medium (RoosterBio, Inc.).

Human aortic smooth muscle cells (SMCs) were purchased from Lifeline Cell

Technologies and expanded in VascuLife growth medium (Lifeline) according to manufacturer

instructions.

9.2.2. Ring fabrication

hMSCs were seeded in agarose molds (2 mm i.d. posts) at a concentration of 600,000

cells/ring. Wells were flooded after 2 hours of cell aggregation with growth medium

(RoosterBio, Inc.), then switched to a custom medium after 24 hours containing DMEM, 5%

FBS, 1% l-glutamine, 1% ITS, 1% penicillin-streptomycin, and 50 µg/ml ascorbic acid.

Human aortic SMCs were pre-loaded with CellTracker Red dye as described in Chapter

5.2.5. Cells were then seeded at a concentration of 400,000 cells/ring in growth medium

(Lifeline). After 24 hours, rings were flooded with growth medium. Rings were kept in growth

medium until tube fabrication. Medium for both hMSC and aortic SMC rings was changed daily.

9.2.3. hMSC tube fabrication

For preliminary hMSC tube fabrication experiments, hMSCs rings were threaded over

silicone mandrels after 3 days of culture. Rings were pushed into contact with one another, and

polycaprolactone (PCL) cuffs ([5], Chapter 4) were pushed onto tube ends. hMSC rings were

cultured in hMSC differentiation medium described in section 9.2.2. After 4 days of fusion

culture (7 days total), 5 ng/ml TGF-β1 and 2.5 ng/ml BMP-4 were added to the medium to

stimulate differentiation to a SMC phenotype. Tubes were fixed after 7 days of fusion culture (10

days total) for immunohistochemistry and histology.


9.2.4. Fabrication of tubes with focal regions of

human aortic SMCs

To fabricate tubes with localized SMC

regions, 3-day old rings were threaded over

silicone mandrels with a central region of 3 human

aortic SMC rings and 4 hMSC rings on either side.

Control tubes were fabricated from human aortic

SMC rings (8 rings per tube), or hMSC rings (5

rings per tube). Three tubes per group were

fabricated with polycaprolactone (PCL) cuffs [5]

on tube ends. A schematic of experimental groups

is shown in Figure 9.1.

All tubes were cultured in hMSC differentiation medium as described in section 9.2.2,

but with added 5 ng/ml FGF for the first 4 days of fusion culture. After 4 days of fusion (3-4

tubes; 3 days of ring culture – 4 days of fusion culture), 5 ng/ml TGF-β1 and 10 ng/ml PDGF

were added to culture medium in all groups, and FGF supplementation was stopped. After 7 days

total of fusion (3-7), tubes were fixed for histological analysis.


After fixing for 1 hour in 10% neutral buffered formalin, samples were processed and

embedded in paraffin. Longitudinal sections 5 µm thick were adhered to positively-charged

slides. Hematoxylin and Eosin staining was used to examine tube morphology and Picrosirius

red/fast green was used to examine collagen deposition.

Immunohistochemistry was performed as described in Chapter 5.2.8. Briefly, antigen

retrieval was performed on samples to be stained for smooth muscle alpha actin (SMA), smooth

muscle protein 22 alpha (SM22-α), and calponin. Slides were blocked in 1.5% normal rabbit

serum for 30 minutes, and were incubated overnight at 4ºC in SMA (Dako, clone 1A4, 1:100),

SM22-α (BioRad VPA00048, 1:100), or calponin (Dako, CALP, 1:100) antibodies. Negative

control samples were incubated with mouse or goat immunoglobulin G (Vector). Samples were

incubated in a secondary antibody (Invitrogen; Alexa Fluor 488 goat anti-rabbit, rabbit anti-

Figure 9.1: Schematic of focal lesion

experimental setup. Pink = hMSC rings, Red =

human aortic SMC rings. Tubes of each cell type,

and hMSC tubes with focal region of aortic SMCs,

were cultured in static conditions on silicone tubing

mandrels for 7 days.


mouse, or mouse anti-goat) at a 1:400 dilution for 1 hour at room temperature. Hoescht dye was

used to visualize nuclei (Invitrogen, 1:6000 in DI water for 6 minutes).

9.3. Results

9.3.1. Fabrication of tissue tubes from hMSC rings

When fabricating engineered blood vessels, it is important that cells express smooth

muscle proteins; indicative of a healthy, contractile SMC phenotype. We did not observe

contractile protein expression in rings or tubes fabricated from primary aortic SMCs (Appendix

F Fig F.4, Chapter 6 Fig 6.9). Therefore, we decided to explore alternative smooth muscle cell

sources. Here, we demonstrated that rings fabricated from bone marrow-derived hMSCs can be

fused into tubes, which were strong enough to handle after 7 days of fusion. Histological analysis

shows that tubes are well

fused, although newly

secreted extracellular matrix

on the tube abluminal surface

is much less dense (Fig 9.2).

Figure 9.3 shows contractile

protein expression of tubes

after 7 days of fusion. SMA

and SM22-α expression is

clearly visible, although

calponin appears

limited.

Contractile

proteins are

predominantly

localized to the

tube outer edge.

Figure 9.4 shows

and H&E stain of

a tube cross

Figure 9.3: Contractile protein expression in fused hMSC tubes. Tubes were fabricated

from 3-day old hMSC rings, which were allowed to fuse for 7 days in static conditions.

Green = SMA (A), SM22-α (B), or calponin (C), blue = nuclei. Lumen on bottom of

image. Scale = 100 µm.

Figure 9.2: Tubes fabricated from hMSCs after 7 days of fusion. Tubes

were stained with Hematoxylin and Eosin (A) or Picrosirius Red/Fast Green

(B; red = collagen, green = counterstain). Lumen on bottom. Scale = 100

µm.


section. Cells appear circumferentially aligned on

the outer edge, but not in the middle region of the

tissue.

9.3.4. Focal region of synthetic SMCs

As a proof-of-

concept experiment, we

fused human aortic SMCs

in a central region of tubes,

between regions of

hMSCs. We have previously observed that human aortic SMCs do not

express contractile proteins in 3D, even with TGF-β1 treatment (not

shown). These tubes were compared to tubes fabricated entirely from

human aortic SMC rings, or entirely from hMSC rings. A schematic of

the groups is shown in Figure 9.1. Tubes were allowed to fuse for 7 days

prior to fixing for histology. We observed that holes began to form in hMSC tubes and regions of

hMSCs in tubes with aortic SMC lesions after 3 days of fusion (Figure 9.5). Despite this, we

continued to culture tubes for the full 7 day fusion period.

Histological analysis shows a high nuclear density in all groups. Picrosirius red/fast green

staining showed very limited collagen deposition in hMSC tubes (Figure 9.6) compared to our

previous experiments, and compared to human aortic SMC tubes. This was also apparent in the

human aortic SMC lesion group, where a region of slightly more dense collagen deposition is

clearly visible within the aortic SMC lesion. Contractile protein expression is shown in Figure

9.7. A thin layer of SM22-α was visible on the abluminal surface of all groups, with slightly

more in the hMSC only group. Almost no SMA or calponin was visible, with no clear

differences between groups.

9.4. Discussion

In Chapter 8, we demonstrated the potential of hMSCs as an alternative cells source for

fabricating vascular tissue rings. Here, we demonstrated in preliminary studies the potential of

hMSC rings for fabricating modular tissue tubes. Tubes fabricated from hMSCs expressed the

Figure 9.4: Alignment of hMSCs within hMSC

tubes. Radial cross-section of hMSC tube after 4

days of fusion. Hematoxylin and Eosin stain.

Lumen on bottom of image. Scale = 100 µm.

Figure 9.5: hMSC tube

with hole. Image at 3

days of fusion culture.

Arrow points to hole.


contractile proteins SMA and SM22-α (Fig 9.3 A, B), although calponin is not visible. The

absence of late- stage differentiation markers is unsurprising, due to the short duration of the

experiment. In an earlier study, we found that when TGF-β1 and BMP-4 were added at the

beginning of ring and tube culture, rings did not fuse successfully and formed holes (not shown).

Thus, we developed the protocol presented in here, where the differentiation factors TGF-β1 and

BMP-4 were added after 4 days of fusion. However, this resulted in only 3 days total of growth

factor treatment for tubes in this experiment, which may explain the limited SMA and SM22 and

absence of calponin (Fig 9.3 C, D).

Cell alignment is also important for creating functional vessels, as radially aligned cells

give tubes the ability to constrict and dilate to regulate blood flow in vivo. In an hMSC tube

cross-section, it is apparent that nuclei are radially aligned around tube outer edges, but not the

middle region of the tissues (Fig 9.4). Alignment may improve in future studies with mechanical

stimulation [6].

