The Roles of the High and Low Molecular Weight Isoforms of Fibroblast Growth Factor 2
in Ischemia-Induced Revascularization
A dissertation submitted to the
Graduate School
of the University of Cincinnati
in partial fulfillment of the
requirements for the degree of
Doctor of Philosophy
in the Department of Pharmacology and Cell Biophysics
of the College of Medicine
by
Adeola Adeyemo B.S. University of Illinois at Chicago
March 2016
Committee Chair: Jo El J. Schultz, Ph.D.
ABSTRACT
Cardiovascular diseases are the underlying cause for majority (>30%) of deaths worldwide.
They coronary heart disease and peripheral artery disease, conditions characterized by limited
blood flow and inadequate oxygenation. Treatment strategies include management of symptoms
and risk factors, reduction of oxygen demand and surgical revascularization to increase circulation.
Therapeutic revascularization is a potential alternative for patients who are poor candidates for
these interventions due to advance disease or co-morbidities. Revascularization involves the
genetic or pharmacologic stimulation of vascular growth processes to facilitate tissue perfusion
and promote functional recovery. Adaptive vascular growth occurs via capillary vessel growth
(angiogenesis) and growth or remodeling of collateral arteries (arteriogenesis). Delivery of angiogenic factors such as fibroblast growth factor 2 (FGF2) to ischemia tissues can stimulate blood vessel growth.
FGF2 consists of two classes of protein isoforms generated from alternative translation of the Fgf2 gene, high molecular weight (HMW) FGF2, and low molecular weight (LMW) FGF2.
Proof-of-concept studies in animal models of chronic ischemia provided evidence for the therapeutic potential of exogenous LMW FGF2. This promise, however, did not translate into successful clinical use. Currently, the functions of the endogenous FGF2 isoforms in ischemia- induced revascularization are not well understood. Elucidating the function(s) of the FGF2 isoforms in vascular growth is of great clinical importance and may lead to the development of novel pharmacological therapies for ischemic diseases.
Mice with a targeted deletion of all FGF2 isoforms (Fgf2-/-), HMW FGF2 (FGF2 LMW- only) and LMW FGF2 (FGF2 HMW-only) were employed to identify the distinct role(s) of the
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FGF2 isoforms in chronic ischemia. Revascularization was evaluated in mice subjected to chronic
hindlimb ischemia using measures of limb function, tissue viability, and vessel growth. FGF2
HMW-only mice had a faster recovery of limb function compared to wildtypes. Fgf2-/- and FGF2
LMW-only mice, however, had a significantly slower recovery of function. Additionally, no limb
necrosis was detected in FGF2 HMW-only mice while Fgf2-/- and FGF2 LMW-only mice had significant necrosis of their ischemic limbs. The early recovery of limb function in FGF2 HMW- only limbs was preceded by improved revascularization (angiogenesis and arteriogenesis). Vessel growth was significantly decreased in FGF2 LMW-only muscles and not different from wildtypes in Fgf2-/- mice, indicating that the presence of LMW FGF2 inhibited vascular growth. The effect of the HMW FGF2 isoforms on revascularization and protection from ischemic injury was not associated with activation of FGF receptor (FGFR). Angiogenesis-related proteins including
IGFBP-3, IGFBP-10 and CX3CL1 were increased in HMW FGF2-only muscles and are likely involved in the protective effect of HMW FGF2. Another possible mechanism mediating HMW
FGF2-induced recovery from ischemic damage is myocyte regeneration. Preliminary results indicate the presence of amplified satellite cell activation in HMW FGF2 muscles, which was represented by upregulation of Pax7 expression. Pax7 expression was coupled with increased expression of the myoblast differentiation genes, Myogenin and MRF4.
Together, these data, for the first time, show that the HMW FGF2 isoforms had a beneficial role in salvaging skeletal muscle from ischemic injury. This dissertation uncovered a biological role for HMW FGF2 isoforms in vascular growth. Though the mechanisms that mediate this function of HMW FGF2 isoforms remain to be thoroughly characterized, this dissertation provides significant and novel evidence for the role of the FGF2 HMW isoforms in ischemia-induced revascularization and preservation of muscle function.
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ACKNOWLEDGMENTS
As I start to pen the final section of this thesis and with the final steps toward completion in sight, my prior belief that this part would be the easiest has turned out not to be true. So, it is with mixed emotions that I continue writing. I am both happy to have arrived at the end of this long journey and sad because of the strong attachments I feel to the people and the place that has nurtured my transition from student to scientist.
First and foremost, I am enormously indebted to my thesis advisor, Dr. Jo El Schultz for her unwavering support throughout my time as a graduate student. She always provided whatever
I needed to succeed including training, support, guidance, friendship and patience. I could always count on her to be available whenever I needed her, to be an attentive ear when I needed to vent as well a shoulder to cry on at times of frustration. She has and will continue to be a mentor, a role model and friend.
I would like to extend my gratitude to my dissertation committee, Drs. James Hoying, W.
Keith Jones, Ronald Millard, and Daria Narmoneva for their valuable time, assistance and encouragement. As a group, they have been an incredible resource for experimental advice, technical support and cardiovascular expertise.
I am also grateful to past and current members of the Schultz lab including Dr. Craig Bolte and Dr. Janet Manning for being amazing graduate student mentors; Dr. Yu Zhang for collaborating on some of my projects, Chahua Huang for being my lab buddy and teaching me a few words in Chinese and Colleen York for helping with our animal colony. Special thanks to honorary Schultz lab member, Stephanie Kelly for her insight into the treadmill experiment setup and her support from behind the scenes.
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I am sincerely indebted to many colleagues and collaborators at the University of
Cincinnati and Cincinnati Children’s Hospital, including Drs. Swathi Balaji and Daria Narmoneva
for their assistance in developing the vessel staining protocol; Dr. Zhihua Qi, Dr. Lisa Lemen, and
Kati LaSance of the Vontz Core Imaging Lab for their guidance in the collection and analysis of
the micro-CT images; Dr. Craig Bolte and Robin Gear for constantly helping me to troubleshoot
histology experiments. In addition, members of the laboratories of Drs. Belcher, Fan, and Kranias,
who were all very generous with their reagents, lab equipment and time.
I am extremely grateful to the undergraduate and graduate students who I have had the
privilege to mentor during their rotations through the Schultz lab including Mykea Ruffin, Allyson
Hamlin, Allison Dixon, Jordan Redfield, Chang Zeng, Chris Johnson, and Andrew Stein. In
different ways, each of you made significant contributions to my research projects.
I owe a tremendous amount of gratitude to members of the Department of Pharmacology and the Molecular, Cellular, and Biochemical Pharmacology (MCBP) program. Drs. Scott
Belcher, Guochang Fan, Evangelia Kranias, Andrew Norman, Robert Rapaport, and Jo El Schultz for their steadfast belief in my capabilities as a scientist and helping to guide me towards my degree completion. A special thanks to Dr. Rapoport and Nancy Thyberg who towards the end were instrumental in keeping me on task, making sure paperwork was filed and deadlines were met.
Finally, I would like to thank family for their love, patience and support. To my wonderful parents for their encouragement, sacrifice, and unconditional support. I would like to express my profound appreciation for my brother Wunmi, and my sisters Tola and Toun, for always listening when I need to vent, never letting me wallow and always giving the best advice.
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TABLE OF CONTENTS Page
Abstract ii
Acknowledgments v
List of Figures and Tables xi
List of Abbreviations xv
Introduction
1. Coronary artery disease (CAD) 1
2. Peripheral artery disease (PAD) 2
3. Vascular growth 4
A. Therapeutic revascularization 4
B. Mechanisms of vascular formation and growth 4
Vasculogenesis 4
Angiogenesis 5
Arteriogenesis 6
Collateral remodeling 6
Vessel network formation 7
Ischemia-induced vascular growth 9
Mechanisms regulating post-ischemic vascular growth 10
4. Fibroblast Growth Factors (FGFs) 15
A. FGF receptors (FGFRs) 18
Heparan Sulfate Proteoglycans (HSPGs) 18
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FGFRs 19
B. FGF/FGFR signaling 22
C. Angiogenic FGFs 24
5. FGF2 28
A. Fgf2 gene and tissue expression 28
B. Phenotypes of Fgf2-/- mice 29
C. FGF2 protein isoforms 31
D. Biological functions of the FGF2 isoforms 34
E. Phenotypes of FGF2 isoform-specific knockout mice (role of endogenous FGF2
isoforms) 39
F. FGF2 in revascularization (Preclinical studies) 44
G. FGF2 in revascularization (Clinical studies) 56
6. Dissertation focus and hypotheses 55
Materials and Methods 64
Animals and exclusion criteria 64
Mouse generation and breeding 64
Generation of Fgf2-/- mice 64
Generation of FGF2 HMW-only mice 65
Generation of FGF2 LMW-only mice 68
Hindlimb ischemia surgery 69
Assessment of necrosis 71
Assessment of limb function 71
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Exercise treadmill testing 71
Fgf2 and Fgfr gene expression 72
RNA isolation 72
Reverse transcription and real-time polymerase chain reaction (PCR) 73
Myogenic satellite cell marker gene expression 75
FGF2 protein isoform expression (Western immunoblot) 78
FGFR expression and phosphorylation (Western immunoblot) 79
Expression of angiogenesis-related proteins 81
Immunohistochemistry 83
Griffonia simplicifolia (Bandeiraea) isolectin B4 83
Alpha smooth muscle actin 84
Myeloperoxidase (MPO) 85
Mac-3 (CD107b) 85
Micro-computed tomography (micro-CT) 86
Perfusion of contrast agent 87
Image acquisition 87
Comparison of contrast agents 88
Image analysis 93
Laser Doppler Perfusion Imaging 95
Statistical Analysis 95
Chapter 1: HMW and LMW isoforms of FGF2 in ischemia-induced revascularization and tissue recovery
Results I 96
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Discussion I 141
Chapter 2: Potential mechanisms of the FGF2 isoforms in adaptive revascularization during chronic ischemia
Results II 166
Discussion II 208
Chapter 3: FGF2 isoform-induced myogenesis during chronic hindlimb ischemia
Results III 233
Discussion III 248
Conclusions and Clinical Relevance 253
Future Directions 258
References 264
Appendix 345
x
LIST OF FIGURES AND TABLES
FIGURES
1. The mechanisms governing post-ischemic revascularization 8
2. Mouse/Human FGF2 gene and protein expression 34
3. LMW and HMW FGF2 isoform expression in non-ischemic skeletal muscles 67
4. The mouse hindlimb ischemia model 70
5. Representative 2-D slice views of a barium sulfate-perfused limb 90
6. Representative 2-D slice views of a lead oxide-perfused limb 91
7. Representative 2-D slice views of a Microfil-perfused limb 92
8. Representative 2-D slice views of a Microfil-perfused limb segmentation 94
9. Limb impairment scores 98
10. Treadmill exercise performance 100
11. Incidence of limb necrosis in ischemic limbs 103
12. Representative micro-CT images of sham and ischemic hindlimbs 108
13. Vessel volume, density and spacing calculated from 3-D micro-CT images 109
14. Histograms of vessel thickness distribution in sham and ischemic limbs analyzed from
micro-CT images 112
15. Volume of small-sized (34 -200μm) and medium-sized (200-400μm) vessels analyzed
from micro-CT images 115
16. Capillary vessel density ratio after 14 days of ischemia 118
17. Representative photomicrographs of GSI-Lectin stained vessels and capillary density in
sham and ischemic muscles after 42 days of ischemia 121
18. Capillary density of sham and ischemic muscles after 42 days of ischemia 122
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19. Gracilis, semi-membranosus, semi-tendinosus and gastrocnemius muscle capillary density
after 42 days of ischemia 123
20. Arteriole vessel density ratio after 14 days of ischemia 126
21. Representative photomicrographs of α-SMA stained vessels, venule and arteriole density
in sham and ischemic muscles after 42 days of ischemia 129
22. Venue and arteriole vessel density of sham and ischemic muscles after 42 days of
ischemia 130
23. Gracilis, semi-membranosus, semi-tendinosus and gastrocnemius muscle venule density
in sham and ischemic muscles after 42 days of ischemia 132
24. Gracilis, semi-membranosus, semi-tendinosus and gastrocnemius muscle arteriole density
in sham and ischemic muscles after 42 days of ischemia 134
25. Wet weights of sham and ischemic gracilis, semi-membranosus, semi-tendinosus and
gastrocnemius muscles 136
26. Representative laser Doppler perfusion images and quantitative analysis of foot perfusion
over time 139
27. Schematic (Chapter 1) 165
28. Fgf2 mRNA expression in non-ischemic muscle 138
29. LMW (18kDa) and HMW (21kDa, 22kDa) FGF2 isoform expression in WT, Fgf2-/-, and
FGF2 LMW-only non-ischemic muscles 167
30. LMW (18kDa) and HMW (21kDa, 22kDa) FGF2 isoform expression in WT and FGF2
HMW-only non-ischemic muscles 169
31. Fgf2 mRNA expression in Fgf2-/- and FGF2 LMW-only sham and ischemic muscles170
32. Fgf2 mRNA expression in FGF2 HMW-only sham and ischemic skeletal muscles 173
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33. LMW (18kDa) and HMW (21kDa, 22kDa) FGF2 isoform expression in WT, Fgf2-/-, and
FGF2 LMW-only sham and ischemic muscles 174
34. LMW (18kDa) and HMW (21kDa, 22kDa) FGF2 isoform expression in WT and FGF2
HMW-only sham and ischemic skeletal muscles 176
35. Fgfr1 mRNA expression in non-ischemic, sham and ischemic muscles 180
36. Fgfr3 mRNA expression in non-ischemic, sham and ischemic muscles 183
37. Fgfr4 mRNA expression in non-ischemic, sham and ischemic muscles 186
38. FGFR activation (phosphorylation) and expression in non-ischemic muscle 189
39. FGFR1 activation (phosphorylation) and expression in ischemic muscles 191
40. FGFR3 activation (phosphorylation) and expression in ischemic muscles 193
41. FGFR4 activation (phosphorylation) and expression in ischemic muscles 195
42. Neutrophil density in non-ischemic, sham and ischemic muscles 203
43. Monocyte density in non-ischemic, sham and ischemic muscles 207
44. Schematic (Chapter 2) 232
45. Schematic of satellite cell activation, differentiation and fusion 235
46. Pax7 mRNA expression in non-ischemic, sham and ischemic muscles 237
47. Myf5 mRNA expression in non-ischemic, sham and ischemic muscles 240
48. MyoD mRNA expression in non-ischemic, sham and ischemic muscles 241
49. Myogenin mRNA expression in non-ischemic, sham and ischemic muscles 243
50. MRF4 mRNA expression in non-ischemic, sham and ischemic muscles 246
51. Schematic (Chapter 3) 252
52. Working model of HMW FGF2 in vascular and skeletal muscle repair 257
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TABLES
1. Phenotypes of Fgf knockout mice 17
2. FGF/FGFR binding specificities 21
3. Phenotypes of Fgfr knockout mice 22
4. FGF2 and other angiogenic growth factors in angiogenesis assays 47
5. Fgf2 gene or LMW FGF2 preclinical studies in models of myocardial ischemia 53
6. FGF2 preclinical studies in models of peripheral ischemia 54
7. FGF2 and other angiogenic growth factors in models of ischemia 55
8. Clinical trials results of FGFs in CAD and PAD patients 58
9. Forward and reverse primers for analysis of Fgf2 and Fgf receptor expression 76
10. Forward and reverse primers for the analysis of MRFs expression 77
11. List of angiogenesis-related proteins analyzed using antibody array 82
12. Descriptive scoring for the assessment of hindlimb use during chronic ischemia 97
13. Expression of angiogenic-related proteins in ischemic muscles 199
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LIST OF ABREVIATIONS
AGENT angiogenic gene therapy
α-SMA alpha smooth muscle actin
Ang angiopoietin
ANOVA analysis of variance
ATE amino-terminal end
BAEC bovine aortic endothelial cell
BCEC bovine capillary endothelial cell
BM-MNC bone marrow mononuclear cell
CAD coronary artery disease
CAM chorioallantoic membrane
CHD coronary heart disease
CHIP chromatin immunoprecipitation
CLI critical limb ischemia
CNS central nervous system
CVD cardiovascular disease
CXCL chemokine (C-X-C) motif ligand
DLL4 delta-like ligand 4
EC endothelial cell
EPC endothelial progenitor cell
ETT exercise tolerance test
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FGF fibroblast growth factor
FGFR fibroblast growth factor receptor
FIF FGF2-interacting factor
FIRST FGF Initiating RevaScularization Trial
GM-CSF granulocyte macrophage-colony stimulating factor
GSL IB4 griffonia simplicifolia Isolectin B4
HB-EGF heparin-binding EGF-like growth factor
HGF hepatocyte growth factor
HIF1α hypoxia inducible factor 1 alpha
HLI hindlimb ischemia
HMW high molecular weight
HSPG heparan sulfate proteoglycan
IC intermittent claudication
IGF1 insulin like growth factor 1
IGFBP insulin-like growth factor binding protein
IL interleukin
INFS integrative nuclear FGFR1 signaling
I/R ischemic/reperfusion kDa kilodaltons
LAD left anterior descending
LDPI laser Doppler perfusion imaging
LMW low molecular weight
MAPK mitogen activated protein kinase
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Micro-CT micro computed tomography
MIP-1 macrophage inflammatory protein-1
MCP-1 monocyte chemoattractant protein-1
MMP matrix metalloproteinase
MPO myeloperoxidase
MRF myogenic regulatory factor
MyoD myogenic determination factor 1
Myf5 myogenic factor 5
NV1FGF nonviral 1 FGF
PAD peripheral artery disease
PAI-1 plasminogen activator inhibitor-1
Pax7 paired box 7
PBS phospho-buffered saline
PCR polymerase chain reaction
PEDF pigment epithelium-derived factor
PDGF platelet-derived growth factor
PD-ECGF platelet-derived endothelial cell growth factor
PIGF placenta growth factor
PI3K phosphatidylinositol 3-kinase
PDGF platelet-derived growth factor
PKC protein kinase C qRT-PCR quantitative real time PCR
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ROI region of interest
SDF stromal-derived factor
SMN survivor of motorneuron protein
TALISMAN Therapeutic Angiogenesis Leg Ischemia Study for the Management of
Arteriopathy and Nonhealing ulcers
TIMP tissue inhibitors of metalloproteinases
TNF-α tumor necrosis factor-alpha
TRAFFIC TheRapeutic Angiogenesis with recombinant FGF2 for Intermittent
Claudication u-PA urokinase plasminogen activator
VEGF vascular endothelial growth factor vSMC vascular smooth muscle cell
VOI volume of interest
WT wildtype
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INTRODUCTION AND BACKGROUND
Approximately 85.6 million individuals in the United States have one or more form of cardiovascular disease (CVD) (Mozaffarian et al., 2015). As recently as 2010, CVDs accounted for over 31% of all U.S deaths. CVD as defined by the International Classification of Diseases includes coronary heart disease (CHD), acute coronary syndrome, high blood pressure, myocardial infarction, angina pectoris, heart failure and stroke (cerebrovascular disease) and peripheral vascular (arterial) disease (Brämer, 1988). This dissertation will focused on coronary heart disease and peripheral vascular disease.
1. Coronary heart disease (CHD)
Coronary heart disease affects over 15.5 million people in the United States. This disease category includes acute myocardial infarction, acute ischemic (coronary) heart disease, angina pectoris, atherosclerotic cardiovascular disease, and chronic ischemic (coronary) heart disease
(Mozaffarian et al., 2015). CHD develops from hardening, narrowing and/or occlusion of one or more of the large coronary arteries of the heart that is caused by atherosclerotic plaque (obstructive lesions) formation. Atherosclerotic lesions are usually an accumulation of lipids, inflammatory cells and fibrous material that are deposited in the inner arterial walls (Falk, Nakano, Bentzon,
Finn, & Virmani, 2013; Libby & Theroux, 2005). In CHD, fixed stenosis of coronary arteries manifests as stable angina pectoris and patients experience reversible ischemic pain during exercise. This type of angina can progress to an unstable state where ischemic pain occurs at rest
(Dragneva, Korpisalo, & Ylä-Herttuala, 2013). Additional risk factors for CHD include cigarette smoking and alcohol use, physical inactivity, hypertension, obesity, diabetes mellitus and aging
(Silvestre, Smadja, & Lévy, 2013). Current therapies for chronic ischemic and coronary heart
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diseases include controlling risk factors (diet, exercise and cessation of smoking), preventive
measures (statins, aspirin, antiplatelets and ACE inhibitors), revascularization procedures
(percutaneous coronary interventions, stents or bypass grafts) and anti-thrombotic agents (Libby
& Theroux, 2005; Rubanyi, 2013).
2. Peripheral arterial disease (PAD)
Peripheral arterial disease (PAD) results in reduction of blood supply to vessels beds other than the heart or the brain. PAD affects an estimated 8.5 million people in the U.S and it has been diagnosed in over 10-15% of adults over age 65 (Mozaffarian et al., 2015). PAD develops from occlusive atherosclerosis and is characterized by loss of blood flow reserve from chronic reduction in the diameter of the lumen of the distal abdominal aorta, pelvic or lower extremity arteries
(Annex, 2013). There are two symptomatic clinical manifestations of PAD; intermittent claudication (IC) and critical limb ischemia (CLI). In IC, patients encounter muscle pain or cramping during walking and exertion which is relieved upon resting, while CLI presents as limited blood flow to the limbs even at rest that can lead to gangrene, ulcers and in some cases amputation (Haas, Lloyd, Yang, & Terjung, 2012). The risk factors for PAD are similar to those for CHD; however, cigarette smoking and diabetes mellitus play a stronger causative role in PAD.
Other risk factors for PAD include hypertension, hyperhomocysteinemia, dyslipidemia, elevated
C-reactive protein, and age (Mascarenhas, Albayati, Shearman, & Jude, 2014; Muller, Reed,
Leuenberger, & Sinoway, 2013). PAD alone can be a strong predictor of CHD and cerebrovascular disease (CBVD) based on vascular effects. The co-diagnosis of CHD with PAD ranges from 35% to 60% of patients and for PAD with CBVD can be as high as 80% of patients. The severity of
PAD correlates with increased CVD-related mortality (Haas et al., 2012). Current treatments for
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PAD are similar to those for CHD and include modification of risk factors like smoking, diet and
exercise (Muller et al., 2013). Smoking cessation slows the progression to critical limb ischemia
and reduces the risk of cardiovascular (CVD) events like myocardial infarction (Jha et al., 2013).
Diet modification and management of diabetes also decreases the risk of CVD events as well as
microvascular complications like retinopathies and neuropathies (Libby & Theroux, 2005; Muir,
2009; Muller et al., 2013). Statins also reduce risk and worsening of symptoms in PAD patients
by lowering serum cholesterol, improving endothelial function, and other atherosclerotic markers.
Antihypertensives like ACE inhibitors or β-AR blockers lower blood pressure and reduce the risk of new CVD events. Secondary prevention of CHD and PAD is conferred by antiplatelet agents which reduce the risks of nonfatal myocardial infarction, ischemic stroke, and vascular-related death (Muir, 2009; Muller et al., 2013; Norgren et al., 2007).
Overall, standard treatments for CHD and PAD are aimed at symptom relief, reducing atherosclerotic risk factors, decreasing oxygen demand and vascular interventions to physically reestablish blood flow to the ischemic regions. Despite these pharmacotherapies, a large and growing number of patients remain symptomatic. The presence of co-morbidities like diabetes and hypertension renders these patients ineligible for the surgical interventions; these patients may eventually develop heart failure or require limb amputations (Ahn, Frishman, Gutwein, Passeri, &
Nelson, 2008; Grochot-Przeczek, Dulak, & Jozkowicz, 2013; Rubanyi, 2013; Van Weel, van
Tongeren, van Hinsbergh, van Bockel, & Quax, 2008). The increasing numbers of these “no- option” patients illustrates the need for alternative treatment approaches for CAD and PAD
(Rubanyi, 2013; van Weel et al., 2008; Annex 2013 ).
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3. Vascular growth
A. Therapeutic revascularization
“Therapeutic revascularization” is an emerging strategy that aims to augment or stimulate
blood vessel growth and remodeling for the improvement of perfusion and function in the ischemic
heart or lower limb (Annex, 2013; Rubanyi, 2013). This approach compensates for the loss of
blood supply that develops from arterial occlusions or stenosis by pharmacological treatment with
promoter(s) of vessel growth.
B. Mechanisms of vascular formation and growth
Blood vessel growth and formation involves four distinct but complementary processes: 1)
vasculogenesis, 2) angiogenesis, 3) arteriogenesis, and 4) collateral growth (Annex, 2013;
Dragneva et al., 2013; Rubanyi, 2013; Silvestre et al., 2013). Vessel growth plays a key role in various physiological and pathological conditions, including the menstrual cycle, embryonic development, wound repair, inflammation, and tumor growth. At present, it is known that all four mechanisms of vascular growth can be triggered in response to ischemia but the relative contributions of each to the restoration of perfusion has yet to be determined (Figure 1).
Vasculogenesis
Vasculogenesis is the de novo formation of the earliest vascular structures (primitive plexus) in the developing embryo. In situ proliferation, migration and differentiation of endothelial progenitor cells (EPCs) (angioblasts in development) or bone-marrow-derived endothelial stem cells (in adult) results in tubules of endothelial cells (ECs) with very little organization (Carmeliet
& Jain, 2011; Rubanyi, 2013). Once considered to be a strictly embryonic process, vasculogenesis in the adult has been shown to occur when nascent vessels formed in response to ischemia were
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proven to contain progenitor cells that mobilize to the site of and are incorporated into the developing vessels (Asahara, 1997; Fischer, Schneider, & Carmeliet, 2006).
Angiogenesis
“Angiogenesis” is frequently used to describe de novo formation or increase in diameter, length, or branching of vessels in any developmental, postnatal or pathological condition (Faber,
Chilian, Deindl, Royen, & Simons, 2014). The use of “angiogenesis” in this dissertation will refer only to capillary growth or formation occurring during embryogenesis or in response to arterial obstructions as described below.
The organization of the nascent primitive plexus formed during vasculogenesis into a network of functional microvessels is termed embryonic angiogenesis. This process involves the sprouting or intussusceptive (splitting into daughter vessels) growth of new vessels from the ends
and walls of the preexisting vascular beds (endothelial tubules) (Carmeliet & Jain, 2011; Fischer
et al., 2006). This expanded nascent network is now made up of narrow microvessels (<10 µm)
which are referred to as capillaries. A variation of this process that occurs postnatally or in the
adult also involves the proliferation of pre-existing vascular endothelial cells into new capillary
sprouts (sprouting) and/or bridging of preexisting vessels into smaller daughter vessels (splitting)
(Fischer et al., 2006; Semenza, 2007). Angiogenesis can be induced by physiological or pathological triggers including organ growth, physical exercise training, wound healing, tumor growth, and vascular occlusions (Carmeliet & Jain, 2011; Fischer et al., 2006; Persson, 2011).
Arteriogenesis
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Arteriogenesis refers to the stabilization and maturation of nascent vascular networks
(Fischer et al., 2006). Mural cells (pericytes and smooth muscle cells) of mesenchymal lineage are recruited to the naked EC sprouts. Deposition of the extracellular matrix and tightening of cell junctions occurs at this time, leading to a more organized and functional network. The origin, type and composition of the mural cells will depend on the site, function and arterial-venous identity of the vessels (Carmeliet & Jain, 2011; Silvestre et al., 2013). A collateral circulation is also formed at this time. Collaterals are artery-to-artery (collateral arteries) or arteriole-to-arteriole
(microvascular collaterals) anastomoses (connections) that form between feed arteries or arterial trees (Faber et al., 2014). They exist in a unique hemodynamic state of very low blood flow and high resistance during normal physiology and are present in the brain, spinal cord, heart, skin and skeletal muscle (Faber et al., 2014; Fischer et al., 2006; J. Liu et al., 2014; Silvestre et al., 2013).
Collateral remodeling
This process often described as an adaptive arteriogenic process, refers to the transformation of pre-existing collateral vessels with previously low blood flow into functional conductance vessels (Faber et al., 2014). Here, remodeling is defined as an anatomic enlargement of lumen diameter and an increase in vascular wall thickness (Heil, Eitenmüller, Schmitz-Rixen,
& Schaper, 2006; Silvestre et al., 2013). This outward remodeling takes place as a result of flow obstructions that induce increased flow in adjacent arterial trees. The pre-existing collateral network across from the occluded vascular tree is recruited to compensate for the change in pressure gradient (J. Liu et al., 2014; Schaper, 2009). The microvascular collaterals are typically
≤100 µm in diameter (in healthy tissues) and are capable of significant outward remodeling by
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increasing their lumen diameter up to 5-10 fold. In the mouse circulation, they can be as small as
30 µm and can increase by 2-5 fold (Faber et al., 2014; Ziegler et al., 2010).
Vessel network formation
During physiological angiogenesis, formation of hierarchally structured three dimensional vascular networks requires more than sprouting of neovessels. Guided maturation, pruning,
regression of the blood vessel network is necessary for the formation of a stable, well-perfused
network (LeBlanc, Krishnan, Sullivan, Williams, & Hoying, 2012). Vessel maturation involves the recruitment and attachment of network-stabilizing perivascular cells like pericytes and vascular smooth muscle cells. The recruitment and differentiation of perivascular cells is mediated by factors released by endothelial cells including platelet endothelial growth factor (PDGF) and transforming growth factor-β (TGF-β).
Capillary regression is regulated by the balance between the expression of positive angiogenic factors like vascular endothelial factors (VEGF) and negative angiogenic factors such as thrombospondin-1 (TSP-1) (Olfert, 2015). This balance is evident in the mammalian corpus luteum. Growth and regression of the cyclic corpus luteum involves distinct phases of angiogenesis, vessel maturation, and vessel regression. Luteolysis leads to the physiologically triggered dissociation of the entire corpus luteum vasculature (Wietecha, Cerny, & DiPietro,
2013). Branch selection for pruning or regression is also regulated by blood flow and perfusion of vessels. The selected branch/vessel constricts until it occludes and the blood flow ceases. The endothelial cells then retract to undergo apoptosis or may migrate away to be re-integrated elsewhere.
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Figure 1: The four distinct but complementary mechanisms governing post-ischemic revascularization. 1) Vasculogenesis, the formation of blood vessels by differentiation of endothelial progenitors cells (EPCs); 2) angiogenesis, the sprouting or budding of new capillaries (Cap) from preexisting vessels; 3) arteriogenesis, the maturation and stabilization of nascent sprouts via recruited smooth muscle cells (SMC) and pericytes (Per) to form arterioles; and 4) collateral (Coll) growth, the remodeling of preexisting arteriolar anastomoses into functional conductance vessels. Adapted from (Silvestre et al., 2013). Reprinted with permission of the American Physiological Society.
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Ischemia-induced vascular growth
The vascular processes described above largely take place during embryonic development but they can be recapitulated in adult tissues under certain pathophysiological conditions like ischemia or physical damage (Ahn et al., 2008; Carmeliet & Jain, 2011; Haas et al., 2012; Silvestre et al., 2013). The physiological responses of cardiac or skeletal muscle to the initial vascular obstruction share some similarities with the wound healing processes observed in tissues like skin.
Tissue healing can be been divided into three chronologically overlapping phases: 1) destruction, 2) inflammatory phase, and 3) remodeling and regeneration phase (Philippou,
Maridaki, Theos, & Koutsilieris, 2012; Smith, Kruger, Smith, & Myburgh, 2008; Weber, Sun,
Bhattacharya, Ahokas, & Gerling, 2013). The destruction phase which begins just after the initial occlusion and persists for up to 2 days is characterized by hypoperfusion, hypoxia and metabolic dysregulation leading to myocyte death by apoptosis and/or necrosis. An acute inflammatory phase follows cellular death and can last for up to 7 days (Silvestre et al., 2013). This phase features degradation of the extracellular matrix to allow homing and migration of inflammatory cells
(neutrophils, monocytes, lymphocytes) into the necrotic tissue for phagocytosis of cellular debris
(Cleutjens, 1999; Matsui, Morimoto, & Uede, 2010). The remodeling and regeneration phase which lasts up to 21 days is comprised of migration and differentiation of fibroblasts, myofibroblasts, and macrophages; synthesis and deposition of extracellular matrix proteins; and scar tissue formation to replace necrotic or apoptotic cells. The final phase is also associated with an immune-inflammatory balance, secretion of chemokines, growth factors and other cytokines which promote myocyte hypertrophy/regeneration and vessel formation or remodeling (Silvestre et al., 2013; Smith et al., 2008).
9
Mechanisms regulating post-ischemic vascular growth
There are many driving forces that initiate adaptive angiogenesis and collateral growth;
they include inflammatory balance, ischemia-induced hypoxia, and fluid shear stress (Heil et al.,
2006; Silvestre et al., 2013).
i. Inflammatory response in post-ischemic vascular growth
Acute inflammation is a crucial component of the repair/remodeling phase of the tissue
healing that occurs after ischemia (Silvestre et al., 2013; Smith et al., 2008). Vessel growth and
inflammation are linked in pathophysiological conditions of wound healing, diabetic retinopathy,
rheumatoid arthritis, and atherosclerosis (Frantz, Vincent, Feron, & Kelly, 2005; Mueller, 2008;
Serhan & Savill, 2005). Inflammation is capable of modulating vessel growth and vice-versa. The endothelium when it is activated by hypoxia secretes chemoattractant cytokines and adhesion molecules that serve as signals for inflammatory cell recruitment and activation (Stockmann,
Schadendorf, Klose, & Helfrich, 2014). In turn, the recruited inflammatory cells (leukocytes) stimulate the formation of new vessels in a paracrine fashion by secreting angiogenic growth factors, pro-and antiangiogenic cytokines and proteases (Smith et al., 2008). The recruitment and
transmigration of immune cells across the sites of ischemia and vascular occlusion is regulated by the expression and surface presentation of specific cell adhesion and chemotactic pathways. These include the diverse group of cell adhesion molecule, chemokine and interleukin protein families.
The types of inflammatory cells that found infiltrate the vasculature and muscle during
ischemic injury include monocytes, neutrophils, and lymphocytes (Cochain, Channon, & Silvestre,
2012; Mueller, 2008; Smith et al., 2008). The level of monocyte infiltration has been shown to
correlate with the extent of neovascularization. Monocytes are rapidly mobilized from the bone
10
marrow, spleen and bloodstream to the site of ischemia (Silvestre et al., 2013). Monocytes
differentiate into macrophages once they have extravasated into the tissue. Tissue macrophages
are made up of two subpopulations of cells: the classically-activated M1 macrophages which
express pro-inflammatory molecules and alternatively-activated M2 macrophages which express anti-inflammatory molecules and promotes vascular and tissue regeneration (Fung & Helisch,
2012; Jetten et al., 2014; Mueller, 2008; Silvestre et al., 2013). The phagocytic M1 cells are localized to inflamed tissues immediately after ischemia to promote phagocytosis and the M2 cells are largely present during tissue healing (Silvestre et al., 2013; Smith et al., 2008).
Neutrophils, unlike monocytes, are predominantly antiangiogenic (Naldini & Carraro,
2005; Nguyen & Tidball, 2003). Since they trigger the release of free radicals and lysosomal proteinases, neutrophils have been linked to the secondary tissue damage that occurs after ischemia especially to the endothelium (Frangogiannis, Smith, & Entman, 2002; Silvestre, Mallat, Tedgui,
& Lévy, 2008; Silvestre et al., 2013). They produce reactive oxygen species which can stimulate revascularization at low levels but can also damage endothelial cells and inhibit angiogenesis at high concentrations (Cochain et al., 2012; Silvestre et al., 2008; Sun, 2009). Lysosomal proteinases such as proteinase 3 and elastase are also generated by neutrophils. These proteinases can induce endothelial detachment and apoptosis (Cochain et al., 2012; Mueller, 2008; Silvestre et al., 2008).
However, neutrophils may contribute to angiogenesis by their secretion of vascular endothelial
growth factor (VEGF) (Cochain et al., 2012; Mueller, 2008).
T-lymphocytes, divided into two subtypes, CD4 or CD8, based on their expression of
membrane antigens, are also involved in post-ischemic inflammatory response (Silvestre et al.,
2013). Mice that are deficient in CD4 or CD8 (CD4-/- or CD8-/-) or both (CD4-/-/CD8-/-) have
impaired vascular responses to ischemia (Stabile et al., 2003, 2006). CD8 T-lymphocytes promote
11
vascularization by the release of growth factors and cytokines including VEGF, interleukins 10
and 16 (IL-10 and IL-16) that will also attract other inflammatory cells like CD4 lymphocytes and
monocytes (la Sala, Pontecorvo, Agresta, Rosano, & Stabile, 2012; Silvestre et al., 2013).
The overall role of inflammation on ischemia-induced revascularization is dependent on
the temporal and spatial balance between the activation of pro- and anti-inflammatory cells.
Additionally, the intensity of pro-/anti-angiogenic molecules secreted by the recruited immune
cells is also important for vascular remodeling (Carmeliet, 2003; Silvestre et al., 2008).
ii. Hypoxia in ischemia-induced angiogenesis
The primary functions of the vascular system include the delivery of O2 and supply of
nutrients to tissues and the removal of toxic metabolic wastes (Semenza, 2007). The reduction or
complete loss of blood supply that occurs in ischemic tissues disrupts this oxygen homeostasis,
and leads to hypoxia. Hypoxia is the local decrease in O2 partial pressure (pO2) that results from
either decreased supply of O2 or increased metabolic demand from tissues (Silvestre et al., 2013).
In hypoxic environments, angiogenesis is stimulated to re-establish the equilibrium between local oxygen need and supply and is mediated by the hypoxia-inducible factor (HIF) (Krock, Skuli, &
Simon, 2011). HIF is a heterodimeric protein composed of a regulatory (oxygen-sensitive) α- subunit and a constitutively expressed β-subunit. There are three isoforms of the HIF α-subunit
(1α, 2α and 3α) that are subjected to rapid polyubiquitination and proteosomal degradation under normal oxygen conditions. HIF1α is universally expressed while HIF2α is predominantly expressed in endothelial cells during development and in the adult (Hickey & Simon, 2006; Krock et al., 2011; Silvestre et al., 2013). HIF3α has been detected in the cerebral cortex, hippocampus and lung epithelial cells (Heidbreder et al., 2003; Q. F. Li, Wang, Yang, & Lin, 2006). During
hypoxic stress when tissue pO2 decreases below 5% (40 mmHg), the HIF1α subunit becomes
12
stabilized and translocates to the nucleus (Krock et al., 2011; Persson, 2011). After dimerization
with the β-subunit, HIF1 binds to hypoxia response elements within promoter regions of target
genes and activates gene transcription (Semenza, 2007). Pathways targeted by the active HIF1
include other transcription factors (NFκB and ETS1), vasodilatory nitric oxide (NO), glycolytic
enzymes, cell cycle inhibitors and angiogenic cytokines (Krock et al., 2011).
Angiogenic cytokines induced by hypoxia and HIF1 exhibit either pro- or anti-angiogenic
behavior. Endogenous and ischemia-induced vascular growth are both tightly regulated and are
governed by the balance between local expression of angiostatic and proangiogenic molecules
(Hickey & Simon, 2006; Krock et al., 2011). Some of these proangiogenic genes include
angiopoietin-1/2 (Ang-1/2), matrix metalloproteinases (MMPs), tissue inhibitor of
metalloproteinases (TIMP), tyrosine kinase with immunoglobulin-like and EGF-like domains 1
(Tie2), tumor necrosis factor-α (TNF-α), interleukins, chemokines, stromal-derived growth factor
(SDF), hepatocyte growth factor (HGF), placenta growth factor (PIGF), platelet-derived growth
factors (PDGFs, vascular endothelial growth factors (VEGFs), and fibroblast growth factors
(FGFs) (Krock et al., 2011; Murakami & Simons, 2008; Persson, 2011; Rubanyi, 2013). This dissertation will focus on the biological functions of fibroblast growth factor 2 (FGF2) in ischemia- induced angiogenesis.
iii. Fluid shear stress-induced collateral remodeling
Unlike ischemia-induced angiogenesis, collateral remodeling has been shown to be largely stimulated by inflammation and is not likely to involve hypoxia (Heil et al., 2006; Heil & Schaper,
2007). Evidence for this includes the lack of HIF1α upregulation in the collateral zone, localization of collaterals in tissue regions that are distal to the ischemic zone, and the occurrence of collateral
13
development much later than the appearance of capillary growth in the ischemic regions (Heil et
al., 2006; van Oostrom, van Oostrom, Quax, Verhaar, & Hoefer, 2008; Van Weel et al., 2008).
During arterial occlusion, the luminal surface of vessels are exposed to hemodynamic
forces such as tensional stretch and shear stress which have been shown to induce vessel growth
by activation of endothelium (Schirmer, van Nooijen, Piek, & van Royen, 2009; Van Weel et al.,
2008). In particular, collateral remodeling of pre-existing arterioles is triggered by an increase in
fluid shear stress (FSS) and circumferential wall stress. FSS results from the steep pressure
gradient between the preexisting arterioles (high pressure) connected upstream of the occlusive
region and those downstream (low pressure) from this point (van Oostrom et al., 2008). The
increased flow through the vessels leads to functional activation of the previously quiescent
endothelial cell layer. The shear stress is also transmitted to the smooth muscle cell layers and the
cytoskeleton via communication between the cells and with extracellular matrix proteins
(Semenza, 2007; Silvestre et al., 2013). The shift in phenotype from quiescent to “activated”
endothelial cells is followed by gene transcription of several genes which have shear stress
response elements (SSREs) in their promoters; these include integrins, chemokines, endothelial
adhesion molecules, and growth factors (Haas et al., 2012; Heil & Schaper, 2007). The
upregulation of chemokines and cytokines induces the attraction and adhesion of bone marrow-
derived and circulating leukocytes particularly monocytes (Van Weel et al., 2008). Monocytes
accumulate in the perivascular space and then transmigrate across the vessels walls, by secretion
of MMPs and other protease that create gaps within the elastic lamina and basal membrane.
Monocytes also secrete additional growth factors like FGF2 that lead endothelial and smooth
muscle cells of the vessel wall to undergo a phenotypic shift from a contractile to a
proliferative/secretory state. The enzymatic breakdown of the extracellular matrix and basal
14
lamina also releases matrix-bound growth factors. Vascular cells proliferate, migrate and rearrange
causing thickening of the intima layer of the vascular wall (Fung & Helisch, 2012; Haas et al.,
2012; Heil & Schaper, 2004).