Figure 9.6: Morphology and matrix deposition of vascular tissue tubes. Tubes were fabricated from

hMSC rings (A, B) human aortic SMC rings (hAoSMC; C, D), or from hMSC rings with a central region

of human aortic SMCs (E, F). Tubes were allowed to fuse for 7 days prior to fixing and staining for H&E

(A, C, E) or Picrosirius Red/Fast Green (red = collagen, green = counterstain; B, D, F). Scale = 100µm.

Lumen on bottom.


Our ultimate goal is to

create focal regions of a

synthetic SMC phenotype

within vascular tissue tubes,

in order to create a focal

region of vascular disease.

Our original approach was to

utilize gelatin microspheres to

locally deliver growth factors

within the tube. However, due

to the challenges of

microsphere-mediated growth

factor delivery discussed in

Chapter 8, we evaluated an

alternative approach. We

fused primary aortic SMC

rings between hMSC rings,

with the same goal of creating

a focal region of synthetic

SMCs. We have not observed contractile proteins in aortic SMC rings in 3D despite treatment

with TGF-β1. Thus, these rings may also serve as the region of synthetic SMC phenotype,

between contractile protein-expressing hMSC rings. This method may be advantageous for

creating an intimal hyperplasia model, because it ensures that the focal lesion consists of SMCs

with a diseased synthetic phenotype, rather than un-differentiated hMSCs, which may not

respond to therapeutics in the same way as synthetic SMCs.

In these experiments, hMSC and hMSC-lesion tubes began to form holes on day 3. This

is surprising, as our modified protocol with delayed growth factor treatment worked successfully

in preliminary experiments (additional replicate of initial experiment not shown). Despite this,

tubes remained in culture until day 7 of fusion (3-7 tubes), when they were fixed and analyzed

histologically. hMSC tubes produced very limited collagen based on Picrosirius Red/Fast Green

staining. This is highly unusual, as we typically observe large amounts of collagen, even with

Figure 9.7: Contractile protein expression in hMSC and human aortic

SMC tubes. Tubes were fabricated from hMSC rings (A-C), human aortic

SMC rings (hAoSMC; D-F), or from hMSC rings with a central region of

human aortic SMCs (G-I). Tubes were allowed to fuse for 7 days prior to

fixing and staining for SMA (A, D, G), SM22 (B, E, H), or calp (C, F, I).

Green = contractile protein, red = human aortic SMCs dyed with

CellTracker Red dye, blue = nuclei. Scale = 100µm. Lumen on bottom.


delayed growth factor treatment. However, the same results were observed when the experiment

was repeated. If tubes did not have sufficient extracellular matrix to support their structure, it

may explain the hole formation.

Contractile protein expression is limited to a thin layer of SM22 along the tube abluminal

edge, which is visible in both aortic SMC and hMSC regions of the tubes. This may be because

tubes were only subjected to 3 days of growth factor treatment, as hole formation prevented

longer culture times. However, the aortic SMC regions can still be distinguished by their larger

amounts of collagen deposition, which is visible by picrosirius red/fast green staining. Thus, we

have still created a distinct region of aortic SMCs within an hMSC tube. However, more tests

may be necessary to determine why hMSCs in these experiments produced so little collagen and

fewer contractile proteins than previous tests. It is possible there is some experimental variation

between batches of frozen hMSCs. If this problem continues to occur, biochemical stimuli such

as increased ascorbate concentration may enhance collagen deposition.

Human MSCs are highly proliferative, reliably differentiate to an SMC phenotype in 3D

cultures. Additionally, hMSCs may allow for the fabrication of patient-specific tissue tubes to

model vascular disease, as shown in principle in previous work with induced pluripotent stem

cell-derived vascular smooth muscle cells [7]. Future work will include optimizing culture

protocols to improve collagen and contractile protein deposition in hMSCs. Overall, these studies

show the feasibility of creating distinct regions within a vascular tissue tube. This work is a

critical step towards fabricating an intimal hyperplasia model.

9.5. References

1. Liu, J.Y., D.D. Swartz, H.F. Peng, S.F. Gugino, J.A. Russell, and S.T. Andreadis,

Functional tissue-engineered blood vessels from bone marrow progenitor cells.

Cardiovasc Res, 2007. 75(3): p. 618-28.



3. Liang, M.S. and S.T. Andreadis, Engineering fibrin-binding TGF-beta1 for sustained

signaling and contractile function of MSC based vascular constructs. Biomaterials, 2011.

32(33): p. 8684-93.


4. Cho, S.-W., S.H. Lim, I.-K. Kim, Y.S. Hong, S.-S. Kim, K.J. Yoo, H.-Y. Park, Y. Jang,

B.C. Chang, C.Y. Choi, et al., Small-Diameter Blood Vessels Engineered With Bone

Marrow Derived Cells. Annals of Surgery, 2005. 241(3): p. 506-515.




6. Standley, P.R., A. Camaratta, B.P. Nolan, C.T. Purgason, and M.A. Stanley, Cyclic

stretch induces vascular smooth muscle cell alignment via NO signaling. Am J Physiol





Chapter 10: Conclusions and future work 163

Chapter 10: Conclusions and future work

10.1. Summary

Cardiovascular disease is the leading cause of death in the United States [1]. There is a

strong need for new therapeutics to treat these diseases. Tissue engineered bloods vessels

(TEBVs) may serve as tools for disease modeling and high-throughput drug screening, which

may ultimately accelerate the development of lifesaving therapies. Most existing methods for

fabricating TEBVs create homogenous tissue tubes, which are not conducive to modeling focal

vascular diseases, such as intimal hyperplasia or aneurysm. In contrast, our lab has developed a

modular system for fabricating TEBVs from individual tissue ring units. Our ability to customize

individual ring sub-units allows us to introduce spatial heterogeneities along the TEBV length,

and potentially model such focal vascular diseases.

Here, we first demonstrated our ability to incorporate degradable gelatin microspheres in

SMC rings to customize ring properties. Microspheres were loaded with TGF-β1, and caused

increases in contractile protein expression comparable to exogenous TGF-β1 treatment. This

demonstrated our ability to use microspheres to delivery bioactive growth factors within tissue

rings and modulate SMC phenotype.

Next, we fused rings into modular tissue tubes, and verified that cells maintain their

spatial positioning along the length of fused tubes. This is critical for creating focal disease

models, as diseased cells must stay in the diseased region of the tube. We then developed a

custom luminal flow bioreactor, and demonstrated that SMC tubes could be dynamically

cultured, with the aid of PCL cannulation cuffs, at physiological shear stresses. We also

demonstrated that we could create a focal region of microsphere incorporation, by fusing rings

with microspheres between rings without microspheres. This demonstrated our ability to

fabricate tubes with spatially-controlled heterogeneities.

Due to the challenges of differentiating primary human SMCs into a contractile

phenotype, we evaluated induced pluripotent stem cells (iPSCs) as an alternative cell source.

Initial experiments yielded rings that expressed smooth muscle contractile proteins. However,


batch-to-batch variability remained a problem. Next, we evaluated human mesenchymal stem

cells (hMSCs) as an alternative cell source. hMSCs strongly expressed smooth muscle

contractile proteins in response to TGF-β1 and BMP-4. We evaluated the effects of exogenous

and microsphere-mediated delivery of PDGF-BB, FGF-2, and TGF-β1 on hMSC thickness,

proliferation, and contractile protein expression. The effects of microsphere-mediated growth

factor delivery on hMSCs were limited compared to exogenous growth factor treatment, possibly

due to the rapid degradation of microspheres in this system. In the future, modifications to

microspheres will be necessary to achieve a longer degradation time and sustained release.

Our final goal was to create distinct phenotypic regions along the length of tissue tubes.

To do this, we fused human aortic SMC rings, which do not produce contractile proteins, in a

central region of the tube between hMSC rings, which do produce contractile proteins. However,

in this experiment, both cell types produced only very small amounts of contractile proteins.

Still, aortic SMC rings were clearly visible due to increased collagen deposition. This indicates

that we were successful in creating a structurally distinct region within our tissue tubes. Overall,

this work has led to the development of a modular platform technology that may be further

developed to fabricate focal vascular disease models.

10.2. Other applications of the ring system

While the focus of these studies is on ring fusion and the fabrication of modular tubular

structures, rings alone also have great potential as tools for disease modeling and drug screening.

The ISO standard for mechanically testing vascular grafts is to cut them into individual ring-

segments, and test each segment [2]. By fabricating individual rings instead of whole tubular

grafts, we may be able to perform more high-throughput experiments to screen the effects of

culture conditions or potential therapeutics on tissue mechanical properties, contractility,

morphology, and matrix deposition, as rings can be used for mechanical and functional testing,

biochemical analysis, or histological and immunohistochemical analysis. As discussed in

Chapter 7, the Qyang lab at Yale University has used our tissue ring system to create rings from

iPSC-vSMCs derived from patients with SVAS, and showed reduced contractile protein

expression and contractile function with these rings compared to rings made with healthy cells

[3]. Thus, rings alone may be sufficient to model some aspects of vascular diseases, and evaluate

the effects of potential therapeutics.


This modular system serves as a platform technology that can be used to fabricate other

tubular tissues, such as trachea [4, 5]. Trachea consists of alternating cartilage and smooth

muscle tissue, with an inner epithelial layer. Experiments performed in the Alsberg lab at Case

Western Reserve University, in collaboration with our lab, have focused on applying the self-

assembled ring system technology to engineer living human tracheae. Microsphere-mediated

growth factor delivery was used to support the formation of two different tissue ring types,

cartilage and smooth muscle, from hMSCs. ECs were co-seeded with hMSCs in smooth muscle

rings, which formed pre-vasculature structures within the rings. These pre-vascular smooth

muscle rings were alternated and fused with cartilage rings [5]. This work demonstrates the

potential of the ring-microsphere system to create complex multi-tissue constructs with other

tissue types.

10.3. Limitations

The presented work has potential as a tool for disease modeling and high-throughput drug

screening. However, there are several important limitations. The tissue tubes in these studies

consist only of SMCs. Endothelial cells (ECs) play a critical role in maintaining a healthy blood

vessel, and damage to the endothelium is a primary cause of IH [6-8]. ECs secrete nitric oxide

(NO) which prevents SMC proliferation, maintains a contractile SMC phenotype, and serves as a

vasorelaxant [6, 9, 10]. Thus, any vascular disease model is incomplete without both functional

ECs and SMCs.

We also did not include inflammatory or immune cells in this preliminary model, which

play a role in the initiation and progression of intimal hyperplasia [11, 12], and many other

vascular diseases. Following endothelial injury, a cascade of growth factors and inflammatory

cells respond to the injury. Any in vitro model will be limited in its’ ability to evaluate the effect

of these complex interactions and chain reactions. However, this simplicity is also an advantage,

as it may allow researchers to study the effects of single or a controlled number of molecules on

disease progression.

Finally, the model must be validated before it can be used to screen new compounds.

Existing drugs that are known to prevent intimal growth, and drugs that have failed in clinical


trials, must be tested. This is necessary to ensure that the in vitro model will respond in a similar

manner to in vivo human blood vessels.

10.4. Future work

Future work will focus on addressing the above limitations and creating an intimal

hyperplasia model. First, challenges with locally modulating hMSC phenotype must be

addressed. As discussed in Chapter 8, optimizing hMSC differentiation protocols and de-

differentiation protocols, to obtain rings in both contractile and synthetic smooth muscle

phenotypes, will be critical. Modifications to microspheres, such as coatings to delay degradation

and growth factor release, may aid in achieving this goal.

Next, a confluent endothelial layer must be

established. In a preliminary experiment, we demonstrated

attachment of human coronary artery ECs to a tissue tube

fabricated from human aortic SMCs. SMC rings were

cultured for 3 days, then allowed to fuse for 7 days to form

a tube. The tube was mounted onto a luminal flow bioreactor

(described in Chapter 5), and a suspension of ECs was

seeded onto the tube lumen. The bioreactor was mounted

onto a custom hexagonal stand that maintained the horizontal

position of the bioreactor, to prevent ECs from flowing out

of the tube (Figure 10.1). The stand was rotated hourly for 6

hours, then maintained in static conditions overnight in an

incubator to further allow for EC attachment. The tube was

then fixed, processed, sectioned, and stained for the EC

marker von Willebrand Factor. Figure 10.2 shows a layer of

positive staining along the luminal surface of the tube.

However, positive staining was only observed on one side of

the lumen, suggesting uneven attachment around the

circumference of the luminal surface. Further experiments

are needed to optimize cell seeding density and the

frequency of bioreactor rotations in order to achieve a uniform luminal EC coating. Additionally,

Figure 10.2: Endothelialization of

SMC tubes. Green = von Willebrand

Factor, Blue = Hoechst. Lumen on top.

Scale = 100µm.

Figure 10.1: Luminal flow bioreactor

in custom stand for endothelialization.


this experiment was also only performed in static conditions. More experiments are needed to

verify that ECs remain attached in the presence of shear forces, and can form a stable, confluent

monolayer.

After a successful EC layer is established, endothelial and smooth muscle function must

be measured. Smooth muscle tubes must constrict and dilate in response to vasoactive

substances. Molecules such as KCl can be used to assess non-receptor-mediated smooth muscle

contraction, and others such as phenylephrine can be used to measure receptor-mediated

contraction. Acetylcholine testing is a standard method for evaluating endothelial function [13-

15]. In the presence of a healthy endothelial layer, acetylcholine will cause ECs to produce NO,

causing the cells to relax. In the absence of a functional endothelial layer, acetylcholine will bind

to activate muscarinic receptors on SMCs and instead trigger them to contract [15, 16]. We aim

to flow acetylcholine through tissue tube lumens, and measure the change in tube outer diameter.

This will indicate if the tube is constricting or dilating in response to acetylcholine, and

determine if the endothelial layer is functional. SMC function can also be verified with calcium

imaging, to ensure that calcium signaling occurs in response to vasoactive substances.

The system must be further validated by evaluating the effects of known therapeutics on

lesion formation in tissue tubes. Statins (3-Hydroxy-3 methylglutaryl CoA reductase inhibitors)

have been shown in clinical studies to prevent IH, and are one of few drugs approved for this

purpose. While they are commonly prescribed to lower patients’ cholesterol, they also

independently inhibit SMC proliferation, and thus have been shown to prevent excessive intimal

growth [17-21]. We would anticipate that statin treatment would also inhibit proliferation and

lesion formation in our system. Edifoligide is a drug that inhibits the E2F transcription factor,

interrupting the cell cycle and preventing cell proliferation [22]. The drug showed success in a

rabbit model [23] but was not effective in clinical trials when compared to a placebo [24, 25].

Thus, we would anticipate Edifoligide to be ineffective at preventing intimal growth in our tissue

tubes. These are just two examples of drugs that could be tested to validate that our in vitro

intimal hyperplasia model responds to treatment similarly to in vivo human tissues.

The effects of the inflammatory system could be evaluated with the addition of

inflammatory molecules, such as IL-6 or IL-1, or platelets, neutrophils or macrophages to

medium flowing through the isolated vessel lumen. Alternatively, the tube may be implanted into


a mouse model with a human immune system [26], in order to assess lesion initiation and

progression in the context of a fully competent human immune system.

Future work may also include using this system as a platform to create other vascular

diseases, such as aneurysm. Rings may be fabricated from elastin-deficient cells, which may be

fused between rings with normal elastin expression. This could create a localized region of

elastin deficiency, which is a primary cause of aneurysm formation [27]. As discussed earlier, we

have also been using the ring-microsphere system to fabricate engineered trachea [5].

Ultimately, ring and tube production will need to be automated in order to scale up

production for use in high-throughput experiments. Through a collaboration with the Robotics

Engineering Program at WPI, we fabricated a custom well-plate for cell seeding and a robotic

punch, that can automatically remove rings from individual wells and stack them on a mandrel.

This system allows us to fabricate tubes 3.5 times faster than manual tube fabrication. The

custom well-plate and robotic punch are described in detail in Appendix H (manuscript in

review).

The work presented has led to the fabrication of a modular platform technology, that can

be modified in the future to model focal vascular diseases, including intimal hyperplasia or

aneurysm. Such in vitro disease models may serve as tools for screening new therapeutics to treat

cardiovascular diseases, and accelerate the development of new, lifesaving drugs.

10.5. References




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in American National Standard, A.f.t.A.o.M. Instrumentation, Editor. 2010.











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8. Ando, J. and K. Yamamoto, Vascular Mechanobiology Endothelial Cell Responses to

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14. Hoenicka, M., K. Lehle, V.R. Jacobs, F.X. Schmid, and D.E. Birnbaum, Properties of the

human umbilical vein as a living scaffold for a tissue-engineered vessel graft. Tissue

Eng, 2007. 13(1): p. 219-29.