4. Fibroblast Growth Factors (FGFs)
Fibroblast growth factor 2 (FGF2) is a classical member of the FGF family. The FGFs are signaling polypeptides that are conserved among vertebrates and play an important role in a multitude of many processes including development, organogenesis, metabolism, neurobiology, tumorigenesis, and tissue homeostasis (Beenken & Mohammadi, 2009; Itoh & Ornitz, 2011). The
22 structurally-related proteins that make up the mammalian FGF family share 30-60% identity and contain ~150-300 amino acids. Crystal structures of the FGFs have revealed a core domain of
~140 amino acids with a conserved β-trefoil fold that consists of 12 anti-parallel β-strands (Belov
& Mohammadi, 2013; Zhu et al., 1991). The human FGF gene family has been grouped into seven subfamilies based on their sequence homology and phylogeny: Fgf1 subfamily (Fgf1/2), Fgf4 subfamily (Fgf4/5/6), Fgf7 subfamily (3/7/10/22), Fgf8 subfamily (Fgf8/17/18), Fgf9 subfamily
(Fgf9/16/20), Fgf19 subfamily (19/21/23), and the Fgf11 subfamily (Fgf11/12/13/14) (Belov &
Mohammadi, 2013; Itoh & Ohta, 2013).
The FGFs can be classified as intracrine, endocrine or paracrine/autocrine factors based on their respective mechanisms of action. The intracrine FGFs (FGF11-FGF14) are generally referred to as FGF homologous factors (FHF) because they share high sequence homology with other FGFs but do not activate the FGF receptors (FGFR) (Belov & Mohammadi, 2013; Goldfarb, 2005). FHFs have been shown to interact with the intracellular domains of calcium channels and voltage-gated sodium channels (Hennessey, Wei, & Pitt, 2013; Laezza et al., 2007; Xu Zhang, Bao, Yang, Wu,
15
& Li, 2012). The endocrine FGF 15, 19, 21 and 23 are hormone-like in their actions, FGFR- dependent and regulate several metabolic pathways (Beenken & Mohammadi, 2009; Krejci,
Prochazkova, Bryja, Kozubik, & Wilcox, 2009; Laestander & Engström, 2014). Endocrine FGFs
(FGF 15, 19, 21, 23) also require additional co-receptors, the Klotho receptors (α or β) to form
active signaling complexes due to their extremely low affinities for heparan sulfates (Beenken &
Mohammadi, 2009; Belov & Mohammadi, 2013; Laestander & Engström, 2014). Fgf1(FGF1/2),
4(FGF4/5/6), 7(3/7/10/22), 8(8/17/18), and 9(9/16/20) subfamilies are the paracrine and/or
autocrine signaling FGFs. They are mostly secreted proteins that bind to and activate the cell
surface tyrosine kinase FGFRs with high affinity and with low affinity to heparan sulfate
proteoglycans (HSPGs) (Beenken & Mohammadi, 2009; Itoh & Ohta, 2013; Itoh & Ornitz, 2011).
FGF 1 and 2 are prototypes of the entire FGF family as well as the paracrine subgroup despite
lacking the classical N-terminal secretion signal peptide present on the other paracrine FGFs (Itoh
& Ohta, 2013; Powers, McLeskey, & Wellstein, 2000). Elucidation of the biological functions of
the individual Fgf genes using knockout mice have begun and are listed in Table 1.
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Gene Knockout mouse phenotype Physiological role
Fgf1 Normal Not established Viable, Loss of vascular tone, Slight loss of cortex Described in detail in section titled Fgf2 neuron, Reduced bone mass (Montero et al., 2000) “Phenotypes of Fgf2-/- mice” Inner ear development (agenesis) in humans Fgf3 Viable, Tail defects (Alvarez et al., 2003) (Tekin et al., 2008)
Embryonic lethal at E4-5 (Feldman, Poueymirou, Cardiac valve leaflet formation, Fgf4 Papaioannou, DeChiara, & Goldfarb, 1995) Limb development Abnormally long hair (Hébert, Rosenquist, Götz, & Fgf5 Martin, 1994) Hair growth cycle regulation Fgf6 Defective muscle regeneration Myogenesis
Fgf7 Matted hair, Reduced nephron branching in kidney Branching morphogenesis Fgf8 Embryonic lethal at E8 Brain, eye, ear and limb development Postnatal death, Gender reversal, Gonadal development and Fgf9 Lung hypoplasia Organogenesis
Fgf10 Failed limb and lung development Branching morphogenesis Fgf11 N/A Not established Neurotransmission of motor function (Xu Fgf12 Viable, Muscle weakness Zhang et al., 2012) Microtubule stabilization (Q.-F. Wu et al., 2012) X-linked mental retardation Fgf13 Impaired learning and memory (Xu Zhang et al., 2012) Neurotransmission of motor function (Xu Fgf14 Ataxia, paroxysmal hyperkinetic movement disorder Zhang et al., 2012) Fgf16 Embryonic lethal Heart development Fgf17 Abnormal brain development Cerebral and cerebellar development Fgf18 Delayed long-bone ossification, Lethal at PD0 Bone development Bile acid homeostasis, Lipolysis, Gall Fgf15/19 Increased bile acid pool, Lethal at E13.5-PD7 bladder filling Fgf20 Viable Neurotrophic factor Fasting response, Glucose homeostasis, Fgf21 Viable Lipolysis and lipogenesis Fgf22 Viable Presynaptic neural organizer Impaired phosphate and Vitamin D metabolism, Phosphate homeostasis, Vitamin D Fgf23 Immature sexual organs, Lethal PW4-13 homeostasis
Table 1: Biological functions of the FGF family. Adapted from (Beenken & Mohammadi,
2009). E: embryonic day; PD: postnatal day; PW: post-wean day.
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A. FGF Receptors
The paracrine and endocrine FGFs mediate diverse cellular response by interactions with
the low affinity heparan sulfate proteoglycan (HSPG) co-receptors and high affinity FGF receptors
(FGFRs). i. Heparan Sulfate Proteoglycans (HSPGs)
Heparan sulfate proteoglycans (HSPGs) are heavily glycosylated proteins with a core
protein and one or more covalently attached heparin/heparan sulfate glycosaminoglycan (GAGs)
chains (Bishop, Schuksz, & Esko, 2007). GAGs are linear polysaccharides consisting of repeating
N-acetylated or N-sulfated glucosamine units (N-acetylglucosamine or N-sulfoglucosamine) and
uronic acids (glucuronic acid or iduronic acid). HSPGs are classified into subfamilies based on
their cellular locations: membrane-spanning HSPGs (syndecans 1-4, betaglycan, neuropilin-1,
CD44v3), glycophosphatidylinositol-anchored HSPGSs (glypicans 1-6), ECM proteoglycans
(perlecan, agrin, type XVIII collagen) and secretory vesicle HSPGs (serglycin) (Sarrazin,
Lamanna, & Esko, 2011). The functions of HSPGs in cell physiology include acting as co-
receptors for growth factors and their tyrosine kinase receptors, serving as endocytic receptors for
the removal of bound ligands, protecting growth factors, cytokines, and chemokines from
proteolysis, and cooperating with integrins to support cell adhesion to ECM and cell motility
(Bishop et al., 2007; Sarrazin et al., 2011). Some HSPGs like members of the syndecan family are
able to function outside the FGF-FGFR signaling complex to activate signaling pathways in
response to growth factor ligand binding (Murakami, Elfenbein, & Simons, 2008). Syndecan-4
independently regulates activation and localization of protein kinase Cα (PKCα) at sites of focal
adhesion (Elfenbein & Simons, 2013; Keum et al., 2004).
18
HSPGs are abundantly expressed on cell surfaces and in basement membranes of almost
all cell types including endothelial cells and fibroblasts (Presta et al., 2005; Sarrazin et al., 2011).
HSPGs bind to a variety of ligands via the heparan sulfate chains: these include growth factors,
cytokines, enzymes, chemokines and ECM proteins. They function in regulating FGF binding to
and activation of FGF receptors (Moosa Mohammadi, Olsen, & Ibrahimi, 2005). HSPGs are co- receptors that facilitates proper presentation of FGFs to FGFR and increases the formation of stable
FGF/FGFR complexes as well as the half-life of the signaling complex (Belov & Mohammadi,
2013; Bishop et al., 2007; Presta et al., 2005). HSPGs of the ECM can act as reservoirs for FGFs, prevent proteolytic degradation and increase local gradients of FGF during stimulation of endothelial cells. Alternatively, free heparin/HS can trap or sequester FGFs in the blood and other extracellular spaces and inhibit FGF activity (Presta et al., 2005, 2007). The affinity of FGFs and other growth factors for the highly negatively-charged heparin/heparan sulfates has been crucial in the discovery and purification of these proteins and is still in use today in affinity matrices and other “pull-down” assays.
ii. Fibroblast growth factor receptors (FGFRs)
The FGF receptors are a group of tyrosine kinase receptors that are encoded by four genes;
FGFR1, FGFR2, FGFR3 and FGFR4. They share up to 70% sequence identity (Eswarakumar,
Lax, & Schlessinger, 2005). These highly conserved receptors have the classic tyrosine kinase structure with an extracellular ligand-binding domain, a single helical transmembrane domain and a split intracellular tyrosine kinase domain (Belov & Mohammadi, 2013; Eswarakumar et al.,
2005). The extracellular domain is composed of three extracellular immunoglobulin (Ig) domains
(I, II and III) and an unique eight acidic residue sequence in the linker connecting domains I and
II called the “acid box”. The domain II and III fragments are necessary for ligand and heparin
19
binding and IgI and the acid box may both be important for autoinhibition (Beenken &
Mohammadi, 2009). Alternative RNA splicing of Ig domain III in the FGFR1-3 genes yields
several isoforms; FGFR1-3IIIa, FGFR1-3IIIb and FGFR1-3IIIc and this determines ligand specificity and tissue expression (Turner & Grose, 2010)(Belov & Mohammadi, 2013). The IIIa
forms have no known signaling capability while the IIIb and IIIc forms are predominantly
expressed in epithelial and mesenchymal cells, respectively (Xiuqin Zhang et al., 2006). Fgfr4 is
not known to be alternatively spliced (Fon Tacer et al., 2010; Ornitz & Itoh, 2015). The specificity
of FGF-FGFR interaction is such that, for example, FGFs expressed in epithelial tissues bind
specifically to the FGFR2b and do not bind to FGFR2c (Eswarakumar et al., 2005) (Table 2).
Several gain-of-function (GOF) and loss-of-function (LOF) mutations in FGFR 1, 2 and 3 have been linked to human diseases of skeletal dysplasia, dwarfism and cancer (Du, Xie, Xian, &
Chen, 2012; Eswarakumar et al., 2005). FGFR2 GOF mutations have been identified in some cases of craniosyntosis (premature fusion of cranial sutures) and FGFR1 GOFs are present in glioblastoma brain tumors and 8P11myeloproliferative syndrome (Beenken & Mohammadi,
2009). Transmembrane mutations in FGFR3 are highly prevalent in all cases of achondroplasia
(dwarfism) (Eswarakumar et al., 2005). The pathophysiological functions of the individual Fgfr genes are beginning to be elucidated using knockout mouse models. The viability and phenotypes of these mouse models are listed in Table 3.
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FGFR isoform Ligand specificity
FGFR1 IIIb FGF1, -2, -3, -7, -10, -22 FGFR1 IIIc FGF1, -2, -4, -5, -6, -19, -21, and -23
FGFR2 IIIb FGF1, -3, -7, -10, -22
FGFR2 IIIc FGF1, -2, -4, -5, -6, -9, -19, -21, and -23
FGFR3 IIIb FGF1, -8, -9, -16, -17, -18, -20
FGFR3 IIIc FGF1, -2, -4, -5, -6, -8, -9, -16, -17, -18, -19, -20, -21, and -23
FGFR4 FGF1, -2, -4, -5, -6, -19, -21, and -23
Table 2: Binding specificities of FGF ligands (except for intracrine FGFs) for the FGF receptors. Adapted from (Eswarakumar et al., 2005; Xiuqin Zhang et al., 2006).
21
FGF Knockout mouse phenotype Biological role receptor
Mesoderm/endoderm differentiation
Fgfr1 Embryonic lethal at E9.5-12.5 (Deng et al., 1994; Yamaguchi, Harpal, Henkemeyer, & Rossant, 1994)
Cell migration through primitive streak (Partanen, Fgfr1b Viable Schwartz, & Rossant, 1998)
Cell migration through primitive streak (Partanen et al., Fgfr1c Embryonic lethal at E9.5 1998)
Placenta and limb bud formation (Laestander & Fgfr2 Embryonic lethal at E4-5 Engström, 2014)
Development of the lung, limbs and teeth (Itoh & Fgfr2b Embryonic lethal at E0 Ornitz, 2011; Ornitz & Itoh, 2015)
Bone and skull development (Beenken & Mohammadi, Fgfr2c Viable 2009; Ornitz & Itoh, 2015)
Long bone overgrowth, Fgfr3 Early and postnatal bone development (Du et al., 2012) osteopenia
Cholesterol metabolism and bile acid synthesis Not Fgfr4 Viable clear (Eswarakumar et al., 2005; Ornitz & Itoh, 2015)
Table 3: Biological roles of the FGF receptors based on knockout mice phenotypes. Adapted
from (Beenken & Mohammadi, 2009; Eswarakumar et al., 2005; Itoh & Ornitz, 2011). E:
embryonic day
iii. FGF/FGFR signaling It is widely believed that the formation of a stable FGF-FGFR complex for intracellular
signaling requires the additional interaction with heparin or HSPGs (Belov & Mohammadi, 2013;
Schlessinger et al., 2000). There is growing evidence, however, to suggest that in the absence of heparin, functional FGF-FGFR complexes are still formed. Recently, Zakrzewska and colleagues
22 generated FGF1 mutants that had reduced affinity for heparin but retained their mitogenic activity due to the presence of stabilizing mutations (Zakrzewska et al., 2009; Zakrzewska, Marcinkowska,
& Wiedłocha, 2008). Once bound to a FGF ligand, the cell surface HSPG:FGF:FGFR and/or
FGF:FGFR units dimerize which facilitates transphosphorylation of the kinase domains on the intracellular side of the cell and activation of the receptor (Eswarakumar et al., 2005; Murakami et al., 2008; Schlessinger, 2000; Xiuqin Zhang et al., 2006). On FGFR1, Tyr653 and Tyr654, which reside in the activation loop, are the first residues to be phosphorylated and initiate an autophosphorylation cascade of other tyrosine residues (M Mohammadi et al., 1996).
Phosphorylated Tyr766 is a binding site for the SH2 domain of phospholipase Cγ (PLCγ) and the adaptor protein, Shb (Murakami et al., 2008). Tyr463 serves as a binding site for phosphotyrosine binding proteins, FGFR substrate 2α, β (FRS2α and FRS2β). FRS2α and β are docking proteins, that are also phosphorylated during receptor activation, and they assemble other downstream signaling molecules such as Src homolog (SH2) domain-containing adaptors Grb2 or Shp2. Grb2 recruits the docking protein Gab1 (Grb2-associated binder 1) and the resulting complex activates
Ras and the MAP kinase (ERK1/2, p38, JNK) pathways (Eswarakumar et al., 2005; Presta et al.,
2005; Turner & Grose, 2010). ERK1/2 activation is involved in endothelial cell proliferation, migration and differentiation while the p38 kinase is linked to FGF-mediated endothelial cell differentiation (Matsumoto, Turesson, Book, Gerwins, & Claesson-Welsh, 2002; Pintucci et al.,
2002; Presta et al., 2007). Gab also recruits and activates the cell survival phosphoinositide 3- kinase (PI3K)/Akt pathway via SH2 domains (Eswarakumar et al., 2005).
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B. Angiogenic Fibroblast Growth Factors
Of the 22 proteins that make up the FGF family, FGF1, FGF2, FGF4 and FGF5 have been
shown to play a role in vascular growth and formation: (Presta et al., 2005). FGF1 and FGF2 are
the most extensively studied and are considered the prototypes of the vascular FGFs. FGF2 will
be discussed in further detail in the sections below.
FGF1 was first isolated from cultured tumor-derived endothelial cells as a mitogen for 343 fibroblasts (Folkman, Merler, Abernathy, & Williams, 1971). It was purified from bovine brain and pituitary using heparin affinity chromatography and found to have an isoelectric point of 5.6 which helped to distinguish it from basic FGF (FGF2, isoelectric point 9.6) which also had a high affinity for heparin and was mitogenic (Böhlen, Esch, Baird, & Gospodarowicz, 1985; Lobb &
Fett, 1984; Thomas, Rios-Candelore, & Fitzpatrick, 1984). Sequencing of bovine brain FGF1 and bovine pituitary FGF2 helped to further characterize FGF1 and to differentiate it from FGF2 though the two growth factors were found to have significant structural homology (Böhlen et al.,
1985; Esch et al., 1985). FGF1 is the “universal” FGF due to its ability to bind and activate all subtypes of the FGFRs (Eswarakumar et al., 2005; Itoh & Ornitz, 2011; Xiuqin Zhang et al., 2006).
The 17.5 kDa FGF1 protein is widely distributed including in primary ECs and vSMCs (Antoine et al., 2005; Fon Tacer et al., 2010). The biological activity of Fgf1 was examined by the generation of mice lacking FGF1. These mice were found to be viable and possess normal neuronal physiology and wound healing capabilities (Miller, Ortega, Bashayan, Basch, & Basilico, 2000).
The mitogenic and angiogenic properties of exogenous FGF1 have been observed in cultured human umbilical vein ECs, the chick chorioallantoic membrane and subcutaneous matrigel plugs (Lobb, Alderman, & Fett, 1985; Lobb & Fett, 1984; Mühlhauser et al., 1995;
Thomas et al., 1985). Topical application of FGF1 was shown to increase the healing rate of
24
excisional dermal wounds in healing-impaired diabetic mice (Mellin et al., 1995). Administration
of FGF1 DNA plasmids or recombinant protein to experimental models of PAD produced positive
results and highlighted the potential for the use of FGFs in treatment of ischemic diseases. Rabbits
(Pu et al., 1993, 1994; Tabata, 1997; Witzenbichler et al., 2006), rats (Rosengart et al., 1997) and
hamsters (Caron et al., 2004) subjected to unilateral femoral artery ligation and treated with recombinant or plasmid FGF1 had increased numbers of angiographically visible collateral vessels and enhanced blood flow. The positive results from these animal studies and a phase I safety study of recombinant FGF1 in 20 patients with three-vessel coronary artery disease confirmed the
clinical utility of growth factors in arterial stenoses (Schumacher, Pecher, von Specht, &
Stegmann, 1998). An open-label phase I safety study of NV1FGF (nonviral recombinant human
FGF1 DNA plasmid) added to the growing list of proof-of-concept (safety) studies and further encouraged the development of larger double-blind placebo controlled II/III trials (Comerota et al., 2002). The Therapeutic Angiogenesis Leg Ischemia Study for the Management of Arteriopathy
and Non-healing ulcers (TALISMAN), phase II trial of 125 CLI patients was the largest of the
phase II studies. NV1FGF intramuscular treatment did not produce differences from the primary
endpoint of ulcer healing. However, secondary endpoints (occurrence of amputation or death) were significantly reduction (Nikol et al., 2008). This study was followed by the TAMARIS trial, a phase III double-blind placebo-controlled multinational clinical trial (Belch et al., 2011). The trial enrolled 525 CLI patients with ischemic lesions and is the largest gene therapy that has been carried out for PAD. The NV1FGF treated-group did not differ from the placebo patients in the primary endpoint of time to major amputation or death. Despite the disappointing results of NVIFGF treatment, subsequent follow-up studies have revealed no adverse ischemic events, retinopathies or malignancies (Belch et al., 2011; Silvestre et al., 2013).
25
The Fgf4 and Fgf5 genes were both discovered during screenings of human tumor for genes with a transforming effect on 3T3 fibroblasts (Bovi et al., 1987; Yoshida et al., 1987; Zhan,
Bates, Hu, & Goldfarb, 1988; Zhan, Culpepper, Reddy, Loveless, & Goldfarb, 1987). They were classified as oncogenic proteins and fibroblast growth factors based on 43% (FGF4) and 50%
(FGF5) amino acid sequence homology to FGF2 (Yoshida et al., 1987; Zhan et al., 1988). The functions of the Fgf4 and Fgf5 were investigated with the development of mice containing targeted deletion of the genes. Fgf4 was found to be necessary for development as embryos died after uterine implantation at embryonic day 4-5 (Feldman et al., 1995). Fgf5-/- mice were viable but had abnormal long hair and suggests an inhibitory role for FGF5 in regulation of the hair growth cycle (Hébert et al., 1994). Fgf4 and Fgf5 are both widely expressed during development while adult expression of Fgf4 is low and limited to the intestines and testes (Feldman et al., 1995; Fon
Tacer et al., 2010; Haub & Goldfarb, 1991). High Fgf5 expression is observed in the adult central nervous system with moderate expression in muscle and bone (Fon Tacer et al., 2010; Haub,
Drucker, & Goldfarb, 1990).
The mitogenic activity of recombinant FGF4 protein has been established in cultured human umbilical vascular endothelial cells (Miyagawa et al., 1988), rat avascular cornea assay and in the chick chorioallantoic membrane (Dell’Era et al., 2001; Yoshida et al., 1994). In cDNA- transfected murine aortic endothelial cells, endogenous Fgf4 increased proliferation but had no effect on morphogenesis in anchorage-dependent models such like 3D fibrin and EC matrix gels.
The proliferative capacity of FGF4 and affinity for FGFRs was also reduced compared to FGF2
(Dell’Era et al., 2001; Rissanen et al., 2003). The therapeutic angiogenic potential of FGF4 for treatment of ischemic disease was also investigated similarly to FGF1 in animal models of hindlimb and myocardial ischemia. Replication-incompetent serotype 5 adenoviruses (Ad5) were
26
implemented to deliver human Fgf4 (Ad5FGF4) to a rabbit lower limb ischemia model. The
Ad5FGF4 therapy promoted angiogenesis and arteriogenesis and increased popliteal blood flow
and muscle perfusion (Rissanen et al., 2003). The vascular effects of Ad5FGF4 were reported to
be direct via FGF4-induced EC proliferation and indirect via increased expression of endogenous
VEGF (Deroanne et al., 1997). Similar improvements in regional tissue perfusion and function
were observed in stress-induced porcine myocardial ischemia models with intracoronary delivery
of Ad5FGF4 (Gao et al., 2004; Roth et al., 2006).
The preclinical results served as the basis for the Angiogenic Gene Therapy (AGENT) trial,
a first-in-man study of intracoronary administration of Ad5FGF4. The randomized placebo- controlled study enrolled only 79 patients with chronic stable angina and after 12 weeks,
improvements from baseline in the primary endpoint (exercise tolerance time, ETT) were observed
(Grines et al., 2002). Another small study, the Phase II AGENT 2 trial, was performed to determine if Ad5FGF4 could improve myocardial perfusion in 51 patients with stable angina and reversible ischemia. The study proved to be too small and underpowered for conclusive results; however, additional safety and feasible data were obtained (Grines et al., 2003). The safety profile of
Ad5FGF4 and the results of the small AGENT/AGENT2 trials propelled the initiation of two nearly-identical double-blind place-controlled phase III trials, AGENT3 and AGENT4. The former was carried out exclusively in the United States while the later was conducted in Europe,
Latin America and Canada (Henry et al., 2007). Together the AGENT3 and AGENT4 trials enrolled 532 recurrent stable angina patients who received intracoronary infusions of placebo or
Ad5FGF4. Single pooled analysis of results from both trials revealed no significant improvements in the primary endpoint of change from baseline of total ETT at 4 weeks, 12 weeks or 6 months.
27
5. Fibroblast Growth Factor 2 (FGF2)
FGF2 was originally isolated from bovine pituitary in the mid-1970s as a 146 amino acid protein that was mitogenic for 3T3 fibroblasts and endothelial cells (Y. Cao et al., 2011; Ribatti,
2014). FGF2 was also referred to as basic FGF (bFGF) due to its isoelectric point of 9.6 and the loss of biological activity at acidic pH values (Gospodarowicz, Bialecki, & Greenburg, 1978;
Gospodarowicz, 1975). In 1984, a growth factor derived from a rat chondrosarcoma was discovered and purified by Shing and co-workers based on the high binding affinity for heparin
(Shing et al., 1984). This factor was described as a cationic polypeptide with a molecular weight
of about 18 kilodaltons (kDa) with the ability to stimulate proliferation of BALB/C 3T3 fibroblasts
and capillary endothelial cells in vitro as well in vivo angiogenesis in the chorioallantoic membrane
(CAM) and yolk sac membrane of the developing chick (Shing et al., 1985). A heparin-binding
angiogenic factor was also purified from human placenta and a human hepatoma cell line and
shown to stimulate protease production, DNA synthesis, and migration capillary endothelial cells
(Moscatelli, Presta, & Rifkin, 1986; Presta, Moscatelli, Joseph-Silverstein, & Rifkin, 1986). The
factor was then sequenced and identified as fibroblast growth factor 2 (FGF2) (Esch et al., 1985).
Soon after, it was discovered that additional FGF2 polypeptides were translated from the human
Fgf2 gene and these proteins were described as high molecular weight forms of FGF2 (Robert Z
Florkiewicz & Sommer, 1989; Prats et al., 1989). FGF2 has since been studied extensively and
shown to have pleiotropic and mitogenic effects on myriad cell types from endoderm, mesoderm,
and neuroectoderm lineages (Okada-Ban, Thiery, & Jouanneau, 2000). The functions of the low
(18kDa) and high molecular weight isoforms (21, 22, 24 and 34kDa) of FGF2 will be discussed in
further detail below.
28
A. Fgf2 gene and its tissue expression
The single copy human Fgf2 gene is mapped to q26 on chromosome 4, and is composed of three exons separated by two 16 kilobase introns and extends to 40 kilobases (Okada-Ban et al., 2000; Ornitz & Itoh, 2001). The gene is highly conserved across many species including chimpanzee, Rhesus monkey, dog, cow, mouse, rat, chicken, zebrafish, and frog (Okada-Ban et al., 2000).
FGF2 is widely distributed in cells of mesenchymal, mesodermal and neuroectodermal lineages and is expressed during development and in the adult. In situ hybridization and RT-PCR analysis detected expression of Fgf2 in all stages of Xenopus and rodent embryonic development
(Gonzalez, 1990; Lea, Papalopulu, Amaya, & Dorey, 2009; Yaylaoglu et al., 2005). In adult murine tissues, Fgf2 mRNA is expressed in the brain, heart, liver, aorta, gut, skin, bone, skeletal muscle, reproductive organs and blood vessels (Fon Tacer et al., 2010). A similar expression pattern was observed in adult mouse tissue sections that were analyzed by immunostaining. In the vasculature, FGF2 was present in the basement membrane and smooth muscle of the tunica media of mid-size vessels while capillary vessels showed expression in the basal lamina and endothelium
(Cordon-Cardo, Vlodavsky, Haimovitz-Friedman, Hicklin, & Fuks, 1990). Primary co-cultures of human umbilical vascular endothelial and arterial smooth muscle cells also had strong expression of Fgf2 (Antoine et al., 2005).
B. Phenotypes of Fgf2-/- mice (Role in development, homeostasis and injury)
Several, independently developed lines of mice deficient in the Fgf2 gene expression have been produced to describe the roles of FGF2 in development and physiology. House and colleagues have shown that normal growth of capillary and smooth muscle-containing vessels occurs in FGF2
29
knockout hearts of mice bred on a 50:50 129/Sv and Black Swiss background (House et al., 2003).
A similar result was observed by the Murray group in Fgf2-/- mice of a different strain, C57BL/6J
(Virag et al., 2007). Mice bred on a C57BL/6J x 129/Sv mixed background, however, had impaired myocardial capillary density when Fgf2 gene expression was partially (+/-) or completely (-/-)
disrupted (Amann et al., 2006). Fgf2-deficient mice have normal retinal vascularization and
neovascularization responses to retinal or choroid layer injury (Ozaki et al., 1998; Tobe et al.,
1998). Ablation of Fgf2 delayed healing of dorsal skin excisional wounds (Ortega, Ittmann, Tsang,
Ehrlich, & Basilico, 1998). Sullivan and investigators showed that in Fgf2-/- hindlimbs, vascular
growth occurs to a similar degree as wildtypes in response to chronic ischemia (Sullivan,
Doetschman, & Hoying, 2002). FGF2 deficiency decreased the effect of the sex hormone 17β- estradiol (E2) on the proliferation and migration of cultured endothelial cells (Garmy-Susini et al.,
2004). After carotid arterial vessel injury, E2-mediated acceleration of re-endothelialization and
endothelial progenitor cell (EPC) mobilization was attenuated in the absence of FGF2 (Fontaine
et al., 2006), while flow-dependent remodeling was not changed (Sullivan & Hoying, 2002) and
normal neointimal hyperplasia was present (Zhou et al., 1998).
Other cardiac phenotypes of the Fgf2-/- mice include resting hypotension, decreased portal
vein vascular smooth muscle contractility, normal basal cardiac function and impaired
baroreceptor reflex (Dono, Texido, Dussel, Ehmke, & Zeller, 1998; Ortega et al., 1998; Pellieux et al., 2001; Schultz et al., 1999; Zhou et al., 1998). Normotensive Fgf2-/- mice exhibited baseline dilated cardiomyopathy with increased left ventricular chamber diameter at systole or diastole
(Pellieux et al., 2001). When subjected to different experimental hypertension models, including angiotensin-induced hypertension (Pellieux et al., 2001), pressure-overload (Schultz et al., 1999), or chronic β-adrenergic stimulation (House et al., 2010); Fgf2-/- hearts were protected from
30 compensatory cardiomyocyte hypertrophy. This blunted hypertrophic response was also present in Fgf2-ablated hearts after permanent occlusion of the left-descending coronary artery, a model of chronic ischemia (Virag et al., 2007) and in response to a closed-chest model of cardiac ischemia-reperfusion injury (House et al., 2015). In both models, cardiac cell death (measured as infarct size) was significantly greater relative to wildtype hearts. The later study also reported reduced cardiac vessel density after 7 days of reperfusion (House et al., 2015). Infarct size was also increased in response to low-flow I/R injury and this resulted in reduced recovery of post- ischemic cardiac function in Fgf2-/- hearts (House et al., 2003).
Non-cardiovascular phenotypes in the FGF2-deficient mice were observed, suggesting a role for FGF2 in bone and nerve patho(physiology). For example, bone mass and formation are decreased in Fgf2-/- femora and tibiae and this osteopenia progressed with age (Montero et al.,
2000). Reduced neuron density and disorganization has been shown in the cerebral cortices of
FGF2-deficient mouse brains (Dono et al., 1998; Korada, Zheng, Basilico, Schwartz, & Vaccarino,
2002; Ortega et al., 1998; Raballo et al., 2000). The neuronal deficit in Fgf2-/- mice was not a factor of defective neural stem cell (NSC) proliferation during embryonic or adult neurogenesis, but rather an impairment in the differentiation of the NSCs into mature neurons (Werner, Unsicker,
& von Bohlen und Halbach, 2011). In response to peripheral nerve crush injury, increased macrophage invasion, regeneration of myelinated axons and greater axon and myelin diameters resulted from an absence of Fgf2 (Jungnickel, Claus, Gransalke, Timmer, & Grothe, 2004).
Additionally, sciatic nerve-injured knockout mice exhibited faster mechanosensory recovery
(Jungnickel et al., 2010).
C. FGF2 protein isoforms
31
The Fgf2 gene produces several protein isoforms from a single mRNA. The protein
isoforms are grouped into two classes: low molecular weight (LMW) and high molecular weight
(HMW). A single LMW FGF2 (18kDa) is translated from a classical AUG codon while the HMW
isoforms are translated from in-frame alternative translational sites at CUG codons that are located
upstream of the LMW start site (Touriol et al., 2003). The 18kDa LMW FGF2 isoform is present
in mice and humans. Humans express four HMW isoforms (22, 22.5, 24, and 34kDa) while mice
express two HMW isoforms (21.5 and 22kDa) (Okada-Ban et al., 2000) (C. H. Chen, Poucher, Lu,
& Henry, 2004; Liao et al., 2009; Okada-Ban et al., 2000). Structurally, the HMW isoforms are co-linear extensions of the LMW isoform with amino terminal nuclear localization signals that localize them to the cell nucleus (Chlebova et al., 2009; Yu, Ferrari, Galloway, Mignatti, &
Pintucci, 2007) (Figure 2). In neuronal cells, HMW isoforms appear in a punctuate pattern within the nucleoplasm, around nucleoli and associated with chromatin (Claus et al., 2003; Förthmann,
Grothe, & Claus, 2015). LMW FGF2 is mostly a cytosolic protein but has been found to localize to nucleoli in the nucleus and is stored in the extracellular matrix (Claus et al., 2003; Förthmann et al., 2013; Liao et al., 2010; Quarto, Fong, & Longaker, 2005).
The precise manner of secretion of LMW FGF2 is still unknown because like FGF1, it lacks a conventional export signal sequence for transport via the canonical endoplasmic reticulum
(ER)/Golgi secretory pathway (Mignatti, Morimoto, & Rifkin, 1992). Proposed mechanisms for release of FGF1 and FGF2 into the extracellular space include an ATP-dependent manner that implicates Na+/K+-ATPase, vesicular shedding from plasma membranes, or from non-lethal
damage or disruption of membranes (Florkiewicz, Anchin, & Baird, 1998; Sørensen, Nilsen, &
Wiedłocha, 2006; Taverna et al., 2003). Specific HMW or LMW FGF2 isoform detection has been
performed using Western immunoblotting. Several groups have reported expression of the LMW
32
(18kDa) and HMW (21, 22kDa) FGF2 isoforms in mouse brain, liver, heart, lung and spleen
(Azhar et al., 2009; Claus, Werner, Timmer, & Grothe, 2004; Coffin et al., 1995; Garmy-Susini et
al., 2004; House et al., 2003; Miller et al., 2000). Selective expression of LMW FGF2 was
observed in the kidney and skeletal muscle (Coffin et al., 1995). In rat embryonic whole brain,
only 18kDa and 21kDa FGF2 were expressed until birth when the 22kDa could then be detected.
A similar pattern was observed in embryonic spinal cord and cortex extracts while adult extracts
expressed higher levels of 21 and 22kDa FGF2 (S. Giordano, Sherman, Lyman, & Morrison,
1992). In the adult cerebellum, however, 18kDa FGF2 was the predominant form and the
expression of the 21kDa protein disappears at birth (S. Giordano et al., 1992). Adult rat heart
ventricles had selectively higher expression of the 18kDa protein in contrast with neonatal
ventricles where the 22kDa isoform was preferentially expressed (L. Liu, Doble, & Kardami,
1993).
33
Figure 2: Canonical (AUG, LMW) and alternative (CUGs, HMW) FGF2 protein isoform translation. Adapted from (Liao et al., 2009).
D. Biological functions of the FGF2 isoforms
FGF2 has been shown to play a pathophysiological role in diverse biological processes, including development, tumorigenesis, wound healing, hematopoiesis, neuronal development, limb formation, cardiac hypertrophy, and vascular growth (C. H. Chen et al., 2004; Fei,
Gronowicz, & Hurley, 2013; Kardami et al., 2007; Liao et al., 2009; Ortega et al., 1998; Presta et al., 2005; Schultz et al., 1999; Shing et al., 1985; Zhou et al., 1998). The pleiotropic effects of
FGF2 in a multitude of biological systems can be linked to the alternative translation of its protein isoforms and their different subcellular localization.
i. Vascular Biology
The initial discovery and characterization of FGF2 was based on its isolation from and mitogenic effects on tumor-derived endothelial cells (ECs) and vascular smooth muscle cells
34
(vSMCs) (Gospodarowicz et al., 1978; Shing et al., 1985). The role of FGF2 has since expanded to include complex cellular processes that contribute to physiological and/or tumor-associated angiogenesis (Auguste, Javerzat, & Bikfalvi, 2003; Javerzat, Auguste, & Bikfalvi, 2002; Presta et al., 2007). These include EC and vSMC proliferation, chemotactic migration, protease production, extracellular matrix (ECM) degradation, morphogenesis and assembly of nascent or remodeled vascular structures (Carmeliet & Jain, 2011; Fischer et al., 2006; Presta et al., 2005). In blood vessels, FGF2 is produced and secreted by vascular ECs and its mechanism of action can be intracrine, paracrine and/or autocrine. Expression of similar levels of Fgfr1IIIc and 3IIIc mRNA
have been shown in ECs and vSMCs, while lower amounts of Fgfr3IIIc and no Fgfr4 were detected
(Antoine et al., 2005). In MCF-7, aortic endothelial and 3T3 cells, only FGFR1 protein expression was detected (Piotrowicz, Ding, Maher, & Levin, 2001). FGF2 is also found in the ECM where it is bound to matrix components including fibronectin, αvβ3 integrins, vimentin, collagen type I,
thrombospondin-1 and HSPGs (Presta et al., 2005, 2007). The ECM-immobilized FGF2 is a
stimulus for angiogenesis when it is freed by proteases and glycosidases and it promotes cell
adhesion, activation and motility via recruitment of αvβ3 receptors and focal adhesion kinases
(Folkman et al., 1988; Presta, Maier, Rusnati, & Ragnotti, 1989; Tanghetti et al., 2002).
To date, only the LMW FGF2 (18kDa) isoform is commercially available in recombinant
form and this protein has been applied exogenously to determine the functions of FGF2. Due to
the distinct subcellular locations and properties of the FGF2 isoforms, it has become critical to
employ genetic models where only one class of the FGF2 isoforms are expressed to examine the
specific functions of the endogenous FGF2 proteins. These mouse models will be described in the
following sections “Phenotypes of FGF2 isoform knockout mice”.
35
ii. Non-vascular effects of exogenous FGF2 isoforms
Studies on the mitogenic functions of exogenous FGF2 isoforms extend beyond their effects on vascular-derived cells to include cells/tissues of mesodermal origins including cardiomyocytes and myofibroblasts in the heart (Fujiu & Nagai, 2014; Kardami et al., 2007;
Santiago et al., 2011). Exogenous administration of recombinant LMW FGF2 to intact rodent hearts undergoing surgical corornary artery occlusion or ex vivo hearts being subjected to acute ischemia significanlty reduced the dysfunction associated with these models of myocardial infarction (MI) (Jiang et al., 2002; Jiang, Srisakuldee, Soulet, Bouche, & Kardami, 2004; Sheikh,
Sontag, Fandrich, Kardami, & Cattini, 2001). Futhermore, myocyte injury (infarct size) was decreased in the FGF2-treated hearts. The protective effect of the LMW isoform was maintained for up to 6 weeks (in vivo studies) and was not dependent of the timing of the ischemic injury; i.e., exogenous LMW FGF2 was beneficial if given to the hearts before, during or after the development of ischemia (during reperfusion) (Jiang et al., 2002, 2004; Kardami et al., 2007). A mutant form of the LMW isoform (S117A) lacking mitogenic or angiogenic ability still retained the potential for cardioprtection (Jiang et al., 2004).
When injected into infarcted rat hearts, recombinant HMW FGF2 (23kDa) alone was also effective in preventing myocyte death and improving functional recovery (Jiang et al., 2007). This level of protection observed was similar to hearts that had been treated with the LMW isoform.
The LMW FGF2 group had sustained cytoprotective effects and protection from contractile dysfunction for up to 6 weeks after the initial treament (Jiang et al., 2007). Increased small vessel
(arteriole) density was also detected after 6 weeks of MI in the LMW FGF2-infarcted hearts. HMW
FGF2 had no effect on vessel growth but increased heart dimensions (LV mass) and cardiomyocyte size were observed after 8 week of MI. This hypertrophic response to HMW FGF2 was confirmed
36 in cultured neonatal cardiomyocytes and non-cardiac myocytes treated with the exogenous isoform (Jiang et al., 2007; Kardami et al., 2004; Santiago et al., 2011). The acute protection by exogenous LMW or HMW FGF2 was found to be dependent on activation of FGFR1 and related downstream signaling proteins including PKCα, ε, δ, and ζ (Jiang et al., 2002, 2004). Several studies using Fgf2-/- mice (described in “Phenotypes of Fgf2-/- mice”) have documented a role for the Fgf2 gene in chronic adrenergic stimulation or pressure overload-induced cardiac hypertrophy. The actions of the recombinant HMW isoform would imply that nuclear signaling by
HMW FGF2 and not LMW FGF2 is responsible for the role of Fgf2 in development of cardiac hypertrophy.
The in vitro and in vivo studies described here to characterize the properties of FGF2 have utilized exogenous (recombinant) forms of the proteins. While LMW FGF2 is exported from the cell and can signal via the cell surface FGFRs, HMW FGF2 isoforms are nuclear proteins which share considerable sequence homology with LMW FGF2 including receptor binding sites. This would suggest that activities of exogenous HMW FGF2 may mimic that of LMW FGF2 signaling, however, physiological functions of HMW FGF2 would have to be confirmed in studies with endogenous HMW FGF2 (Chlebova et al., 2009; Gualandris, Urbinati, Rusnati, Ziche, & Presta,
1994).
iii. Effect of overexpression of FGF2 isoforms
The functional effects of the FGF2 isoforms have also been studied in models of Fgf2 gene overexpression in vitro and in vivo. Mouse aortic endothelial (MAE) cells transfected with human
Fgf2 cDNA displayed an angiogenic phenotype when plated on ECM gels and in the avascular rabbit cornea. This activity was inhibited by a neutralizing FGF2 antibody and suggests an autocrine mechanism for this phenotype (Gualandris et al., 1996). Aortic vascular smooth muscle
37
cells from transgenic mice expressing all the human FGF2 isoforms, all the HMW FGF2 isoforms
or only the 18kDA isoform had enhanced DNA synthesis with increased proliferation observed
when only the nuclear HMW isoforms were overexpressed (Davis et al., 1997). Preservation of
the increased cell growth in presence of antibody neutralization, established an intracrine role for
FGF2 in vascular cell growth (Davis et al., 1997). When 3T3 fibroblasts overexpress only LMW
or only HMW FGF2, different cellular phenotypes emerge. The HMW-expressing cells had
impaired growth relative to wildtype cells, while the LMW FGF2 cells resembled transformed
cells with enhanced saturation density and growth in low serum (Quarto, Talarico, Florkiewicz, &
Rifkin, 1991; Wieder et al., 1997).