15. Deanfield, J.E., J.P. Halcox, and T.J. Rabelink, Endothelial function and dysfunction:

testing and clinical relevance. Circulation, 2007. 115(10): p. 1285-95.

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muscle cell proliferation and migration. Journal of Vascular Surgery, 2002. 36(1): p.

150-157.

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and T.F. Luscher, HMG-CoA Reductase Inhibition Improves Endothelial Cell Function

and Inhibits Smooth Muscle Cell Proliferation in Human Saphenous Veins. Journal of the

American College of Cardiology, 2000. 36(5): p. 1691–7.

19. Kulik, A., P. Voisine, P. Mathieu, R.G. Masters, T.G. Mesana, M.R. Le May, and M.

Ruel, Statin therapy and saphenous vein graft disease after coronary bypass surgery:

analysis from the CASCADE randomized trial. Ann Thorac Surg, 2011. 92(4): p. 1284-

90; discussion 1290-1.

20. Une, D., A. Kulik, P. Voisine, M. Le May, and M. Ruel, Correlates of saphenous vein

graft hyperplasia and occlusion 1 year after coronary artery bypass grafting: analysis

from the CASCADE randomized trial. Circulation, 2013. 128(11 Suppl 1): p. S213-8.

21. Indolfi, C., A. Cioppa, E. Stabile, E.D. Lorenzo, G. Esposito, A. Pisani, A. Leccia, L.

Cavuto, A.M. Stingone, A. Chieffo, et al., Effects of Hydroxymethylglutaryl Coenzyme A

Reductase Inhibitor Simvastatin on Smooth Muscle Cell Proliferation In Vitro and

Neointimal Formation In Vivo After Vascular Injury. Journal of the American College of

Cardiology, 2000. 35: p. 214–21.

22. Ehsan, A., M.J. Mann, G. Dell'Acqua, and V.J. Dzau, Long-term stabilization of vein

graft wall architecture and prolonged resistance to experimental atherosclerosis after

E2F decoy oligonucleotide gene therapy. J Thorac Cardiovasc Surg, 2001. 121(4): p.

714-22.

23. Mann, M.J., G.H. Gibbons, P.S. Tsao, H.E.v.d. Leyen, J.P. Cooke, R. Buitrago, R.

Kernoff, and V.J. Dzau, Cell Cycle Inhibition Preserves Endothelial Function in

Genetically Engineered Rabbit Vein Grafts. J. Clin. Invest., 1997. 99: p. 1295–1301.





JAMA, 2005. 294: p. 2446-2454.

25. Conte, M.S., D.F. Bandyk, A.W. Clowes, G.L. Moneta, L. Seely, T.J. Lorenz, H. Namini,

A.D. Hamdan, S.P. Roddy, M. Belkin, et al., Results of PREVENT III: a multicenter,

randomized trial of edifoligide for the prevention of vein graft failure in lower extremity

bypass surgery. J Vasc Surg, 2006. 43(4): p. 742-751; discussion 751.

26. Brehm, M.A., L.D. Shultz, and D.L. Greiner, Humanized mouse models to study human

diseases. Curr Opin Endocrinol Diabetes Obes, 2010. 17(2): p. 120-5.


27. Campa, J.S., R.M. Greenhalgh, and J.T. Powell, Elastin Degradation in Abdominal

Aortic Aneurysms. Atherosclerosis, 1987. 65.

Appendix A: Reprint permission for Chapter 2.7 172

Appendix A: Reprint permission for Chapter 2.7

Appendix B: Reprint permission for Chapter 3 173

Appendix B: Reprint permission for Chapter 3

Appendix C: Chapter 3 supplemental data 174

Appendix C: Chapter 3 supplemental data

Supplemental methods

Cell culture

For supplementary experiments, testing was repeated with human coronary artery cells

from a different manufacturer (Lonza). These cells were cultured in SmGm-2 complete medium

(Lonza), containing 5% FBS, 0.1% EGF, 0.2% FGF-B, 0.1% Insulin, 0.1% gentamicin sulfate

amphotericin-B, and was also supplemented with 1% penicillin-streptomycin (Mediatech).

Supplemental figures

.

Figure C.1: Mechanical properties of rings treated with TGF-β1. Sample groups included

untreated rings with no microspheres, rings treated with 10ng/ml exogenous TGF-β1, rings with

unloaded gelatin microspheres untreated or treated with exogenous TGF-β1, and rings with

TGF-β1-loaded microsphere incorporation, but no exogenous TGF-β1. Mean values for (A)

UTS, (B) MTM, (C) failure load, and (D) failure strain were calculated from stress-strain curves

for each sample. *p<0.05. Values are mean ± SD, sample size for each group shown on bars


Figure C.2. Effects of TGF-β1 treatment in smooth muscle cell rings sourced from a

different donor. Rings were seeded in growth medium switched to differentiation medium at

day 1 and cultured for a total of 14 days. Rings were photographed (A-E) before and (F-J)

after removal from the agarose posts to measure changes in ring inner diameter and wall

thickness. (A, F) Untreated control ring with no microspheres. (B, G) Tissue rings treated with

10 ng/ml soluble exogenous (exo) TGF-β1 in the culture medium. Tissue rings with unloaded

gelatin microspheres (0.6 mg/million cells) (C, H) untreated or (D, I) treated with 10 ng/ml

exogenous TGF-β1. (E, J) Tissue rings with microspheres loaded with TGF-β1, but no

exogenous TGF-β1 in the medium. Tissue rings contracted after they were removed from

agarose posts, resulting in changes in (K) inner diameter and (L) thickness. Initial images and

thicknesses were measured using the DVT imaging system (A-E), while secondary

measurements were taken with the stereoscope (F-J). Scale = 1mm. *p<0.05. Values are mean

± SEM, sample size for each group shown on bars.


Figure C.3: Effects of TGF-β1 treatment on smooth muscle cell protein expression in rings self-assembled

from human SMCs from a different donor. Rings were seeded in growth medium switched to differentiation

medium at day 1 and cultured for a total of 14 days. (A, F) Control (untreated) rings. (B, G) Rings cultured with

exogenous TGF-β1 (10 ng/ml) added to the medium. Rings with unloaded microspheres (0.6 mg per million cells)

(C, H) untreated or (D, I) treated with 10 ng/ml exogenous TGF-β1. (E, J), Rings with TGF-β1-loaded

microspheres (0.6 mg microspheres per million cells) but without exogenous TGF-β1. Rings were stained for (A-

E) smooth muscle alpha actin and (F-J) calponin (green fluorescence). Nuclei are shown in blue (Hoechst). Scale

= 100µm.

Appendix D: Reprint permission for Chapter 4 177

Appendix D: Reprint permission for Chapter 4






Appendix E: Chapter 4 supplemental data 183

Appendix E: Chapter 4 supplemental data

Taken from part of: H. A. Strobel, E. L. Calamari, A. Beliveau, A. Jain, and M. W. Rolle, “Fabrication

and characterization of electrospun polycaprolactone and gelatin composite cuffs for tissue engineered

blood vessels.” JBMR Part B, 2018. 106B(2): p. 817-826. Reprinted with permission (Appendix D)

Figure E.1: Assembly of custom grips for longitudinal pull to failure test. (A) 3D printed base. (B) Base pieces

[1] are connected by a screw with PDMS spacers [2] to maintain stability during loading. (C) Next, the bottom of

the clamp [3] is placed over pins on the base piece [1]. (D) Then a tube (represented here with silicone tubing), [5]

mounted over a custom cannula [4] is set in the grooved clamp bottom [3]. (E) After the top part of the clamp [6] is

put in place, it is tightened to the bottom clamp [3] with screws. (F) C-clamps on the Instron tensile testing machine

are attached to the 3D printed clamps [3, 6] while still mounted on the base piece [1]. (G) Then, the orange base

piece [1] is removed and (H) tubes can be pulled to failure. When using the 1N load cell, a binder clip was fastened

to the load cell instead of a C-clamp, due to the sensitivity of the load cell.

Appendix F: Chapter 5 supplemental data 184

Appendix F: Chapter 5 supplemental data

H. A. Strobel, T. A. Hookway, M. Piola, M. Soncini, G. B. Fiore, E. Alsberg, and M. W. Rolle, “Assembly

of tissue engineered blood vessels with spatially-controlled heterogeneities.” Tissue Engineering Part A.

In Press.


Cell culture

Human coronary artery SMCs (Lonza) were cultured and maintained in complete SMC

growth medium (SmGM-2; Lonza) containing 5% FBS, 2ng/ml fibroblast growth factor-basic,

0.5ng/ml epidermal growth factor, insulin, 30µg/ml gentamicin, and 15ng/ml amphotericin B.