In response to transient overexpression of LMW FGF2 or HMW FGF2 cDNA in rat
neonatal ventricular myocytes, similar increases in myocyte DNA synthesis and proliferation were
observed (Pasumarthi, Kardami, & Cattini, 1996). When HMW FGF2 (22/21.5kDa)
overexpression and not LMW FGF2 was present, there was increased binucleation of the myocytes
that was not inhibited by FGF2 neutralizing antibodies. Bi- or multi-nuclear phenotype has also
been reported in HMW FGF2-overexpressing neuronal cells (Nindl et al., 2004). These studies
provide further evidence for an intracrine (nuclear) pathway of HMW FGF2 signaling. Mice with
cardiac-specific overexpression of LMW FGF2 had unchanged heart weight-to-body weight ratios and arteriolar vessel density as well as normal expression of cardiac differentiation markers
(Sheikh et al., 2001). Capillary density and expression of MAP kinases (JNK, p38) and PKC isoforms (α, ε) were increased in unstressed LMW FGF2 transgenic hearts. After ex vivo global ischemia-reperfusion (I/R) injury, overexpression of LMW FGF2 was shown to elevate intra- and extracellular FGF2 expression in addition to preventing myocyte death (Sheikh et al., 2001). These results are partially consistent with a study in which cardiac specific overexpression of all human
38
FGF2 (18, 22-34 kDa) isoforms likewise produced no basal changes in vessel densities in the heart
but resulted in protection from I/R injury-induced myocyte loss (House et al., 2003). Conversely,
House and co-workers reported improvements in recovery of myocardial function with normal
expression of MAP kinases and PKC isoforms with the exception for PKCε whose expression was
reduced (House et al., 2003; House, Branch, Newman, Doetschman, & Schultz, 2005; House,
Melhorn, Newman, Doetschman, & Schultz, 2007). Alterations in the activation of p38 and JNK
kinases only occurred in response to I/R injury as overexpression of all human FGF2 isoforms
already had modified p38 and JNK activation independent of ischemia (House et al., 2005). The
human FGF2 transgenic hearts exhibited an exaggerated hypertrophic response to isoproterenol
stimulation in the form of decreased fractional shortening, increased heart weight-to-body weight
ratios and myocyte cross-sectional area.
E. Phenotypes of FGF2 isoform knockout mice (role of endogenous FGF2 isoforms)
FGF2 is a widely expressed growth factor and promotes mitogenic activity in a variety of
cell types including vascular cells (Davis et al., 1997), fibroblasts (Gospodarowicz, 1975),
cardiomyocytes (Pasumarthi et al., 1996), skeletal myocytes, neurons (Grothe, Haastert, &
Jungnickel, 2006), and osteoblasts (Xiao et al., 2003). LMW FGF2-deficient (FGF2 LMWKO) or
HMW deficient (HMWKO) mouse models are viable and fertile. Using microvascular endothelial
cells isolated from ovariectomized LMWKO mice, Garmy-Susini and co-workers observed that
HMW FGF2 was necessary for 17 β-estradiol (E2)-mediated migration and angiogenesis (Garmy-
Susini et al., 2004). This effect of HMW FGF2 was FGF receptor-independent but involved the
anti-apoptotic protein FGF2-interacting Factor (FIF), a putative intracellular signaling partner of
HMW FGF2 (Chlebova et al., 2009; Krejci et al., 2007). To date, this in vitro work by the Arnal
39
group represents the only study undertaken to examine the in vivo effects of the endogenous HMW
FGF2 isoforms in vascular biology. FGF2 isoform knockouts have provided an opportunity to further examine the unique functions of the endogenous FGF2 isoforms in the vascular pathophysiology. Three different groups have performed experiments using these unique mouse models to distinguish the biological roles of the FGF2 isoforms in the heart, bone and brain.
Findings from these studies are summarized below. Taken together, they confirm that when expressed endogenously, the HMW and LMW isoforms of FGF2 play different roles in tissue homeostasis and pathophysiology.
Heart
In parallel with the characteristics of Fgf2-/- mice (described in “Phenotypes of Fgf2-/-
mice”), FGF2 LMWKO and FGF2 HMWKO mice exhibit normal vascular growth and altered
FGF2 expression. Capillary and arteriole density is not significantly different between FGF2
LMWKO (HMW expressing) or FGF2 HMWKO (LMW expressing) and WT hearts. While the
expression of the 21 and 22kDa FGF2 (HMW) isoforms are unchanged in the LMWKO hearts
(Liao et al., 2007), 18kDa (LMW) FGF2 is increased by 1.5 fold in HMWKO hearts (Liao et al.,
2010). The subcellular localization of the HMW isoforms was confirmed to be exclusively nuclear
and LMW FGF2 was expressed in the cytoplasm and nucleus (Azhar et al., 2009; Liao et al., 2009).
The increased level of LMW FGF2 in the absence of HMW FGF2 played a protective role in
ischemia–reperfusion injury
When subjected to an ex vivo work performing model, the knockout hearts displayed no
changes in baseline contractile and relaxation parameters (Liao et al., 2007, 2010). After induction
of low-flow ischemia and reperfusion (I/R), impaired recovery of post-ischemic contractile
40 function was observed in LMWKO hearts (Liao et al., 2007). Conversely, improved recovery of function occurred in HMWKO hearts (Liao et al., 2010). The cardioprotective effect of LMW
FGF2 was shown to be dependent on the activation of FGFR1 and inhibition of JNK signaling
(Liao et al., 2007). Creatine kinase release and myocardial infarct size, measures of cell injury were not different from WT indicating a requirement of LMW and HMW FGF2 isoforms in the protection from cell death (Liao et al., 2007, 2010).
Recently, the Doestchman group has performed echocardiographic measurements to characterize the interactions between sex and the FGF2 isoforms in regulating cardiac physiology and modulating remodeling in (Nusayr & Doetschman, 2013; Nusayr, Sadideen, & Doetschman,
2013). In unstressed hearts, male and female mice LMWKO (HMW expression only) hearts had impaired basal diastolic function (compliance and relaxation) while only female LMWKO hearts had abnormal cardiac development with smaller LV masses (Nusayr & Doetschman, 2013).
HMWKO (LMW expression only) males had significantly higher systolic functional parameters
(cardiac output, stroke volume and fractional shortening) than their female cohorts as well as in comparison to male WTs or LMWKOs. After chronic β-adrenergic stimulation, only female
LMWKO hearts had an elevated hypertrophic response while their male cohorts had increased fibrosis and cardiac remodeling (Nusayr et al., 2013). Male HMWKOs had enhanced expression of the cardiac stress marker, atrial natriuretic factor (ANF) which protected them from the higher cardiac fibrosis that was present in WT and LMWKO male hearts (Nusayr et al., 2013). These observations suggest that in addition to the isoform-specific effects on injury response, there is a sex-specific regulation of cardiac patho(physiology) by FGF2.
Bone growth and remodeling
41
FGF ligands and FGF receptors play important roles in bone formation, homeostasis, repair
and remodeling. In humans, several gain-of-function and loss-of-function point mutations in
FGFRs (FGFR1, 2 and 3) result in deficits of digital, cranial, and skeletal development including craniosynostosis and skeletal dysplasias (Belov & Mohammadi, 2013; Eswarakumar et al., 2005;
Marie, Miraoui, & Sévère, 2012). Phenotypes of FGFR knockout mouse models (summarized in
Table 3) are consistent with the human disorders. Mutations in several FGFs including FGF3,
FGF8, FGF9, FGF10 and FGF23 have also been shown to cause human skeletal diseases (Du et al., 2012).
The role of the FGF system in bone regulation is also evident in mouse models with altered expression of FGF2. Abnormal bone development occurs when LMW and HMW FGF2 are overexpressed globally. These mice have moderate macrocephaly and dwarfism (shortening of the femora and humeri) (Coffin et al., 1995). Additionally, bone mineral density (BMD) and trabecular bone volume were reduced in the femora of the transgenic mice (Sobue et al., 2005). Targeted overexpression of LMW FGF2 in cells of osteoblastic lineages established a role for the LMW isoform as a positive regulator of bone mass. BMD, tibial and femoral bone volume and trabecular and cortical bone thickness were increased in the LMW FGF2 transgenic mice (Xiao et al., 2014).
Conversely, global LMW FGF2 knockout mice have decreased BMD and bone mineral content
(Xiao et al., 2009). When the HMW isoforms are selectively overexpressed in mouse osteoblasts, dwarfism, decreased BMD, osteomalacia and hypophosphatemia were observed (Xiao et al., 2010;
Xiao, Esliger, & Hurley, 2013). This hypophosphatemia phenotype is also present in the ostoeblast-specific overexpression of another FGF, FGF23 (Xiao et al., 2010). FGF23 in the serum and bone mRNA and protein were increased in the HMW transgenic mice. HMW FGF2 regulation of bone phosphate metabolism and inhibition of bone formation was shown to be dependent on
42
FGF23 activation of FGFRs and the MAPK pathway (Xiao et al., 2013). In cultured calvarial
osteoblasts (COBs) isolated from LMW FGF2 or HMW FGF2 newborn transgenic mice,
differential regulation of bone morphogenic protein-2 (BMP-2) was found to mediate the disparate
effects of the FGF2 isoforms on bone growth (Sabbieti et al., 2013). BMP-2 is potent inducer of
osteoblast differentiation and bone formation. When added to COBs, BMP-2 promoted or inhibited
bone differentiation in response to LMW FGF2 or HMW FGF2 overexpression, respectively
(Naganawa et al., 2008; Sabbieti et al., 2013).
Altered expression of FGFs and FGFRs have been found in bone during fracture healing
and bone regeneration (Du et al., 2012; Fei et al., 2013). Bone mRNA expression of Fgf1, Fgf2,
Fgf5, Fgfrs and other markers of bone development are increased in rodent models of tibial and femoral fracture (Du et al., 2012; Schmid, Kobayashi, Sandell, & Ornitz, 2009). When subjected bilateral calvarial defects, male LMW transgenic mice had enhanced healing compared with non- transgenic mice (Xiao et al., 2009). Treatment with low doses of BMP-2 further improved the bone healing at later stage and led complete healing of the defect (Fei et al., 2013).
Neurogenesis and neurodegeneration
FGF2 has been shown to be important for the development of the central nervous system
(CNS) and in adult neurogenesis. FGF2 and FGFR expression is temporally and spatially regulated during development (Woodbury & Ikezu, 2013; Yu et al., 2007). Furthermore, Fgf2 mRNA and
protein expression is present in spinal ganglia and is up-regulated in response to peripheral nerve
injury (Grothe, Meisinger, Hertenstein, Kurz, & Wewetzer, 1996). The FGF2 isoforms were
differentially elevated in the nerve lesions with increased 18kDa isoform expression 5 hours after
injury, while the HMW FGF2 isoforms had a greater increase at 7 days (Grothe, Heese, et al.,
43
2000; Meisinger & Grothe, 1997a). Conversely, no changes in the expression of the FGF2 protein
isoforms or FGF receptor transcripts were observed in neurotoxin-induced lesions of dopaminergic
neurons (Claus, Werner, et al., 2004). Transfection of immortalized Schwann cells (SC) and cultured PC 12 cells with modified rat cDNAs expressing endogenous 18kDa or 21/23 kDa resulted in differential growth and morphological phenotypes (Grothe, Meisinger, Holzschuh,
Wewetzer, & Cattini, 1998). Proliferation of the 18kDa overexpressing Schwann and PC12 cells was decreased compared to the HMW FGF2 cells (Müller-Ostermeyer, Claus, & Grothe, 2001).
The HMW FGF2 isoforms emerged as neurotrophic factors when HMW FGF2 released from implanted SCs lead to improvements in the lengths and numbers of regenerating myelinated axons in a rat model of sciatic nerve axotomy (Timmer, Robben, Müller-Ostermeyer, Nikkhah, &
Grothe, 2003). In Parkinson’s-diseased rat brains, co-transplantation of dopaminergic grafts with
HMW FGF2 SCs also enhanced re-innervation, survival and functional recovery compared to the
18kDa SCs (Timmer et al., 2004). In vivo, grafted FGF2 isoform-modified SCs had differential effects on nerve regeneration across long gaps. The 18kDa grafts were inhibitory for myelination of regenerating axons while the 21/23kDa expressing SCs stimulated long-distance axon myelination and early recovery of sensory nerve functions (Haastert, Lipokatic, Fischer, Timmer,
& Grothe, 2006).
F. Exogenous LMW FGF2 in revascularization in vivo (Preclinical studies)
Models of angiogenesis
Exogenous FGF2 is a potent stimulator of angiogenesis in experimental angiogenesis assays (Presta et al., 2005, 2007; Shing et al., 1985). These include in vitro (EC proliferation,
Boyden chamber migration, EC tubule formation assays, 3-D collagen matrix gels); ex vivo organ
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cultures (rodent aortic ring) or in vivo (chick embryo chorioallantoic membrane (CAM), subcutaneous ECM gel sponges, dorsal skin fold chamber or avascular rodent cornea) assays
(Baker et al., 2012; Staton, Reed, & Brown, 2009). In early studies characterizing the activities of tumor-derived factors, an 18kDa protein stimulated proliferation in capillary bovine capillary endothelial cells (BCECs) and angiogenesis in chorioallantoic membranes and rat cornea (Shing et al., 1984, 1985). This protein, which was soon named FGF2, also induced DNA synthesis, motility, protease production in BCECs (Moscatelli et al., 1986; Presta et al., 1986) and capillary- like tubule formation in BCECs cultured on 3D collagen matrix gel (Montesano, Vassalli, Baird,
Guillemin, & Orci, 1986). After expression and purification from transformed Escherichia coli cells, exogenous treatment with recombinant 24kDa FGF2 was shown to have a similar effect as
18kDa FGF2 in promoting proliferation, protease production and chemotaxis of cultured bovine aortic endothelial cells (BAECs). Induction of angiogenesis in the rabbit cornea and binding of
FGF receptors was also similar in response to 18kDa or 24kDa FGF2 (Gualandris et al., 1994).
LMW FGF2-stimulated angiogenesis in cultured BCECs and in CAMs requires activation of the phosphoinositide 3-kinase (PI3K) pathway and is negatively regulated by the p38 MAP kinase pathway (Matsumoto et al., 2002; Qi et al., 1999).
Levin and colleagues observed that human HMW FGF2 isoforms have an inhibitory effect on BAECs and mammary carcinoma cells (MCF-7) in a Boyden chamber type assay (Piotrowicz,
Maher, & Levin, 1999). When both cell types were cultured in a monolayer and subjected to a wounding assay, recombinant HMW FGF2 inhibited migration of only BAECs (Piotrowicz et al.,
1999). HMW FGF2 had a similar proliferation-promoting effect on cells as the 18kDa. The inhibitory function of HMW FGF2 on cell migration was shown to be dependent on estrogen receptor α and regulated by the amino-terimal extension (ATE) of HMW FGF2 (Ding et al., 2002;
45
Piotrowicz et al., 2001). The inhibitory role of the ATE of HMW FGF2 on cell migration was also observed in vivo and found to be sufficent to suppress tumor angiogenesis in murine dorsal skin- fold model (Levin, Sikora, Ding, Rao, & Sriramarao, 2004). In another study utilizing exogenous
FGF2 in vivo, rat hearts had increased arteriole density in the peri-infarct after permanent coronary occlusion when treated with LMW FGF2. Conversely, HMW FGF2 did not have any effect on microvesels numbers (Jiang et al., 2007).
These in vivo models of angiogenesis have also been used to compare the efficacy of FGF2 with other proangiogenic growth factors. In particular, the corneal pocket assay has been employed to examine the vascular response to exogenous FGF2 relative to vascular endothelial growth factor
(VEGF) and platelet-derived growth factors (PDGF). The cornea is an avascular transparent tissue in the body, so any vessels penetrating into the corneal stroma are newly formed and easily quantified. The results from a number of studies have been summarized in Table 4. These have been limited to studies where FGF2 was compared directly with other growth factors to eliminate the possibility that difference in efficacies are due to variations in experimental parameters.
Overall, FGF2 induced corneal angiogenesis to a similar degree as PDGF-BB. Both FGF2 and
PDGF-BB were more efficacious in stimulating capillary network growth relative to other PDGF isoforms, PDGF-AA and PDGF-AB.
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Delivery Neovascularization Assay FGF2 PDGF-AA PDGF-AB PDGF-BB VEGF-A Reference (protein) endpoint Corneal Slow release Vascular area (H. Lu et +++ + ++ +++ NT micropocket polymer (mm2) al., 2007) Corneal Vascular area (R. Cao et Micropellets +++ + +++ +++ ++ micropocket (mm2) al., 2003) Corneal Slow release Vascular area (Nissen et +++ NT NT +++ NT micropocket polymer (mm2) al., 2007) Corneal Vascular area (J. Zhang et Micropellets +++ + ++ ++ NT micropocket (mm2) al., 2009)
Table 4: Angiogenic responses induced by recombinant LMW FGF2 protein relative to other proangiogenic growth factors in corneal micropocket assay. For the endpoint (vascular area), VEGF and PDGF are responses were estimated relative to the
FGF2 response .
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The early in vitro, ex vivo, and in vivo assays highlighting the proliferative effects of FGF2
and other growth factors served as the rationale for animal studies to evaluate the potential
pharmacological application of these “angiogenic” factors to stimulate angiogenesis and/or
collateral vessel formation in models of myocardial or peripheral ischemia. Experimental
myocardial and peripheral ischemia models have been beneficial in testing the efficacy of
angiogenic gene or protein delivery. For myocardial ischemia, a majority of initial experiments
focused on the vascular responses to progressive occlusion of coronary artery or arterial stenosis
in canine and porcine models (Hearse, 2000; Rubanyi, 2013; Teunissen, Horrevoets, & van Royen,
2012). Since then, other species including mouse, rats, and rabbits have also been studied.
Animal models of myocardial ischemia
The extent of the pre-existing collateral circulation has proved to be an important determinant of the appropriate animal model for analyzing post-ischemic revascularization
(Schaper, 2000). An extensive study by Maxwell, Hearse and Yellon has shown that a wide range
exists between mammalian species in the extent of the pre-formed collateral network (Maxwell,
Hearse, & Yellon, 1987). This network provides collateral flow upon coronary artery occlusion
and is responsible for outward vascular remodeling. Pigs, baboons, and rabbits have little to no
collateral flow (0-6% of pre-ischemia) after ischemia; dogs and cats have intermediate baseline
collateral flow (12-17%) whereas guineas pigs have total compensation (100%) of flow after
arterial occlusion of a single coronary artery. The canine models were the most commonly utilized
models, and the work by the Yellon group and others revealed an extensive collateral network in
dogs that was capable of providing 40% of baseline perfusion after acute coronary artery occlusion
(G. C. Hughes, Post, Simons, & Annex, 2003; Maxwell et al., 1987; Teunissen et al., 2012). These
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studies prompted a re-evaluation of the most appropriate experimental model for use in researching
arterial disease therapeutics for human patients. The minimal pre-existing collateral network present in pigs (1% of normal flow), which is similar to that found in humans, renders it a more suitable model of proof-of-concept studies (Hearse, 2000; Maxwell et al., 1987; Rubanyi, 2013).
Additionally, the heart weight-to-body weight ratio, functional hemodynamics and vascular response to myocardial ischemia of the pig is similar to that of humans (Elmadhun et al., 2013;
Rubanyi, 2013; Teunissen et al., 2012).
The ameroid model of myocardial ischemia developed in both dogs and pigs is widely used and is considered the gold standard for pre-clinical studies of pro-angiogenic therapeutics (G. C.
Hughes et al., 2003; Radke et al., 2006; Rubanyi, 2013). In this model, an ameroid constrictor is placed around the left circumflex coronary artery (LCx) which supplies 20% of the left ventricle in pigs and 25% in dogs (G. C. Hughes et al., 2003; Reimer, Ideker, & Jennings, 1981). The ameroid constrictor is a metal device with an inner ring of casein that is encased in a stainless steel sheath. Casein, a hygroscopic compound, gradually swells as it absorbs moisture in the body (G.
C. Hughes et al., 2003; Slatter, 2003). The steel ring is designed to force inward swelling of the casein and which gradually occludes the enclosed vessel over time (3-7 weeks) (Becker, Schuster,
Jugdutt, Hutchins, & Bulkley, 1983; F. J. Giordano et al., 1996; G. C. Hughes et al., 2003; Laham,
Rezaee, et al., 2000; Radke et al., 2006; Unger et al., 1994). The ameroid placement recapitulates the gradual occlusion of coronary arteries that occurs in human patients of CAD.
Animal models of peripheral ischemia
The capacity of the pre-formed collateral network was also determined in the periphery
(limbs) of animal models. As was observed in the heart, the extent of the pre-existing collateral
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circulation of the hindlimb is heterogeneous between animal species (Teunissen et al., 2012;
Waters, Terjung, Peters, & Annex, 2004). After acute femoral artery occlusion, mice, rabbits, and
dogs have low collateral flow (1-7%) when compared to pigs and rats (10-30%) (Hoefer, van
Royen, Buschmann, Piek, & Schaper, 2001; MacGabhann & Peirce, 2010; Rakue et al., 1998;
Teunissen et al., 2012). The slight baseline collateral formation in mice, rabbits and dogs is similar
to that observed in human PAD patients. A large proportion of preclinical and proof-of-concept
studies targeting therapeutic revascularization in PAD has focused on rodents (Haas et al., 2012;
Waters et al., 2004). Surgical ligation and/or excision of the common femoral or iliac arteries have
been reported to induce limb ischemia in these models. Due to the considerable level of pre-formed
collaterals in the rat, ligation of the femoral artery does not affect resting blood flow but does limit
muscle perfusion upon exercise activity (G. L. Tang, Chang, Sarkar, Wang, & Messina, 2005;
Teunissen et al., 2012). This model mimics the intermittent claudication in patients with PAD
whose symptoms are only observed during activity but not at rest (Mascarenhas et al., 2014; Muir,
2009; Muller et al., 2013). The rabbit and mouse are well-characterized models of resting vascular insufficiency that have been employed to test therapeutics for PAD (Couffinhal et al., 1998;
Dragneva et al., 2013; Haas et al., 2012; Madeddu et al., 2006; Pu et al., 1994).
LMW FGF2 gene or protein delivery in chronic ischemia
The potential of FGF2 as an agent of therapeutic revascularization has been examined and reported in several preclinical models of myocardial and limb ischemia. The results of these studies are summarized in Tables 5 and 6.
For the myocardial ischemia studies, canine or porcine models were treated with human recombinant LMW FGF2 protein exogenously administered by injection to the pericardium or
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myocardium or by infusion through the coronary arteries or veins (Laham, Rezaee, et al., 2000;
Lazarous et al., 1995; Rajanayagam et al., 2000; Unger et al., 1994; Watanabe et al., 1998). In all
but one of these cases, LMW FGF2 was delivered to the ischemic myocardium 7-14 days after the
ameroid placement surgery. The effects of the growth factor on infarct size, regional blood flow
and left ventricular (LV) function, were evaluated from 2 to 4 weeks after growth factor treatment.
The exception to this course of study was the work done by Yanagisawa-Miwa and co-workers where intracoronary infusion of LMW FGF2 was performed within hours of the occlusion of the coronary arteries in dogs which more closely resembles an acute ischemia model and not the human disease more accurately modeled by use of the ameroid occluders (Yanagisawa-Miwa et al., 1992). Furthermore, improvements in LV function and reduction in infarct size were observed after 7 days of ischemia. As previously established in “Mechanisms of vascular formation and growth” section, the vascular repair response in chronically ischemic tissues typically begins after
7-10 days lasting up to 21 days. The positive effects of LMW FGF2 observed only 1 week after treatment suggests an acute non-mitogenic effect of FGF2 on vasodilation and may not be a true revascularization response which requires 1-2 weeks to develop. A study by Tiefenbacher and colleagues showed that exogenous LMW FGF2 stimulates vasodilation of isolated coronary arterioles in a dose-dependent manner (Tiefenbacher & Chilian, 1997). Similar results of FGF2- induced vasoactivity was present in isolated cremaster muscle (H. M. Wu, Yuan, McCarthy, &
Granger, 1996) or intact pial arterioles (Rosenblatt, Irikura, Caday, Finklestein, & Moskowitz,
1994) and perfused whole hearts (Hampton et al., 2000). These responses were FGFR-mediated and were dependent on endothelium-derived NO production (Z. Huang, Chen, Huang, Finklestein,
& Moskowitz, 1997; Tiefenbacher & Chilian, 1997).
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In canine or rodent models, LMW FGF2 protein was exogenously administered via intra-
arterial, intravenous or intramuscular injections into the ischemic hindlimbs for a period ranging
from 1 week to 4 weeks after the hindlimb ischemia surgery. The effect of LMW FGF2 on indices
of angiogenesis and/or arteriogenesis was evaluated at 7-42 days, postoperatively. These include
the number of angiographically-visible vessels (collaterals), collateral ischemic/normal limb
perfusion (blood flow) ratio, and microvascular density determined by histology.
The studies described above and in Table 5 and 6 demonstrated a positive role for FGF2
(the LMW isoform) in promoting vascular growth/remodeling and improving post-ischemic blood
flow. In the decade between the late 1980s and late 1990s, many studies of similar nature were
undertaken for a variety of angiogenic factors, including other members of the FGF family,
vascular endothelial growth factor (VEGF), hepatocyte growth factor (HGF), and platelet-derived growth factor (PDGF), [reviewed here (Haas et al., 2012; G. C. Hughes et al., 2003; Katoh, 2013;
Madeddu et al., 2006; Waters et al., 2004)]. Several studies have performed comparisons between
FGF2 and other angiogenic factors using myocardial and peripheral ischemia models. The results of these studies are summarized in Table 7. Taken together, these studies suggested that FGF2 and
HGF stimulated the development of angiographically-visible or α-SMA-positive vessels to a similar degree. Furthermore, FGF2 was more efficacious at inducing collateral vessels compared to VEGF or several PDGF isoforms.
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Assessments of vascular FGF Model Species Delivery Findings Reference growth and tissue function
Thrombus injection in LVG, CAG, Histology, ↑LVEF, ↓Infarct size, (Yanagisawa- FGF2 Dog IC coronary artery Infarct size after 1 week ↑Capillaries and arterioles Miwa et al., 1992)
↑BF at 2 or 4 weeks, ↑Vessels Ameroid constriction Serial collateral BF, (Unger et al., FGF2 Dog IC ≥20µm, ↔Capillaries, ↑EC of the LCx Histology after 4 weeks 1994) proliferation, acute vasodilation
Ameroid constriction Serial collateral BF, ↑BF at 2 weeks, ↓MAP, (Lazarous et al., FGF2 Dog IC of the LCx Histology after 9 weeks ↔Infarct area, ↑Vessels >10µm 1995)
Ameroid constriction Single Bolus, IV, ↑BF in IC group, ↔BF in bolus, (Rajanayagam et FGF2 Dog BF, infarct size of the LCx Pericardial, or IC pericardial or IV al., 2000)
(Watanabe et al., FGF2 LAD embolization Pig Myocardial Injection Histology after 4-5 weeks ↔BP, ↔Capillaries, ↑Arterioles 1998)
↔MAP, ↑BF, ↑LV function, Ameroid constriction BF, CAG, MRI, Histology (Laham, Rezaee, FGF2 Pig Pericardial Injection ↑Collaterals, ↓Infarct zone, of the LCx after 4 weeks et al., 2000) ↑Capillaries
Table 5: Preclinical studies with the Fgf2 gene or LMW FGF2 in models of myocardial ischemia. BF, blood flow measured by microspheres;
CAG, Coronary angiography; IC, intracoronary; LAD, left anterior descending coronary artery; LCx, left circumflex coronary artery;
LVEF, left ventricular ejection fraction; LVG, left ventriculography; MAP, mean arterial pressure. ↓, Decrease; ↑, Increase; or ↔, No
change from control.
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Methods for assessing FGF Model Species Delivery Findings Reference vascular growth
Bilateral caudal and TcPO2, Angiography, ↑Angiographic collaterals, ↑Calf or (Baffour et al., FGF2 Rabbit IM injection iliac artery ligation Histology thigh TcPO2, ↑Capillaries 1992) ↑BPR, ↑Angiographic score, Unilateral femoral Doppler probe, FGF2 Rabbit IA injection ↔Collateral diameter, ↑Capillaries (Asahara 1995) artery excision Angiography, Histology at day 30 BF, Muscle function, ↑Calf BF, ↑Muscle function, Bilateral femoral artery FGF2 Rat IA infusion Vascular casting, ↑Collateral vessels, ↑Capillaries at (Yang et al., 1996) ligation Histology day 28 IM injection Unilateral femoral and slow Angiography, ↑Angiographic collaterals, (J. Zhang et al., FGF2 Rat artery ligation release Histology, LDPI ↑Arterioles, ↔ BF at day 42 2009) polymer Angiography, ↑Angiographic collaterals, Unilateral femoral Histology, (Rakue et al., FGF2 Dog IV injection ↑Collateral diameter, ↑Collateral artery ligation Hemodynamic flow 1998) flow volume at day 28 probe
Table 6: Preclinical studies on the effects of FGF2 in models of peripheral ischemia. BF, muscle blood flow measured by microspheres;
BPR, ischemic limb/normal limb systolic pressure ratio; IA, intra-arterial; IM, intramuscular; Laser Doppler Perfusion Imaging, LDPI;
TcPO2, transcutaneous oximetry. ↓, Decrease; ↑, Increase; or ↔, No change from control.
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Assessment of Model Species FGF2 PDGF-AA PDGF-AB PDGF-BB VEGF-A HGF Reference vascular growth (H. Lu et LAD ligation Pig α-SMA+ vessels +++ NT NT ++ NT NT al., 2007) (Hao et al., LAD ligation Rat α-SMA+ vessels ++ NT NT +++ NT NT 2004) Femoral artery (R. Cao et Rat α-SMA+ vessels ++++ NT NT +++ NT NT ligation al., 2003) Femoral artery Angiographic (J. Zhang et Rat +++ +++ +++ NT NT NT ligation collateral vessels al., 2009)
Femoral artery (J. Zhang et Rat α-SMA+ vessels +++ + ++ NT NT NT ligation al., 2009) Femoral artery Angiographic (R. Cao et Rabbit +++ NT NT ++ NT NT ligation collateral vessels al., 2003) Femoral artery (J. Li et al., Rabbit α-SMA+ vessels +++ NT NT +++ NT NT ligation 2010) Femoral artery (Masaki et Mouse α-SMA+ vessels ++++ NT NT NT ++ NT ligation al., 2002) Femoral artery (Marui et Mouse α-SMA+ vessels +++ NT NT NT NT +++ ligation al., 2005)
Table 7: Efficacy of LMW FGF2 gene or recombinant protein relative to proangiogenic growth factors in models of myocardial and peripheral ischemia. LAD, left anterior descending coronary artery; α-SMA, alpha smooth muscle actin; NT, not tested
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F. FGF2 therapy in PAD and CAD (Clinical studies)
The plethora of preclinical data revealing the beneficial effects of treating animal models
of ischemia with FGF2 and other growth factors propelled the field to attempt to translate these
results to CAD and PAD patients. To date, six human trials for recombinant FGF2 (LMW isoform)
have been conducted (Aviles, Annex, & Lederman, 2003). Four of these trials were Phase I/II
designed to evaluate the safety profile of FGF2 after intra-arterial, intracoronary, or intramuscular
administration (Laham et al., 1999; Laham, Chronos, et al., 2000; Lazarous et al., 2000; Unger et
al., 2000). None of these trials had more than 60 patients, and no signs of non-target organ
angiogenesis, malignancy, or altered vascular permeability were observed. However, in two of the
Phase I/II trials, low rates of proteinuria and resolving incidents of hypotension were observed
(Laham, Chronos, et al., 2000; Unger et al., 2000).
The two larger randomized, double-blinded, placebo-controlled studies were each aimed
at patients of PAD or CAD (Table 8). The Therapeutic Angiogenesis with FGF-2 for Intermittent
Claudication Trial (TRAFFIC) tested the efficacy of bilateral intra-arterial infusion placebo, single
bolus or double bolus of 30 µg/kg of FGF2 in 190 patients with moderate-to-severe IC (PAD)
(Lederman et al., 2002). The primary endpoint was change in peak walking time (PWT) at 90 days.
In a non-parametric intention-to-treat analysis (not the pre-specified ANOVA) of the results, single
bolus treatment with FGF2 increased PWT and no change was observed with double bolus FGF2
(Aviles et al., 2003; Lederman et al., 2002). Secondary outcomes of the study, claudication onset
time or quality of life measures, did not show improvement with FGF2 treatment. The FGF
Initiating RevaScularization Trial (FIRST) included 337 patients with CAD that were not eligible
for surgical revascularization (Simons et al., 2002). Patients received placebo or one of three doses
(0.3, 3 or 3 µg/kg) of FGF2 by intracoronary injection. The primary outcome for the study was
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change (improvement) in exercise tolerance test (ETT) time from baseline at 90 days after
treatment. While ETT was the standard clinical endpoint for CAD, it is not ideal as it does not
measure myocardial function but skeletal, pulmonary and vascular function and serves as a clinical
surrogate for vascular growth and reduction in myocardial infarction (Annex & Simons, 2005;
Aviles et al., 2003; Silvestre et al., 2013). ETT times for any FGF2 dose group were not
significantly different from placebo. Secondary endpoints, which include change in ETT time at
180 days and change in stress perfusion defect (measured by single-photon emission computed tomography, SPECT), were also not changed relative to placebo at any dose of FGF2.
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Primary FGF Drug Clinical Disease n Delivery Endpoint, Outcomes Reference (trial name) Formulation Phase followup
FGF2 Hr LMW IC Intra-arterial PWT and ABI, 90 ↑in PWT w/ single bolus, ↔ (Lederman et II 190 (TRAFFIC) FGF2 (PAD) infusion days w/ double bolus al., 2002)
FGF1 NV1 DNA II CLI Intramuscular Ulcer healing, 25 (Nikol et al., 125 No diff from placebo (TALISMAN) plasmid DB, R (PAD) injection weeks 2008)
FGF1 CLI Intramuscular Major amputation (Belch et al., NV III 525 No diff from placebo (TAMARIS) (PAD) injection or death, 1 year 2011)
Hr LMW II (Simons et al., FGF2 (FIRST) CAD 337 Intracoronary ETT, 90 days No diff from placebo FGF2 DB, R 2002)
FGF4 Angina Treadmill exercise (Henry et al., Ad5FGF4 III/III 532 Intracoronary No diff from placebo (AGENT3/4) (CAD) time, 12 weeks 2007)
Table 8: Summary of clinical trial results of FGFs in CAD and PAD patients. ABI, ankle brachial index; Ad5, adenovirus type
5; CLI, critical limb ischemia; DB, R double blind, randomized; ETT, exercise treadmill test time; IC, intermittent
claudication; Hr, Human recombinant; NV, Non-viral. ↑, Increase; ↔, No change from placebo
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6. Dissertation focus and hypotheses
FGF2 is a potent stimulator of wound healing, tissue repair and vascular growth processes
(Grothe, Haastert, & Jungnickel, 2006; Masahiro Murakami & Sakurai, 2012; Ortega, Ittmann,
Tsang, Ehrlich, & Basilico, 1998; Presta et al., 2005). These properties put forward FGF2 as
prototypic growth factor for use in the study of therapeutic revascularization (Katoh, 2013;
Murakami & Simons, 2008). Use of exogenous LMW FGF2 in in vitro and in vivo vascular growth studies has revealed a function for the growth factor in regulating extracellular matrix remodeling, blood vessel destabilization, endothelial and vascular smooth muscle cell migration, proliferation, and tube formation; all of which are critical aspects of the vascular growth processes including angiogenesis and arteriogenesis (Laham et al., 1999; Presta et al., 2005).
The central hypothesis of this dissertation is that the two classes of FGF2 isoforms have
distinct roles in chronic ischemia-induced vascular and tissue repair/remodeling. There is recent,
and still emerging, evidence to suggest that the two classes of FGF2 isoforms mediate cellular
pathways that both converge and diverge in their modulating effects on vascular biology, cardiac
injury, bone development/remodeling, and neuronal development/homeostasis (Coffin et al., 1995;
Garmy-Susini et al., 2004; Grothe et al., 2006; Jiang et al., 2007; Liao et al., 2009). Further
evidence to support the over-arching hypothesis that the isoforms have functionally distinct roles
include their different translational mechanisms, subcellular and subnuclear distribution, factors
controlling release/secretion/trafficking from cells, gene expression profiles and receptor
activation (Claus et al., 2003; Liao et al., 2009; Ma et al., 2007; Quarto et al., 2005). This
dissertation project contains three working hypothesis.
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Hypothesis 1: The HMW and LMW FGF2 isoforms promote differential levels of
revascularization and tissue recovery in response to chronic ischemia of skeletal muscle
The FGF2 isoforms have been shown to have contrasting functions in protecting the heart
from cardiac dysfunction but have similar roles in preserving cardiomyocyte viability during acute
ischemia/reperfusion injury (Liao et al., 2007, 2010). The opposing activities of the FGF2 isoforms have also been observed in estrogen-stimulated endothelial cell proliferation and migration
(Garmy-Susini et al., 2004). To establish the specific roles of endogenous LMW FGF2 and HMW
FGF2 isoforms in chronic ischemia-induced revascularization, wildtype (WT) mice, mice with a targeted deletion of the Fgf2 gene (Fgf2-/-), mice with specific ablation of the 21kDa and 22kDa
FGF2 isoforms (FGF2 LMW-only) and mice deficient in 18kDa expression (FGF2 HMW-only) will be subjected to a model of unilateral hindlimb ischemia. This is an in vivo model where vascular perfusion of the lower hindlimb is interrupted leading to chronic ischemia of the underlying skeletal tissue. Surrogate outcome measures for functional recovery (active movement scores and the extent of toe necrosis) of the ischemic limb were assessed. Revascularization
(angiogenesis and arteriogenesis; collateral vessel formation) will be evaluated by histological quantification of capillary and arteriole vessel density, respectively and micro-computed topography. Functional determinants of post-ischemic recovery of skeletal muscle perfusion and arterial conductance were determined via serial Doppler imaging measurements of lower limb
(foot) blood flow.
Hypothesis 2: The LMW and HMW isoforms differentially modulate FGF2 expression and
FGF receptor activation in addition to stimulating a unique set of angiogenic growth factor and cytokines in response to chronic hindlimb ischemia
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The expression of Fgf2 gene and protein isoforms has been shown to be tissue- and context- specific (Claus, Werner, et al., 2004; Coffin et al., 1995; S. Giordano et al., 1992; Liao et al., 2009).
In the peripheral nervous system (PNS), Fgf2 gene and/or protein expression as well as distribution are altered in response to nerve lesion injury. FGF2 protein expression is also upregulated under cardiac stress conditions including low-flow ischemia and adrenergic overstimulation (House et al., 2003; Padua & Kardami, 1993). To demonstrate the effects of deletion of a specific FGF2 isoform in the absence or presence of chronic ischemia on overall Fgf2 expression in skeletal muscle, the expression levels of Fgf2 transcript and protein, LMW FGF2 and HMW FGF2 were evaluated in non-ischemic, sham-operated or ischemic WT, Fgf2-/-, FGF2 LMW-only (HMWKO) and FGF2 HMW-only (LMWKO) hindlimbs.
In the heart and brain, the differential effects of the FGF2 isoforms have been studied under
acute and chronic injury, respectively. These studies have revealed that the activation of FGF2
signaling pathways are different in the presence of only the LMW or only the HMW isoforms
(Grothe et al., 2006; Kardami et al., 2007; Liao et al., 2009; Nindl et al., 2004). These processes
are paracrine, autocrine and receptor-dependent (LMW) or nuclear/intracrine and receptor-
independent (HMW). To identify the role of FGF receptor expression and activation in HMW and
LMW FGF2 responses to chronic ischemia, skeletal muscle isolated from non-ischemic, sham-
operated and ischemic hindlimbs of WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice
will be analyzed for Fgfr1b, Fgfr1c, Fgfr2b, Fgfr2c, Fgfr3b, Fgfr3c, or Fgfr4 mRNA and protein
expression levels.
In addition to its own effect on vascular growth, FGF2 can regulate the expression and
release of other angiogenic cytokines, which in turn, have their own specific roles in
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revascularization (Murakami & Simons, 2008). This positions FGF2 as a “master regulator” of ischemia-induced revascularization. In cultured NIH 3T3 fibroblasts, HMW FGF2-transfected cells display a phenotype distinct from LMW FGF2-expressing cells by inducing the expression of a unique transcriptome (Quarto et al., 2005). To characterize the repertoire of angiogenic cytokines whose protein expression is altered in response to FGF2 isoform-stimulated vascular growth and remodeling, WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only sham-operated or ischemic muscle were evaluated using an angiogenesis proteome array.
During ischemia, angiogenic factors activate the microvascular endothelium to express cell-adhesion molecules and chemokines. These chemoattractants then recruit and activate inflammatory cells to invade the site of injury (Silvestre et al., 2008). Equally, inflammatory cells release angiogenic cytokines that can also regulate the revascularization process (Heil & Schaper,
2004; la Sala et al., 2012). A cross-talk exists between inflammatory and angiogenic responses during FGF2-mediated revascularization (Andrés et al., 2009; Presta et al., 2005; Presta, Andrés,
Leali, Dell’Era, & Ronca, 2009). Inflammatory cells can also play a role that is counter to vascular reparation by releasing oxidative metabolites that are cytotoxic and cytolytic to ischemic muscle
(Tidball, 2011). To establish the role of inflammatory cell infiltrates in FGF2 isoform-induced repair and regeneration, markers of inflammation were measured during several timepoints of ischemia in WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only non-ischemic, sham or ischemic skeletal muscle.
Hypothesis 3: The effect of the HMW and LMW isoforms FGF2 on ischemia-induced revascularization and tissue recovery is dependent upon differential modulation of tissue regeneration pathways
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In the early periods following skeletal muscle injury (chronic ischemia), necrotic tissue
clearance is required for muscle healing and the re-establishment of muscle homeostasis. This stage of injury also coincides with activation and proliferation of quiescent resident progenitors called satellite cells that differentiate into mature myofibers (Le Grand & Rudnicki, 2007; Meisner,
Annex, & Price, 2014; Tedesco, Dellavalle, Diaz-Manera, Messina, & Cossu, 2010). The stages of the satellite cell activity (proliferation, migration and differentiation) are controlled by a family of muscle-specific transcription factors known as the myogenic regulatory factors (MRFs)
(Ceafalan, Popescu, & Hinescu, 2014). FGF2 and other growth factors have been shown to function as “wound hormones” when released from degenerating myocytes where they can stimulate the satellite cell activation which leads to muscle regeneration (Allen & Boxhorn, 1989;
Anderson, Liu, & Kardami, 1991; DiMario, Buffinger, Yamada, & Strohman, 1989). To examine
the contribution of the endogenous LMW and HMW isoforms of FGF2 to satellite cell activation
and differentiation in ischemic muscle injury and recovery, expression of several myogenic
regulatory factors (MRFs) , markers of satellite cell activity was determined. Transcript levels of
Pax3, Pax7, MyoD, myogenin, Myf5 and MRF4 was quantified in non-ischemic, sham-operated or
ischemic WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only hindlimbs.