Rings were seeded as described in section 2.1.


Figure F.1: Fusion of human SMC rings. Fusion kinetics from duplicate experiments with human aortic

SMCs shown in (A-C), and with human coronary artery SMCs shown in (D-F). Fusion angles (B, D), tube

length (C, E) and thickness (D, F) as a function of time for tubes fabricated from rings cultured for 3, 5, or 7

days prior to 7 days as tubes. * P<0.05 for 7-7 vs 3-7 and 5-7, ** p<0.05 for 5-7 vs 7-7, x = P<0.05 for 3-7 vs 5-

7 and 7-7, and # = P<0.05 for 3-7 vs 7-7 groups, n=3. Dashed line = 180º.


Figure F.2: Fusion of human coronary artery SMC rings. Phase contrast images of 3-7 (A), 5-7 (B), and

7-7 (C) tubes over a 7 day fusion period. Scale = 1 mm. Images representative of n=3 samples per group.

Figure F.3: Fluorescent images of human coronary artery SMC ring fusion. Rings with alternating red and green

cell tracker were allowed to fuse for 7 days (A) prior to sectioning and Hoechst staining. Samples were sectioned after

2 (B-E) or 7 (F-I) days to determine whether cells within ring units maintain their spatial position. Blue = nuclei (B,

F). Red = CellTracker Red (C, G), green = CellTracker Green (D, H), and merged image shown in (E, I). Lumen on

left. Scale = 1 mm (A) or 100 µm (B-I). Images representative of n=3 samples per group per time point.


Figure F.4: Contractile protein expression in aortic SMC tubes. Tubes fabricated from

human aortic SMCs were either kept in static conditions for 7 days (A), or kept in static

conditions for 7 days followed by 7 days of dynamic culture with approximately 12 dyne/cm2

of applied shear stress (B). Green = smooth muscle alpha actin, blue = nuclei. Lumen on

bottom of image. Scale = 100 µm.

Appendix G: Microsphere characterization 187

Appendix G: Microsphere characterization

For all experiments presented in this dissertation, cross-linked gelatin microspheres were

prepared and characterized by the Alsberg Lab at Case Western Reserve University. Microsphere

size and crosslink density analysis are presented in Table F.1. Microspheres were prepared and

characterized according to published protocols described in:

Dikina AD, Strobel HA, Lai BP, Rolle MW, Alsberg E. “Engineered cartilaginous tubes for

tracheal tissue replacement via self-assembly and fusion of human mesenchymal stem cell

constructs.” Biomaterials. 2015;52:452-62.

Batch

Average

Diameter (µm) Crosslink Density

(%) Experiments

1 47.5 ± 42.7 32.6 ± 6.1 Chapter 3 (Microsphere incorporation)

2 48.4 ± 41.9 35.7 ± 15.4

Chapter 3 (TGF-β1 delivery), Chapter 6

(Microsphere lesion with coronary artery

SMCs), Chapter 7

3 59.3 ± 28.8 32.3 ± 15.3

Chapter 6 (Microsphere lesion with aortic

SMCs, PDGF release)

4 51 ± 16 15

Chapter 6 (Fusion comparison), Chapter 8

(PDGF release)

5 50 ± 32 60 ± 7 Chapter 8 (FGF and TGF- β1 release)

Table G.1: Characterization of gelatin microspheres used for each experiment

Appendix H: Supplemental data for Chapter 8 188

Appendix H: Supplemental data for Chapter 8


Rings were prepared and analyzed as described in Chapter 8. Rings contained either

unloaded microspheres, with or without 5 ng/ml exogenous FGF (Peprotech, instead of Cell

Signaling as in Chapter 8), or with microspheres pre-loaded with 200 ng FGF per mg

microsphere.


Figure H.1: Effect of FGF treatment on ring thickness. Thickness measurements of rings at 3, 7, and 14 days

with unloaded microspheres and no FGF treatment (No FGF) or 5 ng/ml exogenous FGF (Ex FGF), or containing

microspheres loaded with 200 ng FGF per mg microspheres (FGF MS, A). Representative DVT images of rings in

each group shown in (B). * P<0.05 compared to no FGF and FGF MS groups within time point. Two-way ANOVA

with Holm-Sidak Post Hoc test. N = 3 for days 3 and 7, n = 5 for day 14. Bars are mean ± SEM. Scale bar = 1 mm


Figure H.2: Collagen deposition in rings with FGF treatment. Rings were

cultured for 3 (A-C), 7 (D-F), or 14 (G-I) days. Rings containing unloaded

microspheres were treated with no FGF (No FGF, A, D, G), 10ng/ml exogenous

FGF (Ex PDGF, B, E, H), or contained microspheres loaded with 400 ng FGF per

mg microspheres (FGF MS, C, F, I). Picrosirius Red/Fast Green stain (red =

collagen, green = counterstain). Lumen on bottom/right. Scale = 100µm.

Figure H.3: Proliferation in rings with FGF treatment. Rings containing

unloaded microspheres were treated with no FGF (No FGF, A-C), 10ng/ml

exogenous FGF (Ex FGF, D-F), or contained microspheres loaded with 400 ng FGF

per mg microspheres (FGF MS, G-I). Rings were stained at day 3 (A, D, G), 7 (B,

E, H), or 14 (C, F, I). Ki67 immunostain stain (brown = Ki67, purple = nuclei).

Lumen on bottom/right. Scale = 100µm.


Figure H.4: Contractile protein expression in FGF treated rings. Rings containing

unloaded microspheres were treated with no FGF (No FGF, A-C), 5ng/ml exogenous

FGF (Ex FGF, D-F), or contained microspheres loaded with 200 ng FGF per mg

microspheres (FGF MS, G-I). Rings were stained at day 14 for SMA (A, D, G), SM22-

α (B, E, H), or calponin (C, F, I). Green = contractile protein, blue = nuclei. Lumen on

bottom/right. Scale = 100µm. Rings are 14 days old.

Appendix I: Automation and scale-up of tissue tube production 191

Appendix I: Automation and scale-up of tissue tube production

C. J. Nycz, H. A. Strobel, K. Suqui, G. S. Fischer, M. W. Rolle, “A Method for High-Throughput

Automated Assembly of 3-Dimensional Vascular Tissue.” In Review.

Authorship contributions: CJN designed and built robotic punch, aided in design and fabrication of well-plates,

made schematic figures, co-wrote and edited manuscript. HAS contributed to design of robotic punch and well-

plates, performed fusion experiments, supervised KS and co-wrote manuscript. KS performed ring testing

experiments, contributed to well-plate design, and edited manuscript. GSF and MWR contributed to experimental

design, supervised data collection, data analysis, and preparation of the manuscript, and edited the manuscript.

CJN and HAS share first author credit.

Abstract

An essential step towards commercializing engineered tissues is to scale-up and automate

their production. This presents a challenge for self-assembled tissue systems, which are fragile at

early time points and difficult to handle using automated systems. The goal of this study was to

automate tissue engineered blood vessel (TEBV) fabrication by creating a custom cell seeding

and self-assembly system that is conducive to robotic manipulation, coupled with a robotic

system to assemble smooth muscle cell ring units into tissue tubes. To generate self-assembled

tissue ring units manually, cells are seeded at a high density into custom agarose wells that have

center posts (2 mm inner diameter), around which cells aggregate and contract to form rings.

Agarose is well-suited for cell seeding and ring self-assembly because it is non-cell adhesive, can

be autoclaved, and reproducibly cast to form wells using silicone templates. However, agarose

gel is soft, which makes reliable robotic manipulation challenging. To solve this problem, we

designed a custom ring self-assembly plate utilizing a polyetherimide (PEI) well plate with

MED610 3D-printed center posts and a MED610 well negative that allows the casting of

individual, annular agarose cell-seeding troughs within the PEI plate. Rings cultured in the new

plate system were morphologically similar to rings cultured in control agarose gels, and had a

slightly higher failure load. We created a unique robotic punch system to push tissue rings out of

the PEI-MED610 plate onto a stainless-steel mandrel to enable tube fusion. Tubes fabricated via

manual or automated ring removal and placement demonstrated similar morphology after tube

fusion, and the automated system substantially reduced the time required to fabricate tubes

manually. In summary, we developed a novel robotic assembly system to precisely manipulate

self-assembled tissue rings and enable scale-up and automation of TEBV biofabrication.