This dissertation research project was designed to provide novel evidence of the specific
roles of the endogenous FGF2 isoforms in ischemia-induced revascularization, in particular, HMW
FGF2 in ischemic-induced revascularization. Additionally, the molecular signaling pathways that
mediate the effect of the FGF2 isoforms on vascular and tissue regeneration was explored. These
findings contribute to the understanding of the FGF2 isoforms in vascular biology and
pathophysiology. Furthermore, they may one day lead to the development of novel therapies for
the treatment of coronary and peripheral arterial diseases.
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MATERIALS AND METHODS
Animals and Exclusion Criteria
Mice were housed in a pathogen-free facility and handled in accordance with standard use
protocols, animal welfare regulations, and the NIH Guide for the Care and Use of Laboratory
Animals. All protocols were approved by the University of Cincinnati Institutional Animal Care
and Use Committee. All mice were bred on a (50%) Black Swiss and (50%) 129/Sv mixed
background. Any animals with severe necrosis and/or auto-amputation of tissue in the surgical
limb were excluded from the study and prematurely euthanatized.
Generation of Fgf2-/- mice
Mice with a targeted ablation of the Fgf2 gene (Fgf2-/-) were generated using a tag and
exchange construct by Ming Zhou in the laboratory of Dr. Thomas Doetschman as previously
described (Zhou et al., 1998). Briefly, a 0.5-kb NarI/XbaI portion of the Fgf2 gene containing the
proximal promoter region and the entire first exon was replaced with an Hprt (hypoxanthine
phosphoribosyl transferase) minigene in E14TG2a embryonic stem (ES) cells derived from 129
strain mice. A herpes virus thymidine kinase gene inserted outside of the homologous region at
the 3’ end of the targeting construct was used for negative selection. ES cells with the target vector
inserted were selected with HAT (hypoxanthine, aminopterin, thymidine) medium. Loss of the
thymidine kinase gene was selected with the use of gancyclovir. Double-resistant cells were confirmed for the targeted allele with polymerase chain reaction (PCR) and Southern blotting.
The targeted ES cells were then microinjected into C57BL/6 blastocysts and chimeric mice were generated from foster mothers. Germline transmission was tested by breeding the chimeric mice with 50% 129 and 50% Black Swiss mice. Wildtype (+/+), heterozygous (+/-) and knockout (-/-)
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offspring produced from this breeding and other subsequent pairings were identified with PCR
analysis and loss of FGF2 expression was confirmed with Southern, Northern and Western blotting
(Figure 3).
For genotyping with PCR, DNA was isolated from mouse pup tail clips that were placed
in a digestion buffer (100 mM NaCl, 10 mM Tris-Cl pH 8, 25 mM EDTA, 0.5% SDS) along with
100 µg proteinase K (Roche) at 60oC overnight. After centrifugation to remove cellular debris, the
DNA was precipitated in 100% isopropyl alcohol and washed with 70% ethanol. The pelleted
DNA was then air dried and dissolved in TE buffer overnight or at 55oC for 2 hours. PCR was
performed with two sets of primers using Taq DNA polymerase (Roche): wildtype allele, 5’- CGA
GAA GAG CGA CCC ACA C - 3’, 5’- CCA GTT CGG GGA CCC TAT T - 3’ and knockout
allele, 5’- AGG AGG CAA GTG GAA AAC GAA - 3’, 5’- CCC AGA AAG CGA AGG AAC
AAA - 3’. 1 µL of DNA was added to a 19 μL PCR master mixture (Roche 1.5 mM Mg PCR
buffer, 2.5 mM dNTP, 5X Cresol Red, H2O, and Taq DNA polymerase) and the following
amplification reaction was carried out. 1 cycle of (95°C for 90 seconds), 35 cycles of (95°C for 30
seconds for denaturing, 58 °C for 50 seconds for annealing and 72°C for 90 seconds for
elongation), followed by 1 cycle of 95°C for 30 seconds, 58°C for 50 seconds and 72°C for 10
minutes. The PCR products were run on a 1% agarose gel for 30 minutes at 140 mV. A band at
1299 base pairs (bp) signifies the presence of the targeted (knockout) allele while the wildtype
allele was indicated by a 185bp product.
Generation of FGF2 HMW-only (LMWKO) mice
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FGF2 HMW-only (LMWKO) mice were generated in the laboratory of Dr. Thomas
Doetschman as previously described (Azhar et al., 2009; Garmy-Susini et al., 2004). Briefly, the
Hprt minigene in the targeting vector described for the ablation of the Fgf2 gene was exchanged
by gene targeting with a mutated exon 1 where the wildtype CCATGC sequence is replaced with
CTGCAG. This replaces the ATG translational start site for FGF2 LMW protein with a CGA as
well introducing a PstI site that serves as a diagnostic for the exchanged allele. After selecting for
loss of the Hprt gene with 6-thioguanine (6-TG), ES cells carrying the exchanged allele were
microinjected into C57BL/6 blastocysts and chimeric mice were generated from foster mothers.
Germline transmission was tested by breeding male chimeric mice with 50% 129 and 50% Black
Swiss mice. Wildtype (+/+), heterozygous (+/-) and knockout (-/-) offspring produced from this
breeding and other subsequent pairings were identified with PCR analysis and absence of LMW
isoform expression was confirmed with Western blotting (Figure 3). The colony was subsequently
maintained on a mixed (50%) 129/Sv x (50%) Black Swiss background (Garmy-Susini et al.,
2004).
For genotyping with PCR, DNA was isolated from mouse pup tail clips that were placed
in a digestion buffer (Qiagen Puregene Cell Lysis Solution) along with 30µg proteinase K (Roche) at 60oC overnight. After treatmet with Qiagen Puregene Protein Precipitation Solution to remove
protein and cellular debris, the DNA was precipitated in 100% isopropyl alcohol and washed with
70% ethanol. The pelleted DNA was then air dried and dissolved in Qiagen Puregene DNA
Hydration solution overnight or at 55oC for 2 hours. PCR was performed with Epicentre Tfl
polymerase and the following primer set: 5’- CCC GCA CCC TAT CCT TAC ACA - 3’and 5’-
GCC GCT TGG GGT CCT TG - 3’ to amplify both the wildtype and mutated alleles. 1 µL of
DNA was added to a 19 μL PCR master mixture (Epicentre 20X buffer, Epicentre 25mM Mg2+,
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Epicentre 10X Enhancer, 2.5 mM dNTP, 5X Cresol Red, H2O, and Tfl DNA polymerase) and the following amplification reaction was carried out. 1 cycle of 97˚C for 5 minutes, 35 cycles of 97˚C for 60 seconds for denaturation, 58˚C for 60 seconds for annealing and 72˚C for 2 minutes for elongation and followed by 1 cycle of 72oC for 10 minutes. The PCR products were incubated for
1 hour at 37oC with PstI restriction enzyme, Buffer 3 and BSA (New England Bio Labs). The digested PCR products were run on a 2% agarose gel for 40 minutes at 140 mV. A band at 566 bp signifies the presence of the wildtype allele while two bands at 476 bp and 90 bp indicate the presence of the knockout allele.
Figure 3. FGF2 protein isoform expression in non-ischemic skeletal muscles. Representative immunoblots of LMW (18 kDa) or HMW (21 kDa and 22 kDa) FGF2 isoforms in non- ischemic wildtype (WT), Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only skeletal muscles.
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Generation of FGF2 LMW-only (HMWKO) mice
FGF2 LMW-only (HMWKO) mice were generated in the laboratory of Dr. Tom
Doetschman as previously described (Azhar et al. 2009). Briefly, the Hprt minigene in the
targeting vector described for the ablation of the Fgf2 gene was exchanged by insertion of a 14 bp
oligonucleotide (CTAGTCTAGACTAG) that contains a XbaI into a SmaI site located upstream
of the LMW FGF2 translational start site. This insertion introduced stop codons in all reading
frames and a frameshift mutation of all translation at the alternative start sites that lie upstream of
the LMW FGF2 AUG start codon. Loss of the Hprt gene was selected for in ES cells resistant to
6-TG and was confirmed with Southern blotting, DNA sequencing, PCR, and XbaI digestion. The
cells were then microinjected into C57BL/6 blastocysts and chimeric mice were generated from
foster mothers. Germline transmission was tested by male breeding the chimeric mice with female
Black Swiss mice. Wildtype (+/+), heterozygous (+/-) and knockout (-/-) offspring produced from
this breeding and other subsequent pairings were identified with PCR analysis. Insertion of the
14bp oligonucleotide and the XbaI was confirmed by Southern blotting and DNA sequencing
respectively. Loss of HMW FGF2 expression was confirmed by Western blotting (Figure 3).
For genotyping with PCR, DNA was isolated from mouse pup tail clips that were placed
in a digestion buffer (Qiagen Puregene Cell Lysis Solution) along with 30µg proteinase K (Roche) at 60oC overnight. After treatmet with Qiagen Puregene Protein Precipitation Solution to remove
protein and cellular debris, the DNA was precipitated in 100% isopropyl alcohol and washed with
70% ethanol. The pelleted DNA was then air dried and dissolved in Qiagen Puregene DNA
Hydration solution overnight or at 55oC for 2 hours. PCR was performed with Epicentre Tfl
polymerase and the following primer set: (Forward 5’- CCC AAG AGC TGC CAC AG - 3’ and
Reverse 5’-CGC CGT TCT TGC AGT AGA G -3’) to amplify both the wildtype and knockout
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alleles. 1 µL of DNA was added to a 19 μL PCR master mixture (Epicentre 20X buffer, Epicentre
2+ 25 mM Mg , Epicentre 10X Enhancer, 2.5mM dNTP, 5X Cresol Red, H2O, and Tfl DNA
polymerase) and the following amplification reaction was carried out. 1 cycle of 97˚C for 5
minutes, 35 cycles of 97˚C for 60 seconds for denaturation, 58˚C for 60 seconds for annealing and
72˚C for 2 minutes for elongation and followed by 1 cycle of 72oC for 10 minutes. The PCR products were run on a 4% agarose gel for 2.5 hours at 75mV. A band at 166 bp signifies the presence of the knockout allele while a 152 bp band indicates the presence of the wildtype allele.
Hindlimb ischemia surgery
Age- (10 -12 weeks) and sex-matched WT, Fgf2-/-, FGF2 LMW-only, and FGF2 HMW- only mice were anesthetized with 2.5% Avertin (2, 2, 2-tribromoethanol and tertiary amyl alcohol in 0.9% saline) at a dose of 0.02 mL/g, i.p and placed on a heating pad at 37oC. Unilateral hindlimb
ischemia was induced surgically as previously described (Couffinhal et al. 1998; Sullivan et al.
2002). A longitudinal incision was made in the skin overlying the middle portion of the left
hindlimb to expose tissue . The femoral artery and vein were then exposed and dissected free of
the associated nerve (Figure 4). Preservation of the femoral nerve was important to prevent atrophy of the muscle fibers. The vessel pair was ligated proximal to the external iliac artery and distally
at the bifurcation into the saphenous and popliteal branches. The intervening vessel segment was
then excised. The overlying skin was then closed with 5-0 silk sutures and tissue adhesive. A sham
procedure was performed on the contra-lateral (right) hindlimb, wherein a longitudinal skin incision and sutures were placed without ligation.
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A
B.
Femoral artery, Nerve vein & nerve
R L
Figure 4: The mouse hindlimb ischemia model. A) Schematic for the surgical induction of
hind limb ischemia. Modified from (Sullivan et al., 2002) B) Photograph of mouse lower
limbs after surgical excision of femoral artery-vein pair with intact nerve in ischemic right
hind limb (R) and intact femoral artery-vein-nerve in the sham limb (L).
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Assessment of necrosis
Necrosis was defined as the apparent darkening of one or more toes on the ischemic foot.
Darkening of ischemic feet began as a redness and in some cases, discoloration progressed to black
and ischemic toe(s) were lost. These mice were characterized as having necrotic limbs.
Assessment of limb function
The incidence of hindlimb tissue necrosis and limb function was assessed daily up to 14
days of ischemia and then weekly up to 42 days of ischemia. Functional use of the ischemic limb
was measured on the following scale modified from (Stabile et al., 2003) (4 = dragging of foot, 3
= no dragging of foot but no plantar flexion, 2 = plantar flexion but no flexing of toes and 1=
normal function, with flexing of toes to resist gentle traction on the tail).
Treadmill exercise testing
Mice were exercised on a modified OmniPacer LC4 motor-driven treadmill apparatus
(Omnitech, Columbus, OH) (Fewell et al., 1997; Lerman et al., 2002). The four-lane mouse treadmill was equipped with belts of adjustable speeds (0-100 m/min) and shock bars of adjustable amperage (0-2 mA) at the rear of each belt to stimulate the mice to run. Two pairs of infrared (IR) emitters/detectors were placed across from one another at the head of the shock bars. When the double IR beam was blocked by a mouse, a trip signal was generated and recorded. The trip signals
(beam breaks) were recorded every second for the length of each mouse’s exercise run. For each run, the number of breaks in the detection beam is a measure of the exercise performance of the animal. Therefore, the higher the number of beam breaks per minute, the less able the mice are to keep pace with the belt speed. Before each exercise test, the treadmill was placed at a 7o incline
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and the mice were acclimated over a 2-day period. On the first day of acclimation, mice were left undisturbed in the treadmill for 15 minutes with the instrument powered on, shock bars turned off and the belt speed set at 0 m/min. The belt was then turned on and speed was set at 2 m/min for 2 minutes, 5 m/min for 5 minutes and 7 m/min for 10 minutes. On the second day of acclimation, the shock grid was turned on, set at 1 mA, and the protocol from the previous day was repeated.
After the acclimation, exercise treadmill testing was performed at baseline (1-3 days before hindlimb ischemia surgery) and 14 days after surgery. Each test began with mice placed in the treadmill for 15 minutes and left undisturbed with the shock grid turned on to reduce any effect of stress on the exercise performance. The belt speed was then set at 7, 10 and 15 m/min for 5 minutes and at a maximum speed of 20 m/min for either 15 minutes or the length of time before which the animal fails to keep up the running pace. Failure to sustain the speed was indicated by the mouse lagging at the rear of the belt on the metal bars despite the stimulus of the shock current. If the animal received >10 shocks in a row, the test was halted and the mouse allowed to rest. The beam breaks were calculated over each 5 minute period of treadmill speed and over the final 15 minute run for animals able to complete the test. For animals that failed to reach 15 minutes, the beam breaks were calculated over their respective run times.
Assessment of gene expression in skeletal muscles
RNA isolation
Skeletal muscles were harvested from non-ischemic, sham or ischemic hindlimbs of WT,
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice. Sham or ischemic thigh and calf muscles
were dissected free of the femur and tibia/fibula, respectively and snap-frozen in liquid nitrogen
at 1, 3, 7 and 14 days of ischemia. For quantitative real-time polymerase chain reaction (qRT-
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PCR), 50-80 mg of skeletal muscle was placed in 1 mL of Tri-Reagent (Molecular Research
Center) for RNA isolation. The tissue was immediately homogenized with a benchtop motorized homogenizer, after which 1 mL of the homogenate was transferred to 1.5 mL microcentrifuge tube and centrifuged at 12,000x g at 4oC for 10 minutes. A phase separation to extract the RNA was performed with 100 µL of 1-bromo-3-chloropropane (BCP, Molecular Research Center) followed with centrifugation at 12,000x g at 4oC for 15 minutes. The aqueous phase was transferred into a new 1.5 mL microcentrifuge tube where the RNA was precipitated with 500 µL of isopropanol and centrifuged again at 12,000x g at 4oC for 10 minutes. The RNA pellet was then washed twice with 1 mL of 75% ice-cold ethanol followed by centrifugation at 12,000x g at 4oC for 4 minutes.
After air drying on ice for 5 minutes, the pellets were resuspended the RNA pellet in 10µL of nuclease-free water. To assess the integrity of the RNA, 1 µL of each RNA sample was added to
1 µL of 6X bromophenol blue and 4 µL of H2O and the solution was run on a 1% agarose gel at
90V for 40 minutes. The presence of two ribosomal bands at ~5 kilobases (28S) and at 2 kilobases
(18S) indicates intact RNA while a smear or the absence of RNA suggests that the RNA has been degraded. To determine the purity and the concentration of RNA, a 1:10 dilution of the sample in nuclease-free water was measured using a Nanodrop ND1000 spectrophotometer. The total concentration of RNA in each sample was calculated with the absorbance at 260 nm, and the purity was determined from the 260 nm/280 nm absorbance ratio. Samples with a 260/280 ratio of at least 1.8 or above were saved and stored at -80oC and utilized for reverse transcription.
Real-Time Quantitative Reverse Transcription PCR
Prior to reverse transcription of RNA to cDNA, 500 ng each of the isolated RNA was purified using RQ1 RNase-Free DNase (Promega). For reverse transcription with the SuperScript
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II Reverse Transcriptase (Life Technologies), 250 ng RNA was incubated with 10 mM dNTPs,
o random hexamers and H2O and at 65 C for 5 minutes and placed immediately on ice. The RNA
mix was then added to 10X RT buffer, 25 mM MgCl2, 0.1 M DTT and RNAseOUT and incubated
at room temperature for 2 minutes. 1 µL of SuperScript II reverse transcriptase was added to the
mixture which was then incubated at room temperature for 10 minutes 42oC for 50 minutes
followed by 15 minutes at 70oC, 1 minute at 4oC where the reaction was paused for addition of 1
µL of RNAse H to each sample. This step degrades any remaining template RNA. To confirm the
presence of cDNA, a control RNA sample that was converted to cDNA is amplified with
conventional PCR. A PCR reaction for each of four dilutions of the cDNA (1:1000, 1:10,000,
1:100,000 or 1:1,000,000) was evaluated using Taq Polymerase (Roche) and the following reaction
mixture: 10X + Mg buffer, 2.5 mM dNTP, Primer A (5’- GAC ATG GAA GCC ATC ACA GAC
- 3’), Primer B (5’- AGA CCG TTC AGC TGG ATA TTA C - 3’) 5X Cresol Red, and H2O.
For FGF2, qRT-PCR analysis of mRNA was performed in triplicates using the comparative
cycle time (ΔΔCT) method and Step One Plus ABI Real Time PCR system. 40 cycles of
amplification was performed in a 25 μL volume containing SYBR GREEN PCR master mix (Life
Technologies) and a 10μM primer mix, containing primers for the Fgf2 gene or for the
housekeeping gene, 18S. The following PCR parameters were used to amplify the cDNA (95o for
10 minutes, 95o for 15 seconds and 60o for 60 seconds) after determining that 18S (reference gene)
expression was not altered in the Fgf2-/-, FGF2 LMW-only or FGF2 HMW-only skeletal muscle relative to WT.
For FGF receptor (FGFR) expression, qRT-PCR analysis of mRNA was performed in triplicates using the ΔCT method and Step One Plus ABI Real Time PCR system. 40 cycles of
amplification was performed in a 25 μL volume containing SYBR GREEN PCR master mix (Life
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Technologies) and a 10μM primer mix, containing primers for one of the following Fgfr1b, Fgfr1c,
Fgfr2b, Fgfr2c, Fgfr3b, Fgfr3c, or Fgfr4 or for the 18S housekeeping gene (Table 9). The
following PCR parameters were used to amplify the cDNA (95o for 10 minutes, 95o for 15 seconds
and 60o for 60 seconds) after determining that 18S (reference gene) expression was not altered in
the Fgf2-/-, FGF2 LMW-only or FGF2 HMW-only skeletal muscle relative to WT.
For expression of the myogenic regulatory factors (MRFs), qRT-PCR analysis of mRNA
was performed in triplicates using the ΔCT method and Step One Plus ABI Real Time PCR system.
40 cycles of amplification was performed in a 25 μL volume containing SYBR GREEN PCR master mix (Life Technologies) and a 10μM primer mix, containing primers for one of the following Pax3, Pax7, Myf5, MyoD, MyoG or Mrf4 or for the 18S housekeeping gene (Table 10).
The following PCR parameters were used to amplify the cDNA (95o for 10 minutes, 95o for 15 seconds and 60o for 60 seconds) after determining that 18S (reference gene) expression was not altered in the Fgf2-/-, FGF2 LMW-only or FGF2 HMW-only skeletal muscle relative to WT.
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Primer Sets Forward, 5’- AGT CCC TGC CCT TTG TAC ACA -3’ 18S Reverse, 5’- CCG AGG GCC TCA CTA AAC - 3’ Forward, 5’- CAA CCG GTA CCT TGC TAT GA -3’ Fgf2 Reverse, 5’- TCC GTG ACC GGT AAG TAT TG -3’ Forward 5’- CAA CTT GCC GTA TGT CCA GAT C -3’ Fgfr1b Reverse 5’- CTC CGC ATC CGA GCT ATT AA -3’ Forward 5’- GCC AGA CAA CTT GCC GTA TG -3’ Fgfr1c Reverse 5’- ATT TCC TTG TCG GTG GTA TTA ACT C-3’ Forward 5’- GGG CTG CCC TAC CTC AAG -3’ Fgfr2b Reverse 5’- CTT CTG CAT TGG AGC TAT TTA TCC -3’ Forward 5’- CCC GGC CCT TCA -3’ Fgfr2c Reverse 5’- GTT GGG AGA TTT GGT ATT TGG TT -3’ Forward 5’- GCA CGC CCT ACG TCA CTG TA -3’ Fgfr3b Reverse 5’- GCG TCT GCC TCC ACA TTC T -3’ Forward 5’- ACG GCA CGC CCT ACG T -3’ Fgfr3c Reverse 5’- CTC CTT GTC GGT GTT AGC -3’ Forward 5’- CGC CAG CCT GTC ACT ATA CAA A -3’ Fgfr4 Reverse 5’- CCA GAG GAC CTC GAC TCC AA -3’
Table 9. List of forward and reverse primers for analysis of Fgf2 and Fgf receptor expression in WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only non-ischemic, sham and ischemic skeletal muscle tissues.
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Primer Sets Forward 5’- CAA CTT GCC GTA TGT CCA GAT C -3’ Pax3 Reverse 5’- CTC CGC ATC CGA GCT ATT AA -3’ Forward 5’- GCC AGA CAA CTT GCC GTA TG -3’ Pax7 Reverse 5’- ATT TCC TTG TCG GTG GTA TTA ACT C-3’ Forward 5’- GGG CTG CCC TAC CTC AAG -3’ Myf5 Reverse 5’- CTT CTG CAT TGG AGC TAT TTA TCC -3’ Forward 5’- CCC GGC CCT CCT TCA -3’ MyoD Reverse 5’- GTT GGG AGA TTT GGT ATT TGG TT -3’ Forward 5’- GCA CGC CCT ACG TCA CTG TA -3’ MyoG (myogenin) Reverse 5’- GCG TCT GCC TCC ACA TTC T -3’ Forward 5’- ACG GCA CGC CCT ACG T -3’ Mrf4 Reverse 5’- CTC CTT GTC GGT GGT GTT AGC -3’
Table 10. List of forward and reverse primers for analysis of myogenic regulatory factors
(MRFs) expression in WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only non-ischemic, sham and ischemic skeletal muscle tissues.
Gene expression (data analysis)
To quantify gene expression, the comparative CT method was used to measure the fold
changes in expression of Fgf2, Fgfr1-4, Pax3, Pax7, Myf5, MyoD, MyoG or Mrf4 in Fgf2-/-, FGF2
LMW-only or FGF2 HMW-only muscles relative to WT tissues (Bookout et al., 2006). Relative
expression of each gene of interest was determined relative to expression of 18S rRNA, an internal
control gene. Briefly, relative expression (ΔCT) = CTgene of interest – CT18S) and fold changes due to
(-ΔΔCT) selective protein expression = 2 (where ΔΔCT = CTFgf2-/-, FGF2 HMW-only, FGF2 LMW-only –
CTWT)(Livak & Schmittgen, 2001; Schmittgen & Livak, 2008).
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FGF2 protein isoform expression (Western immunoblot)
Skeletal muscles were harvested from non-ischemic, sham and ischemic hindlimbs of WT,
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice. Sham and ischemic thigh and calf muscles were dissected free of the femur and tibia/fibula, respectively and snap-frozen in liquid nitrogen at 1, 3, 7 and 14 days of ischemia. 100-120 mg of skeletal tissues were pulverized and powdered and then placed in an ice-cold glass dounce tissue grinder in an extraction buffer containing 20 mM Tris, 2 mM EDTA, 2 M NaCl, 1% NP40, EDTA-free protease inhibitor cocktail (Roche, 1
tablet/10 mL), and 1 mM PMSF. The homogenate was centrifuged at 15,000g for 15 minutes and
the supernatant was collected. Protein concentration was determined using a Bio-Rad DC protein
assay, and 2 or 5 mg (FGF2 LMW-only) of total protein was added to a 75% heparin sepharose
bead slurry (GE Healthcare). The samples and beads were incubated at 4°C for one hour, pelleted
and washed three times with a wash buffer (0.6 M NaCl and 10 mM Tris-HCl pH 7.4). 30 mL of
10X protein sample buffer was added to the beads and boiled for 10 minutes. The entire protein
sample was then loaded on a 15% SDS-PAGE gel and transferred to nitrocellulose membrane.
Transfer efficiency and loading equality were examined by staining the membrane with 0.1%
Ponceau S in 5% acetic acid. The membranes were blocked in 5% dry milk in 0.1% PBS/Tween
solution for 1 hour to prevent non-specific binding and then incubated with rabbit polyclonal primary antibody to FGF2 (1:1000 dilution, Santa Cruz Biotechnology) and incubated overnight at 4°C. After primary antibody incubation, membranes were washed and then incubated at room temperature in anti-rabbit IgG secondary antibody (1:7000 dilution, Santa Cruz Biotechnology)
two hours. The blots were then visualized using a chemiluminescence kit (Amersham ECL, GE)
and densitometry of protein bands were quantified using a Fluorchem 8800 gel imager.
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FGF2 protein isoform expression (quantitative data analysis)
To evaluate FGF2 isoform protein expression, the intensity of each protein band was
determined as described above. For comparison of expression across all genotypes, FGF2 protein
levels in Fgf2-/-, FGF2 LMW-only, and FGF2 HMW-only were normalized to WT protein
measures. This allows for statistical assessment of the FGF2 isoforms in tissues that required
different amounts of starting protein (2 mg for Fgf2-/- and FGF2 LMW-only; 5 mg for FGF2
HMW-only).
FGFR expression and phosphorylation (Western immunoblotting)
Skeletal muscles were harvested from non-ischemic, sham or ischemic hindlimbs of WT,
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice. Sham or ischemic thigh and calf muscles
were dissected free of the femur and tibia/fibula, respectively and snap-frozen in liquid nitrogen
at 3 and 7 days of ischemia. 2 mg of skeletal tissues were pulverized and powdered and then placed
in an ice-cold glass dounce tissue grinder in an extraction buffer containing 25 mM Hepes, 150
mM NaCl, 1% Triton X-100, 5 mM EDTA, 1% glycerol, 1 mM sodium orthovanadate, 25 mM β–
glycerolphosphate, 50 mM sodium fluoride, 0.5 mM okadaic acid, 100 mM calpain inhibitor,
Pefabloc Stock 1 and 2 (Roche), Roche phosphatase inhibitor (1 tablet/10 mL), Roche complete
mini EDTA-free protease inhibitor cocktail (1 tablet/10 mL), and 1 mM PMSF. Protein
concentration was determined using a Bio-Rad DC protein assay. 200 µg of protein was then loaded on a 8% SDS-PAGE gel and transferred to nitrocellulose membrane. Transfer efficiency and loading equality were examined by staining the membrane with 0.1% Ponceau S in 5% acetic acid. The membranes were blocked in 5% BSA in 0.1% TBS/Tween solution for 1 hour to prevent non-specific binding and then incubated overnight at 4ºC with primary antibodies against phospho-
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Y654 FGFR1 (1:1000 dilution, Abcam), phospho-Y742 FGFR3 (1:50000 dilution, Abcam) or phospho-Y642 FGFR4 (1:500 dilution, Abcam) followed by 1 hour incubation at room temperature in anti-rabbit IgG secondary antibody (1:5000, Santa Cruz). The phosphorylation of
FGFR1, FGFR3, and FGFR4 expression was visualized by ECL or ECL Plus (phospho-FGFR3) and densitometry of the protein bands were quantitated using a Fluorchem 8800 gel imager. The membranes were then stripped of the phospho-antibodies with stripping buffer (62.5 mM TRIS, pH 6.8, 2% SDS, and 100 mM β-mercaptoethanol) for 45 minutes at 56ºC. After washing, the membranes were blocked again in 5% BSA in 0.1% TBS/Tween solution for 1 hour before incubation with primary antibodies against total FGFR1 (1:400, Abcam), FGFR3 (1:1000, Abcam) or FGFR4 (1:500, Santa Cruz) overnight at 4ºC. After primary antibody incubation, membranes then incubated for 1 hour at room temperature with anti-rabbit IgG secondary antibody (1:5000,
Santa Cruz). Total receptor expression of FGFR1, FGFR3, and FGFR4 was visualized by ECL or
ECL Plus (FGFR3) and densitometry of the protein bands were quantitated using a Fluorchem
8800 gel imager.
FGFR phosphorylation (quantitative data analysis)
To evaluate FGFR activation (i.e. phosphorylation), the intensity of each protein band was determined as described above.
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Expression profile of angiogenesis-related proteins
Skeletal muscles were harvested from non-ischemic, sham and ischemic hindlimbs of WT,
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice. Sham and ischemic thigh and calf muscles were dissected free of the femur and tibia/fibula, respectively and snap-frozen in liquid nitrogen at 7 days of ischemia. 100-120 mg of skeletal tissues was pulverized and homogenized on ice in a glass dounce in an extraction buffer containing 20 mM Tris, 2 mM EDTA, 2 M NaCl, 1% NP40,
EDTA-free protease inhibitor cocktail (Roche, 1 tablet/10 mL), and 1 mM PMSF. The homogenate was centrifuged at 15,000g for 15 minutes and the supernatant was collected. After protein concentration was determined using a Bio-Rad DC protein assay, 600 µg of total protein in 1.5 mL of array buffer from the proteome profiler mouse angiogenesis antibody array kit (R&D Systems) was mixed with 15 µL of a detection antibody cocktail and incubated at 25oC for one hour. Each
sample/antibody mixture was then added to a nitrocellulose membrane containing 53 different
capture antibodies (Table 11) dotted in duplicate and incubated at 4oC overnight. After washing,
2 mL of buffered Streptavidin-HRP was added to each membrane and allowed to incubate at 25oC
for 30 minutes. Array membranes were developed using chemiluminescence with the provided
Chem Reagent mix. The densitometry of the antibody/protein dots were quantified using a
Fluorchem 8800 gel imager.
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ADAMTS1 EGF IGFBP-1 MIP-1α/CCL3 Osteopontin Amphiregulin CD105/Endoglin IGFBP-2 MMP-3 PD-ECGF Angiogenin Endostatin IGFBP-3 MMP-8 (pro) PDGF-AA Angiopoietin-1 Endothelin-a IL-1α MMP-9 PDGF-BB Angiopoietin-3 FGF1 TIMP-1 IGFBP-9/NOV Pentraxin-3 Coagulation factor III/Tissue Factor FGF2 IL-10 KGF/FGF7 SDF/CXCL12 CXCL16 Fractalkine/CX3CL1 CXCL10/IP-10 CXCL4/PF4 Serpin E1/PAI-1 IGFBP-10/Cyr61 GM-CSF CXCL1, KC PIGF-2 Serpin F1/PEDF DLL4 HB-EGF Leptin/OB Prolactin Thrombospondin-2 CD26/DPPIV HGF MCP-1/CCL2 Proliferin TIMP-4 VEGF-A VEGF-B
Table 11. List of angiogenesis-related proteins analyzed for relative expression in WT,
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only ischemic skeletal muscle tissues.
Expression profile of angiogenesis-related proteins (data analysis)
To determine the relative expression of each protein (Table 11), the intensity of each array dot was quantified as described above. Expression was determined as the average intensity of the pair of duplicate spots representing each angiogenesis related protein subtracted from an averaged background signal. For each protein, expression levels in ischemic WT, Fgf2-/-, FGF2 LMW-only, or FGF2 HMW-only arrays were normalized to the corresponding sham muscle expression.
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Immunohistochemistry
Animals were sacrificed before (non-ischemic) or 1, 3, 7, 14, or 42 day(s) of ischemia.
Skeletal muscles harvested from the thigh (gracilis, semi-membranosus, and semi-tendinosus) and
calf (gastrocnemius) regions of the ischemic limb were immersion-fixed overnight in 4% formaldehyde followed by immersion in 70% ethanol for processing and embedding.
Griffonia simplicifolia isolectin B4 staining
Five μm thick transverse paraffin-embedded sections were deparaffinized and rehydrated with xylenes and alcohol/water (100%, 95% and 70%) solutions, respectively. A 10 minute antigen retrieval step was performed at 100oC using a pH 6.0 citric acid solution. Endogenous peroxidase
activity was blocked with H2O2 for 12 minutes followed by treatment with 5% normal goat serum
(Vector Labs) and 1% bovine serum albumin (Vector Labs) at room temperature for 1 hour to
reduce nonspecific binding. Serial sections were then incubated overnight at 4oC with biotinylated
Griffonia simplicifolia (Bandeiraea) isolectin B4 (IB4 1:100, Vector Labs) to detect endothelial
cells. IB4 from the African-based black bean Griffonia simplicifolia binds with high affinity to
terminal α-galactosyl residues of glycoproteins on the cell surface of brain, cardiac muscle and
skeletal muscle endothelial cells (Laitinen, 1987; J. E. Lee et al., 2008). The antigen-Lectin
complexes were labeled utilizing the Vectastain ABC reagent kit (Vector Labs) and
immunoperoxidase staining was detected with 3, 3’-diaminobenzidine (DAB, Vector Labs).
Sections were counterstained with Mayer’s Hematoxylin (Life Technologies) to detect nuclei,
dehydrated, and coverslipped with Permount mounting medium (Fisher Scientific). Vessels with
IB4-positive staining (small brown circular structures) and an outer diameter of 7-10 μm were
classified as capillaries. Images were captured under 400X magnification from each section (2
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sections/limb, 40 fields/section). Vessel densities were determined as the total number of vessels
per myofiber number in each field.
Alpha-smooth muscle actin staining
Five μm thick transverse paraffin-embedded sections were deparaffinized and rehydrated with xylenes and alcohol/water (100%, 95% and 70%) solutions, respectively. Endogenous peroxidase activity was blocked with H2O2 followed by treatment with 5% normal goat serum at
room temperature for 1 hour to reduce nonspecific binding. Serial sections were then incubated
overnight at 4oC with rabbit polyclonal antibody against α-smooth muscle actin (α-SMA 1:15000,
Abcam) to detect vascular smooth muscle cells. A biotinylated goat anti-rabbit secondary antibody
(1:500, Vector Labs) was then applied to the sections for 2 hours at room temperature. The antigen- antibody complexes were labeled utilizing the Vectastain ABC reagent kit (Vector Labs) and immunoperoxidase staining was detected with 3, 3’-diaminobenzidine (DAB, Vector Labs).
Sections were counterstained with Mayer’s Hematoxylin (Life Technologies) to detect nuclei, dehydrated, and coverslipped with Permount mounting medium (Fisher Scientific). α-SMA positive vessels with a 20 μm or more outer diameter were identified as arterioles and venules.
Venules were distinguished from arterioles by their collapsed luminal structure and thinner layers of smooth muscle. Images were captured under 200X magnification from each section (2 sections/limb, 20 fields/section). Vessel densities were determined as the total number of vessels per myofiber number in each field.
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Myeloperoxidase (MPO) staining
Five μm thick transverse paraffin-embedded sections were deparaffinized and rehydrated with xylenes and alcohol/water (100%, 95% and 70%) solutions, respectively. Endogenous peroxidase activity was blocked with H2O2 followed by treatment with 5% normal rabbit serum
to reduce nonspecific binding. Serial sections were then incubated overnight at 4oC with goat
polyclonal antibody against MPO (1:5000, R&D Systems) to detect neutrophils. A biotinylated
rabbit anti-goat secondary antibody (1:500, Vector Labs) was then applied to the sections for 2
hours at room temperature. The antigen-antibody complexes were labeled utilizing the Vectastain
ABC reagent kit (Vector Labs) and immunoperoxidase staining was detected with 3, 3’- diaminobenzidine (DAB, Vector Labs). Sections were counterstained with Mayer’s Hematoxylin
(Life Technologies) to detect nuclei, dehydrated, and coverslipped with Permount mounting medium (Fisher Scientific). Images were captured under 400X magnification from each section (2 sections/limb, 40 fields/section). The number of MPO positive cells were counted in each images and presented as the total number of cells per field.
Mac-3 staining
Five μm thick transverse paraffin-embedded sections were deparaffinized and rehydrated with xylenes and alcohol/water (100%, 95% and 70%) solutions, respectively. Endogenous peroxidase activity was blocked with H2O2 followed by treatment with 5% normal goat serum to reduce nonspecific binding. Serial sections were then incubated overnight at 4oC with rat-anti
mouse monoclonal antibody against Mac-3 (CD107b 1:1000, BD Biosciences) to detect macrophages. A biotinylated mouse absorbed rabbit anti-rat secondary antibody (1:500, Vector
Labs) was then applied to the sections for 2 hours at room temperature. The antigen-antibody
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complexes were labeled utilizing the Vectastain ABC reagent kit (Vector Labs) and
immunoperoxidase staining was detected with 3, 3’-diaminobenzidine (DAB, Vector Labs).
Sections were counterstained with Mayer’s Hematoxylin (Life Technologies) to detect nuclei,
dehydrated, and coverslipped with Permount mounting medium (Fisher Scientific). Images were
captured under 400X magnification from each section (2 sections/limb, 40 fields/section). The
number of Mac-3 positive cells were counted in each images and presented as the total number of
cells per field.
Micro-Computed Tomography Imaging of Vasculature
Micro-CT (Perfusion of contrast agent)
During preliminary perfusion experiments, three different contrast agents with high radio
density were tested for their ability to provide the X-ray attenuation necessary for micro-CT
imaging of vascular structures: barium sulfate, lead oxide and lead chromate. To prepare the
barium sulfate or lead oxide for perfusion, 105% w/v barium sulfate (E-Z-EM Canada Inc) or
100% w/v lead oxide (Sigma Aldrich) and 5% porcine gelatin (Sigma Aldrich) were added to
saline and heated with stirring to 60oC until the solution was completely dissolved. The solution was cooled to and maintained at 37oC in a water bath until a 1 mL syringe was filled and injected
at 1 mL/h. Samples were immediately stored at 4oC overnight to induce polymerization of the
gelatin suspensions within the vasculature until they were ready for CT scanning. For Microfil
perfusion, a curing agent (5% v/v) was added to catalyze the components (compound and diluent
mixed in equal amounts, according to the manufacturer recommendations; Flow-Tech Inc) to give
a working time of 20 minutes before the start of polymerization. 0.5 mL of the prepared compound
was drawn in a 1 mL syringe and injected at 1 mL/h with care taken to remove any perfusate that
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contaminated the surrounding tissue, skin or fur. Perfused specimens were allowed to sit at room
temperature for 30 minutes and then stored at 4oC overnight to allow for contrast agent
polymerization.
For all perfusion experiments, a hand-pulled catheter (made from polyethylene [PE] tubing,
1/4" ID, 3/8" OD) was attached to a three-way tubing connector via 23-gauge needle adapters.
Standard PE-50 tubing were attached to the Y branches of the tubing connector; one side to deliver
the perfusates via a syringe pump and the other to connect to a pressure transducer and blood
pressure analyzer. The pressure across the tubing was monitored and maintained between 95 and
100 mmHg with adjustments made to the pump speed if pressure exceeded this range. Non-
ischemic (in preliminary experiments) and ischemic mice were anesthetized and overdosed with
sodium pentobarbital at a dose of 80 mg/kg, i.p. A horizontal midline incision was made right
below the diaphragm to avoid pneumothorax and to ensure blood flow during catheter insertion.
The abdominal aorta was cannulated, the catheter secured with 5-0 suture and a 6 mL heparinized
(100 U/mL) saline solution containing a mixture of vasodilators (10 µM acetylcholine, 10 µM
adenosine, 100 µM papaverine and 200 µM sodium nitroprusside; Sigma-Aldrich) was infused at
1 mL/min with a 3 mL plastic syringe (Becton Dickson #309659) for blood washout. The inferior
vena cava was severed to allow for drainage of blood and the infusing solution.
Micro-CT image acquisition
Non-ischemic, sham and ischemic limbs were dissected at the head of the femur for individual scanning at high resolution using the micro-CT scanner in the Tri-modal Siemens
Inveon (Siemens Healthcare) and the Inveon Acquisition Workplace (IAW) (version 1.5.28) software. Each limb was placed in a field of view of 18 x 25 mm, positioned on the dorsal side,
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and scanned from the thigh to the ankle. CT scanning parameters were 80 kVp and 300 μA (voltage
and current) with 0.5 mm Al filtration. A 192o half scan was performed for 384 steps with an exposure time of 2100 ms for each step. Center offset and dark/light calibrations were performed immediately prior to scanning. The acquisition protocol included using high system magnification with 2x2 binning to generate an isotropic voxel size of 17 μm3. For the raw data reconstruction,
beam hardening was applied and was accurately Hounsfield Unit (HU) calibrated. Hounsfield units
are values assigned to voxels (3-D pixels) within a CT image that reflect the range of electron
densities and is proportional to the degree of X-ray attenuation (Kline & Ritman, 2012). Siemens
Inveon Research Workplace, (IRW version 4.0) and the 3-D Visualization tool were used to
generate 2-D cross-sections in yz-(sagittal), xy-(coronal), and xz-(axial) planes and 3-D volume
rendered images (Figures 5-7).
Comparison of contrast agents
Limbs perfused with either barium or lead contrast agents were not suitable for vascular
visualization or analysis. In the case of barium sulfate perfused-limbs, bone was very clearly
visualized and distinguishable from tissue but no vessels could be seen (Figure 5). Deposits of the
contrast agent were visible near the top of the limbs (i.e. adjacent to the site of perfusion)
suggesting that perfusion of the vessels was not successful. The lead oxide-perfused limbs produced images showing clear distinction of the bone and vessels (perfused with contrast agent) from muscle. The range of the HU values of the lead oxide-containing perfused vessels exceeded the upper limits of the viewing software and resulted in oversaturated voxels that could be not be analyzed (Figure 6). The Microfil (lead chromate with silicone rubber) contrast agent also produced images with clear distinctions of perfused vessels from bone and tissue (Figure 7). These
88 images had HU values within the range of the viewing software and were suitable for vascular visualization and quantification.