I.1. Introduction

Tissue engineered constructs have enormous potential as both implantable grafts, or as

tools for disease modeling and high throughput drug screening. A key step for translating these

technologies into marketable products is automating and scaling-up their production. Automation

can not only significantly reduce production time and increase output, but can also reduce human

error and improve product consistency. Technology has been developed for automated cell

seeding, passaging, and medium changes for 2D cell cultures, which are the first steps to creating

engineered tissue [1]. However, there are limited technologies available for scaling-up and

automating the fabrication of 3D engineered tissues.

Bio-printing is commonly used in additive manufacturing to create 3D tissue units [2-5].

With bio-printing, a wide range of scaffold-based cell-laden bio-inks [5, 6] or scaffold-free cell

spheroids [4] can be precisely patterned directly onto a surface. Layer-by-layer printing

approaches may allow for even more complex shapes, or multi-tissue structures [4, 7]. A major

disadvantage of bio-printing is that most machines have a limited resolution, and may not be able

to print small or micro-tissues [7]. In addition, bio-printing frequently relies on the fusion of

smaller tissue units into larger structures after printing. Spheroids are the most commonly used

tissue sub-units for both bio-printing and other additive manufacturing techniques because they

are relatively simple to fabricate [4, 8, 9]. However, constructs fabricated from even tightly

packed spheroids often have remaining gaps after fusion, as spheroidal units cannot be pushed

into complete contact with one another [4, 10, 11].

A major challenge with additive manufacturing of tissue engineered constructs is the

ability to lift individual tissue units and precisely place them together to build a complex

composite tissue structure. This may be especially challenging at early time points in culture,

when tissue is fragile. However, tissue units harvested at earlier time points are known to have

improved tissue fusion compared to more mature tissue units [12, 13], so it is essential that tissue

building blocks can be manipulated early in culture. Recently, robotic systems have been

developed that can precisely pick up and place self-assembled tissues [14, 15]. However, the

system is not completely automated, and the user must still manually place the tissue using a

series of positioning knobs. Additionally, it was difficult to stack toroidal tissue units precisely


enough for seamless tissue fusion, and individual tissue unit borders were still visible after 3

days of culture [14].

Additive manufacturing may be advantageous for fabrication of scaffold-free, self-

assembled tissues. Without any scaffold material, entirely cell-based tissues may not have the

structure needed to support a large construct, until they have secreted sufficient extracellular

matrix proteins. Creating smaller sub-units, and later fusing those sub-units, may aid in the

production of larger tissues. Cellular self-assembly approaches have advantages over scaffold-

based approaches, due to their biocompatibility, enhanced cell-cell and cell-matrix interactions,

and physiologically relevant mechanical properties [16-19]. These characteristics make cellular

self-assembly ideal for fabricating many different tissue types, including tissue engineered blood

vessels (TEBVs) [19-22]. Our lab has developed a unique system for creating self-assembled

TEBVs from individual tissue-ring subunits [23, 24]. Briefly, cells are seeded into an agarose

well with a center post, where they aggregate together to form vascular tissue rings. Vascular

tissue ring units can then be threaded onto a mandrel, pushed together, and fused into tubes [23,

25], as shown schematically in Figure I.1. Rings may fuse more seamlessly than spheroids, as

greater portions of ring surface area are in contact with one another, making them more

conducive to TEBV

fabrication.

Our overall goal

is to develop a

completely automated

robotic assembly

system to scale up the

fabrication of TEBVs

for commercial

purposes (Figure I.2), for use as either implantable grafts or as tools for high-throughput drug

screening. In this study, we focus on developing a means to automatically remove rings from

agarose gels and thread them onto mandrels for fusion. There are currently no other systems

available that are suitable for this purpose.

To do this, we first re-designed the agarose gels used for ring self-assembly. Agarose is

well-suited for enabling cellular aggregation due to its non-cell-adhesive properties, but the

Figure I.1: Manual method for fabrication of self-assembled vascular tissue

rings and tubes. Cells are seeded into individual ring-shaped agarose wells, where

they aggregate to form self-assembled tissue rings. Tissue rings are manually

harvested and threaded onto silicone tubing mandrels, where they remodel and fuse

together to form a tissue tube.


softness of agarose makes it challenging to grip and manipulate consistently using conventional

robotic systems. The first goal of this study was to develop a new cell seeding well system that

would enable both self-assembled ring formation, and robotic manipulation. We used a

combination of traditional subtractive machining and additive 3D printing techniques to

prototype a unique multi-component plate system with a rigid base that is conducive to both cell

aggregation and automated handling. The plate system contains a polyetherimide (PEI, trade

name Ultem 1000) base machined to the dimensions of a standard 96 well plate, with individual

wells. The bottom of the plate was 3D printed from a MED610 photopolymer (Stratasys Ltd.),

which contained posts (2 mm ID) that fit into the center of each PEI well. A MED610 negative

was also printed, which enabled agarose troughs to be cast in each well to facilitate cellular self-

assembly. The complete plate assembly is shown schematically in Figure I.3.

We evaluated the effect of the custom plate on tissue ring morphology and mechanical

properties as compared to rings cultured in typical agarose gel wells. Then, we used the PEI-

MED610 plates to test the feasibility of using a custom robotic punch to harvest tissue rings

directly from the plates and stack them onto a stainless-steel mandrel, a normally labor and time-

intensive process when performed by hand. The morphology of tubes cultured in PEI-MED610

plates stacked using the automated system was then compared to manually stacked tubes.

Overall, this novel plate and robotic punch system enabled automated fabrication of tissue tubes

Figure I.2: Overview of proposed method of automated assembly of TEBVs fabricated from self-

assembled vascular tissue rings.


from individual ring sub-units.

I.2. Methods

I.2.1. Mold design

The new PEI-MED610 plate system consists of a custom 96-well plate, a center-post

plate inserted from the bottom, and a removable well negative for casting agarose cell seeding

troughs (Fig. I.3). The 96-well plate is machined from PEI (UltemTM 1000, SABIC,) plastic, with

dimensions based on

ANSI SLAS 4-2004

(R2012) standards for a

96 well microplate. PEI

plates contain bottomless

wells with a 6mm

diameter to match the

dimension of our control

agarose gel system [23,

24], that flare out to

9mm at the top of the

well to match the 96 well

microplate footprint. An array of 2mm center posts that fit within the wells of the PEI plate was

3D printed from MED610 (Stratasys Ltd.) material using a photopolymer printer (Object Connex

260, Stratasys Ltd.). The well negative is also printed from MED610 material, which, when

inserted in the PEI plate, allows agarose troughs to be cast within the PEI plate wells. For the

prototype testing presented in this study, the 96-well plate format was scaled down to 16 wells (2

rows) to reduce the amount of reagents and cells required.

I.2.2. Cell culture

Rat aortic smooth muscle cells (WKY-3M22 [26]) were cultured in medium comprised of

Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum, 1%

non-essential amino acids, 1% L-glutamine, 1% sodium pyruvate, and 1% penicillin-

streptomycin.

Figure I.3: PEI-MED610 plate system. A MED610 center post plate fits under

the open-bottomed PEI 96 well plate. A MED610 negative sits on top of the 96

well plate, to allow casting of rounded-bottom agarose troughs in each well.


I.2.3. Agarose gel preparation

For the control configuration in this study, agarose gels were fabricated as described

previously [24]. Two percent agarose (Lonza) in DMEM was autoclaved and poured into PDMS

templates. When cooled, each agarose gel was removed from the PDMS, placed into a well of a

6 well plate, and allowed to equilibrate overnight in culture medium prior to use. Each agarose

gel contained wells for 5 individual rings (2mm ID; Figure I.1) [24].

To prepare PEI-MED610 plates for the proposed automated approach, the assembled

plate was ethylene oxide sterilized and allowed to de-gas 48 hours prior to use. A solution of 3%

Low Melting Temperature Agarose (Lonza) in DMEM was autoclaved and allowed to

equilibrate for 30 minutes in a 45ºC water bath. This allowed the agarose to cool to a temperature

below 52°C to avoid damaging the 3D printed MED610 pieces. Agarose was then carefully

pipetted into the assembled PEI-MED610 plate system and allowed to solidify. The well

negative was removed, and a hydrophobic 1% Pluronic F-127 (Thermo Fisher) solution was

pipetted into wells to reduce cellular adhesion to the PEI components in the plate wells. After

incubating 15 minutes at room temperature, the Pluronic was aspirated and the well plate was

equilibrated overnight in culture medium prior to use.

I.2.4. Ring fabrication

Rings were seeded as described previously [24]. Briefly, cells were trypsinized and

resuspended at a final concentration of 10 million cells per ml. 500,000 cells per ring were

seeded into each well of either control agarose gels or PEI-MED610 plates. On day 1, wells are

flooded with fresh culture medium. Culture medium was changed after two days.

I.2.5. Mechanical testing

After 4 days of culture, rings were removed from molds and placed in a dish filled with

phosphate-buffered saline (PBS). Rings were placed under a machine vision system (DVT series

600-Model 630), and thickness measurements were taken in 4 locations around the ring

circumference with edge detection software (Framework 2.4.6., DVT Corp.) [21]. Average ring

thickness was used to calculate ring cross-sectional area. Rings were then loaded onto a custom

grip setup submerged in PBS, subjected to 8 pre-cycles from 1-5 mN, and then pulled to failure


at a rate of 10mm/min using an ElectroPuls E1000 (Instron). Failure load and ultimate tensile

stress (UTS, failure load/ring cross sectional area) were recorded.

I.2.6. Histology

After 4 days of culture, rings were fixed for 1 hour in 10% neutral buffered formalin,

processed, and paraffin embedded. 5µm sections were prepared and adhered to charged glass

slides. A Hematoxylin and Eosin stain was used to visualize tissue morphology, and a Picrosirius

Red/Fast Green stain was used to visualize extracellular matrix deposition. Images of stained

tissue sections were acquired with an upright microscope (Leica DMLB2 with DFC 480 digital

camera).

I.2.7. Robotic punch design

To extract self-assembled tissue rings from the PEI plates and assemble them into a

tubular construct, we developed an automation platform controlled by a simple graphical user

interface. The system indexes to each well of the PEI-MED610 plate, lowers a 316 stainless steel

mandrel over the well's center post, and punches 4 titanium rods up through the agarose troughs

to thread the ring onto the mandrel. Steel mandrels have a thin coating of USP class VI epoxy

(EP42HT-2MED, Master Bond Inc.) applied to the tip to help prevent rings from sliding off after

automated stacking. A diagrammatic representation of the ring extraction process is shown in

Figure I.4. The platform uses two high resolution linear ball screw stages with ±3μm

repeatability (404100XRMS,

Parker Hannifin Corporation)

to position a bottomless tray

holding the PEI-MED610 plate

between punch and mandrel.

The punch and mandrel are

mounted to two pneumatic

linear slides (13-MXS16-30,

SMC Corporation) allowing

motion in the vertical direction.

To ensure rigidity, the frames

Figure I.4: Schematic of robotic process to remove tissue rings from a

PEI-MED610 plate. A 4-prong punch pushes through a cast agarose

trough and moves self-assembled tissue rings onto a stainless steel mandrel.

As the mandrel is lifted away from the well, an aluminum agarose remover

removes any remaining agarose while the tissue ring stays in place.


supporting the linear slides and PEI-MED610 plate were machined from aluminum with system

components mounted to a thick polycarbonate base. All components fit inside a standard tissue

culture hood. Images of the robotic assembly system are shown in Figure I.5. All components

that come in direct contact with the PEI-MED610 plate system are removable and autoclavable.

After autoclaving, parts are re-attached inside the cell culture hood with sterile surgical gloves.

To control the system, a graphical user interface created in Matlab2016a (Mathworks

Inc.) outputs tray positions and punch commands over RS-232 serial communication to a 2-axis

motion controller (Compumotor 6K2, Parker Hannifin Corp.). Magnetic limit switches are used

for the homing calibration process of the positional stages and for checking the current positions

of the pneumatic slides. Absolute position commands are given to the controller based on a

measured offset between the home position and the first well. Pneumatic slide positions are used

for error checking, locking out the system from indexing while punch or mandrel are not clear of

the PEI-MED610 plate.

I.2.8. Tube fabrication

Smooth muscle cell rings were cultured for 4 days in either control agarose gels or PEI-

Figure I.5: Robotic assembly system in cell culture hood. A positional stage moves the PEI-MED610 plate

over a 4-prong punch. The punch pushes rings out of the well and up onto a stainless steel mandrel. For testing,

a shortened 16-well plate format was used to reduce the amount of reagents and cells required to evaluate

iterative prototype changes.


MED610 plates prior to harvesting for tube fabrication. PEI-MED610 plates were loaded onto

the robotic platform inside a cell culture hood, and automatically punched out of the plate and

onto a stainless-steel mandrel. A brief 10 second dwell period was included between the removal

of each ring from its well and the punch retraction, to allow rings to contract around the mandrel.

Rings in the PEI-MED610 plate and on the mandrels were periodically hydrated with culture

medium using a pipet. Mandrels with stacked rings were then loaded into custom polycarbonate

holders for fusion culture [21]. For control tubes, rings are manually threaded with forceps onto

either silicone mandrels as previously described [21], or onto the stainless steel mandrels that

were used for the automated procedure and secured in custom polycarbonate holders, to evaluate

the effect of mandrel material on ring fusion. All tubes were then allowed to fuse in culture for 4

days prior to fixing 1.5 hours in 10% neutral buffered formalin, followed by processing and

sectioning for histological staining.

I.2.9. Statistics

Statistical analysis was performed using SigmaPlot Software (Systat, version 12.5) on

ring thickness, failure load, and ultimate tensile stress. A student’s t test was performed to

analyze statistical differences between mechanically tested rings on data with a normal

distribution. A Mann-Whitney test was used to compare data that failed a normality test.

I.3. Results

I.3.1. Ring fabrication in PEI-MED610 plate system

Because agarose gels are soft and challenging to manipulate with robotic systems, we

developed a new 3-part plate system utilizing MED610 3D printed posts and plate negative, and

PEI machined 96-well plate. To ensure the change in well material and format did not adversely

affect cellular self-assembly, ring morphology, and tissue mechanical strength, we compared

rings cultured in control agarose gels to rings cultured in the PEI-MED610 plates (Fig I.6A). We

observed no significant differences in ring thickness (0.42 ± 0.02 mm and 0.44 ± 0.04 mm, rings

cultured in agarose vs PEI-MED610 plates, respectively; Fig I.6B). There was a slight increase

in ultimate tensile stress (UTS; 30.5 ± 7.9 and 45.7 ± 14.9 kPa for rings cultured in agarose vs.

PEI-MED610 molds; Fig I.6C) although this was not significant. There was significant increase

in maximum load at failure (failure load) when rings were cultured in the PEI-MED610 plates


(8.6 ± 2.1 and 14.2 ± 4.6 mN for

rings cultured in agarose vs.

PEI-MED610 plates, Fig I.6D).

When examined

morphologically, no visible

differences were observed

between rings cultured in

control agarose gels or PEI-

MED610 plates, either

macroscopically (Fig I.7 A, B)

or by Hematoxylin and Eosin

staining (Fig I.7 C, D). Collagen

deposition was visible in both

groups, with no apparent

differences (Fig I.7 E, F).

I.3.2. Automation system

For testing the robotic punch, two PEI-MED610 plates were used. When preparing PEI-

MED610 plates, 10 out of 16 agarose troughs successfully formed on the first plate, and 14 out

of 16 on the second. Rings were seeded in all plate wells with agarose troughs, although one ring

on each plate broke within 24 hours of seeding. From the first plate, 6 out of 9 self-assembled

tissue rings were successfully pushed onto the mandrel; 5 out of 13 were successfully pushed

onto the mandrel from plate 2. During pilot testing, we determined that a 10 second pause after

pushing the ring onto the mandrel, before retracting the punch, helped prevent rings from sliding

off the mandrel when the remnant agarose gels are removed. Still, some rings did slide off the

mandrel, which contributed to this reduced yield. Additionally, some rings broke during removal

from the PEI-MED610 plate. Four out of 28 rings failed during manual stacking of control rings

and could not be used. Ring failure rates are summarized in Table I.1.

Figure I.6: Structure and strength of tissue rings grown in PEI-

MED610 plates compared to control agarose gels. Images of 4 day-old

rings cultured in agarose gels (left) or PEI-MED610 plates (right) shown

in (A). Comparisons of ring thickness (B), ultimate tensile stress (C), and

maximum load at failure (D). * P<0.05. A non-parametric t-test was used

for (B) and (C), and a Mann Whitney test in (D). Scale bar = 1mm. N = 6

for rings cultured in agarose controls, n = 8 for rings cultured in PEI-

MED610 plates.


Table I.2: Failure rates of tissues rings stacked manually or automatically.

Total rings

seeded

Ring failures

during culture

Ring failures during

stacking

Time required to

stack 96 rings

Control agarose gels 28 0 4 3.49 hours

PEI-MED610 plates 24 2 11 0.45 hours

After initial setup, the robotic assembly system

reduced the time required to remove rings from their

cell seeding molds and thread them onto a mandrel to

fabricate tissue tubes. To manually stack 28 rings onto

2 silicone mandrels (including 4 rings that broke during

manual placement) took approximately 61 minutes,

which averages to approximately 2.2 minutes per ring.