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A AXIAL
C D
SAGITTAL 3D MIP
B CORONAL
Figure 5: Representative 2-D cross-sectional axial (A), coronal (B) and sagittal (C) slice views of a non-ischemic WT limb perfused with a barium sulfate and gelatin suspension which did not provide vascular contrast due possibly to improper perfusion or vasodilation. (D) A high resolution 3-D maximum intensity projections (MIP) of the same limb where the contrast agent was visible at the top of the image but did not perfuse the vessels (red arrows).
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A AXIAL
D C 3D MIP SAGITTAL
B
CORONAL
Figure 6: Representative 2-D cross-sectional axial (A), coronal (B) and sagittal (C) slice views of a non-ischemic WT limb perfused with a lead oxide and gelatin suspension which perfused the vessels and provided contrast. (D) A high resolution 3-D maximum intensity projections (MIP) of the same limb. The high electron density of the contrast agent resulted in images that could not be analyzed due to oversaturation (red arrows) of the vascular structures.
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A AXIAL
C D SAGITTAL 3D MIP
B CORONAL
Figure 7: Representative 2-D cross-sectional axial (A), coronal (B) and sagittal (C) slice views of a non-ischemic WT limb perfused with a lead chromate silicone rubber suspension (Microfil) which perfused vessels and provided contrast. (D) A high resolution 3-D maximum intensity projections (MIP) of the same limb. This contrast agent produced images with adequate contrast of vessels from tissue and 3-D volumes suitable for visualization and quantitative analysis of vascular structures. 92
Micro-CT image analysis
For 3-D vessel morphology analysis, bones were digitally removed and the vasculature was
segmented from the surrounding tissue (background) based on the density of the X-ray attenuated-
voxels of the Microfil. Briefly, manual segmentation or automatic segmentation was performed to designate regions of the scanned volumes as bone, vessel or non-vascular tissue (Figure 8). The manual segmentation entailed manually drawing volumes of interest (VOI) that surrounded the bone regions in each of the serial 2-D sections (~500 slices in the coronal plane) that make up the
3-D volumes. This was followed by automated segmentation based on the density of the voxels in the Microfil-filled vessels (thresholding). A threshold range of 1000 -7000 HU was chosen and
used to evaluate all the scanned volumes. This range was selected based on comparing the ability
of different ranges of HU values to effectively separate tissue and vessels into distinct volumes
(binarization) in post-segmentation images.
Histomorphometric parameters including vessel density (number of vascular
structures/mm), volume (volume of segmented voxels normalized to limb volume), diameter (local
thickness of voxels), and spacing (average distance/separation between vessels quantified from
background voxels) were calculated for each defined volume of interest (sham or ischemic calf or
thigh) using the Siemens IRW Bone Morphometry Tool. The values were reported as a percent
ratio of ischemic to non-ischemic limbs for each animal to minimize any variations in the vascular
morphology parameters that could be attributed to the contrast agent perfusion.
To determine and compare the vessel thickness (diameter) distribution in the mouse limbs,
bone-subtracted 2-D slices were analyzed for local thickness in Fiji (NIH). For each 3-D data set,
an in-house Matlab (Mathworks; Natick, MA) subtraction program was applied to generate images where all bone had been removed. These stacked data were converted to DICOM files and
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evaluated for “local thickness” in Fiji. The local thickness tool defines a volume-based local
thickness at every voxel in the limb volume as the diameter of the largest sphere that both contains
the voxel and is entirely within the volume of structure (Zagorchev et al., 2010). Histograms were
generated to display the volume and range of vessel sizes diameter (34-400 μm) across the sham and ischemic limbs.
Figure 8: Representative 2-D cross-sectional coronal (A), sagittal (B), and axial (C) slice views of a non-ischemic limb WT after manual (to subtract bones) and automatic (HU-based
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thresholding of vessels) segmentation. The calf region is highlighted in red with bones
segments digitally deleted (white open arrows) and vessels segmented based on HU voxel
intensity (white arrowheads).
Laser Doppler Perfusion Imaging
Perfusion was measured using images acquired with an infra-red laser Doppler perfusion imager (Moor LDI2, Moor Instruments). Anesthetized animals were placed ventrally with legs outstretched on a heating pad at 37oC and positioned to allow for scanning of the both lower
hindlimbs (sham and ischemic). Scanning of the lower hindlimbs (paws) was performed before
surgery (baseline) and on days 1, 14 and 42 after induction of chronic ischemia. Three consecutive
color-coded images representing the microcirculation flow distribution were obtained per animal
for the calculation of perfusion values. In each image, high perfusion is signified by the red colors,
yellow and green depict the mid-range and the blues indicate low perfusion. Relative perfusion
was expressed as the ratio of the mean perfusion values of the ischemic to sham (non-ischemic)
feet.
Statistical Analysis
Data are expressed as mean ± S.E.M. Limb use/function data were compared with two-
way repeated measures analysis of variance (ANOVA). Necrosis data are presented as percentages
and were analyzed using Chi-Square test for categorical data. All other analyses were performed
using one-way or two-way ANOVAs. When ANOVA evaluations produced significance (p≤
0.05), Bonferroni post-hoc tests were carried out for multiple or pair wise comparisons.
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Chapter 1. HMW and LMW isoforms of FGF2 in ischemia-induced revascularization and
tissue recovery
RESULTS I
The roles of the HMW and LMW FGF2 protein isoforms in embryonic vascular
development of the heart have been previously studied (Amann et al., 2006; Liao et al., 2009).
Liao and colleagues analyzed non-ischemic hearts from adult FGF2 HMWKO (LMW expression
only) and FGF2 LMWKO (HMW expression only) mice for capillary and smooth muscle- containing vessels (arterioles). Those experiments produced no differences from WT in the number of cardiac capillaries and arterioles in either knockout group. The purpose of this dissertation was to evaluate the roles of the FGF2 isoforms in ischemia-induced vascular growth and remodeling.
To that end, WT (all FGF2 isoforms expressed), Fgf2-/- (no LMW or HMW FGF2 expression)
FGF2 HMWKO (no HMW FGF2 expression), and FGF2 LMWKO (no LMW FGF2 expression), mice were subjected to a surgical model of hindlimb ischemia (Couffinhal et al., 1998; Murohara et al., 1998; Sullivan et al., 2002). This is a well-characterized model where a section of the superficial femoral artery and vein pair is ligated and then excised (Figure 4). A majority of the mice survived the surgery and all deaths that occurred were related to anesthesia and not the procedure. The mice were allowed to recover and monitored daily for any health or procedure-
related issues. For enhanced clarity of the following figures and results, the FGF2 HMWKO and
FGF2 LMWKO mouse groups will, henceforth, be referred to as FGF2 LMW-only and FGF2
HMW-only, respectively.
Accelerated recovery of ischemic limb use in the presence of HMW FGF2
Evaluation of ischemic hindlimb use was performed at specific time points after induction
of ischemia to address the roles of the FGF2 isoforms in recovery of the ischemic limb. A semi-
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quantitative scoring scale modified from Stabile and colleagues (Helisch et al., 2006; Stabile et al.,
2003) was utilized to perform daily and weekly assessments of ischemic limb impairment or use
(Table 12). Mice from all groups experienced impaired limb use immediately after the ischemia surgery which presented as dragging of the ischemic foot and this observation persisted for the next three days of ischemia (Figure 9), most likely due to the ischemic pain and/or surgical procedure. Soon after, all groups of mice began to exhibit improved use of their ischemic limbs to varying degrees. By 28 days, WT had resumed normal limb function and their ischemic limbs could no longer be differentiated from sham limbs. Mice expressing only LMW FGF2 had a more prolonged recovery time and ischemic limb use did not return to normal until day 42. Conversely, the FGF2 HMW-only limbs had a faster recovery (2-3x) of the ischemic limb with normal use regained at 14 days compared to the other groups (p<0.05, Figure 9). Mice with no expression of
FGF2 (Fgf2-/-) did not completely recover ischemic limb use by the end of the study at 42 days.
Score Hindlimb Use
4 Dragging of ischemic foot
No dragging of ischemic foot but no plantar 3 flexion
2 Plantar flexion but no flexing of toes
Normal use of ischemic limb with flexing of 1 toes to resist gentle traction on the tail
Table 12: Descriptive scoring for the assessment of hindlimb use during chronic ischemia.
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WT 5 Fgf2 -/-
4 FGF2 LMW-only FGF2 HMW-only
3
2 *†‡ Limb Impairment Score Impairment Limb 1 *†‡ *†‡ †‡ †‡
0 0 3 7 10 14 21 28 35 42
Days post-surgery
Figure 9: Average limb impairment scores of WT (n=35), Fgf2-/- (n=26), FGF2 LMW-only
(n=31) and FGF2 HMW-only (n=25) ischemic limbs after induction of hindlimb ischemia.
A descriptive scoring system listed in Table 12 was used to assign scores daily for up to 14 days of ischemia and then weekly for up to 42 days of ischemia. Data are presented as mean
± SEM. *p<0.05 vs. WT, †p<0.05 vs. Fgf2-/-, ‡p<0.05 vs. FGF2 LMW-only.
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Loss of FGF2 expression negatively influences limb functional capacity
As a physiological alternative to the hindlimb use assessment, the effect of FGF2 isoform
expression on functional recovery of the ischemic was also determined by measuring the forced
exercise capacity of the mice in this study before and 14 days after the ischemia surgery was
performed. At 1-3 days prior to the surgery (baseline test), a cohort of WT, Fgf2-/-, FGF2 LMW-
only and FGF2 HMW-only mice were subjected to exercise treadmill testing at multiple belt
speeds and the same group was again tested 14 days later. The speed was gradually increased over
time and mice were allowed to run for 5 minutes at the 7, 10 and 15 m/min speeds. When a speed
of 20 m/min was reached, the mice were allowed to run for either 15 minutes or length of time
before which the animal was unable to keep the pace with the motorized treadmill belt; as indicated
by a continuous break in both of the infrared beams located at the end of each treadmill belt (Fewell
et al., 1997; Q. Yang et al., 2001). For the entire length of the test, the number of breaks in the dual
infrared beam by a part of the animal (usually tail and/or hindlimbs) was detected and recorded
(Fewell et al., 1997; Lerman et al., 2002).
In absence of ischemia (baseline), no differences in the levels of exercise performance was detected between WT and Fgf2-/- mice at lower treadmill speeds (7 or 10 m/min) (Figure 10A).
When the speed was increased to 15 m/min, the Fgf2-/- mice performed poorer than WT as
evidenced by the increased number of beam breaks/minute. When the treadmill speed was further
increased to 20 m/min, the performance of Fgf2-/- mice was unchanged from WT levels. After two weeks of ischemia, the performance of Fgf2-/- mice was unchanged from WTs at 7, 10 or 15 m/min. However, exercise activity appears to be decreased when the maximum speed of 20 m/min was reached. For either baseline or post-ischemic exercise performance of FGF2 LMW-only or
FGF2 HMW-only mice, levels of exercise capacity similar to WT mice were observed at all speeds
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of the treadmill testing protocol. These results provide preliminary evidence suggesting that in the
absence of Fgf2 expression, there is an impaired functionality of the musculoskeletal system occurring in the absence or presence of ischemic injury.
A. Performance at baseline 50 WT Fgf2-/- 40 FGF2 LMW-only FGF2 HMW-only 30
20 Beam breaks/min Beam 10
0 7 10 15 20 Speed (meters/min)
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B. Performance after 14 days of ischemia 50 WT Fgf2-/- 40 FGF2 LMW-only FGF2 HMW-only 30
20 Beam breaks/min Beam 10
0 7 10 15 20
Speed (meters/min)
Figure 10. Treadmill exercise performance of WT (n=1), Fgf2-/- (n=2), FGF2 LMW-only
(n=2) and FGF2 HMW-only (n=2) ischemic limbs before (A) and 14 days after (B) induction of hindlimb ischemia. Performance data are presented as mean beam breaks ± SEM.
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HMW FGF2 promotes while LMW FGF2 abrogates gross post-ischemic tissue viability
In conjunction with recovery of ischemic limb function evaluation, daily examinations
were performed to assess the FGF2 isoform-induced gross tissue viability of the ischemic limb.
None of the mice had any necrosis of the sham-operated limb. Of the 124 male and female mice included in the study, ~16% (20/124) exhibited signs of tissue necrosis in the ischemic limb.
Necrosis was defined as the darkening of one or more toes or part of the footpad within 1-3 days of surgery but did not include limb loss (auto-amputation). Only 4 of the 124 mice with necrosis experienced auto-amputation and these mice were immediately euthanized and excluded from the rest of the study. In the mice with necrotic limbs, the tissue necrosis was not progressive and they were healed within 2 weeks of the surgical induction. WT mice had a 22.5% (7/31) incidence of necrosis (Figure 11). Fgf2-/- and FGF2 LMW-only mouse groups had increased incidences that
were not statistically different, 21.9% (7/32) and 23.16% (6/26), respectively. Conversely, mice
with expression of only the HMW FGF2 had no signs of tissue necrosis (0/26, Figure 11, p<0.05)
compared to the other groups. Interestingly, the mice that had to be excluded from the study due
to limb auto-amputation were from the WT or FGF2 LMW-only groups (2 of each). This fact
coupled with the observations that limbs expressing only HMW FGF2 had no necrosis and those
expressing only LMW FGF2 were more susceptible to necrosis suggests a protective role for
HMW FGF2 in tissue viability and a deleterious function for LMW FGF2.
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100
80
60
40 % ANIMALS
20
0 WT Fgf2-/- FGF2 LMW- FGF2 HMW- only only NECROSIS NO NECROSIS
B. MALE MICE * † ‡ 100
80
60
40 % ANIMALS
20
0 WT Fgf2-/- FGF2 LMW- FGF2 HMW- only only NECROSIS NO NECROSIS
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FEMALEAll mice MICE C. 140 * † ‡ 100 WT 120 Fgf2 -/- 80 100 FGF2 LMW-only FGF2 HMW-only 60 80
60
40% Animals
% ANIMALS 40
20 20
0 0 NECROSIS NO NECROSIS WT Fgf2-/- FGF2 LMW- FGF2 HMW- only only NECROSIS NO NECROSIS
Figure 11. (A) Incidence of limb necrosis (defined as discoloration of toes and/or foot observed daily for up to 14 days of ischemia compared to the sham-operated) in WT,
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only ischemic limbs. (B) Incidence of limb necrosis in male ischemic limbs. (C) Incidence of limb necrosis in female WT (n=14), Fgf2-/-
(n=13), FGF2 LMW-only (n=18) and FGF2 HMW-only (n=10) ischemic limbs. For each sex/genotype, the presence or absence of necrosis is expressed as a percentage of the number of animals in which hindlimb ischemia surgery was performed. Data is presented as percentages of all animals in the study. *p<0.05 vs. WT, †p<0.05 vs. Fgf2-/-, ‡p<0.05 vs. FGF2
LMW-only.
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Expression of only HMW FGF2 enhanced post-ischemic vascular network
To identify the role of the individual FGF2 isoforms in ischemia-related vascular and collateral growth and remodeling, both contrast-enhanced micro-CT imaging and immunohistochemistry were performed. The next set of figures (Figure 12-15) depict the outcomes from the micro-CT experiments. This imaging technique is relies on the radiopacity of material and resultant X-ray attenuation to generate sequential tomographic images (slices) which are then reconstructed to produce three dimensional (3-D) volumes of the scanned object (Bouxsein et al.,
2010a; Downey et al., 2012). The low inherent contrast of vessels and muscle (due to low atomic numbers of carbon and oxygen) requires the use of a radiodense contrast medium to increase image contrast (A. S. Lin et al., 2007; Zagorchev et al., 2010). In this case, a lead-based material, Microfil was perfused into the hindlimbs of WT, Fgf2-/-, FGF2 LMW-only, and FGF2 HMW-only mice to visualize and quantitatively analyze the vascular network at 42 days of ischemia (Figure 12).
The 3-D images of the sham and ischemic limbs reveal the presence of perfused superficial femoral arteries in the former limbs and the absence in the latter. After digitally removing the bone structures and automatic segmentation of tissue from vessels in the 3-D volumes, morphometric analysis was performed to determine the volume, number and spacing of the vessels in the calf or thigh regions. The delineation of the calf from the thigh allows for the distinction of angiogenesis from arteriogenesis in ischemic vascular repair (Limbourg et al., 2009). Femoral artery ligation and excision has been shown to predominantly induce angiogenesis in the calf muscles of the lower hindlimb and collateralization in the upper thigh regions (Oses et al., 2009; G. Tang, Charo, Wang,
Charo, & Messina, 2004; Tirziu et al., 2005). To minimize the possibility of vascular over- or
under-estimation resulting from variations in contrast-agent perfusion, the vascular morphology
parameters of each ischemic limb was reported as the percent change from contralateral, non-
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ischemic (sham) limb. The increase in vessel volume (volume of segmented vessels normalized to
calf volume) in the ischemic calf of WT mice was similar to Fgf2-/- limbs (47% vs. 47%) (Figure
13A). Expression of only LMW FGF2 produced a significantly smaller change in vascular volume
(18%, p<0.05 vs WT). The FGF2 HMW-only ischemic calves had the greatest increase in vessel
volume (69%, p<0.05 vs WT). The change in vessel density (number of vessels/mm of tissue) and
intervessel spacing (distance between vessels) were also determined (Figure 13B-C). Consistent
with the vascular volume results, the ischemic limbs, expressing only LMW FGF2, had the least
change in vessel density and was significantly different from WT or HMW-only limbs (p<0.05).
A decrease in intervessel spacing in the ischemic limbs relative to the sham limb is indicative of
an increase in the portion of the tissue occupied by vessels. This factor was only decreased by 10%
in FGF2 LMW-only calves while WT and FGF2 HMW-only had significantly “higher” (i.e. more negative) changes in vessel separation with 40.68% and 40.73%, respectively (p<0.05).
Simultaneous measures of vessel volume, density and spacing were also obtained in the thigh regions of the sham and ischemic limb volumes (Figure 13). The thigh vessel volume followed the same pattern as was observed in the ischemic calf where the increase in ischemic vascular volume of the Fgf2-/- was similar to WT limbs but the increase in the FGF2 HMW-only
muscles was higher than WT. The thigh vessel numbers in the HMW FGF2 expressing limbs were
also not different from WT. Surprisingly, the vascular density in the ischemic Fgf2-/- thighs was
decreased compared to WT despite similar vessel volumes. The FGF2 LMW-only thighs had reduced vessel densities relative to WT which would be expected given the lower vessel volumes that was also observed. The percent change in the thigh intervessel spacing of the WT, Fgf2-/- and
FGF2 HMW-only limbs were not different from each other but they were significantly greater than
the spacing of the FGF2 LMW-only thighs (p<0.05).
106
Taken together, these micro-CT data demonstrate that the HMW isoforms of FGF2 possess a greater capacity for promoting collateral arteriogenesis in both ischemic calf and thigh muscles.
107
FGF2 FGF2 WT Fgf2-/- LMW-only HMW-only
SHAM
HLI
Figure 12. Representative micro-computed tomography (micro-CT) images of sham (top row) and ischemic (bottom row) hindlimbs
(HLI) from WT, Fgf2-/-, FGF2 LMW-only, and FGF2 HMW-only mice at 42 days of ischemia. Perfused femoral arteries (red arrows)
are present only in the sham limbs.
108
A. 100 WT Fgf2 -/- FGF2 LMW-only 80 FGF2 HMW-only *
60 * † *
40 Vessel Volume Vessel
% % ChangeSham from * † 20
0 CALF THIGH
B. 100
80
* 60 * * †
40 Vessel Density Vessel
% Change from Sham from Change% * † 20
0 CALF THIGH
109
C. WT -100 Fgf2 -/- FGF2 LMW-only -80 FGF2 HMW-only
-60
*† -40 Vessel Spacing Vessel % Change from Sham from Change% -20 *† *
0 CALF THIGH
Figure 13. Vessel volume (normalized for tissue volume) (A), vessel density (B) and vessel spacing (C) was calculated from 3-D micro-CT images of WT (white bars), Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars) limbs. Volume, density and spacing of the ischemic limb vasculature in the calf or thigh are expressed as change (%) from sham limb. n=4 to 6. *p< 0.05 vs. WT, †p<0.05 vs. FGF2 HMW-only.
110
Expression of only HMW FGF2 increases the number of small and medium-sized collaterals in the post-ischemic vascular network
In order to provide additional characterization of the HMW or LMW FGF2-induced vascular network, the range of vessel diameters present in sham and ischemic hindlimbs was determined. 3-D limb volumes generated by micro-CT were analyzed and quantified for the distribution of vessels thicknesses (diameter). Histograms of vessel sizes present in the mouse skeletal muscle showed vessels that ranged from 34µm to 408µm. 34µm was the smallest vessel able to be visualized and quantified (i.e. the highest resolution possible for this study) and 408 µm was the upper limit above which the vascular data proved to be unreliable for analysis.
A similar distribution of vessels was observed in the (sham) non-ischemic limbs of
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice relative to WT (Figure 14). Also, the volume of the vessels at each thickness category (vessel frequency) was not different from WT. In
WT and FGF2 HMW-only ischemic limbs, the vessels on the lower half of the thickness range
(34-200µm, smaller collaterals) were increased in volume compared to their respective sham limbs
(Figure 15A, p<0.05). Additionally, the number of 34-200μm-sized collaterals was significantly higher in the FGF2 HMW-only group than WT (p<0.05). Conversely, the 34-200µm vessel volume of FGF2 LMW-only ischemic limbs was unchanged from the sham limb and significantly lower than WT or FGF2 HMW-only volumes (p<0.05).
Volumes of medium-sized (200-400μm) collateral vessels in WT and FGF2 HMW-only
ischemic limbs were also increased compared to the contralateral sham limb (Figure 15B, p<0.05).
Expression of only HMW FGF2 resulted in significantly higher volumes of 200-400μm vessels
relative to WT (p<0.05), while the presence of only the LMW isoform did not change the post-
111
ischemic development of the medium-sized collaterals. As was observed with the smaller ≤200µm
vessels, medium sized vessels in FGF2 LMW-only muscles were significantly decreased
compared to WT and HMW FGF2-only muscles (p<0.05). Overall, these results suggest that the
development of small (34-200µm) and medium (200-400µm, to a lesser degree) sized vessels are
responsible for the increased vascular volume present in ischemic HMW FGF2-expressing muscle.
A. 30 Vessel Diameter (µm)
# WT Sham WT HLI # # # 20 voxels) 4 4
10 Vessel Volume (x10 Volume Vessel
0 34 68 102 136 170 204 238 272 306 340 374 408
112
B.
30 Fgf2 -/- Sham
Fgf2 -/- HLI
voxels) 20 4 # #
10 Vessel Volume (x10 Volume Vessel
0 34 68 102 136 170 204 238 272 306 340 374 408 Vessel Diameter (µm)
C. 30 FGF2 LMW-only Sham
FGF2 LMW-only HLI
20 voxels) 4
10 Vessel Volume (x10 Volume Vessel
0 34 68 102 136 170 204 238 272 306 340 374 408 Vessel Diameter (µm) 113
D. 40 FGF2 HMW-only Sham # # # # FGF2 HMW-only HLI 30 voxels) 4 20 #
10 Vessel Volume (x10 Volume Vessel 0 34 68 102 136 170 204 238 272 306 340 374 408 Vessel Diameter (µm)
Figure 14. Histogram of vessel thickness distribution in sham (white bars) and ischemic
(black bars) limbs of WT (A), Fgf2 -/- (B), FGF2 LMW-only (C), and FGF2 HMW-only (D)
mice analyzed from micro-CT images at 42 days of ischemia. Data are presented as mean ±
SEM, n=4 to 6. #p<0.05 vs. SHAM cohort.
114
200 34-200μm diameter vessels A. WT 160 Fgf2 -/- FGF2 LMW only # * FGF2 HMW only voxels)
4 120 # † 80
40 Vessel Volume (x10 Volume Vessel
0 SHAM HLI B.
100 200 - 400µm diameter vessels
WT # * 80 Fgf2 -/- FGF2 LMW only #
voxels) FGF2 HMW only 4 60
* † 40
Vessel Volume (x10 Volume Vessel 20
0 SHAM HLI
115
Figure 15. The volume of small-sized (34 -200μm) vessels (A) and medium-sized (200-400μm) vessels (B) analyzed from micro-CT images of WT (white bars), Fgf2-/- (light gray bars),
FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars) limbs. Data are presented as mean ± SEM, n=4 to 6. #p<0.05 vs. SHAM cohort, *p< 0.05 vs. WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
116
Loss of LMW FGF2 perturbs while HMW FGF2 augments capillary growth after 14 days
of ischemia
To elucidate the effect of altered FGF2 isoform expression on the initial vascular response to ischemic injury, skeletal muscles from sham-operated or ischemic WT, Fgf2-/-, FGF2 LMW- only and FGF2 HMW-only mice were harvested at 14 days of ischemia. Immunohistological analysis was performed to determine the level of post-ischemic microvessel growth. Angiogenesis, referring solely to capillary vessel development, was measured by quantitative analysis of muscle cross-sections stained with Griffionia Simplicifolia Isolectin B4 (GSI Lectin). GSI Lectin-positive
endothelial cells (ECs) were present on the inner layer (endothelium) of all blood vessels (Figure
17). Capillary vessels were identified as vessels with an outer diameter ≤10 µm and positive
staining for ECs. Capillary vessel densities were determined as the ratio of the total number of
capillary vessels to myofiber numbers. This was done to address the possibility of under or
overestimation of vessel numbers due to ischemia-related edema or atrophy.
Capillary density was measured in muscle regions of the mouse hindlimb that are proximal
(calf) or distal (thigh) to the excised portions of the femoral artery and vein pair (Figure 16). After
14 days of ischemia, the capillary vessel density ratio (ischemic/sham) of WT, Fgf2-/- and FGF2
HMW-only limbs were greater than 1 indicating the presence of increased vessel numbers in the
ischemic muscles. FGF2 LMW-only ratios were scarcely above 1 at (1.07 ± 0.05) and (1.01 ±
0.06) for calf and thigh muscles respectively, and were decreased compared to WTs. Fgf2-/-
capillary density ratios were similar to WT ratios in either the calf or thigh. In FGF2 HMW-only
limbs, the ratio of capillary vessels of the thigh muscles was significantly greater than WT while
the calf muscle ratio had a trend towards an increase. Additionally, FGF2 HMW-only capillary
ratios were elevated relative to the FGF2 LMW-only ratios in both calf and thigh muscles. These
117 results suggest that modulation of FGF2 isoform expression alters the vascular response to ischemia. Furthermore, the data show the opposing roles for HMW FGF2 (stimulation) and LMW
FGF2 (suppression) in angiogenesis (capillary growth).
Capillary Density WT 3 Fgf2-/- FGF2 LMW-only 2.5 FGF2 HMW-only
2 *
1.5 * † * † 1
0.5 Vessel Density Ratio (Ischemic/Sham) Ratio Density Vessel
0 CALF THIGH
Figure 16. Capillary density (capillary number/myofiber number) in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars) calf or thigh expressed as a ratio of ischemic to sham muscle vessel density after 14 days of ischemia. Data are presented as mean ratio ± SEM, n= 4 to 5. *p< 0.05 vs. WT,
†p<0.05 vs. FGF2 HMW-only.
118
LMW FGF2 isoform did not increase post-ischemic vascular (capillary) growth while HMW
FGF2 improved capillary vascularization after 42 days of ischemia
To further elucidate the effect of altered FGF2 isoform expression on the vascular response to ischemic injury, skeletal muscles from sham-operated or ischemic WT, Fgf2-/-, FGF2 LMW- only and FGF2 HMW-only mice were also harvested at 42 days of ischemia. No significant differences in capillary density were observed in non-ischemic (sham) muscles between WT and
Fgf2-/- hindlimbs (Figure 18). Limbs with FGF2 HMW-only expression were also similar to WTs while the FGF2 LMW-only limbs had decreased capillaries in the sham muscles. Capillary density in the WT, Fgf2-/-, and FGF2 HMW-only ischemic muscles were significantly increased relative to their respective sham limbs (p<0.05). FGF2 LMW-only ischemic limbs did not have any change capillary density compared to their non-ischemic, sham limbs. Moreover, the proportional increase in capillary density was highest in the HMW FGF2 expressing muscles and the density of vessels in the ischemic FGF2 HMW-only limbs was greater than that of the WT, Fgf2-/-, or FGF2 LMW- only ischemic limbs (p<0.05).
To determine the contributions of the specific calf and thigh muscle groups to the overall changes in ischemic vascular density, capillary density was evaluated in the sham and ischemic gracilis, semi-membranosus, semi-tendinosus and gastrocnemius. The latter muscle makes up the calf muscle and the former three groups are found in the thigh. Increased vascular density in the gracilis and gastrocnemius muscles had the greatest contribution to the capillary growth (Figure
19 A, D). In WT, Fgf2-/- and FGF2 HMW-only gracilis and gastrocnemius muscles, capillary density was significantly higher than the contralateral sham muscles (p<0.05). FGF2 LMW-only ischemic muscles had no change in the vascular density relative to the sham limbs in the gracilis, semi-membranosus, semi-tendinosus or gastrocnemius muscles (Figure 19 A-D). Furthermore, the
119
capillary densities in WT or FGF2 HMW-only ischemic muscles were consistently greater than in
FGF2 LMW-only muscles (p<0.05). Overall these data, for the first time, indicate that HMW
FGF2 isoform expression is important for capillary growth after ischemia and supports the
anatomical vascular density results obtained with micro-CT.
120
WT Fgf2-/- FGF2 LMW-only FGF2 HMW-only
SHAM
HLI
Figure 17. Representative photomicrographs of GSI-Lectin stained vessels (all vessels) in WT, Fgf2-/-, FGF2 LMW-only, and
FGF2 HMW-only sham and ischemic muscle sections harvested at 42 days of hindlimb ischemia. Scale bar: 10μm, arrows
(capillaries), arrowheads (non-capillary GSI-Lectin+ vessels).
121
Capillary Density 1 WT # * Fgf2 -/- 0.8 FGF2 LMW-only FGF2 HMW-only # 0.6 #
*† 0.4 Vessel Myofiber# / # Vessel 0.2
0 SHAM HLI
Figure 18. Capillary density (expressed as capillary number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light gray bars),
FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs. WT HLI, †p<0.05 vs. FGF2
HMW-only HLI.
122
A. B. Semi-membranosus 1
0.8
0.6
0.4 Vessel # / Myofiber # Vessel 0.2
0 SHAM HLI
C. Semi-tendinosus D. Gastrocnemius 1 1 WT Fgf2 -/- WT FGF2 LMW-only 0.8 Fgf2 -/- 0.8 FGF2 HMW-only FGF2 LMW-only * FGF2 HMW-only * * 0.6 * 0.6 †
0.4 0.4
# # Vessel / Myofiber Vessel # Vessel / # Myofiber 0.2 0.2
0 0 SHAM HLI SHAM HLI
Figure 19. (A) Gracilis muscle capillary density (expressed as vessel number/myofiber
number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF HMW-only (black
123
bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs.
WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
(B) Semi-membranosus muscle capillary density (expressed as vessel number/myofiber
number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black
bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs.
WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
(C) Semi-tendinosus muscle capillary density (expressed as vessel number/myofiber number)
of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light
gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs. WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
(D) Gastrocnemius muscle capillary density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs. WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
124
Loss of LMW FGF2 affects the early arteriogenic response to chronic ischemia
To address the function of selective FGF2 isoform expression in ischemia-induced vascular
growth, the level of arteriole (arteriogenesis) and venule growth were examined. To detect
arteriolar or venule vessels, α-smooth muscle actin (α-SMA) antibody was used to perform
immunohistological labeling of serial sections of sham-operated or ischemic WT, Fgf2-/-, FGF2
LMW-only and FGF2 HMW-only mice. α-SMA is a marker for pericytes and vascular smooth
muscle cells (vSMCs) present in the walls of venules, veins, arterioles and arteries (Skalli et al.,
1989). Arterioles/arteries were defined as α-SMA positive vessels with ≥20 µm outer diameter.
They had several layers of positive staining and a round lumen. Veins/venules were
morphologically distinguishable from arterioles/arteries by their collapsed luminal structure and
thinner layers of smooth muscle (Figure 21).
Arteriole density was determined as a ratio of the number of myofibers. Arteriole density
at 14 days of ischemia was evaluated in sham or ischemic muscle and with the ischemic density
expressed as a ratio of the later (Figure 20). The ratios of WT, Fgf2-/-, FGF2 LMW-only, FGF2
HMW-only limbs were higher than 1 indicating the presence of increased vessel numbers in the ischemic muscles. However, in the presence of only LMW FGF2, ratios were only slightly greater than 1 at (1.16 ± 0.06) and (1.04 ± 0.08) for calf and thigh muscles respectively. These ratios were decreased compared to WT or FGF2 HMW-only limbs. WT, Fgf2-/- and FGF2 HMW-only limbs all had similar arteriole density ratios (calf or thigh). These data suggest that unlike the HMW
FGF2 isoforms, LMW FGF2 does not stimulate collateral vessel remodeling (arteriogenesis) in response to ischemia.
125
Arteriole Density 3 WT Fgf2-/- 2.5 FGF2 LMW-only FGF2 HMW-only 2
1.5 * † * † 1
0.5 Vessel Density Ratio (Ischemic/Sham) Ratio Density Vessel
0 CALF THIGH
Figure 20. Arteriole density (arteriole number/myofiber number) in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars) calf or thigh expressed as a ratio of ischemic to sham muscle vessel density after 14 days of ischemia. Data are presented as mean ratio ± SEM, n= 4 to 5. *p< 0.05 vs. WT,
†p<0.05 vs. FGF2 HMW-only.
126
Expression of only HMW FGF2 has greater impact on ischemic arteriolization
(collateralization) 42 days after ischemia compared to WT or LMW isoform expression
Arteriole or venule density was also determined at 42 days and expressed as a ratio of the
number of myofibers (Figure 21). Venule or arteriole density was evaluated in sham or ischemic
muscles. No differences in venule or arteriole density were detected in non-ischemic (sham)
muscles between WT and Fgf2-/- hindlimbs (Figure 22A). Similarly, FGF2 LMW-only or FGF2
HMW-only sham muscles were not different from WT. After 42 days of ischemia, venule densities
in ischemic limbs of all groups were significantly increased compared to their respective sham
limbs (Figure 22A, p<0.05). To identify the changes in specific in calf (gracilis, semi-
membranosus, and semi-tendinosus) or thigh (gastrocnemius) muscles, the density of venule
vessels in the respective muscle layers was also determined. In the thigh, gracilis venule density
was increased above sham levels in all mouse groups except FGF2 LMW-only (Figure 23A,
p<0.05). The gastrocnemius muscle had elevated venule densities in all mouse groups (Figure 23D,
p<0.05). However, the venule densities in the ischemic gracilis, semi-membranosus, semi- tendinosus or gastrocnemius muscles were not different.
Arteriole density was significantly increased in ischemic muscles of WT, Fgf2-/-, FGF2
LMW-only or FGF2 HMW-only mice (Figure 22B, p<0.05). While expression of only LMW
FGF2 in ischemic hindlimbs induced a similar increase in arteriole density as the WT or Fgf2-/- limbs, expression of only the HMW isoforms promoted an even greater amount of revascularization compared to the three other groups (p<0.05). Ischemic FGF2 HMW-only arteriole density was elevated nearly five-fold relative to non-ischemic limbs but WT, Fgf2-/- and
FGF2 LMW-only muscles had only a two-fold increase. To assess the roles of individual calf and thigh skeletal muscles in the arteriole density changes observed during ischemia, vessel density in
127 gracilis, semi-membranosus, semi-tendinosus, or gastrocnemius muscle alone was calculated.
Unlike the angiogenesis results where the ischemia-induced capillary growth could be attributed to only the gracilis and gastrocnemius muscles, arteriogenesis was observed in all calf and thigh muscles analyzed (Figure 24A-D). Additionally, the expression of only the HMW FGF2 isoforms produced consistently higher levels of arterioles across all the muscle groups (p<0.05). Taken together, these results provide additional evidence that arteriogenesis is compromised in the LMW
FGF2-expressing limbs and confirms the increased of capacity of the HMW isoforms for inducing collateral growth.
In order to eliminate the likelihood that vessel density measurements were over- or under- estimated due to ischemia-related edema (tissue swelling) or atrophy (muscle shrinking), the size of the sham and ischemic muscles was determined. Muscles from the calf (gastrocnemius) or thigh
(semi-membranosus, semi-tendinosus, or gracilis) were weighed when harvested at the end of the study (Day 42 of ischemia). In general, the masses of the ischemic muscles were decreased relative to the sham limb but the differences were not significant. For all genotypes, the weights of the ischemic gracilis, semi-membranosus, semi-tendinosus, or gastrocnemius muscle was not different from the sham group (Figure 25).
128
WT Fgf2-/- FGF2 LMW-only FGF2 HMW-only
SHAM
HLI
Figure 21. Representative photomicrographs of α-smooth muscle actin (α-SMA) stained vessels (venules or arterioles) in WT,
Fgf2-/-, FGF2 LMW-only, and FGF2 HMW-only sham and ischemic muscle sections harvested at 42 days of hindlimb ischemia.
Scale bar: 10μm.
129
A. Venule Density 0.05 WT 0.04 Fgf2-/- FGF2 LMW-only FGF2 HMW-only 0.03
0.02 * * Vessel # / # Myofiber Vessel * * 0.01
0 SHAM HLI
B. Arteriole Density
0.05 WT Fgf2-/- *# 0.04 FGF2 LMW-only FGF2 HMW-only 0.03
*† * * 0.02
Vessel # / Myofiber # # Myofiber / Vessel 0.01
0 SHAM HLI
130
Figure 22. (A) Venule density (expressed as vessel number/myofiber number) of sham and
ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light gray bars),
FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort.
(B) Arteriole density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light gray bars), FGF2 LMW- only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ±
SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs. WT HLI, †p<0.05 vs. FGF2 HMW- only HLI.
131
A. B. Gracilis Semi-membranosus 0.05 0.05 WT Fgf2-/- 0.04 0.04 FGF2 LMW-only FGF2 HMW-only 0.03 0.03 * 0.02 0.02 * * Vessel # / Myofiber # Vessel Vessel # / Myofiber # Vessel 0.01 0.01
0 0 SHAM HLI SHAM HLI
C. D. Semi-tendinosus Gastrocnemius 0.05 0.05
0.04 0.04
0.03 0.03 * 0.02 0.02 * * * Vessel # / Myofiber # Vessel
Vessel # / Myofiber # Vessel * 0.01 0.01
0 0 SHAM HLI SHAM HLI
Figure 23. (A) Gracilis muscle venule density (expressed as vessel number/myofiber number)
of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light
gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are
presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort.
132
(B) Semi-membranosus muscle venule density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8.
(C) Semi-tendinosus muscle venule density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort.
(D) Gastrocnemius muscle venule density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort.
133
A. Gracilis B. Semi-membranosus
0.05 0.05 *# WT Fgf2 -/- 0.04 * 0.04 * # FGF2 LMW-only FGF2 HMW-only 0.03 0.03 † * * # 0.02 0.02 * † * * Vessel # / Myofiber # Vessel Vessel # / Myofiber # Vessel 0.01 0.01
0 0 SHAM HLI SHAM HLI
C. D. Gastrocnemius Semi-tendinosus 0.05 0.05 WT * # Fgf2 -/- *# 0.04 FGF2 LMW-only 0.04 FGF2 HMW-only * 0.03 * * 0.03 * † * * 0.02 0.02 Vessel # / Myofiber # Vessel
0.01 # / Myofiber # Vessel 0.01
0 0 SHAM HLI SHAM HLI
Figure 24. (A) Gracilis muscle arteriole density (expressed as vessel number/myofiber
number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black
134
bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs.
WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
(B) Semi-membranosus muscle arteriole density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs.
WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
(C) Semi-tendinosus muscle arteriole density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars),
Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs.
WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
(D) Gastrocnemius muscle arteriole density (expressed as vessel number/myofiber number) of sham and ischemic muscles after 42 days of ischemia in WT (white bars), Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars). Data are presented as mean ± SEM, n= 6 to 8. *p<0.05 vs. SHAM cohort, #p<0.05 vs. WT HLI, †p<0.05 vs. FGF2 HMW-only HLI.
135
G racilis Semi-membranosus 100 200
75 150
50 100
25 50 W et m uscle w eight (m g) W et m uscle w eight (m g) 0 0 SHAM HLI SHAM HLI
Semi-tendinosus G astrocnem ius WT 250 200 Fgf2 -/- FG F2 LM W -only FG F2 H M W -only 200 150
150 100 100
50 50
W m et weights uscle (m g) W m et weights uscle (m g) 0 0 SHAM HLI SHAM HLI
Figure 25. Wet weights of sham and ischemic gracilis (A), semi-membranosus (B), semi-
tendinosus (C) and gastrocnemius (D) muscles. Sham or ischemic WT, Fgf2-/-, FGF2 LMW- only and FGF2 HMW-only muscles were collected at 42 days of ischemia.
136
Recovery of ischemic foot perfusion is unaffected by targeted deletion of LMW or HMW
FGF2
To determine the role of differential levels of ischemia-induced vascularization on limb perfusion, blood flow in the sham and ischemic limbs was analyzed over time. The non-invasive nature of the laser Doppler imaging allowed for baseline measurements before surgery and several serial measurements up to 42 days after hindlimb ischemia surgery from the non-ischemic (sham) and ischemic lower limbs (paws) (Leahy, de Mul, Nilsson, & Maniewski, 1999; Wårdell,
Jakobsson, & Nilsson, 1993). The lower hindlimbs (paws, and below the site of ischemic injury) of anesthetized WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice were scanned in triplicates and color-coded images displaying the resting microcirculation flow distribution were generated (Figure 26A). Perfusion (an index of blood flow) was calculated from images and expressed as the ratio of the mean perfusion values of the ischemic to non-ischemic (sham) feet.
Basal resting perfusion ratios for all the mouse groups were similar and approximately equal to 1 (Figure 26B). This confirms the histological evidence presented previously that ablation of LMW, HMW or all FGF2 alone does not alter basal vascularization (capillaries, specifically) in skeletal muscle. After induction of hindlimb ischemia (day 1), perfusion ratios were reduced drastically to between 6 and 14% of pre-ischemia values. Perfusion improved gradually over time and at day 14 of ischemia, up to 46% of baseline recovery had been recovered. At day 42 of ischemia, all groups had similar recovery of perfusion with perfusion ratios between 52 and 56% of baseline values. No significant differences were observed between the perfusion ratios of
Fgf2-/-, FGF2 LMW-only or FGF2 HMW-only mice relative to WT at either day 1, 14 or 42 of ischemia. In conclusion, these results illustrate that the LDPI method was not able to detect any
137 changes in post-ischemic resting microcirculatory flow in the presence of only HMW FGF2, only
LMW FGF2 or no FGF2.
138
POST ISCHEMIA SURGERY A. BASELINE DAY 1 DAY 14 DAY 42
WT
Fgf2-/-
FGF2 LMW- only
FGF2 HMW- only
HLI SHAM
PERFUSION DISTRIBUTION
MAX MIN
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B. WT 1.2 Fgf2 -/- FGF2 LMW-only 1.0 FGF2 HMW-only
0.8
0.6 Perfusion Ratio Ratio Perfusion (Ischemic/Sham) 0.4
0.2
0.0 Baseline Day 1 Day 14 Day 42
Figure 26. (A) Representative laser Doppler perfusion images of lower limbs (i.e., footpad) of WT, Fgf2-/-, FGF2 LMW-only, and FGF2 HMW-only mice before (Baseline) and serially
(Days 1, 14 and 42) after surgery. The ischemic (HLI) limb is pictured on the left in each image and the non-ischemic limb (SHAM) is on the right. In each image, maximum (high) perfusion is signified by the red colors, yellow and green depict the mid-range and the blues indicate minimum (low) perfusion. (B) Recovery of footpad perfusion during chronic ischemia of WT (blue), Fgf2-/- (red), FGF2 LMW-only (green), and FGF2 HMW-only
(black) mice. Quantitative analysis of foot perfusion over time computed from color-coded images. Perfusion ratio is average perfusion value of the ischemic to sham (non-ischemic) limbs. Data are presented as mean ± SEM, n=4.
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DISCUSSION I
The results presented in this dissertation demonstrate, for the first time, that the endogenous
HMW FGF2 isoforms are important and necessary for the protection of skeletal muscle from
chronic ischemia. In the presence of only the HMW isoforms, limb impairment and necrosis were
abolished in the ischemic mouse hindlimb. The salvage of the ischemic muscles was associated,
in part, with enhanced FGF2 expression and revascularization (angiogenesis, arteriogenesis and
collateralization). Early preservation of ischemic limb function occurred in parallel with increased
remodeling/development of small and medium sized collateral vessels which suggests a protection
of skeletal muscle that was independent of the recovery of muscle perfusion (Figure 27).
Research into therapeutic interventions with the potential to ameliorate/treat the morbidity
and poor quality of life associated with coronary or peripheral arterial disease has necessitated the
use of pre-clinical models. In vivo experimental models with a three-dimensional representation
of the whole organ (muscle and vasculature) that could reproducibly recapitulate symptoms of the human disease conditions are the ideal choice (Hoefer, van Royen, & Jost, 2006; Limbourg et al.,
2009; Lotfi et al., 2013). The hindlimb ischemia (HLI) model has been used to successfully investigate the tissue response to PAD and to assess the efficacy of candidate therapies (Dragneva et al., 2013; Madeddu et al., 2006; Waters et al., 2004). This is a murine model where unilateral
ligations and/or excisions are performed on the common femoral, common iliac or external iliac
arteries that perfuse the lower periphery (Hoefer et al., 2006; Lotfi et al., 2013; Madeddu et al.,
2006; Waters et al., 2004). In this dissertation, dual ligatures were inserted at proximal and distal
ends of superficial femoral artery/vein pair. The proximal suture was placed at the origin of the
deep femoral branch while the distal ligation occurred at the bifurcation of the saphenous and
popliteal branches (Figure 4). This approach routinely results in a reduction in foot perfusion (as
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low as 20% of pre-surgery flow) of the ligated limb (Limbourg et al., 2009; Peng et al., 2011;
Sullivan et al., 2002; Westvik et al., 2009). The unligated contralateral side served as an internal
control for comparison. The immediate responses to the dramatic loss of blood supply in this model
range from tissue necrosis, ischemic limb impairment, muscle atrophy to auto-amputation of toes
and/or feet. In order to prevent sufferance of the animals, mice with spontaneous auto-amputation
were euthanized and excluded from the rest of the study.
This dissertation for the first time demonstrated that the endogenous FGF2 isoforms play
opposing functions in protecting ischemic skeletal muscle from necrosis and preserving functional
capacity. Necrosis was largely confined to the lower limbs and comprised of (dis)coloration and/or
loss of one or more toe or toenails. Mice experiencing auto-amputation of toes and/or entire limbs
were excluded from the analysis of necrosis and all other endpoints. Deficiency of HMW FGF2
(expression of only LMW FGF2) had significant necrosis of ischemic limbs (Figure 11). While
extent of necrosis was not different compared with WT limbs (22.6% vs 17.1%), it was
significantly increased compared with HMW FGF2 expressing limbs which had no incidences of
necrosis. Total loss of FGF2 expression (Fgf2-/-) also produced no changes in macroscopic
necrosis relative to WT. Functional recovery of the ischemic hindlimbs was assessed with
treadmill testing. This measure of forced exercise performance was performed at baseline before
the induction of hindlimb ischemia and at 14 days after ischemia surgery (Figure 12). Mice were
tested at a 7o incline using an exercise protocol that began with an initial speed of 7 meters/min
and ended with a maximal speed of 20 meters/min. In mice where only LMW FGF2 or HMW
FGF2 was expressed, exercise performance was not different from WT at baseline or during
ischemia recovery. In the absence of any FGF2 expression (Fgf2-/-), exercise capacity was decreased at baseline and during ischemia recovery (at 14 days).These results are preliminary due
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to the small sample sizes (≤ 2 mice per group). Nevertheless, they provide some evidence that Fgf2
is important for maintaining hindlimb functional capacity in the presence or absence or ischemia.
As an additional measure of the FGF2-dependent functional response to ischemia, mice
were graded for the spontaneous or stimulated (tail traction) use of their ischemic limbs. Movement
of the ischemic limbs was restricted immediately following surgical procedure and continued for
the first three days following the ischemia surgery where mice gait was altered and presented as
dragging and no weight bearing of the ischemic foot (Table 12). This was observed in the ischemic feet of all WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice (Figure 9). HMW FGF2 expression resulted in a more rapid (~3x) recovery of active foot and limb movement (plantar flexion with flexing of toes and full weight bearing) in contrast to WT mice (10 vs. 28 days). In the presence of only LMW FGF2, mice had a sustained impairment of ischemic limbs and were delayed in regaining full use of their ischemic limbs until 42 days after ischemia surgery. Mice with a complete loss of Fgf2 gene expression (Fgf2-/-) only had a partial recovery of ischemic limb function by the end of the study.
These results suggest that the HMW FGF2 isoforms have a pronounced effect on preserving tissue viability and limb salvage. These data support the hypothesis that HMW FGF2 promotes cell protection and/or survival pathways in injured tissue. Evidence for this supposition was first observed in mice with global loss of Fgf2 expression. In addition to an altered cardiac
phenotype, these mice had delayed wound healing and neuronal deficits (Dono et al., 1998; Miller
et al., 2000; Ortega et al., 1998). The deficits included changes in the architecture of the neocortex
and a decrease in the neuronal density of the motor cortex layers. Additionally, exogenous 21 or
23kDa HMW FGF2 mediated the survival and neurite promoting activities of cultured
mesenphalic dopaminergic neurons and protected them from 6- hydroxydopamine (6-OHDA)-
143 induced neurotoxicity, a model of Parkinson’s disease (Grothe, Schulze, et al., 2000). When
Schwann cells were transfected to overexpress the HMW FGF2 isoforms, they stimulated the survival and functional recovery of co-transplanted dopaminergic micrografts (Timmer et al.,
2004). Also, apomorphine-induced survival of midbrain dopaminergic neurons was associated with increased trafficking of HMW FGF2 (A. Li et al., 2006). HMW FGF2 also displayed survival promoting properties in non-neuronal cell types. Specifically, overexpression of the HMW isoforms and not 18kDa has been found to reduce the cytotoxic effect of lentiviral gene transfer in corneal endothelial cells (Valtink et al., 2012). Transfection of 24kDa FGF2 confers an increased resistance to radiation-induced cell death in NIH3T3 fibroblasts and HeLa cells (Cohen-Jonathan et al., 1997; Delrieu, Arnaud, Ferjoux, Bayard, & Faye, 1998). 24kDa FGF2 also enhances the survival of rat bladder carcinoma cells (Thomas-Mudge et al., 2004).
The presence of only HMW FGF2 isoforms is necessary to protect chronically ischemic hindlimb muscle from necrosis. In the heart, however, a requirement of both LMW and HMW
FGF2 for protection from cell death has been observed in response to acute cardiac ischemia- reperfusion injury (Liao et al., 2007, 2009, 2010). Correspondingly, hearts overexpressing all of the human FGF2 isoforms were protected from myocardial infarction (MI) (House et al., 2003).
Similar decreases of infarct sizes were detected at 24 hours post-MI in hearts treated with exogenous LMW or HMW FGF2 (Jiang et al., 2007). Taken together, these results paint a picture of a tissue-dependent (cardiac vs. skeletal muscle) and stimulus-dependent (acute vs. chronic ischemia) regulation by the FGF2 isoforms. Examples of the context-dependent properties of the
FGF2 isoforms include the differing (opposing) functions in peripheral never repair, bone homeostasis and fracture repair (Haastert et al., 2006; Homer-Bouthiette, Doetschman, Xiao, &
Hurley, 2014; Xiao et al., 2014).
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One caveat to note when comparing the results of one study with other rodent models of
ischemia is the dependence of the level of necrosis and recovery of limb use on the age, gender
and strain of the animals utilized. Investigators often utilize only young, male mice in their studies.
However, age and gender have been shown to significantly alter the response to hindlimb ischemia
in rodent models. Aged mice (≥13 months) subjected to ischemia have a slower recovery of foot
movement and increased necrosis relative to younger mice (3 months) (Bosch-Marce et al., 2007;
Faber et al., 2011; Rivard et al., 1999; Westvik et al., 2009). At 7 days after ischemia surgery, male mice have been shown to have improved hindlimb use and similar extent of necrosis compared to their females cohorts (Peng et al., 2011). All the hindlimb ischemic surgeries for the experiments in this dissertation were performed on age-matched (~3 months) male and female mice. The results of the necrosis experiments were stratified by gender and compared for differences (Figures 10B,
C). Within each genotype, no differences in necrosis were present between male and female mice.
Genetic backgrounds should also be considered during comparisons of ischemic mouse models (Limbourg et al., 2009; Lotfi et al., 2013). Several studies have shown that differences in inbred strains influence the severity and incidence of tissue necrosis and ischemic limb recovery.
When comparing the two most commonly utilized strains in limb ischemia studies (C57BL/6 and
BALB/c), the Schaper group observed that wildtype C57BL/6 mice had a more rapid recovery of hindlimb use function compared to the latter strain (Helisch et al., 2006). These findings were independently confirmed in two other studies and the latter group identified a similar pattern of susceptibility in the appearance of ischemic damage (Chalothorn, Clayton, Zhang, Pomp, & Faber,
2007; Dokun et al., 2008). This group also observed that A/J strain mice had an increased impairment of limb use and necrosis than BALB/c mice (Chalothorn & Faber, 2010). Overall, the
C57BL/6 and BALB/c strains can be considered to represent two extremes of the strain-dependent
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responses to chronic hindlimb ischemia where the C57BL6 strain are ischemia-protected and the
BALB/C mice are ischemia-vulnerable. The 129/Sv strain is less widely utilized in ischemia studies; Helisch and colleagues compared this strain to C57BL/6 and BALB/c mice and found their level of tissue necrosis to be intermediate between the two extremes (Helisch et al., 2006).
This serves as a template for comparison of the mouse models used in this dissertation which were
bred on a mixed background of 50% 129/Sv and 50% Black Swiss (BS) (Azhar et al., 2009;
Garmy-Susini et al., 2004; Zhou et al., 1998). Several ischemia studies have been performed in mice bred on a 50% 129/Sv and 50% C57BL/6 background. The average incidence of ischemic damage in WT hindlimbs across these studies was ~14% (Chalothorn, Zhang, Clayton, Thomas,
& Faber, 2005; Lizotte et al., 2013; Rey et al., 2009). The levels ranged from a complete absence of necrosis to an incidence as high as 33%. Recovery from functional impairment of ischemic limbs was also observed in 129 mice with normal use of the ischemic limb returning at 28 days after ischemic surgery (Bosch-Marce et al., 2007; Rey et al., 2009). These incidences of necrosis
are consistent with the results observed in the WT 129SV/BS mice studied for this dissertation.
Moreover, the complete recovery of functional use of the WT ischemic limbs occurs within a
similar time frame as that observed by others in 129SV mixed background mice.
The impairment and necrosis of the ischemic limbs examined serve as clinically relevant
endpoints for assessing the role of the FGF2 isoforms in limb salvage and function. These
observations in the mouse ischemic limb are similar to those observed in human patients of PAD
whose symptoms present clinically as either intermittent claudication (IC) or critical limb ischemic
(CLI) (Lotfi et al., 2013; Rutherford et al., 1997; Waters et al., 2004). Another relevant endpoint
is the quantitative evaluation of ischemia-induced of vascular adaptation and regeneration.
Vascular growth in response to arterial obstruction occurs via two distinct processes that differ in
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their vessel type and triggering stimulus (Limbourg et al., 2009; Schaper, 2009). Angiogenesis,
the growth (sprouting or intussception) of capillary networks from pre-existing capillaries is initiated by hypoxia in ischemic tissues while arteriogenesis, the enlargement (lumen area and wall thickness) of pre-existing collateral arterioles into larger conductance vessels is stimulated by the increase of shear forces (Fischer et al., 2006; Heil et al., 2006). The type(s) of vascular growth processes that occurs during hindlimb ischemia is highly dependent on the nature of the vascular injury. The surgical ligation (proximal and distal ligatures) and excision of the intervening segments of the femoral artery as performed for this study induces simultaneous occurrence of angiogenesis and arteriogenesis in the ischemic hindlimb (Hoefer et al., 2006; Lotfi et al., 2013;
Waters et al., 2004). In light of this, ischemic muscle tissue from the thighs where arteriogenesis
predominates and tissue from distal limb (calf) where angiogenesis occurs were collected and
analyzed (Hoefer et al., 2006; Limbourg et al., 2009). Muscles of the hindlimb from the pelvic
region (near the inguinal ligament) to the knee were defined as thigh muscles and muscles from
the knee to the ankle were designated as calf muscles (Hedrich & Bullock, 2004; Suckow,
Weisbroth, & Franklin, 2006). The gracilis, semitendinosus, and semimembranosus muscles from
the thigh and the gastrocnemius/soleus bundle from the calf were isolated from each ischemic and
sham-operated contralateral limb. In calf or thigh muscles, the extent of angiogenesis and arteriogenesis was quantified by histomorphometric assessment of tissues isolated at 42 days after
the ischemia surgery and presented as the density of capillary and arteriolar vessels respectively.
Results presented in Chapter 1 correlate with studies in the literature where a combination of
angiogenesis and arteriogenesis was present in the ischemic hindlimbs. While capillary vessel
growth was not entirely confined to the gastrocnemius (calf) muscle, greater increases from sham
densities were observed in ischemic distal limb muscles. Likewise, ischemia-induced changes in
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arteriole densities were higher in the thigh muscles. Mouse hindlimb models have shown that
increases in vascular density of ischemic limbs occur as early 7 days (Couffinhal et al., 1998;
Scholz et al., 2002; Sullivan et al., 2002) and persist up to days 28, 35, (Boerckel et al., 2014;
Meisner, Song, Annex, & Price, 2013; Ruifrok et al., 2009) and 42 (J. Zhang et al., 2009). The vascular densities presented here were calculated from skeletal muscles isolated at 14 and 42 days postoperatively to allow for stabilization of vascular remodeling in the ischemic tissues.
These data, for the first time, represent novel evidence of a beneficial role for HMW FGF2 in stimulating vascular growth, repair and remodeling during chronic ischemia. Mice with expression of only HMW FGF2 had an increased level of post-ischemic capillary density suggesting that the HMW isoforms have angiogenic properties in response to ischemic injury
(Figures 16, 18). The effect of endogenous HMW FGF2 on angiogenesis has rarely been examined.
In fact, the sole report on the role of endogenous HMW FGF2 in angiogenesis is from the Arnal group who showed that the endogenous HMW isoforms mediate the effect of estradiol (E2) on
angiogenesis in vitro (Garmy-Susini et al., 2004). Specifically, HMW FGF2 was necessary for the
stimulating effects of E2 on endothelial cell migration. This important study was limited in its use
of in vitro angiogenesis models as well as its focus on the indirect effects of HMW FGF2. The effect of exogenous recombinant HMW FGF2 on in vitro angiogenesis has also been examined.
Administration of HMW FGF2 (24kDa) to cultured endothelial cells was shown to promote cell
proliferation and inhibit migration (Piotrowicz et al., 1999). This dissertation serves to address the
paucity of research of the direct and in vivo roles of the endogenous HMW isoforms of FGF2.
The expression of only LMW FGF2 in ischemic limbs did not improve capillary growth.
In fact, the density of capillaries in FGF2 LMW-only limbs were similar compared to control
(sham) level while the FGF2 HMW-only muscles had the greatest proportional increase of
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capillary vessel numbers. The observed angiogenic response to LMW FGF2 in ischemic hindlimbs
is not consistent with some previous research showing that when given exogenously, the LMW
isoform promotes angiogenesis in vitro and in in vivo models of ischemia. Vascular cells (EC and
vSMCs) respond to treatment with the recombinant LMW isoform in several angiogenesis assays
including cell proliferation, migration or tube formation (Davis et al., 1997; Piotrowicz et al., 1999;
Presta et al., 2005, 2007; Shing et al., 1985). In in vivo models of peripheral artery ligation,
infusion of exogenous FGF2 (low molecular weight form) into ischemic muscle has been shown
to induce capillary growth (Asahara et al., 1995; Baffour et al., 1992; Bush et al., 1998; H.-T. Yang
et al., 1996). In contrast, no angiogenesis was detected in FGF2 LMW-only ischemic limbs. This
corresponds well with data from intra-arterial infusion of recombinant LMW FGF2 to ischemic
rabbit limbs and/or injection into infarcted porcine hearts where angiogenesis was inhibited by the
growth factor (Scholz et al., 2002; Watanabe et al., 1998). The incompatibility between the results presented here and those that show increased capillary growth with LMW FGF2 treatment could be due to several reasons. First, it cannot be overstated that the cellular response to a growth factor
in cell culture is vastly different from the conditions that exist in vivo where cell phenotype is
dependent upon a coordinated physiological response. Cell culture experiments may likely
overestimated the functions of the growth factor. Second, exogenous LMW FGF2 was freely
administered to the ischemic limbs in these studies via osmotic mini-pumps over a period of 7-10
days while the endogenous expression of FGF2 is likely under a large degree of gene regulation
and/or protein sequestration that is only lessened in response to specific stimuli. Third, the level
of endogenous protein released in response to the ischemic stress may not be comparable to the
pharmacologic (supra-physiological) levels of the exogenous LMW FGF2. Fourth, it is worth
noting that while recombinant LMW FGF2 is capable of inducing angiogenesis in vitro and in
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vivo, endogenous LMW FGF2 may not possess the potent “angiogenic” ability that has been
assigned to it. The field of recombinant protein production is well established and a plethora of
studies have been undertaken that demonstrate recombinant proteins behaving in a similar manner
as their endogenous “equivalents” by showing comparable receptor binding, activation and
downstream signaling in target cells. In spite of this, it is still conceivable that endogenous LMW
FGF2 does not share the angiogenic potential of its exogenous counterpart because of the presence
of in vivo mechanisms that the recombinant proteins are not subjected to. Some evidence for this
discrepancy may exist in the fact that recombinant growth factors including LMW FGF2 failed to
show efficacious benefit in the treatment of human ischemic disease. Promising results in
multitude of small and large animal models of peripheral and coronary artery diseases (PAD,
CAD) propelled the investigation of LMW FGF2 in several placebo-controlled clinical trials
(Laham, Chronos, et al., 2000; Lederman et al., 2002; Molin & Post, 2007). Thus far, the use of
recombinant LMW FGF2 protein for the treatment of CAD or PAD has not been successful (Y.
Cao, Hong, Schulten, & Post, 2005; van Royen, Piek, Schaper, & Fulton, 2009). While there may
be a number of reasons for the failure of these trials including the insufficiency of single growth
factor administration, the mode of and route of growth factor delivery, the presence of
comorbidities in advanced age patients and the selection of appropriate clinical endpoints (Y. Cao,
2009; Dragneva et al., 2013), it is not out of the realm of possibility that the recombinant proteins
employed in these studies are the true culprits. Lastly, the differences in species, surgical ischemia
models and data interpretation may also explain the conflicting results. This dissertation provides
novel evidence of the in vivo roles of endogenous LMW FGF2 isoforms in ischemia-induced angiogenesis.
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When there was no expression of FGF2 (in Fgf2-/- mice), similar increases in capillary
numbers were detected in response to ischemia (Figures 16 and18). These results are in agreement
with a similar study by Sullivan and colleagues where the absence of Fgf2 gene did not affect post-
ischemic capillary density (Sullivan et al., 2002). Normal injury-induced angiogenesis was also
observed in the eye when Fgf2-/- mice were subjected to ischemic retinopathy or choroidal injury
(Ozaki et al., 1998; Tobe et al., 1998). Postnatal cardiac capillary growth is also affected by loss
of the Fgf2 gene. Complete (Fgf2-/-) or partial (Fgf2+/-) disruption of FGF2 gene expression
impairs adult myocardial capillarogenesis (Amann et al., 2006). On the other hand, our laboratory
and others detected no alterations to baseline capillary development in Fgf2-/- cardiac or skeletal
muscles (House et al., 2003; Liao et al., 2009; Sullivan et al., 2002). The present study also did not discover any changes to capillary vessel development in non-ischemic skeletal muscle (Figures
16, 19). Compensation for the loss of Fgf2 by FGF1, the most closely related FGF was thought account for the absence of a phenotypic response to the genetic deletion of FGF2. However, when
Fgf2-/- mice were crossed with Fgf1 knockouts, the resultant double knockout offspring displayed similar phenotypes as the Fgf2-/- mice (Miller et al., 2000). By the same token, Fgf5/Fgf6,
Fgf6/Fgf7, and Fgf5/fgf6/Fgf7 double and triple mutants did not exhibit any defects beyond that observed in Fgf6-/- skeletal muscles that could account for the residual regeneration ability of Fgf6 mutant myotubes (Armand et al., 2005; Floss, Arnold, & Braun, 1997; Neuhaus et al., 2003). The advent of mouse models in which total Fgf2 expression is not merely disrupted but where specific expression of only one class of its protein isoforms are deleted allows for a more nuanced examination of the functions of FGF2.
In addition to its role in ischemia-induced capillary growth, the effect of expression of
HMW FGF2 on arteriole density was also examined (Figures 20, 22B). The arteriole density of
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ischemic HMW FGF2-expressing muscles was nearly two-fold greater than WT ischemic levels
(Figure 22B). This indicates that in the presence of shear and ischemic stress, the HMW isoforms
have a profound stimulatory effect on arteriogenesis and represents a previously unrecognized
function of endogenous HMW FGF2. While no other direct evidence of the role of HMW isoforms
on ischemia-induced revascularization exists, findings from Jiang and coworkers showed the effect
of exogenous HMW FGF2 administration in infarcted rat hearts (Jiang et al., 2007). Specifically,
recombinant 23kDa produced no changes to arteriole vessel density in the infarct area. The
differential outcomes provide additional proof to distinguish the properties of endogenously
produced FGF2 from the exogenous isoforms. Nonetheless, the possibility that the conflicting
results could be because of differences in ischemia models and species remains. The effect of
expression of only the LMW isoforms on arteriogenesis was somewhat intriguing in light of the
total lack of angiogenic response observed in our model. LMW FGF2 stimulated an increase
arteriole and venule density in ischemic muscle (Figure 22A). The degree of arterial growth was
identical to that observed in WT limbs where arteriole numbers were elevated almost four-fold in the presence of ischemia. These results are consistent with other hindlimb ischemia studies performed in mice (Scholz et al., 2002) and rats (J. Zhang et al., 2009) where LMW FGF2
treatment stimulated arteriolar growth. Porcine (Watanabe et al., 1998) and murine (Jiang et al.,
2007) hearts undergoing chronic MI also had increased arteriole density in response to LMW FGF2
administration. It is worth noting that while the LMW FGF2 was capable of promoting
arteriogenesis in ischemic limbs, the HMW isoforms produced a greater effect on collateral vessel
growth. Resultantly, the number of arteriolar vessels in the ischemic FGF2 HMW-only muscles
was nearly two-fold greater than in the LMW FGF2-only cohorts.
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As previously mentioned, ischemic limbs of Fgf2-/- mice had no deficits in angiogenesis
(capillary density). Arteriogenesis was also found to be normal in these mice (Figures 20, 22B).
These findings paralleled work by Sullivan et al. whose study also revealed similar arteriole
density in Fgf2-/- ischemic muscle as in WT (Sullivan et al., 2002). Conversely, Virag and co- workers have reported reduced density of smooth muscle actin containing-vessels in Fgf2-/- hearts after 4 weeks permanent coronary artery ligation (Virag et al., 2007). The discrepancies in these outcomes could be related differences in the site of ischemia induction or animal strain. Overall, these results might be surprising when they are viewed in the context of some of the earlier studies performed that identified FGF2 as a “potent angiogenic” molecule. Furthermore, the absence of vascular abnormalities/changes in Fgf2-/- mice at birth or in response to hindlimb ischemia led to questioning of the earlier in vitro work establishing FGF2 in this role. In fact, it suggested that the functions of FGF2 cannot always be gleamed from the use of Fgf2-/- where all of the isoforms are absent as this mode presumes that all the isoforms have similar functions. The availability of animal models in which total Fgf2 expression is not only deleted, but where selective expression of only HMW or LMW FGF2 is present will provide a clear picture of the functional roles of
FGF2. When only LMW FGF2 (in FGF2 HMW-only mice) or only HMW FGF2 (in FGF2 LMW- only mice) is deleted, there appears to be greater vascular physiology effect than in Fgf2-/- mice.
In the presence of only the HMW isoforms, FGF2 has a positive effect on revascularization and the opposite occurs in presence of only the LMW isoform. The results presented in this dissertation serve as evidence for the antithetical roles of the FGF2 isoforms in ischemia-induced angiogenesis or arteriogenesis.
In light of the previously addressed differences that are observed in the response to ischemia by mice of different background strains, the angiogenic and arteriogenic responses of the
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WT mice in this study were compared to other studies of hindlimb ischemia. This also ensured that all changes observed were due only to selective or total deletion of FGF2 expression and not based on differences or inconsistencies in the surgical ischemia method. In five studies utilizing the surgical hindlimb ischemia method in WT mice of similar strain (50% 129SV), the average capillary and arteriole density in ischemic muscles was increased by 1.4- and 1.8-fold respectively
(Chalothorn et al., 2005; Lizotte et al., 2013; Scholz et al., 2002; Sullivan et al., 2002; Tilan et al.,
2013). The changes in WT capillary and arteriole density observed in this dissertation were 1.3- and 4.0-fold, respectively. The source of the differences in arteriole density between our work and published results may lie in the subjective criteria each group uses to defined arterioles. Positive staining of a smooth muscle cell marker (α-smooth muscle actin) and morphology was used in the identification of arterioles/arteries as well as to distinguish them from venules/veins. This classification is not always performed by others and the arteriole density reported is actually a measure of all vessels that are positive for smooth muscle. Another technical detail that might confound comparisons with other published work is the choice of ischemic muscle bed chosen for histological assessment of vascular growth. The importance of distinctions between calf and thigh vessel growth type has been addressed previously. Overall, the level of angiogenesis and arteriogenesis present in the WT mice were consistent with similar hindlimb ischemia studies.
In the absence of ischemia, the level of vessel density in FGF2 LMW-only and FGF2
HMW-only limbs were similar as WT (as seen in sham-operated skeletal muscle). The lack of baseline differences was also present in hearts with specific FGF2 isoform deletions (Liao et al.,
2007, 2010). Similarly, mice with chronic overexpression of all FGF2 isoforms or only HMW
FGF2 had no differences in baseline cardiac vessel density (House et al., 2003; Liao et al., 2010;
Virag et al., 2007). However, the Kardami group detected an increase (20%) in the capillary
154 density of LMW FGF2-overexpressing hearts (Sheikh et al., 2001). On the whole, perturbations in
FGF2 expression alone does not appear to produce vascular growth defects. This dissertation along with a study by the Murry group where Fgf2 overexpression induced increased vessel density during chronic ischemia, has shown that the effect of FGF2 isoforms on vascular growth is dormant at baseline and manifests in response to chronic vascular injury (Virag et al., 2007). This suggests that a latent phenotype of the FGF2 isoforms exists and is only activated during tissue pathophysiology.
The central pathophysiology to manifest in human diseases of arterial occlusion is the loss or reduction of tissue perfusion. Accordingly, preclinical studies modeled after these human conditions are set up to identify efficacious treatments that will increase perfusion to ischemic regions. In the mouse hindlimb ischemia model, adaptive or pharmacologically-induced vascular growth/remodeling via angiogenesis, arteriogenesis or collateral growth can all contribute to the restoration or augmentation of muscle blood flow. To establish the role of the FGF2 isoforms in the recovery of limb flow, perfusion of ischemic skeletal muscle was determined using laser
Doppler perfusion imaging (LDPI). Serial perfusion of the sham and ischemic feet (paws) was measured as a surrogate for skeletal muscle blood flow (Figure 26). In FGF2 HMW-only mice, recovery of resting foot perfusion was unchanged from WT. No differences from WT in relative foot perfusion (ischemic-to-sham ratio) were detected at baseline (absence of ischemia), immediately after (day 1 of ischemia), at 2 weeks or at 6 weeks after ischemia surgery. In FGF2
LMW-only mice, relative perfusion was normal in the absence of ischemia. After ischemia surgery, the trend of perfusion recovery described for the HMW FGF2 feet was also present in
LMW FGF2-expressing limbs (Figure 26B). The apparent lack of effect of LMW FGF2 is in alignment with a study where rats receiving LMW FGF2 treatment had no changes in foot
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perfusion recovery after 6 weeks of ischemia (J. Zhang et al., 2009). Conversely, several studies
in myocardial (Laham, Rezaee, et al., 2000; Lazarous et al., 1995; Rajanayagam et al., 2000; Unger
et al., 1994) and hindlimb (Rakue et al., 1998; Scholz et al., 2002; H.-T. Yang et al., 1996) ischemia
models have reported improved regional blood flow after administration of recombinant LMW
FGF2.
The presence of species-to-species variations in blood flow recovery after myocardial or
peripheral arterial occlusion has been previously reported (Maxwell et al., 1987; Teunissen et al.,
2012; Waters et al., 2004). Similarly strain-to-strain variations exist within the same species and
have been detected in several mouse genetic strains (Scholz et al., 2002). To address changes in perfusion that could be strain dependent, the level of foot perfusion recovery in WT mice was compared to similar reports in 129SV WT mice. Perfusion ratios typically reached a plateau at ~28 days and the recovery of resting perfusion in the ischemic feet was as low as 50% (Landázuri,
Joseph, Guldberg, & Taylor, 2012; Rey et al., 2009; Tirziu et al., 2005) and as high as 90%
(Sullivan et al., 2002) of baseline levels. When averaged across several published studies, the recovery of resting perfusion in 129SV WT was ~60% (Bosch-Marce et al., 2007; Chalothorn et al., 2005; Heil et al., 2002; Helisch et al., 2006; Tilan et al., 2013). A similar recovery of pre- ischemia perfusion levels was present in the WT mice studied in this dissertation (Figure 26). An interesting observation was the similarity between WT, FGF2 LMW-only and FGF2 HMW-only mice in resting perfusion recovery of ischemic limbs despite their differential levels of angio- and/or arteriogenesis. The prevailing paradigm in the study of peripheral ischemia using experimental models suggests that increases or decreases in ischemic muscle vascular growth present should lead to improved or reduced limb perfusion, respectively (Couffinhal, Dufourcq,
Barandon, Leroux, & Duplàa, 2009; Limbourg et al., 2009; Silvestre et al., 2013; Waters et al.,
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2004). While this holds true for some studies, there are published reports where changes in limb
perfusion do not always follow enhanced vascularization (L. S. Brevetti et al., 2001; Chalothorn et al., 2005; Shireman & Quinones, 2005; J. Zhang et al., 2009). The apparent discrepancy between
muscle vascular density and perfusion could be attributed to several factors including the effect of
ischemia on blood flow control, regulation of vasomotor tone by FGF2 expression, and the
perfusion measurement methodology.
Chronic ischemia alone has been shown to negatively impact blood flow control. This
phenomenon is present in animal models as well human patients of arterial occlusive disease
(Dragneva et al., 2013). Intermittent claudication patients have normal muscle perfusion at rest but
can experience ischemic pain during walking or exercise (Annex, 2013; Brass, 2013; Muir, 2009).
The impaired flow control manifests as altered vascular tone and reactivity where the
repairing/remodeled vasculature exhibits reduced vasodilation and/or enhanced vasoconstriction
(Colleran, Li, Yang, Laughlin, & Terjung, 2010; Kelsall, Brown, & Hudlicka, 2001; LeBlanc et
al., 2012). The presence of endothelial and smooth muscle dysfunction in ischemic muscles has
been confirmed by decreased reactivity to acetylcholine or sodium nitroprusside (Cardinal et al.,
2011; Kelsall, Brown, Kent, Kloehn, & Hudlicka, 2004). Another possible confounding variable
to consider when interpreting the results of the perfusion measurements is the effect of FGF2 itself
on regulation of vasomotor tone. Administration of LMW FGF2 to isolated arterioles of cerebral,
(Kajita, Takayasu, Yoshida, Dietrich, & Dacey, 2001), coronary (Tiefenbacher & Chilian, 1997)
or skeletal muscle (H. M. Wu et al., 1996) origins and infarcted porcine hearts (Unger et al., 1994)
induced arterial vasodilation. This effect was dependent on the presence of an intact endothelium,
nitric oxide production (NO) and activation of ATP-sensitive potassium channels (Tiefenbacher
& Chilian, 1997). When the Fgf2-/- mice were first produced to characterize the role of the Fgf2
157 gene in vascular physiology, a significant phenotype present in this mice was decreased vascular tone (Zhou et al., 1998) and basal hypotension (Dono et al., 1998; Ortega et al., 1998; Zhou et al.,
1998). The vascular tone control exerted by the presence of Fgf2 (via LMW FGF2) could account for the lack of differences observed in the recovery of ischemic perfusion of FGF2 LMW-only mice compared to WT. This would suggest that increased muscle perfusion in these mice is not reflective of changes in vascularization effects but due to the positive vasodilatory effects of LMW
FGF2. Studies by Sullivan and colleagues in Fgf2-/- mice provide some evidence to support this theory. They observed that despite normal resting perfusion in Fgf2-/- ischemic limbs, the reactive hyperemic response to acute iliac artery occlusion (vasodilation followed by a transient blood flow increase) was diminished in Fgf2-/- ischemic limbs at 14 days of ischemia (Sullivan et al., 2002;
Sullivan, 2002). In light of this, the effect of the specific FGF2 isoforms on vasomotor tone at baseline and during the ischemia repair process would be very important. Potential differences in vasomotor tone control due to the presence of ischemia and/or altered FGF2 expression could be addressed by repeating the LDPI measurements after brief periods of acute ischemic (via balloon cuff inflation) or bouts of exercise to induce reactive or functional hyperemia, respectively. Both challenges sidestep the low resting blood flow of skeletal muscle and serve to increase the demand for O2 in the ischemic tissues and requires the maximal dilation of the ischemic vasculature (L. S.
Brevetti et al., 2001; Ziegler et al., 2010). They can test the reactivity and capacity of the FGF2
LMW or HMW-only repairing vasculature (both the endothelium and smooth muscle) to respond to increased metabolic demand. Treadmill exercise, electrical nerve stimulation and acute ischemia have been shown to expose deficits in regional blood flow that are not present in resting (non- exercised) ischemic skeletal muscle (L. S. Brevetti et al., 2001; Cardinal & Hoying, 2007; Helisch et al., 2006; Hudlicka, Brown, Egginton, & Dawson, 1994). Treadmill exercise testing as
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previously described was performed in a small cohort of mice in this dissertation (Figure 10).
While these experiments were performed to determine limb functional capacity and did not include perfusion measurements, it is noteworthy that in the absence of Fgf2 expression, ischemic limb
function is impaired.
There are several alternative strategies to the present method for determining blood flow
changes in ischemia models including arterial flow probes, X-ray angiography, magnetic
resonance imaging, and radio- or fluorescently-labelled microspheres (de Lussanet et al., 2007;
Lotfi et al., 2013; Madeddu et al., 2006; Waters et al., 2004). The use of microspheres is considered the gold standard for the evaluation of blood flow in hindlimb ischemia models (Lotfi et al., 2013;
Madeddu et al., 2006; Waters et al., 2004). The method utilizes labelled microspheres (usually
15µm) that are injected intra-arterially or into the left ventricle. Regional blood flow is directly
proportional to the number of microspores that becomes lodged in the tissue of interest (Deveci &
Egginton, 1999; Prinzen & Bassingthwaighte, 2000). This approach is highly invasive and
technically challenging in smaller sized animal models as it requires both arterial catheterization
and withdrawal of a reference blood sample at a fixed rate. This blood sampling can lead to
hypotension due to small blood volumes and limits the use of microspheres in longitudinal studies
(Lotfi et al., 2013). Laser Doppler Perfusion Imaging (LDPI) provides a measure of tissue perfusion based on the magnitude and frequency distribution of the Doppler shift in the laser light after it is scattered by moving erythrocytes (Wårdell et al., 1993). The reflected Doppler signal is linearly proportional to the underlying surface tissue perfusion (Leahy et al., 1999). Disadvantages of this method include the intrinsic depth of penetration of the scanning laser (1-2 mm) which limits the flow measured to superficial skin circulation only (Madeddu et al., 2006) and the use of
this cutaneous perfusion as a surrogate measure of blood flow. Another important technical
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limitation of LDPI is the region/portion of the mouse hindlimbs that are scanned and analyzed.
Animal fur and skin pigmentation of the upper hindlimb can interfere with LDPI signal and
eliminates the upper thigh area of the hindlimb as a suitable region for measurement (Hoefer et al.,
2006; Limbourg et al., 2009). Therefore, perfusion data reported in this dissertation and in other
studies with the mouse HLI model were quantified from the plantar sole (feet) of the ischemic and
sham legs (Figure 26). Another important caveat to consider when evaluating perfusion in rodents
using LDPI is the animals’ core body temperature. Animals should be monitored during scanning
to ensure that no fluctuation in core body temperature occurs (≤ 0.5oC). To that end, all animals
were placed on a warming pad and equipped with a rectal temperature probe to monitor body
temperature during the perfusion experiments. Core body temperature for the mice scanned in this
study was an average of 36.8oC.
Despite the outlined shortfalls of the laser Doppler technique for the assessment of skeletal
muscle perfusion, it is still widely accepted as an appropriate tool for testing the vascular/tissue
responsiveness to hindlimb ischemia. It has the advantage of being a non-invasive and repeatable technique. Furthermore, longitudinal experiments can be carried to measure perfusion recovery over a long period (weeks-months). It is extremely useful for identifying blood flow deficits in the ischemic feet relative to the non-ischemic contralateral foot immediately after the ischemia surgery
(1-24 hours) which confirms successful ligation of the femoral artery (Figure 26, Day 1 perfusion ratios) (Madeddu et al., 2006; Waters et al., 2004). However, this method may be less sensitive in detecting subtler changes in flow and this could account for the uncoupling of the changes in vascular density quantified by histology and muscle perfusion.
An alternative quantitative assessment of FGF2 isoform-induced revascularization was performed with the use of micro-computed tomography (micro-CT). This imaging modality
160
addresses the limitations inherent within the previously mentioned techniques for quantifying
vascular growth including the two-dimensional (2-D) nature and small sampling of both
histological and LDPI measurements, the subjectivity of histology and the superficial and resting
perfusion analyzed by the latter (Duvall, Taylor, Weiss, & Guldberg, 2004; Lotfi et al., 2013;
Madeddu et al., 2006; Ziegler et al., 2010). While these other methods are still more widely
utilized, micro-CT is advancing the field of microvascular imaging by providing a means for
obtaining high resolution, objective and volumetric analysis of vascular anatomy (A. S. Lin et al.,
2007; Zagorchev et al., 2010). Unlike bone which has high inherent radiopacity, micro-CT imaging
of soft tissues like blood vessels require the use of contrast agents to enhance the x-ray attenuation of the ischemic vascular network. Several preliminary studies were performed with the perfusion of non-ischemic hindlimbs WT with three different contrast agents; barium sulfate, lead oxide and lead chromate (Figures 5-7). Barium sulfate (suspended in 5% gelatin) did not perfuse vessels well and while lead oxide produced images that were not suitable for quantitative analysis due to the extremely high x-ray attenuation of the perfused vessels. Lead chromate suspended in a silicon polymer solution was selected based on its ease of vascular perfusion, viscosity, and homogeneity.
The hindlimbs were scanned 42 days after ischemia surgery to produce three-dimensional
(3-D) representations of the vasculature in sham and ischemic calf or thigh regions (Figure 12).
The definition of separate regions of interest (ROI) within each 3-D volume allows for evaluation
of angiogenesis and arteriogenesis in the upper limb defined as thigh and lower limb as calf,
respectively. For each ischemic calf or thigh ROI, vessels were analyzed for the relative (% change
from sham) vessel volume, number (density) and spacing. In FGF2 HMW-only mice, vascular volumes in the ischemic calves and thighs were greater than in WT cohorts (Figure 13A). This is in agreement with the results of the immunohistological labeling of vessels described previously
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where capillary and arteriole density was greatly increased in the presence of FGF2 HMW. The
volume of vessels in the FGF2 LMW-only calf or thigh also corroborated the 2-D histological
data, showing that the presence of only LMW FGF2 inhibited production of vessels in response to
ischemia. By the same token, changes in vessel density and vessel spacing due to ischemia were
both decreased (Figures 13B, C). The spacing parameter represents the vessel separation and is a
measure of the volume of the scanned limbs not occupied by vessels (Bouxsein et al., 2010a;
Duvall et al., 2004; A. S. Lin et al., 2007). This means that as the vascular volume increases, there
is a corresponding decrease in the amount of vessel separation.