This is compared to 17 seconds per ring to remove and

stack each ring using the automated system. These

times include time lost due to failed rings. For

reference, on a 96-well plate this translates to 0.45

hours for the automated approach and 3.49 hours for

the manual approach. Because of this increased speed,

even when accounting for the observed 54% ring

failure rate, the PEI-MED610 plates can still stack

approximately 3.5 times as many rings as a human in

the same amount of time.

I.3.3. Tube fusion following automated ring stacking

To evaluate if either the automated stacking

procedure or stainless-steel mandrels affected ring fusion, 5-6 rings/tube (n = 2 tubes per group)

were fused for 4 days following automated stacking or manual stacking onto stainless steel or

silicone mandrels. Photographs of tubes after fusion are shown in Figure I.8A-C. Longitudinal

sections of fixed tubes (Fig I.8D-F) show that rings in all three groups have fused, although ring

Figure I.7: Morphology of self-assembled

tissue rings cultured in agarose gels or PEI-

MED610 plates. Photographs of tissue rings

in cell seeding wells (A) and (B), PEI plate has

been removed from (B), so only rings around

MED610 posts are shown. Ring sections were

stained with Hematoxylin and Eosin (C, D) or

Picrosirius Red/Fast Green (E, F; red =

collagen, green = counterstain). Rings in (A)

and (B) are 2mm ID. Scale = 100µm (C-F).

Sections representative of n = 3 rings. Arrows

point to rings (A, B).


boundaries are still distinct. Rings and tubes appeared slightly longer in the PEI-MED610 group.

A slight growth of cells up the PEI posts was observed, causing a slight increase in ring “height,”

which may have contributed to the increased tube length.

I.4. Discussion

Automation is key to scaling up the production of human tissues for commercial use,

either as implantable grafts or as tools for high throughput drug screening. Here, we developed a

unique tissue ring self-assembly plate, and robotic tube assembly system, and demonstrated the

feasibility of automatic assembly of modular TEBVs fabricated from individual ring units.

The first goal of our study was to develop a new well system for seeding smooth muscle

cell rings that both had a rigid frame conducive to robotic manipulation, and enabled cellular

self-assembly. Thus, we developed a 3-part plate system using a PEI plate with individual open-

bottom wells, 3D printed MED610 center posts, and a MED610 well negative that enabled the

casting of agarose troughs in the PEI plate wells. In addition to providing a non-adhesive cell

seeding and ring self-assembly surface, the agarose troughs also enabled ring harvesting by

punching through the agarose well bottoms to push the rings out of the plate and onto the

stainless steel mandrel in the robotic assembly system.

Figure I.8: Fusion of automatically or manually fabricated tissue tubes. Tissue rings were either

automatically stacked onto stainless steel mandrels using the robotic assembly system (A, D), or manually

stacked onto stainless steel (B, E) or silicone (C, F) mandrels. Photographs of tubes after 4 days of fusion

(A-C). Hematoxylin and Eosin stain of fused tube tissue sections (D-F). Scale = 0.5mm. N = 2.


The described PEI-MED610 plate system is a result of several design iterations. Initially,

all parts were fabricated from MED610 photopolymer, due to the one-step 3D printing process,

and its status as a USP plastic class VI, indicating it is safe to be in contact with tissues [27].

After seeding cells in the MED610-only mold, we observed cells failing to self-assemble into

tissue rings, possibly because they adhered to the material. Thus, we developed a hybrid system,

and used a PEI thermoplastic for the bulk of the plate system. PEI was chosen due to its

availability, machinability, and previous use in culture of other cell types [28]. We continued to

use 3D-printing to generate the MED610 post insert plate, as the shape of this piece could be

made easily with other prototyping methods, and the small amount of MED610 in the posts did

not appear to affect ring formation. A 3D-printed MED610 negative that fit over the plate

assembly enabled us to cast agarose troughs in the bottoms of each well in the assembled plate,

which are non-cell adhesive and enable cellular self-assembly. While rings successfully formed

with this method, they still adhered slightly to the MED610 posts, which made it challenging to

remove them from the wells. This issue was addressed by adding a Pluronic coating in our

current prototype, which prevents cell adhesion to materials [29].

After implementing the PEI-MED610 plate system, we evaluated the effects of the new

plate materials on ring formation, morphology, and mechanical strength. Overall, both groups

had a low failure load and UTS, which is likely due to the short 4-day ring culture duration used

in this study (compared to previous mechanical testing studies of 2 mm rat smooth muscle cell

tissue rings, typically cultured for at least 14 days [21, 23]). With the PEI-MED610 plate system,

we observed a significant increase in maximum load at ring failure compared to rings cultured in

agarose gels. It is possible that the larger quantity of culture medium required for the new culture

system contributed to this improvement. The PEI-MED610 plates are placed in a glass dish for

culture, and required approximately 70 ml of medium to cover a plate containing 16 rings. This

is substantially more than our agarose gel system [30], which uses 4.5ml medium in each well of

a 6 well plate, which covers 5 rings in an agarose gel. Most importantly, this indicates that the

PEI-MED610 plates did not adversely affect ring formation or mechanical properties, and can be

used as an alternative to pure agarose gels for culturing tissue rings for automated assembly.

The presented robotic assembly system is advantageous due to its simple design and

compatibility with existing standards for lab automation. The device requires only a


commercially available 2-axis positioning stage and two commercially available linear slides

with no additional moving linkages, degrees of freedom, grippers, or extruders. Other systems

for manipulating engineered tissue are limited, and may be significantly more expensive and

complex to build.

This design was well-suited for the task of tissue ring extraction and stacking, and

substantially reduced the amount of time required to fabricate tubes compared to stacking by

hand. However, some future optimization is still required to reduce failure rates. For example,

despite the thin epoxy layer on the mandrel tip (to slightly increase friction), failures sometimes

resulted when rings slid off the mandrel, and followed the punch back down into its’ original

well. This could be addressed in future work by optimizing the mandrel shape and surface finish

to prevent slipping. A small number of rings also failed during punching. This may be improved

by optimizing culture conditions to increase ring strength. Despite these failures, we were still

able to precisely move 3.5 times as many rings onto mandrels as a human can manually stack in

the same amount of time.

After automated or manual stacking of rings, rings were allowed to fuse for 4 days in

culture. After 4 days, we observed ring fusion regardless of mandrel type or stacking method.

Ring boundaries are still visible in all groups, which is consistent with our previously published

work with rat aortic smooth muscle cells [23]. Fusion may improve with longer culture times, as

tubes were only allowed to fuse for 4 days (following 4 days of ring culture, for 8 days total

culture). Boundaries are primarily visible on the luminal side, which may improve if tubes are

cultured with luminal flow later in culture. In longitudinal tube sections, rings that had been

cultured in the PEI-MED610 plate appeared longer than rings cultured in agarose gels. While we

did not observe differences in ring thickness when viewed from above, rings did appear to grow

out along the length of the MED610 posts, resulting in slightly increased ring height compared to

agarose wells with agarose center posts. However, this does not appear to have negatively

affected ring fusion, mechanical properties, or our ability to remove rings from the PEI-MED610

plate system. Such rings may even enable us to make longer tubes without the need for

additional cells and reagents.

Another challenge to automated tissue fabrication is maintaining sterility. After 4 days of

fusion culture post-automated stacking, there was no visible contamination, indicating that


autoclaving machined parts that came in direct contact with the tissue rings and culture wells was

sufficient to maintain sterility.

Future work will involve automating other components of the tube fabrication process,

such as cell seeding and medium management. PEI-MED610 plates are designed to mimic the

dimensions of a standard 96 well plate, which will enable the use of these existing automation

technologies for this purpose [1]. The stacking procedure presented here is perhaps the most

challenging step in fully automating production of engineered blood vessels from ring modules.

Here, we demonstrated the feasibility of using a robotic punch to push self-assembled tissue

rings up out of a custom designed hybrid PEI-MED610 plate and onto a steel mandrel, where

they fuse to form a tissue tube. This automatic system substantially reduced time required to

fabricate tubes, and is a critical step for creating a fully automated tissue tube fabrication system.

Scaling-up the tube fabrication process allows for increased production and may improve

product consistency, which is an essential step towards commercializing any tissue engineered

product. The PEI-MED610 plate and robotic punch system is a step towards complete

automation and scale-up of vascular graft fabrication, which may be applied more broadly to

biofabrication of other tubular tissues, such as trachea [31, 32].

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