In Fgf2-/- calf and thigh regions, only the changes in vascular volume were in agreement
with the trend observed in the histology results were the loss of Fgf2 expression did not alter
vascular growth. Vessel density and spacing in the Fgf2-/- limbs were both reduced relative to
WT. This highlights the advantage of the micro-CT method for uncovering alterations in vascular
growth that are beyond the ability of immunohistochemistry. The loss of vascular reactivity and
presence of endothelial dysfunction described previously for ischemic Fgf2-/- muscles could be the reason for these differences. The micro-CT technique is dependent on the utility of contrast agents to enhance the X-ray attenuation of hydrated soft tissues (Bouxsein et al., 2010b; A. S. Lin et al., 2007; Zagorchev et al., 2010). The contrast material is perfused at constant pressure into
vessels previously flushed with a cocktail of endothelium and smooth muscle-responsive vasodilators to produce maximal vasodilatation. The altered vascular tone and reactivity present in Fgf2-/- under hyperemic conditions would suggest that response to this cocktail were limited and produced the observed changes in the 3-D volumes (Cardinal et al., 2011; Sullivan, 2002).
The use of a contrast agent in vessels for micro-CT imaging of vessels highlights both a strength and drawback of this technique. Incomplete perfusion of vessels, artifacts from high or
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inconstant perfusion pressure or vascular leakage can lead to inhomogeneous attenuation and
erroneous quantitative analysis. An additional limitation is the post-mortem nature of the tissue preparation (Bouxsein et al., 2010a; Zagorchev et al., 2010). This prohibits its use for longitudinal studies and increases the cost and number of animals required for each study. The time point of 42 days after the ischemia surgery was chosen to allow for comparison with the histological counts of vascular density. While micro-CT is not as widely used as LDPI for the analysis of the mouse models of ischemic vasculature due to its cost and/or technical difficulty, there have been a number of reports where this methodology was successfully employed (Duvall et al., 2008, 2004; W. Li et al., 2006; Oses et al., 2009). These studies largely focused on the distribution of vessel thickness present in the ischemic hindlimbs. This evaluation provides additional information on trends in the development of the post-ischemic vasculature that may not be evident from the averaged assessments of vascular morphology (volume, number or spacing). To further characterize the observed changes in vascular volume response to specific FGF2 isoform expression, size distribution analysis of the vasculature was performed. Specific isoform deletion or genetic knockout of FGF2 alone did not affect the distribution of vessels in the hindlimb (Figure 14). In the presence of ischemia, expression of the HMW isoforms alone stimulated an increase in the development of small (36-200µm) and medium (200-400µm)-sized collateral vessels.
Contrastingly, the LMW FGF2-expressing limbs had no change in the level of small or medium collateral vessels after 42 days of ischemia. These results closely mirror the responses observed in the histological staining of vascular structures. WT muscles also had greater numbers of collateral vessels in the ischemic muscles but to a lesser degree than the FGF2 LMW-only limbs. Several hindlimb ischemia studies in WT mice have detected similar increases in the volumes of vessels between 50 and 200µm in diameter (Duvall et al., 2004; Landázuri et al., 2012; W. Li et al., 2006;
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Morrison et al., 2014). Likewise, enhancement of vessels ranging in size from 200 to 420µm has been reported by others (Duvall et al., 2004; Landázuri et al., 2012; Westvik et al., 2009).
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Figure 27: Schematic outlining the FGF2 isoform-mediated tissue response to hindlimb ischemia. The nuclear localized HMW FGF2 is protective against necrosis and promotes recovery of limb function. On the other hand, LMW FGF2 is located in the cytosolic and nuclear compartments and may be secreted during ischemia. LMW FGF2 induced arteriogenesis and had no effect on angiogenesis. HMW FGF2 isoforms stimulated a greater degree of arteriogenesis as well as angiogenesis.
Inhibit Activate
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Chapter 2: Potential mechanisms of the FGF2 isoforms in adaptive revascularization
during chronic ischemia
RESULTS II
Loss of HMW FGF2 leads to increased expression of Fgf2 mRNA in non-ischemic muscle.
The absence of HMW FGF2 expression in non-ischemic hearts results in increased LMW
FGF2 protein expression (Liao et al., 2010). It has yet to be determined if ablation of LMW FGF2 or HMW FGF2 affects overall Fgf2 gene and/or protein expression in skeletal muscle. To determine Fgf2 transcript levels, RNA was isolated from non-ischemic WT, Fgf2-/-, FGF2 LMW- only and FGF2 HMW-only thigh and calf muscles. First strand cDNA was amplified by quantitative real-time PCR (qRT-PCR) using primers for Fgf2 and an internal control gene, 18S.
No Fgf2 mRNA was detected in Fgf2-/- mice which confirmed the deletion of the FGF2 gene in skeletal muscle (Figure 28). Fgf2 mRNA was significantly increased by about three-fold in FGF2
LMW-only muscles relative to non-ischemic WTs (p<0.05). Similarly, FGF2 HMW-only mice had upregulated Fgf2 expression, nearly four times greater than WT (p<0.05). mRNA levels were not different between FGF2 LMW-only and FGF2 HMW-only muscles. Overall, deletion of the
LMW or HMW isoform resulted in elevated Fgf2 gene transcription. This would suggest that there is transcriptional regulation of Fgf2 by the presence of either class of FGF2 isoforms.
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Figure 28. Fgf2 mRNA expression in non-ischemic muscles. Quantitative RT-PCR analysis
for Fgf2 mRNA expression in non-ischemic skeletal muscle relative to WT. Fgf2-/-, FGF2
LMW-only or FGF2 HMW-only transcript expression levels are represented as fold- changes relative to WT non-ischemic muscles. Data are presented as mean ± SEM. n=4 for each mouse group. *p<0.05 vs. WT.
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Loss of HMW FGF2 does not affect LMW isoform expression in non-ischemic skeletal
muscle
Genome and proteome-wide studies have shown that mRNA levels in mammalian cells do
not always correlate well with protein abundance (Greenbaum, Colangelo, Williams, & Gerstein,
2003; Vogel & Marcotte, 2012). To determine if selective deletion of either the LMW or HMW
isoforms induced changes in skeletal muscle FGF2 expression at the protein level, Western
blotting was performed on non-ischemic skeletal muscles. As there is currently no specific
antibody for 18kDa, 21kDa or 22kDa FGF2, analysis for the LMW and HMW protein isoforms
was performed simultaneously (Figure 3). Quantitation of the Western blot bands were performed
by densitometry analysis and are presented in Figures 29 and 30. As expected, neither Fgf2-/- nor
FGF2 HMW-only muscles expressed the LMW isoform (Figure 29). There was expression of
18kDa FGF2 in WT and FGF2 LMW-only limbs. There was a trend towards increased expression
of the LMW isoform in the FGF2 LMW-only group relative to WT.
Absence of the LMW isoform leads to increased expression of HMW FGF2 in non-ischemic
muscle
Ablation of the 18kDa LMW FGF2 isoform was confirmed in non-ischemic Fgf2-/- and
FGF2 HMW-only skeletal muscles. WT tissues expressed similar amounts of the 21kDa or 22kDa isoforms. This pattern was also present in FGF2 HMW-only limbs (Figure 30). When compared to WT, expression of each HMW isoform was significantly increased in the FGF2 HMW-only muscles (p<0.05). HMW FGF2 isoform levels were three times higher when LMW is absent than in WT. This would suggest that the FGF2-dependent regulation that is present in transcription of
Fgf2 extends to isoform specific translation.
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FGF2 isoform expression (Non-ischemic skeletal muscle)
4 WT
Fgf2-/- 3 N.S FGF2 LMW-only
2
1
units) (Arbitrary
expression) protein FGF2 0 18kDa 21kDa 22kDa
-1
Figure 29. FGF2 protein isoform expression in non-ischemic skeletal muscles. Quantitative analysis of LMW (18kDa) and HMW (21kDa, 22kDa) FGF2 isoform expression in WT (white bar), Fgf2-/- (light gray bar), and FGF2 LMW-only (dark gray bar) non-ischemic muscles.
Data are presented as mean ± SEM. n=4 for each mouse group at each time point. N.S (Not
significant).
169
FGF2 isoform expression (non-ischemic muscle) 8 WT
7 * FGF2 HMW-only 6
5 * 4 3 Arbitrary units 2
FGF2 protein expression FGF2 protein 1
0 18kDa 21kDa 22kDa
Figure 30. FGF2 protein isoform expression in non-ischemic skeletal muscles. Quantitative
analysis of LMW (18kDa) and HMW (21kDa, 22kDa) FGF2 isoform expression in WT (white
bar) and FGF2 HMW-only (black bar) non-ischemic muscles. Data are presented as mean ±
SEM. n=4 for each mouse group at each time point. *p<0.05 vs. WT for each HMW isoform.
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Loss of HMW FGF2 or ischemia did not affect Fgf2 transcription
To assess the effect of hindlimb ischemia on the expression of Fgf2, calf and thigh skeletal muscles were isolated from sham-operated and ischemic limbs at different time points of ischemia.
RT-PCR was used to determine Fgf2 transcript levels at 3, 7 and 14 days after the ischemia surgery.
Fgf2 mRNA in Fgf2-/- and FGF2 LMW-only muscles was analyzed relative to WT expression
(Figure 31). As expected, no Fgf2 mRNA was detected in Fgf2-/- limbs. In FGF2 LMW-only
muscles, mRNA expression in ischemic limbs was unchanged compared to WT at day 3, elevated
at day 7 and returning to normal at day 14 of ischemia. However, ischemic transcript levels, at any
time point of ischemia, were not different from the sham cohorts.
Expression of only the HMW isoforms increases Fgf2 mRNA during ischemia
In the presence of only HMW FGF2, Fgf2 mRNA was significantly increased in response to ischemia. Fgf2 transcript levels were increased in ischemic FGF2 HMW-only limbs (compared to sham cohorts or WT). Fgf2 mRNA expression was elevated from non-ischemic levels at day 3 and day 7 of ischemia and returned to normal levels at 14 days (p<0.05, Figure 32). Overall, there appears to be a FGF2 protein (HMW)-dependent upregulation of Fgf2 expression that occurs in
the absence of LMW FGF2 and the presence of ischemia.
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Fgf2-/- SHAM Fgf2 mRNA Expression (ischemic muscle) Fgf2 -/- HLI FGF2 LMW-only SHAM 10 FGF2 LMW-only HLI WT
8
6
4 (Arbitrary Units)
2 mRNA Expression mormalized to WT to mormalized Expression mRNA 0 3d 7d 14d
Figure 31. Fgf2 mRNA expression in sham and ischemic skeletal muscles. Time course of
Fgf2 mRNA expression relative to WT. Quantitative RT-PCR analysis was performed on sham (SHAM) and ischemic (HLI) muscles at days 3, 7, and 14 of hindlimb ischemia. For each time point of ischemia, sham (open) or ischemic (filled) fold-changes in Fgf2-/- (square) or FGF2 LMW-only (circle) muscles are expressed as a ratio of the corresponding WT (solid line) transcript levels. Data are presented as mean ± SEM. n=4 for each mouse group at each time point.
172
Fgf2 mRNA Expression (ischemic muscle)
# *
# *
*
Figure 32. Fgf2 mRNA expression in sham and ischemic skeletal muscles. Time course of
Fgf2 mRNA expression relative to WT. Quantitative RT-PCR analysis was performed on
sham (SHAM) and ischemic (HLI) muscles at days 3, 7, and 14 of hindlimb ischemia. For
each time point of ischemia, sham (open) or ischemic (filled) fold-changes in FGF2 HMW- only (circle) are expressed as a ratio of the corresponding WT (solid line) transcript levels.
Data are presented as mean ± SEM. n=4 for each mouse group at each time point. # p<0.05 vs. SHAM cohort, *p<0.05 vs. WT HLI.
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Ablation of HMW FGF2 did not affect LMW isoform expression during ischemia
To elucidate the effect of hindlimb ischemia on the expression of FGF2 protein isoforms,
calf and thigh skeletal muscles were evaluated with Western immunoblotting at 3, 7 and 14 days
of ischemia. There was no expression of the LMW 18kDa isoform in Fgf2-/- muscles. In ischemic
WT tissues, the 18 kDa protein level was not different from sham expression at day 3, day 7 or day 14 (Figures 33A and 34A). A similar pattern was also observed in FGF2 LMW-only limbs.
This suggests the absence of FGF2 LMW-dependent regulation of FGF2 translation during
ischemia.
A. 18kDa isoform expression in ischemic muscle
4.5 4 3.5 3 2.5 2
Arbitrary Units Units Arbitrary 1.5 1 0.5 0 -0.5 3d 7d 14d
174
B. 21kDa isoform expression in ischemic muscle 3 2.5 2 1.5 1 0.5 Arbitrary Units 0 -0.5 -1 3d 7d 14d
C. 22kDa isoform expression in ischemic muscle
2
1.5
1
0.5
Arbitrary Units 0
-0.5
-1 3d 7d 14d
Figure 33. FGF2 protein isoform expression in sham and ischemic skeletal muscle.
Quantitative analysis of 18kDa (A), 21kDa (B) and 22kDa (C) FGF2 expression in WT,
Fgf2-/-, and FGF2 LMW-only sham (dotted line) and ischemic (solid line) muscles at days 3,
7, and 14 of hindlimb ischemia. Data are presented as mean ± SEM. n=4 for each mouse group at each time point.
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Loss of LMW FGF2 altered HMW FGF2 protein expression
The expression of the HMW isoforms in the absence of LMW FGF2 was also examined
during ischemic injury. The trend of 21kDa or 22kDa FGF2 expression in WT muscles mirrored
that of the 18kDa isoform with increased expression of either protein at day 3 and continued
elevation at day 14 (Figure 33-34B and C). In FGF2 HMW-only limbs, isoform expression at 3 days of ischemia had decreased relative to non-ischemic HMW expression (data not shown).
Expression had begun to rise by day 7, remained increased at day 14 to return to baseline (non- ischemic) levels. Despite the reduction in HMW FGF2, expression was still slightly increased relative to WT (for 21/22 kDa, Figure 34B and C). This could indicate that ischemia does not affect
FGF2 protein isoform expression but the absence of one class of FGF2 protein (LMW) is compensated for by increased expression of HMW FGF2 at baseline.
A. 18kDa isoform WT SHAM
10 WT HLI
8
6
4 Arbitrary Units 2
0 3d 7d 14d
176
WT SHAM
21kDa isoform WT HLI B. FGF2 HMW-only SHAM 7 FGF2 HMW-only HLI 6 5
4 * 3
Arbitrary Units 2 1 0 3d 7d 14d C. 22kDa isoform
4
3 *
2
Arbitrary Units 1
0 3d 7d 14d
Figure 34. FGF2 protein isoform expression in sham and ischemic skeletal muscle.
Quantitative analysis of 18kDa (A) 21kDa (B) and 22kDa (C) FGF2 expression in WT and
FGF2 HMW-only sham (dotted line) and ischemic (solid line) muscles at days 3, 7, and 14 of
hindlimb ischemia. Data are presented as mean ± SEM. n=4 for each mouse group at each
time point. *p<0.05 vs. WT HLI.
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Role of FGF2 protein isoforms in FGFR expression
In response to ischemia-reperfusion injury in the heart, the LMW isoform of FGF2 protects against cardiac dysfunction via activation of the FGFR1 receptor tyrosine kinase (Liao et al.,
2010). Additionally, overexpression of HMW FGF2 depresses basal FGFR1 activation (Liao et al., 2010). Yet, it is unknown if ablation of LMW FGF2 or HMW FGF2 alters the expression profile of FGF receptors in skeletal muscle basally or under stress such as ischemia. To determine the gene expression levels of the murine FGF receptors, RNA was isolated from non-ischemic and ischemic WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only limbs. First strand cDNA was amplified by quantitative real-time PCR (qRT-PCR) using primers for the seven FGF receptors,
Fgfr1b, -1c, -2b, -2c, -3b, -3c, and -4 and an internal control gene (18S).
HMW FGF2 expression influences Fgfr1 gene expression at baseline and during ischemia
No Fgfr2b or Fgfr2c mRNA was detected in skeletal muscle. Fgfr1b and Fgfr1c mRNA expression were both unchanged in Fgf2-/- or FGF2 LMW-only mice muscles relative to non- ischemic WTs (Figure 35A). In FGF2 HMW-only limbs, Fgfr1b transcript levels were significantly decreased (p<0.05 vs. WT) while Fgfr1c expression was unchanged.
To characterize the effect of FGF2 isoform and ischemia on Fgfr1 expression, qRT-PCR was used to determine Fgfr1 mRNA expression at 1, 3, 7 and 14 days after the induction of ischemia. Transcript levels are presented as a ratio of sham muscles to address any changes related to the hindlimb ischemia surgery. In all groups, Fgfr1b expression increased with time, was highest after 7 days and at a decline by 14 days of ischemic (Figure 35B). At 3 days, Fgfr1b and Fgfr1c had a trend towards or was significantly increased in Fgf2-/- and FGF2 HMW-only muscles
178 relative to WT (p<0.05 vs. WT; Figures 35B and C). Fgfr1b and Fgfr1c mRNA had a similar expression profile in FGF2 LMW-only tissues relative to WT.
179
Fgfr1 expression in non-ischemic muscle A. 2.5 WT Fgf2-/- 2 FGF2 LMW-only FGF2 HMW-only
1.5
1 (Arbitrary Units)
# $‡ 0.5 mRNA Expression mormalized WT to mRNA
0 Fgfr1b Fgfr1c
B. Fgfr1b expression (ischemic muscle) 5 WT Fgf2-/- 4 FGF2 LMW-only FGF2 HMW-only
3
2 (Arbitrary units)
1 mRNA Expression relative to sham to relative Expression mRNA
0 1 3 7 14 Days after ischemia surgery 180
Fgfr1c expression (ischemic muscle) C. 5 WT Fgf2-/- FGF2 LMW-only 4 FGF2 HMW-only
3 # ‡
2 # ‡ (arbitrary units) (arbitrary
1
sham to relative Expression mRNA
0 1 3 7 14 Days after ischemia surgery
Figure 35. Fgfr1 mRNA expression in non-ischemic, sham and ischemic skeletal muscles. (A)
Quantitative RT-PCR analysis for Fgfr1b and 1c gene expression in non-ischemic skeletal
muscle relative to WT (white bar). Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar)
or FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes
relative to WT non-ischemic muscles. Time course of Fgfr1b (B) and Fgfr1c (C) mRNA
expression. For each timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA
expression in ischemic muscles are expressed relative to corresponding sham levels in WT
(blue), Fgf2-/- (red), FGF2 LMW-only (green), or FGF2 HMW-only (black) limbs. Data are
presented as mean ± SEM. n=4 for each mouse group at each timepoint. #p<0.05 vs. WT,
$p<0.05 vs. Fgf2-/-, ‡p<0.05 vs. FGF2 LMW-only, †p<0.05 vs. FGF2 HMW-only.
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Alterations in FGF2 protein expression did not affect Fgfr3 transcription
Fgfr3 expression in non-ischemic WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only
skeletal muscles was also examined. Fgfr3b and Fgfr3c mRNA expression were both unchanged
in Fgf2-/- mice relative to non-ischemic WTs (Figure 36A). FGF2 LMW-only and FGF2 HMW-
only muscles also had similar levels of Fgfr3b or Fgfr3c mRNA as WTs.
To describe the effect of specific FGF2 isoform expression and hindlimb ischemia on Fgfr3
transcript levels, sham and ischemic muscles were isolated and analyzed for Fgfr3b and Fgfr3c
mRNA expression. In WT, FGF2 LMW-only, and FGF2 HMW-only limbs, no changes in expression of either transcript were detected at 1, 3, 7 or 14 days of ischemia (Figures 36B and C).
While the Fgf2-/- tissues had increased Fgfr3b expression at 7 days compared to WT, this change was not significant (Figure 36B). In all groups, ischemic Fgfr3c expression increased with time, was highest at 7 days and returned to normal at 14 days (Figure 36C).
182
Fgfr3 expression in non-ischemic muscle
A. 3 WT Fgf2-/- 2.5 FGF2 LMW-only FGF2 HMW-only 2
1.5 (Arbitrary Units)
1 mRNA Expression mormalized WT to mRNA 0.5
0 Fgfr3b Fgfr3c
B. Fgfr3b expression (ischemic muscle) 6 WT Fgf2-/- 5 FGF2 LMW-only FGF2 HMW-only 4
3
2 (arbitrary units) (arbitrary
1 mRNA Expression relative sham to relative Expression mRNA
0 1 3 7 14 183 Days after ischemia surgery Fgfr3c expression (ischemic muscle) C. 7 WT Fgf2-/- 6 FGF2 LMW-only
5 FGF2 HMW-only
4
3 (arbitrary units) (arbitrary 2
mRNA Expression relative sham to relative Expression mRNA 1
0 1 3 7 14 Days after ischemia surgery
Figure 36. Fgfr3 mRNA expression in non-ischemic, sham and ischemic skeletal muscles. (A)
Quantitative RT-PCR analysis for Fgfr3b and 3c gene expression in non-ischemic skeletal
muscle relative to WT (white bar). Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar)
or FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes
relative to WT non-ischemic muscles. Time course of Fgfr3b (B) and Fgfr3c (C) mRNA
expression. For each timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA
expression in ischemic muscles are expressed relative to corresponding sham expression
levels in WT (blue), Fgf2-/- (red), FGF2 LMW-only (green), or FGF2 HMW-only (black)
limbs. Data are presented as mean ± SEM. n=4 for each mouse group at each timepoint.
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Expression of only LMW FGF2 affects Fgfr4 expression during ischemia
Fgfr4 expression in non-ischemic WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only skeletal muscles was also examined. For all mouse groups, Fgfr4 mRNA expression was not different from WT (Figure 37A).
To determine the role of FGF2 isoform expression and ischemia on Fgfr4 gene expression, transcript levels were measured at 1, 3, 7 or 14 days of ischemia. FGF2 LMW-only muscles had increased levels at 7 days relative to WTs (p < 0.05, Figure 37B) and returned to normal expression at 14 days. Fgf2-/- and FGF2 HMW-only also trended towards an elevated Fgfr4 expression at 7 days. Both groups had an Fgfr4 expression profile that increased slightly with time, was highest at 7 days and returned to normal at 14 days (Figure 37B).
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Fgfr4 expression (non-ischemic muscle) A. 2.5 WT Fgf2-/- 2 FGF2 LMW-only FGF2 HMW-only
1.5
1 (arbitrary units) (arbitrary
# ‡† Relative mRNA expression expression mRNA Relative 0.5
0
B. Fgfr4 expression (ischemic muscle) 6 WT Fgf2-/- 5 FGF2 LMW-only FGF2 HMW-only 4 #
3 †
* 2 (arbitrary units) (arbitrary
1 mRNA Expression relative sham to relative Expression mRNA
0 1 3 7 14 186 Days after ischemia surgery Figure 37. Fgfr4 mRNA expression in non-ischemic, sham and ischemic skeletal muscles. (A)
Quantitative RT-PCR analysis for Fgfr4 gene expression in non-ischemic skeletal muscle relative to WT (white bar). Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar) or
FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes relative to WT non-ischemic muscles. (B) Time course of Fgfr4 mRNA expression. For each timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA expression in ischemic muscles are expressed relative to corresponding sham levels in WT (blue), Fgf2-/- (red),
FGF2 LMW-only (green), or FGF2 HMW-only (black) limbs. Data are presented as mean ±
SEM. n=4 for each mouse group at each timepoint. *p<0.05 vs. Day 1 cohort, #p<0.05 vs.
WT, †p<0.05 vs. FGF2 HMW-only.
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Basal FGFR activation is not affected by selective FGF2 isoform expression
The FGFRs are typical receptor tyrosine kinases whose activation in response to ligand binding and receptor dimerization is driven by a sequence of tyrosine autophosphorylation and results in intracellular signaling and response (Eswarakumar et al., 2005). To determine whether the trends in Fgfr1, Fgfr3 or Fgfr4 gene expression observed in response to selective deletion of
LMW or HMW FGF2 affected FGFR expression and/or activation, Western blotting was performed on non-ischemic skeletal muscles, sham and ischemic muscles.
As there are currently no specific antibodies for the “b” or “c” forms of the FGFRs, immunoblotting results for FGFR1 actually represents activation and expression of FGFR1b and
FGFR1c protein translation products (Figures 38-41). The level of activation of FGFR1, 3 or 4 was expressed as the ratio of the amount of each phosphorylated receptor to total level of the receptor. In line with the qRT-PCR analysis, FGFR2 expression or activation was not observed in non-ischemic skeletal muscles. Neither deletion of the Fgf2 gene (Fgf2-/-) nor selective ablation of FGF2 isoforms (FGF2 LMW-only or FGF2 HMW-only) affected the level of FGFR1 activation
(Figure 38A). Similarly, FGFR3 and FGFR4 activation were unchanged with any alterations of
FGF2 expression.
Total protein expression of the FGFR1, FGFR3 and FGFR4 in non-ischemic skeletal muscles was not different between the mouse groups (Figure 38B). Overall, manipulation of FGF2 gene or protein isoform expression had no effect on FGF receptor protein or activation under baseline conditions.
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FGFR Activation
A. WT 3 Fgf2-/- FGF2 LMW-only FGF2 HMW-only 2 FGFR/total FGFR - 1 phospho
0 FGFR1 FGFR3 FGFR4
30 B.
25
20
15
10 (Arbitrary Unit)
Total FGFR expression FGFR expression Total 5
0 FGFR1 FGFR3 FGFR4
Figure 38. FGFR protein expression and activation (phosphorylation) in non-ischemic
skeletal muscle. Quantitative analysis of FGFR1, FGFR3 and FGFR4 activation (A) and total
expression (B) in WT (white bar), Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar),
and FGF2 HMW-only (black bars) non-ischemic muscles. Data are presented as mean ±
SEM. n=4 for each mouse group.
189
FGFR1 activation is elevated during ischemic injury in the absence of all FGF2 expression
LMW FGF2 protects against cardiac dysfunction following acute ischemia-reperfusion
injury. This response is non-mitogenic and dependent on activation of the FGF receptor, FGFR1
(Liao et al., 2010). To determine if the vascular response to endogenous LMW or HMW FGF2
expression during chronic ischemia was associated with changes in FGFR activation and/or
expression, WT, Fgf2-/-, FGF2 LMW-only or FGF2 HMW-only mice were subjected to hindlimb
ischemia for 3 or 7 days. Skeletal muscles were isolated from ischemic limbs (HLI) as well as the
sham-operated, contralateral limbs (SHAM). To address variations in protein expression related only to the ischemia surgery procedure, receptor activation and expression in ischemic limbs were presented as a ratio of the corresponding sham limb expression.
In the presence of only HMW FGF2, activation (phosphorylation) of FGFR1 was unchanged from WT after 3 days of ischemia (Figure 39A). Expression (total protein) of the
receptor was also not different at this timepoint (Figure 39B). FGF2 LMW-only or Fgf2-/- muscles also had a similar pattern of FGFR1 activation and expression.
A higher degree of receptor phosphorylation was observed in Fgf2-/- muscles at day 7 of ischemia (p<0.05 vs. WT, Figure 39A). Conversely, expression of only the LMW or only the
HMW isoforms of FGF2 did not affect FGFR activation after a week of hindlimb ischemia. FGFR1 protein expression was also similar to WT in isoform-specific knock out skeletal muscles.
190
FGFR1 activation (ischemic muscle)
A. 4 WT Fgf2-/- # ‡ † FGF2 LMW-only 3 FGF2 HMW-only SHAM/totalSHAM) - 2
1 * FGFR activation relative to sham HLI/total HLI)/(p HLI/total -
(p 0 3d 7d
FGFR1 expression (ischemic muscle) B. 2
1.5
1
(total HLI/total SHAM) SHAM) (total HLI/total 0.5 FGFR expression relative to sham relative expression FGFR
0 3d 7d Figure 39. FGFR1 activation (phosphorylation) and expression at 3 or 7 days of ischemia.
Phosphorylation (ratio vs. total protein) (A) and expression (B) in ischemic WT, Fgf2-/-,
FGF2 LMW-only, and FGF2 HMW-only muscles at relative to sham tissues. Data are presented as mean ± SEM. n=4 for each mouse group. *p<0.05 vs. Day 7 cohort, #p<0.05 vs.
WT, ‡p<0.05 vs. FGF2 LMW-only, †p<0.05 vs. FGF2 HMW-only.
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Ablation of FGF2 and hindlimb ischemia does not affect FGFR3 activation
The degree of FGFR3 activation and protein expression in WT, Fgf2-/-, FGF2 LMW-only
and FGF2 HMW-only skeletal muscles was also examined at 3 or 7 days of ischemia. While the
complete loss of FGF2 expression (Fgf2-/-) produced a trend towards decreased phosphorylation
of FGFR3 at day 3, this change was not significant (Figure 40A). Total protein expression of
FGFR3 was increased in the Fgf2-/- muscles (p<0.05, Figure 40B). In the presence of only LMW
or HMW FGF2, the level of FGFR3 activation at 3 days of ischemia was unchanged. Similarly,
protein expression of the receptor was not different from WT in the isoform knockout mice.
Activation of FGFR3 at 7 days of ischemia in Fgf2-/-, FGF2 LMW-only and FGF2 HMW- only limbs was not significantly different from WT (Figure 40A). Likewise, total protein expression of the receptor was similar among the mouse groups (Figure 40B).
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FGFR3 activation (ischemic muscle)
3 WT Fgf2-/- FGF2 LMW-only FGF2 HMW-only 2 SHAM/totalSHAM) -
1 FGFR activation relative to sham HLI/total HLI)/(p HLI/total -
(p 0 3d 7d
FGFR3 expression (ischemic muscle) 4
# ‡ † 3
2
(total HLI/total SHAM) 1 FGFR expression relative shamto relative expression FGFR
0 3d 7d
Figure 40. FGFR3 activation (phosphorylation) and expression at 3 or 7 days of ischemia.
Phosphorylation (ratio vs. total protein) (A) and total protein expression (B) in ischemic WT,
Fgf2-/-, FGF2 LMW-only, and FGF2 HMW-only muscles at relative to sham tissues. Data are presented as mean ± SEM. n=4 for each mouse group. #p<0.05 vs. WT, ‡p<0.05 vs. FGF2
LMW-only, †p<0.05 vs. FGF2 HMW-only.
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Activation of FGFR4 does not influence FGF2-mediated response to chronic ischemia
To elucidate the effect of hindlimb ischemia and selective FGF2 expression on the
activation of FGFR4, expression of all and only phosphorylated forms of the receptor were
evaluated with Western immunoblotting at 3 and 7 days of ischemia (Figure 41A). In FGF2 LMW-
only or FGF2 HMW-only muscles, FGFR4 activation was not changed from WT at 3 days of
ischemia. FGFR4 protein expression was not different from WT expression (Figure 41B). Fgf2-/-
limbs followed a similar pattern of FGFR4 activation and expression.
By 7 days of ischemia, activation of FGFR4 in Fgf2-/-, FGF2 LMW-only and FGF2 HMW-
only limbs was unchanged relative to WT tissues (Figure 41A). Correspondingly, total protein
expression of the receptor was similar among the mouse groups (Figure 41B).
The absence of changes in FGFR activation and expression in these results confirm
previous data from our laboratory where FGFR1 protein levels were unchanged in non-ischemic
FGF2 HMWKO hearts but contradict data from mice overexpressing HMW FGF2 where basal
FGFR1 activation was depressed (Liao et al., 2010; Manning et al., 2013). Furthermore, FGFR1 activation was similarly activated by LMW FGF2 or HMW FGF2 in skeletal muscles but is upregulated in Fgf2-/- limbs. The elevated receptor activation in Fgf2-/- muscles is most likely an attempt to increased FGF2 signaling to compensate for the loss of the ligand expression. These findings suggest that there is an FGF2-mediated regulation of FGFR in skeletal muscle that is independent of the class of FGF2 protein isoform.
194
FGFR4 activation (ischemic muscle)
A. WT 3 Fgf2-/- FGF2 LMW-only FGF2 HMW-only
2 SHAM/totalSHAM) -
1 FGFR activation relative to sham HLI/total HLI)/(p HLI/total - (p 0 3d 7d
B. FGFR4 expression (ischemic muscle) 2
1 (total HLI/total SHAM) FGFR expression relative shamto relative expression FGFR
0 3d 7d Figure 41. FGFR4 activation (phosphorylation) and expression at 3 and 7 days of ischemia.
Phosphorylation (ratio vs. total protein) (A) and expression (B) in ischemic WT, Fgf2-/-,
FGF2 LMW-only, and FGF2 HMW-only muscles at relative to sham tissues. Data are
presented as mean ± SEM. n=4 for each mouse group.
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Effect of Fgf2 gene or protein isoform deletion on angiogenesis-related protein expression
FGF2 signaling plays a role in the regulation of normal and adaptive vascular development
(Annex & Simons, 2005; Potente, Gerhardt, & Carmeliet, 2011; Presta et al., 2005). There is also recent evidence to suggest that, in addition to its own direct effect on vascular growth, FGF2 has an indirect role in neo-/re-vascularization by promoting the expression of other angiogenic growth factors, cytokines adhesion molecules, and chemokines (Murakami & Simons, 2008; Presta et al.,
2009; X. Yang et al., 2015). Furthermore, the different autocrine, paracrine and intracrine signaling pathways mediated by the FGF2 isoforms suggests that in response to ischemic injury, HMW
FGF2 might induce the expression of a repertoire of angiogenesis-related proteins distinct from that of LMW FGF2 and vice-versa.
To identify the expression profile of angiogenic factors, cytokines and chemokines regulated by the individual FGF2 isoforms in ischemic-induced revascularization of the mouse hindlimb, the level of several angiogenesis-related proteins was determined using a proteome antibody array. The array containing antibodies against 53 different proteins spotted in duplicates was hybridized with equal amounts of protein isolated from sham and ischemic skeletal muscles after 7 days of ischemia. This was chosen because the ischemic-induced upregulation of FGF2 isoform expression is maximal 7 days after surgery as previously described in Chapter 1 of this dissertation. The array dots were quantified by densitometric analysis and the ischemic expression of each protein (after compensation for differences in background intensity) was normalized to its expression in the contralateral (sham) limb as presented in Table 13.
A majority of the dots (antibodies) showed similar intensities between WT, Fgf2-/-, FGF2
LMW-only and FGF2 HMW-only ischemic muscles but changes were observed in 7 different antibodies. Among these was MMP-9, a proteolytic enzyme involved in the remodeling of the
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basement membrane and extracellular matrix (ECM) during neovascularization to allow for
outward migration of ECs and vSMCs (Q. Chen et al., 2013; P.-H. Huang et al., 2009). In the
absence of any FGF2 expression (Fgf2-/-), MMP-9 expression was significantly increased compared to WT after 7 days of ischemia (p<0.05). Serpin E1 (PAI-1), a serine protease inhibitor, was also elevated in Fgf2-/- ischemic muscles (p<0.05 vs. WT). PAI-1 is the principal inhibitor of the urokinase-type plasminogen activator (uPA) system. u-PA catalyzes the conversion of plasminogen to plasmin, another serine protease whose proteolytic activity during vascular growth mirrors that of MMP9 (Medcalf, 2007; Van De Craen, Declerck, & Gils, 2012). Plasmin can also convert inactive MMPs (pro-MMPs) zymogen into their active forms (Q. Chen et al., 2013).
When only HMW FGF2 is expressed, insulin-like growth factor binding protein-3 (IGFBP-
3) protein expression was significantly increased compared to WT (p<0.05). Classically, IGFBPs
are members of the endocrine IGF signaling pathway, where they serve as carriers of IGF and
regulate the turn-over of circulating IGFs (Baxter, 2013; Yamada & Lee, 2009). However, IGF- independent activities on cell growth have also been attributed to these proteins. Expression of
IGFBP-3, in particular has an inhibitory effect on cell proliferation and tumor growth (Oh, Gucev,
Ng, Müller, & Rosenfeld, 1995; Yamada & Lee, 2009). IGFBP-3 can also play a more ambiguous role in cell proliferation. When endogenous IGFBP-3 expression is reduced, EGF-induced proliferation of breast epithelial or cancer cells is also decreased (Baxter, 2015; Butt et al., 2004;
McIntosh et al., 2010). EGF, epidermal growth factor is the prototypic member of the EGFR family characterized by promotion of epidermal cell regeneration and healing of acute or chronic dermal wounds. (Bhora et al., 1995; Peplow & Chatterjee, 2013; Seeger & Paller, 2015). EGF-stimulated wound healing involves the proliferation and migration of keratinocytes and fibroblasts (Haase,
197
Evans, Pofahl, & Watt, 2003; Seeger & Paller, 2015). In Fgf2-/- muscles, EGF was significantly
decreased relative to FGF2 LMW-only and trended towards lower expression compared to WT.
FGF7, another member of the FGF family is associated with wound healing. It is
exclusively expressed by mesenchymal cells and regulates wound healing via proliferation and
migration of epithelial cells (keratinocytes) and activation of the alternatively spliced variant of
the FGF receptor, FGFR2IIIb (Maretzky et al., 2011; Niu et al., 2007; Tsuboi et al., 1993; Yen,
Thao, & Thuoc, 2014). FGF7 had increased expression in FGF2 HMW-only muscles that trended towards significance.
Another IGF binding protein, IGFBP-10 (CCN1) was also analyzed using the antibody array. There was a trend towards increased of expression CCN1 in FGF2 HMW-only ischemic
muscles. IGFBP-10 (CCN1) is not a classical member of the IGF family but is categorized as an
IGFBP based on shared structural homology with IGFBPs1-7 (Malik, Liszewska, & Jaworski,
2015). CCN1 is a secreted matricellular protein of the CCN family that exist at the border of cells and the ECM (Jun & Lau, 2011). While not involved in structural integrity of the ECM, they modulate surface receptors (integrins and HSPGs) to promote pro-angiogenic activity, vascular cell adhesion, migration and proliferation (C.-C. Chen & Lau, 2009; Fataccioli et al., 2002; C. G.
Lin et al., 2003). CCNs can also function as transcription factors under hypoxia conditions to regulate expression of other angiogenic factors including VEGFs and FGFs (Kubota & Takigawa,
2007; Lau, 2011; Malik et al., 2015).
An antibody to a protein (CX3CL1/fractalkine) with both angiogenic and inflammatory roles was also detected on the array. CX3CL1, the sole chemokine (chemotactic cytokine) in the
CX3C subfamily of chemokines is expressed primarily by vascular cells and macrophages and signals through its receptor, CX3CR1 to induce EC proliferation, migration and tube formation
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(Imai et al., 1997; S.-J. Lee et al., 2006; Shireman, 2007). In addition, the CX3CL1/CX3CR1
interaction mediates the chemotaxis and survival of inflammatory leukocytes such as monocytes/
macrophages (Owen & Mohamadzadeh, 2013; White & Greaves, 2009). There was a trend
towards increased expression of CX3CL1 in FGF2 HMW-only ischemic muscles (p=0.06 vs WT).
Overall, the proteome profile in ischemic skeletal muscle suggests that complete loss of
FGF2 expression (in Fgf2-/- limbs) stimulates increased expression of proteins involved in ECM
remodeling and activation. These ECM remodelers might serve to promote the release and
trafficking of other growth factors that are typically bound to the structural proteins that make up
the ECM including proteoglycans and fibronectin.
Functional Group Fold change in pixel intensity
Growth Factors WT Fgf2-/- FGF2 LMW-only FGF2 HMW-only FGF1 0.94 ± 0.13 0.88 ± 0.09 0.85 ± 0.16 1.09 ± 0.24 FGF2 0.84 ± 0.07 0.89 ± 0.09 0.93 ± 0.16 0.87 ± 0.28 FGF7 0.87 ± 0.06 0.91 ± 0.04 0.99 ± 0.11 1.15 ± 0.04 VEGF-A 1.08 ± 0.12 0.83 ± 0.13 0.99 ± 0.08 1.33 ± 0.41 VEGF-B 0.92 ± 0.05 1.13 ± 0.11 1.02 ± 0.12 1.29 ± 0.40 PIGF-2 0.85 ± 0.08 1.05 ± 0.08 1.10 ± 0.13 0.95 ± 0.13 PDGF-AA 0.92 ± 0.09 0.90 ± 0.02 1.04 ± 0.17 1.17 ± 0.09 PDGF-BB 0.91 ± 0.06 0.98 ± 0.14 1.11 ± 0.18 1.11 ± 0.20 HGF 0.99 ± 0.19 1.12 ± 0.06 1.29 ± 0.24 1.06 ± 0.20 EGF 0.87 ± 0.03 0.80 ± 0.07 1.07 ± 0.03 0.98 ± 0.08 HB-EGF 0.84 ± 0.03 0.99 ± 0.10 1.05 ± 0.09 1.05 ± 0.22
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PD-ECGF 0.90 ± 0.19 1.17 ± 0.06 1.00 ± 0.07 1.08 ± 0.01 Angiogenesis Angiogenin 1.07 ± 0.12 1.04 ± 0.02 1.29 ± 0.23 1.03 ± 0.17 Angiopoietin-1 0.89 ± 0.16 1.13 ± 0.15 1.38 ± 0.33 1.01 ± 0.39 Angiopoietin-3 0.93 ± 0.07 0.87 ± 0.05 1.11 ± 0.16 0.89 ± 0.16 CCN1/IGFBP-10 0.91 ± 0.12 1.98 ± 0.30 1.61 ± 0.79 2.05 ± 0.42# CCN3/IGFBP-9 0.82 ± 0.27 3.55 ± 0.63 1.46 ± 0.48 2.38 ± 0.71 IGFBP-2 0.86 ± 0.17 2.03 ± 0.25 1.75 ± 0.66 1.03 ± 0.16 Endothelin-1 0.86 ± 0.03 1.03 ± 0.05 1.10 ± 0.17 1.11 ± 0.32 Leptin/OB 0.90 ± 0.04 0.86 ± 0.05 1.09 ± 0.19 1.01 ± 0.22 Osteopontin 1.18 ± 0.45 1.69 ± 0.34 1.15 ± 0.35 1.13 ± 0.59 Prolactin 0.94 ± 0.12 1.01 ± 0.06 0.99 ± 0.11 1.26 ± 0.41 Proliferin 0.75 ± 0.10 1.05 ± 0.02 0.90 ± 0.05 1.38 ± 0.44 Angiostatic DLL4 0.89 ± 0.10 0.99 ± 0.05 1.17 ± 0.16 1.03 ± 0.07 DPPIV/CD26 1.06 ± 0.34 0.98 ± 0.18 1.09 ± 0.20 1.11 ± 0.31 Endostatin 0.95 ± 0.08 1.02 ± 0.12 0.96 ± 0.15 1.09 ± 0.13 IGFBP-1 0.84 ± 0.10 1.02 ± 0.10 1.11 ± 0.07 0.86 ± 0.22 IGFBP-3 1.19 ± 0.39 3.25 ± 0.34 2.29 ± 0.55 3.9 ± 0.86 # TIMP-1 0.76 ± 0.15 2.97 ± 1.31 2.03 ± 0.92 2.22 ± 0.96 TIMP-4 0.86 ± 0.07 0.91 ± 0.04 0.92 ± 0.13 1.01 ± 0.19 Pentraxin-3 0.80 ± 0.32 1.97 ± 0.18 1.80 ± 0.77 2.84 ± 1.31 Serpin F1/PEDF 1.02 ± 0.19 1.35 ± 0.17 1.29 ± 0.27 1.44 ± 0.26 Thrombospondin-2 0.78 ± 0.17 1.63 ± 0.21 1.37 ± 0.35 1.5 ± 0.41 ECM Remodeling MMP-3 1.10 ± 0.28 2.43 ± 0.38 2.29 ± 0.84 1.74 ± 0.56 MMP-8 0.84 ± 0.05 1.19 ± 0.03 1.07 ± 0.14 0.92 ± 0.21 MMP-9 0.69 ± 0.03 4.471 ± 0.57 #‡† 0.88 ± 0.25 1.92 ± 0.74 ADAMTS1 0.92 ± 0.07 0.93 ± 0.05 1.12 ± 0.23 1.17 ± 0.06 Serpin E1/PAI-1 0.68 ± 0.23 6.43 ± 0.56 #‡† 2.64 ± 0.93 1.38 ± 0.52
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Chemokines MCP-1 0.56 ± 0.17 2.23 ± 0.74 1.38 ± 0.34 1.25 ± 0.29 MIP-1α 0.92 ± 0.12 1.05 ± 0.05 1.12 ± 0.11 0.87 ± 0.16 CXCL1 0.82 ± 0.08 0.90 ± 0.11 1.07 ± 0.16 0.98 ± 0.05 CXCL4/PF-4 0.92 ± 0.10 0.98 ± 0.05 1.09 ± 0.14 1.02 ± 0.08 CX3CL1 0.83 ± 0.04 1.10 ± 0.15 1.05 ± 0.09 1.26 ± 0.08 (Fractalkine) CXCL16 0.83 ± 0.11 1.93 ± 0.39 2.13 ± 0.88 2.25 ± 0.96 CXCL10 0.78 ± 0.07 0.99 ± 0.11 1.34 ± 0.31 1.07 ± 0.03 CXCL12/SDF-1 0.96 ± 0.26 1.81 ± 0.17 1.77 ± 0.20 2.62 ± 0.98 GM-CSF 0.94 ± 0.12 0.87 ± 0.08 0.93 ± 0.04 0.98 ± 0.08 Inflammation IL-1α 0.84 ± 0.12 1.04 ± 0.08 0.82 ± 0.07 1.03 ± 0.17 IL-10 (anti-) 0.89 ± 0.18 1.12 ± 0.05 1.19 ± 0.14 1.12 ± 0.10 Amphiregulin 0.97 ± 0.02 0.84 ± 0.06 0.95 ± 0.09 1.03 ± 0.17 Tissue Factor 0.89 ± 0.22 1.50 ± 0.25 1.51 ± 0.35 1.45 ± 0.48 Endoglin/CD105 0.86 ± 0.16 1.36 ± 0.05 1.17 ± 0.19 1.15 ± 0.41
Table 13: Expression levels of various angiogenic-related proteins were determined in WT,
Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only muscles at 7 days of ischemia using a
protein antibody array. For each group, the fold change in ischemic muscle protein
expression is presented as a ratio of the contralateral (sham) limb. Data are presented as
mean ± SEM. n=4 for each mouse group. Fold changes that appear in red indicate a trend
towards or significant difference. Statistical significance is marked by #p<0.05 vs. WT,
‡p<0.05 vs. FGF2 LMW-only, †p<0.05 vs. FGF2 HMW-only.
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HMW FGF2 promotes neutrophil infiltration during ischemia
In addition to vascular cells, ischemia-induced revascularization also triggers the activation and mobilization of bone marrow derived-mononuclear cells (BM-MNCs) through increased expression of chemokines and adhesion molecules (Frangogiannis et al., 2002; Keeley, Mehrad,
& Strieter, 2008). These cells of myeloid lineage are the first responders and their trafficking starts within minutes following the ischemic injury and can last up to two weeks (Meisner & Price, 2010;
Tidball, 2005). Initially, neutrophils predominate the BM-MNCs that are recruited to the ischemic muscles (Frangogiannis & Entman, 2005; Tidball, 2005). Their numbers have been observed to increase as early as two hours after the injury and are undetectable 7 days later (Hoefer et al., 2005;
Silvestre et al., 2013). Many studies have described a role for inflammatory cell response in FGF2- mediated neovascularization (Andrés et al., 2009; Jih et al., 2001; Leali et al., 2003; Presta et al.,
2007, 2009).
To identify the role of the FGF2 isoforms in the tissue inflammatory response to chronic hindlimb ischemia, neutrophil recruitment in non-ischemic, sham-operated or ischemic calf and thigh was assessed (Figure 42). To that end, neutrophils were identified as myeloperoxidase
(MPO) positive cells. MPO is a peroxidase enzyme and the major component of neutrophil azurophilic granules (Bradley, Priebat, Christensen, & Rothstein, 1982; Oklu, Albadawi, Jones,
Yoo, & Watkins, 2013). Calf (gastrocnemius) and thigh (gracilis, semi-membranosus and semi- tendinosus) muscles were harvested at baseline or 3 days of ischemia from WT, Fgf2-/-, FGF2
LMW-only and FGF2 HMW-only mice. After immunodetection of MPO, no significant differences between genotypes in the number of neutrophils were present in non-ischemic muscles
(Figure 41A). Three days of ischemia showed a significantly higher number of neutrophils in ischemic muscles from Fgf2-/- and FGF2 HMW-only mice relative to their respective sham
202 muscles (p<0.05 vs HLI, Figure 42B). Surprisingly, neutrophil levels in WT and FGF2 LMW-only ischemic muscles were unchanged from their contralateral controls. Furthermore, FGF2 HMW- only muscles and Fgf2-/- had significantly increased neutrophil density relative to WT (p<0.05).
This suggests that while ischemia-induced neutrophil accumulation was inhibited by expression of all the FGF2 isoforms, HMW FGF2 alone can sufficiently mount an inflammatory response.
A. Neutrophil density (non-ischemic muscle)
0.1 WT Fgf2-/- FGF2 LMW-only 0.075 FGF2 HMW-only
0.05
0.025 Number ofcells/field MPO+ Number
0 1
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Neutrophil density (ischemic muscle) B.
5 WT # * ‡ Fgf2-/- 4 FGF2 LMW-only FGF2 HMW-only 3 # *
2
1 Number ofcells/field MPO+ Number
0 SHAM HLI
Figure 42. Neutrophil density expressed as the number of myeloperoxidase (MPO)-positive cells/field in WT (white bars), Fgf2-/- (light gray bars), FGF2 LMW-only (dark gray bars), and FGF2 HMW-only (black bars) non-ischemic (A) and ischemic (B) skeletal muscle after
3 days of ischemia. Data are presented as mean ratio ± SEM, n= 4. *p<0.05 vs. SHAM cohort,
#p< 0.05 vs. WT, ‡p< 0.05 vs. FGF2 LMW-only.
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Selective ablation of FGF2 isoform expression does not alter monocyte infiltration
Subsequent to the accumulation of neutrophils at the site of ischemic injury, a different
population BM-MNCs are recruited (Meisner & Price, 2010). These late arriving BM-MNCs (3
days after injury) are predominantly monocytes/macrophages and they further amplify the
inflammatory response (J. Lu, Pompili, & Das, 2013; Nguyen & Tidball, 2003). Neutrophils also
secrete cytokines and chemokines that attract and activate macrophages. The initial neutrophil
inflammatory infiltrate is replaced by macrophages and this is important for the removal of
necrotic tissue and resolution of inflammation (Serhan & Savill, 2005; Silvestre et al., 2008;
Tidball, 2005). Macrophages also secrete a repertoire of pro-angiogenic growth factors (including
FGF2), cytokines and ECM remodeling proteins that propagate the revascularization process
(Fung & Helisch, 2012; Silvestre et al., 2008, 2013). FGF2 exerts a direct chemotactic effect on
monocytes as well as upregulates the expression of other chemoattractants important for the
adhesion and recruitment of monocytes/macrophages (Andrés et al., 2009; Leali et al., 2003; Presta
et al., 2005, 2009).
To further examine the role of the FGF2 isoforms in the inflammatory response to chronic
hindlimb ischemia, macrophage infiltration was quantified in non-ischemic, sham-operated or
ischemic skeletal muscles. Macrophages/monocytes were identified as Mac-3-positive cells
(Koestler, Rieman, Muirhead, Greig, & Poste, 1984; Meisner et al., 2014; Shireman, Contreras-
Shannon, Reyes-Reyna, Robinson, & McManus, 2006). Calf (gastrocnemius) and thigh (gracilis, semi-membranosus and semi-tendinosus) muscles were harvested at baseline, 3 or 7 days of ischemia from WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only mice. No significant differences between genotypes in the density of macrophages were present in non-ischemic muscles (Figure 43A). By 3 days of ischemia, there was significantly increased macrophage
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infiltration in the ischemic muscles of WT, Fgf2-/-, and FGF2 LMW-only mice compared to their respective sham muscles (p<0.05 vs HLI, Figure 43B). This differed from ischemic FGF2 HMW- only muscles whose macrophages numbers were similar to their sham limb levels.
Monocyte/macrophage infiltration was also evaluated after 7 days of ischemia (Figure
43B). The number of Mac-3-positive cells in the ischemic muscles was significantly increased compared to sham muscles for all mouse groups (p<0.05 vs HLI). Expression of only LMW FGF2 or only HMW FGF2 did not change the level of ischemia-induced macrophage recruitment relative to WT. However, complete absence of FGF2 protein expression (Fgf2-/-) affected the amount of macrophage infiltrate in response to ischemia. At 3 or 7 days of ischemia, Fgf2-/- muscles had higher numbers of macrophages. There was a nearly 3-fold increase in infiltration in the Fgf2-/-
ischemic limbs compared to WT, FGF2 LMW-only or FGF2 HMW-only limbs.
The data presented here suggest that expression of FGF2 protein (either LMW or HMW isoforms) is necessary to induce normal macrophage infiltration during ischemia and loss of total
FGF2 expression results in an exaggerated and potentially harmful inflammatory response.
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Monocyte density (non-ischemic muscle)
A. 0.5 WT Fgf2-/- 0.4 FGF2 LMW-only FGF2 HMW-only 3+ cells/field - 0.3
0.2
0.1 Number of Mac Number
0 1
B. Monocyte infiltration (ischemic muscle) 20 * #‡ †
15 *# ‡ †
10
* * * 5 * *
Number of Mac - 3+ cells/field Number
0 3d SHAM 3d HLI 7d SHAM 7d HLI
Figure 43. Monocyte density expressed as the number of Mac-3-positive cells/field in WT
(white), Fgf2-/- (light gray), FGF2 LMW-only (dark gray), and FGF2 HMW-only (black)
non-ischemic (A) and ischemic (B) skeletal muscle after 3 or 7 days of ischemia. Data are
presented as mean ratio ± SEM, n = 2-4. *p<0.05 vs. SHAM cohort, #p< 0.05 vs. WT, ‡p<0.05
vs. FGF2 LMW-only, †p<0.05 vs. FGF2 HMW-only.
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DISCUSSION II
The results presented in this chapter of the dissertation demonstrate that the endogenous
HMW FGF2 isoforms have an important role in protecting skeletal muscle from ischemia-induced damage. This effect of the HMW isoforms is possibly mediated by FGF receptor-independent signaling, leading to gene expression changes in angiogenic pathways (IGFBPs and CX3CL1) and in conjunction with an altered inflammatory cell response (Figure 44).
Since its discovery in the late 1980s, FGF2 has been classified as a “classic angiogenic factor” (Presta et al., 2007; Ribatti, 2014). In vitro experiments showed a mitogenic effect of FGF2 on numerous cell types including endothelial cells, smooth muscle cells, breast cancer cells, fibroblasts, neuronal cells, and osteoblasts (Bikfalvi et al., 1995; Grothe, Meisinger, Holzschuh,
Wewetzer, & Cattini, 1998; Pasumarthi, Kardami, & Cattini, 1996; Piotrowicz, Maher, & Levin,
1999; Xiao et al., 2003). Moreover, exogenous FGF2 is a potent inducer of proliferation, migration and/or differentiation of endothelial and vascular smooth muscle cells; cellular activities which are critical for vascular growth (Davis et al., 1997; Pintucci et al., 2005; Piotrowicz et al., 2001,
1999). The function of endogenous FGF2 has also been investigated in angiogenesis and other cellular processes. The distinction between the roles of the endogenous or exogenous forms is important in the study of many proteins but is particularly significant when considering FGF2. The existence of several protein isoforms produced from a single Fgf2 gene coupled with discrete cellular localizations of those isoforms prompted the use of the isoform-specific mouse models when deciphering the functions of the classes of FGF2 isoforms.
In the present study, mice with a targeted deletion of the LMW FGF2 or HMW FGF2 were utilized to determine the endogenous roles of the FGF2 isoforms in revascularization of the hindlimb. Characterization of normal skeletal muscle was performed to eliminate the potential of
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confounding results from altered baseline FGF2 expression. Expression of Fgf2 mRNA and all protein isoforms was present in non-ischemic WT muscles (Figures 28 & 29). This is in accordance with the patterning of FGF expression levels in adult mouse tissues that found moderate expression of Fgf2 mRNA in the structural system (skin, bone and skeletal muscle) (Fon Tacer et al., 2010).
When Fgf2 mRNA was examined in tibiae of HMWKO mice (i.e. FGF2 LMW only), expression was significantly decreased (Homer-Bouthiette et al., 2014). In normal skeletal muscle, however,
Fgf2 mRNA expression is significantly increased when only HMW FGF2 or only LMW FGF2 is present. Ablation of either the LMW or HMW protein isoforms also altered FGF2 protein expression. Similar to the changes in mRNA, loss of HMW or LMW FGF2 expression in skeletal muscles resulted in increased FGF2 protein expression. In the case of global deletion of HMW
FGF2, the LMW isoform was significantly increased and vice versa. This phenotype is partly recapitulated in non-ischemic HMWKO hearts, where the LMW isoform is increased in the absence of the HMW isoforms (Liao et al., 2010). Conversely, in LMWKO (i.e. FGF2 HMW only) hearts, expression of the HMW isoforms (21 or 22kDa) remains unchanged (Liao et al., 2007).
These changes in FGF2 expression suggest that under baseline conditions, there exists a FGF2- mediated regulation of FGF2 expression at the transcriptional and translational levels. Previous work by Jimenez had provided evidence that FGF2 regulates its own promoter in cardiac myocytes
(Jimenez et al., 2004). Furthermore, this regulation is selective based on the FGF2 isoform(s) expressed and in which tissues they are expressed (Coffin et al., 1995). The increased expression of the HMW isoforms takes on more importance because the 18kDa LMW FGF2 isoform is the predominant FGF2 that is expressed in both wildtype and FGF2 transgenic skeletal muscle (Coffin et al., 1995). However, it is important to note that there may be a baseline level of FGF2 protein expression irrespective of isoform identity that is required for cellular functions in skeletal
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myocytes. The possibility that this is the driver of the increased FGF2 isoform or mRNA
expression in the absence of LMW or HMW FGF2 cannot be excluded.
In light of the changes to baseline levels of Fgf2 and its protein products when a specific class of protein isoform is deleted, the effect of chronic ischemia on FGF2 expression was also addressed. This dissertation demonstrated that induction of chronic ischemia in the mouse hindlimb produced changes in FGF2 transcript or protein expression. The presence of only LMW
FGF2 significantly increased skeletal muscle Fgf2 mRNA compared to wildtype at day 7 of
ischemia (Figure 31). Similarly in the absence of the LMW isoform (only HMW FGF2 present),
Fgf2 mRNA was significantly elevated. This increase persisted over the course of the tissue repair
and remodeling phases (1-14 days) that characterize the physiological response to ischemia in
muscle (Silvestre et al., 2013; Smith et al., 2008; Tidball, 2011). Studies of ischemia in the rabbit
hindlimb have also reported a similar a pattern of FGF2 transcript expression (Bush et al., 1998;
Walgenbach, Gratas, Shestak, & Becker, 1995). Deindl and colleagues however, did not observe
any changes in FGF mRNA (both FGF1 and FGF2) at any time points of ischemia in the same
model.
When only the HMW isoforms were present, FGF2 expression (21 or 22kDa) was also
increased compared to wildtype expression at day 7 of ischemia (Figures 33B, C). However, the
level of FGF2 protein in ischemic muscles did not rise above that of baseline expression (FGF2
HMW-only non-ischemic) muscles and was similar to expression in sham tissues. This suggests that while HMW FGF2 isoform expression might be changed when the LMW isoform is ablated; there is no additional ischemia-dependent promotion of FGF2 translation in ischemic skeletal muscle. Increased expression of FGF2 protein (all isoforms) has been detected in the urine, serum or ischemic tissues of patients with myocardial or limb ischemia (Campuzano et al., 2002; Cuevas
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et al., 1997; Gu, Santiago, Olowe, & Weinberger, 1997; Hasdai et al., 1997; Rohovsky et al., 1996).
Enhanced FGF2 expression has also been observed in animal models of acute cardiac
ischemia/reperfusion injury or hindlimb ischemia (Arras et al., 1998; Bush et al., 1998; Chleboun
& Martins, 1994; Cohen, Vernon, Yaghdjian, & Hatcher, 1994; X. Liu & Zhu, 1999; Sullivan et
al., 2002; Walgenbach et al., 1995).
The upregulation of FGF2 expression in ischemia models is not limited to cardiac or
skeletal myocytes alone. Rodent models of focal cerebral ischemia have been shown to induce
Fgf2 mRNA and/or protein expression within 24h of the injury (T. N. Lin, Te, Lee, Sun, & Hsu,
1997; Speliotes et al., 1996; X.-C. Zhao et al., 2013). Additionally, optic or peripheral nerve
axotomy initiates release of FGF2 from astrocytes or glial cells (Duprey-Díaz, Blagburn, &
Blanco, 2012; Grothe et al., 1996; Meisinger & Grothe, 1997b). Drug-induced neurotoxicity that
produce lesions of the nigrostratal pathway is another injury condition where increases in Fgf2
mRNA and/or protein are present in astrocytes and microglia (Chadi, Cao, Pettersson, & Fuxe,
1994; Chadi & Gomide, 2004; Leonard et al., 1993; Nakagawa & Schwartz, 2006; Noda et al.,
2014; Yurek, Fletcher, Peters, & Cass, 2010). Furthermore, cultured cortical astrocytes in an
ischemia-like state produce increased FGF2 protein expression that largely accumulates in nuclear bodies (X. Liu & Zhu, 1999). This spatial distribution of FGF2 may provide some evidence for
the identity of the FGF2 isoform that is preferentially upregulated in during brain injury. The
HMW FGF2 isoforms has been shown to be predominantly localized to the nucleus and not
secreted in various cell types including cardiomyocytes, neurons, fibroblasts, and endothelial cells
(Arese et al., 1999; Chadi & Fuxe, 1998; Claus et al., 2003; Garmy-Susini et al., 2004; Liao et al.,
2010).
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Detection of FGF2 protein by Western blotting allows for quantification of the levels of
each isoform and represents a more informative method than histological methods. The currently available FGF2 antibodies do not distinguish between the isoforms types; therefore, immunohisto-
or immunocyto- chemistry techniques utilized in the previously described reports are only useful
for demonstrating changes in overall FGF2. In response to neurotoxin-induced degeneration in the
rat brain, expression of all three isoforms was analyzed for up to 4 weeks after the lesions (Claus,
Werner, et al., 2004). The LMW or HMW isoforms displayed similar expression patterns and were
not different from intact rat brains. Using a similar injury model where only the levels of the HMW
FGF2 was reported, the 21kDa was unchanged while 23kDa expression was elevates relative to
intact brains (Silva, Fuxe, & Chadi, 2009). The distinction made in these few studies between the
endogenous FGF2 proteins is very critical to further assign cellular functions to the HMW and
LMW isoforms. This dissertation, for the first time, demonstrated that in skeletal muscle,
endogenous LMW or HMW FGF2 can regulate overall FGF gene or protein expression under
baseline conditions. Furthermore, this isoform-specific control of FGF2 expression persists under chronically ischemic conditions and is present at both the level of gene transcription and protein translation.
FGFs are widely expressed across several physiological systems where they serve as
ligands to a family of high affinity FGF receptors (FGFRs 1, 2, 3 & 4). FGFR1, 2 and 3 have
additional splice variants of the immunoglobulin-like domain III (IIIb and IIIc) which are important for FGF ligand-binding specificity (Eswarakumar et al., 2005; Itoh & Ornitz, 2011). The
most closely related FGF to FGF2, FGF1 is known to bind and activate all the FGFR splice variants
(Ornitz & Itoh, 2015). FGF2 (the LMW isoform) activates all FGFRc isoforms, FGFR3b and
FGFR4 (summarized in Table 2). To establish whether the phenotypic changes observed in
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response to ischemia and the presence of only LMW FGF2 or HMW FGF2 were associated with
FGF receptor levels and/or activation, receptor gene and protein expression and activation was
determined in normal or ischemic muscles.
Interestingly, neither Fgfr2 mRNA (IIIb or IIIc form) nor protein was detected in the any
of the muscles analyzed. This finding supports previous observations by others in which Fgfr2
transcript expression was below the level of detection or present in very low quantities in myogenic
cells isolated from wildtype mouse limbs (Fon Tacer et al., 2010; Yablonka-Reuveni, Danoviz,
Phelps, & Stuelsatz, 2015). Similarly, very low levels of Fgfr2 was present in adult rat flexor digitorum brevis and tibialis anterior (TA) muscles (Kästner, Elias, Rivera, & Yablonka-Reuveni,
2000). In a study by Yablonka-Reuveni and coworkers, muscle from wildtype mouse limbs were isolated and separated into non-myogenic and myogenic cell populations. Low levels of Fgfr2c
mRNA were detected in the non-myogenic cells but absent from the myogenic population
(Yablonka-Reuveni et al., 2015). These non-myogenic cells are likely to include cells of neuronal
origin as the CNS, including the spinal cord, has high expression of Fgfr2c transcripts and little or no Fgfr2b (Fon Tacer et al., 2010; Hensel et al., 2012; Ratzka, Baron, & Grothe, 2011). These
studies confirm that the RT-PCR and immuoblotting experiments performed in this dissertation
were specific for Fgfr expression in skeletal myocytes and the absence of expression is not a result
of unsuccessful primer design or other errors in experimentation.
Receptor activation assays have shown that FGF2 (LMW isoform) is selective for the IIIc
variant of FGFR1 and has lower binding to FGFR1b (Beenken & Mohammadi, 2009; Xiuqin
Zhang et al., 2006). In studies where the expression of the IIIb isoform of Fgfr1 was distinguished
from the IIIc form, skeletal muscle expression of Fgfr1c was predominant over Fgfr1b (Fon Tacer et al., 2010; Hensel et al., 2012; S. E. Hughes, 1997). Likewise in the CNS (spinal cord), the Fgfr1c
213 transcript is more abundant over Fgfr1b (Fon Tacer et al., 2010; Hensel et al., 2012; Ratzka et al.,
2011). Both splice variants of Fgfr1 were present in all skeletal muscles analyzed from WT or
FGF2 HMW-only mice (Figure 35A). In normal muscle expressing only the HMW isoforms,
Fgfr1b transcript levels were depressed while Fgfr1c was unchanged. While Fgfr1c is the more abundant variant in muscle and vascular cells, FGF2 can still bind and activate the protein product of the Fgfr1b transcript (S. E. Hughes, 1997; Ornitz & Itoh, 2015; Ornitz et al., 1996). The down- regulation of the receptor could be attributed to the presence of increased HMW FGF2 protein expression in non-ischemic muscle. Both variants of Fgfr3 were not differentially expressed from
WT in FGF2 HMW-only mice (Figure 36A). Similarly to Fgfr1, the “b” isoform of Fgfr3 was lower in expression or similar to the level of detection in WT muscle or spinal cord (Fon Tacer et al., 2010; Hensel et al., 2012; Ratzka et al., 2011). The presence of only HMW FGF2 did not alter
Fgfr4 mRNA levels in non-ischemic muscles (Figure 37A). Fgfr4 mRNA has been shown to be present in equal amounts to Fgfr1 transcripts in muscle, brain and bone cells (Fon Tacer et al.,
2010; Han, Xiao, & Quarles, 2015; Hensel et al., 2012; Yablonka-Reuveni et al., 2015).
In response to ischemia, Fgfr1b or Fgfr1c mRNA expression was highest by 7 days in WT muscle, declining to control levels at 14 days (Figures 35B & C). When only the HMW isoforms of FGF2 were present, a similar pattern of Fgfr1 expression was observed with a marked increase in transcript levels at day 3 of ischemia. This effect can be attributed to the loss of LMW FGF2 protein as Fgf2-/- muscles exhibited a comparable pattern of Fgfr1 transcript expression. The results from the WT mice are not consistent with a rabbit arteriogenesis study where increases to
Fgfr1 mRNA in skeletal muscle was rapid, peaked at 6 hours after the femoral artery occlusion, and decreased to control levels by 3 and 7 days (Deindl et al., 2003). The absence of qRT-PCR experiments from ischemic muscles isolated before 24 hours precludes a true comparison between
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the results of this dissertation and the work by the Schaper group but it does suggest that there
might be a bi-phasic (early and late phases) effect on Fgfr1 expression during hindlimb ischemia
(Deindl et al., 2003). Experimental evidence for this phenomenon was present in ischemic rat
extensor digitorum longus (EDL) muscles where VEGF mRNA and protein was elevated within
3 days of the injury, returned to normal by 7 days and a secondary wave of VEGF protein was
observed at 14 days (Milkiewicz, Hudlicka, Shiner, Egginton, & Brown, 2006). Similar to Fgfr1,
two splice variants of Fgfr3 (Fgfr3IIIb and Fgfr3c) were detected in skeletal muscle. In the
presence of ischemia and only HMW FGF2 or only LMW FGF2 expression, neither Fgfr3
transcript was altered (Figures 36B & C). By the same token, the single Fgfr4 mRNA was
unchanged with the loss of either FGF2 protein isoform (Figure 37B). This suggests that the loss
of LMW isoform expression modulates ischemic Fgfr expression, particularly the Fgfr1 isoforms.
It is likely a compensatory measure during a stress condition like ischemia to boost autocrine
FGFR1 signaling in the absence of LMW FGF2. This is not surprising as FGFR1 is the
predominant receptor for FGF2 in skeletal muscle (Fon Tacer et al., 2010; Hensel et al., 2012;
Yablonka-Reuveni et al., 2015), heart (Liao, 2008) and it shares this role with FGFR2 in bone and neuronal development and repair/regeneration (Du et al., 2012; Fei et al., 2013; Grothe et al., 2006;
Masumura, Murayama, Inoue, & Ohno, 1996; Nakajima et al., 2001).
FGFR expression was also evaluated at the protein level along with FGFR activation (via
phosphorylation). Protein expression of FGFR1, 3 or 4 was not affected by the presence of only
the HMW FGF2 isoforms in non-ischemic muscles. Likewise, cultured calvarial osteoblasts with
an overexpression of HMW FGF2 had no change in either FGFR1 or FGFR2 protein expression
(Sabbieti et al., 2013). Overexpression of the human FGF2 isoforms (22, 23 and 24kDa), however,
resulted in amplified Fgfr1c and Fgfr3c mRNA expression as well as increased nuclear
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accumulation of FGFR1 protein in adult osteoblasts or bone marrow stromal cells (Xiao et al.,
2010, 2013). This pattern is also observed in human neuroblastoma cells transfected with human
23kDa FGF2 (Dunham-Ems et al., 2009). The AR4-2J pancreatic cell line also produced increased
Fgfr1 mRNA three-fold in the presence of 22.5kDa FGF2 overexpression (Estival et al., 1996).
On the other hand, FGF receptor protein expression was not altered in cultured NIH 3T3 fibroblasts
in response to HMW FGF2 (22, 22.5 and 24kDa) overexpression (Bikfalvi et al., 1995). There
appears to be a tissue-selective up-regulation of FGFR transcript or protein, especially FGFR1 in
the presence of only HMW FGF2 expression. In this dissertation, activation (tyrosine
phosphorylation) of FGFRs 1, 3 and 4 was also unchanged by HMW FGF2 expression (Figure
38). Basal FGFR1 activation has been shown to be decreased in mouse hearts overexpressing
24kDa HMW FGF2 with no change in the receptor protein expression (Liao et al., 2010; Manning
et al., 2013). These results confirm that in skeletal muscle, the HMW FGF2 isoforms act in an
intracrine manner and are not secreted like their paracrine/autocrine LMW counterpart to interact
with the FGFRs. This joins the increasing evidence suggesting that HMW FGF2 is a nuclear
growth factor with putative transcription factor activity (Chlebova et al., 2009; Liao et al., 2009;
Yu et al., 2007).
After the induction of ischemia (3 or 7 days), FGFR 1, 3, and 4 protein expression or
activation was again evaluated and found to be similar between WT and HMW FGF2 expressing-
limbs (Figures 39-41). The study discussed above by the Schaper group also assessed the
expression of FGFR1 protein in the ischemic rabbit hindlimb (Deindl et al., 2003). The authors
observed an early up-regulation of both FGFR1 phosphorylation (i.e., activation) and protein
expression at 6 and 12 hours, respectively. As with the transcript expression data, these results
propose that the effect of ischemia on FGFR1 expression/activation occurs at an early phase that
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is excluded from the results of this dissertation. The other FGFRs, FGFR3 and 4 might also be
similarly induced in ischemic muscle. Nevertheless, the timeframe chosen for the mRNA and
protein expression analyses is in agreement with other published reports of receptor tyrosine kinase
expression/activation during ischemia. VEGF-R2 (Flk-1), a receptor that positively modulates vascular growth and homeostasis in response to the VEGFs, was elevated within 3 days of ischemia and declining to control levels by 7 - 10 days in rat and rabbit muscles after femoral or iliac artery ligation (Lloyd, Prior, Yang, & Terjung, 2003; Milkiewicz, Hudlicka, Verhaeg, Egginton, &
Brown, 2003; Rissanen et al., 2002). However, Prior and coauthors observed no changes to VEGF-
R2 mRNA at any timepoint (up to 3 weeks) (Prior et al., 2004). Another tyrosine kinase receptor,
PDGFRα was also upregulated in expression at 3 days of ischemic and declining after 10 days
(Moriya et al., 2013).
Further proof of the changes in FGFR expression has also been observed in other muscle injury models. Chronic low frequency stimulation which has been shown to increase capillary growth in fast twitch fibers (Hudlicka et al., 1994; Hudlicka & Brown, 2009; Shen et al., 2009)
amplified Fgfr1 and Fgfr4 mRNA almost two-fold in rat tibialis anterior (TA) muscles
(Düsterhöft, Putman, & Pette, 1999). However, Fgfr1 was shown to be unchanged in rat EDL stimulated for 5 days (Brown et al., 1998). Fgfr1 mRNA/protein and Fgfr4 mRNA were elevated in response to mouse hindlimb suspension and this prevented muscle disuse atrophy (Eash, Olsen,
Breur, Gerrard, & Hannon, 2007). Similarly, both Fgfr1 and Fgfr4 transcripts were up-regulated in a spinal muscle atrophy (SMA) model (Hensel et al., 2012). Transient ischemia in the rat eye or brain selectively induces Fgfr1 mRNA within hours of injury and persists for days (Endoh,
Pulsinelli, & Wagner, 1994; Masumura et al., 1996; Miyashiro et al., 1998). Similarly several
“ischemia-like” insults (serum-free, glucose-free or glutamate addition) in cultured astrocytes
217
increased FGFR1 immunoreactivity (X. Liu & Zhu, 1999). Nerve lesion injury also induces
expression of Fgfr1-3 transcripts and FGFR1 protein at the lesion site (Grothe, Meisinger, & Claus,
2001; Grothe et al., 1996). Temporal expression analyses of skeletal fractures revealed early
upregulation of Fgfrs 1 and 2 at 1 day after injury which increased over the healing period with
maximal expression at 14 days post-fracture (Nakajima et al., 2001; Schmid et al., 2009). In
contrast, expression of Fgfr3 was not observed until 10 days in a rat model of femoral fracture
(Rundle et al., 2002).
At baseline or in response to ischemia, ablation of the HMW FGF2 isoforms (in FGF2
LMW-only mice) did not affect Fgfr expression at the RNA or protein level with the exception of
Fgfr4 (Figures 35 - 41). The presence of LMW FGF2 alone increased the levels of Fgfr4 transcripts after 3 and 7 days of ischemia (Figure 37B) which did not produce any alterations of FGFR4 protein expression or phosphorylation. Consistent with the results presented here, expression of only LMW FGF2 in the heart (HMWKO) did not affect FGFR1 protein expression at baseline
(Liao et al., 2010; Manning et al., 2013). FGF receptor levels were decreased in NIH 3T3 cells overexpressing 18kDa FGF2 (Bikfalvi et al., 1995). Conversely, osteoblastic-specific LMW FGF2 overexpression increased basal Fgfr1 and Fgfr2 mRNA expression (Xiao et al., 2009, 2014). The
AR4-2J pancreatic cell line also produced increased Fgfr1 mRNA two-fold in the presence of
18kDa FGF2 overexpression (Estival et al., 1996). Cultured calvarial osteoblasts with an overexpression of LMW FGF2 had increased FGFR1 protein levels with no change in FGFR2 expression (Sabbieti et al., 2013). These studies suggest that shifting the FGF2 equilibrium towards increased LMW FGF2 results in upregulation of its dominant receptors in the respective tissues.
The existence of FGFR-independent signaling and responses induced by endogenous
HMW FGF2 adds a layer of complexity to the LMW/HMW/FGFR interactions. It is widely
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accepted that of all endogenous FGF2 proteins, it is the 18kDa LMW FGF2 that interacts with the
cell surface FGF receptors to mediate autocrine and paracrine signaling responses (Chlebova et
al., 2009; Yu et al., 2007). This is due to the localization of this isoform to the cytoplasm, cell
surface and the extracellular matrix (Kardami et al., 2007; Presta et al., 2005). It can also be
secreted at times of cellular stress (Piotrowicz, Martin, Dillman, & Levin, 1997; Steringer, Müller,
& Nickel, 2014). On the other hand, the HMW isoforms with NH2 terminal nuclear localization
signals are present exclusively in the nuclear compartment and do not typically interact with the
membrane-bound FGF receptors (Claus et al., 2003; R Z Florkiewicz, Baird, & Gonzalez, 1991).
Studies with exogenous administration of recombinant HMW FGF2 have reported FGFR-
dependent cellular signaling on par with activation by LMW FGF2 (Gualandris et al., 1994; Jiang
et al., 2007; Moscatelli, Joseph-Silverstein, Manejias, & Rifkin, 1987). There is evidence to
suggest that there are differences between the cellular activities of exogenous and endogenous
HMW FGF2. Unlike LMW FGF2, endogenous HMW FGF2 is seldom secreted under basal or
stress conditions (Claus et al., 2003; House et al., 2003; Pintucci et al., 2005). Cardiac
myofibroblasts have been shown to accumulate and export HMW FGF2 after angiotensin II
stimulation (Santiago et al., 2011). The HMW isoforms share considerable sequence homology with LMW FGF2 including the binding domains and affinities for HSPG or FGFR1 so it is likely that the HMW FGF2/FGFR-dependent signaling observed in these studies is not representative of
actions of endogenous HMW FGF2 (Moosa Mohammadi et al., 2005).
In NIH 3T3 cells overexpressing all the HMW isoforms (22, 22.5 and 24kDa FGF2) or
only the LMW isoform (18kDa), similar levels of cell migration were observed and were
subsequently inhibited by the introduction of a dominant negative form of the FGF receptor in
only the cells with transfected with 18kDA cDNA (Bikfalvi et al., 1995). Likewise, HMW FGF2-
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induced alteration of PKCδ, PKCε or ERK1/2 activation in cultured pancreatic cells were not
affected by dominant negative FGFR1 (Gaubert et al., 2001; Hortala, Estival, Pradayrol, Susini, &
Clemente, 2005). FGF2 neutralizing antibodies also did not revert cellular responses induced by
HMW FGF2 indicating that paracrine/autocrine signaling by secreted FGF2 was not responsible for the effect of HMW FGF2 (Estival et al., 1996; Gaubert et al., 2001; Ma et al., 2007; Nindl et al., 2004; Pasumarthi et al., 1996; Xiao et al., 2003). In response to acute I/R injury, Liao and
colleagues reported decreased PKCα activity (translocation), increased MAP kinase activation
(p38, ERK or JNK) and amplified Akt activation in hearts expressing only the HMW isoforms or
overexpressing the human 24kDa FGF2 (Liao, 2008; Liao et al., 2007). These cellular responses likely represent intracrine signaling by the nuclear localized HMW FGF2 that are not mediated by classical interaction with membrane FGFRs.
Integrative nuclear FGF receptor 1 signaling (INFS) is a novel mechanism that involves gene regulation by a nuclear regulatory complex of FGFR1 and its partner CREB binding protein
(CBP) (Fang, Stachowiak, Dunham-Ems, Klejbor, & Stachowiak, 2005; Stachowiak, Maher, &
Stachowiak, 2007). Nuclear FGFR1 (nFGFR1) is a soluble full-length protein that accumulates in the nucleus bypassing Golgi processing and does not represent translocation of cell-surface FGFR1
(Dunham-Ems et al., 2009; Stachowiak et al., 2015). Gene activation by nuclear FGFR1 is not blocked after inhibition of ligand-induced FGFR1 internalization from the cell surface and does not require tyrosine kinase activity which supports the theory of INFS (Stachowiak et al., 2015,
2007). Immunolocalization and co-immunoprecipitation experiments have shown that HMW
FGF2 (23kDa) interacts with nuclear FGFR1 and is suggested to be part of a chromatin-bound complex with the transcription factor co-activator, CBP (Baron et al., 2012; Dunham-Ems et al.,
2009; Stachowiak et al., 2015). In contrast, LMW FGF2 does not interact with nuclear FGFR1
220
(Dunham-Ems et al., 2009; Stachowiak et al., 2015). Evidence for INFS induction has been observed in differentiation of neuronal populations, neurogenesis and neurodevelopment, activation and invasion of pancreatic stellate cells (Förthmann et al., 2013; Kiyota, Ingraham,
Jacobsen, Xiong, & Ikezu, 2011; Woodbury & Ikezu, 2013). Nuclear FGFR1 is also involved in direct regulation of several processes genes including; Fgf2, Fgf23, Nurr1, c-Jun and Nur77
(Baron et al., 2012; Han et al., 2015; Stachowiak et al., 2015; Terranova et al., 2015).
An intriguing phenotype emerged from the expression experiments performed in Fgf2-/-
mice. While basal expression of Fgfr 1b, 1c, 3b, 3c or 4 mRNA was unchanged in skeletal muscles
with deletion of Fgf2, ischemia produced an increase of Fgfr1c transcript at 3 days of ischemia.
The level of Fgfr1 activation observed was in keeping with this change in gene expression and
elevated in response to ischemia after 7 days of ischemia. Fgfr3 expression was unaltered at the
mRNA level but protein expression was increased at 3 days of ischemia. No differences in mRNA
or protein expression of Fgfr4 were present in Fgf2-/- muscles. Characterization of FGF receptor
expression profile in Fgf2-deficient skeletal muscles has never been reported. However, others
have documented the effects of Fgf2 ablation on FGFR expression in other tissues. Ortega and
coworkers described similar levels of Fgfr1 and Fgfr2 mRNA in brain, heart, liver and testes of
Fgf2-/- mice as WT (Ortega et al., 1998). Likewise, the Grothe group detected no changes in any
Fgfr transcripts in Fgf2 knockout spinal cord, striatum and ventral mesencephalon (Ratzka et al.,
2011). However, changes were present when expression of the receptors was determined in either
nuclear or cytoplasmic compartments of dopaminergic (DA) neurons. Fgf2-deficient DA neurons
had increased nuclear Fgfr1 accumulation but similar cytoplasmic Fgfr1 (Ratzka, Baron,
Stachowiak, & Grothe, 2012). The over-activation of INFS signaling (measured by nuclear-
localized FGFR1) is reversed after treatment with recombinant 18kDa which suggests the presence
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of selective FGF2 isoform-induced regulation of the equilibrium between intracrine and
cytoplasmic in FGF2-FGFR signaling.
The role of INFS in cellular responses of the endogenous HMW FGF2 isoforms has yet to
be studied in isoform-specific knockout mice. FGFR1 is currently the only FGF receptor known
to be present in the nucleus, where it is present in its full-length form and in association with HMW
FGF2 (Dunham-Ems et al., 2009; Stachowiak et al., 2003). The experiments to determine the
FGFR expression/activation in non-ischemic and ischemic skeletal muscles did not consider the
apparent compartmentalization of FGFR1 and the other FGFRs into cell-membrane or nuclear factions and represent overall cell/tissue expression of the receptors. Differences in gene regulation by membrane FGFRs and nuclear FGFR in ischemic limbs may represent a novel mechanism for mediating the FGF2 isoform-specific effect on ischemia-induced revascularization.
While whole muscle lysates were analyzed for the FGFR activation and expression results presented here, the levels of receptor phosphorylation and expression in other cells distinct from skeletal myocytes are also important. Skeletal muscle represents a mixture of myofibers and non- myogenic cell types including satellite cells, neurons, fibroblasts, and cells of the vascular compartment; endothelial (ECs) and smooth muscle (vSMCs). Immunodetection of FGFR1 in rodent muscle sections or rat myofiber cultures showed strong expression in myocytes, fibroblasts, pericytes, myogenic cells or vSMCs and no expression in ECs (Brown et al., 1998; Deindl et al.,
2003; Eash et al., 2007; Kästner et al., 2000). Conversely, positive FGFR4 staining was exclusively localized to myogenic cells and absent in non-myogenic cells (Cornelison, Filla, Stanley,
Rapraeger, & Olwin, 2001; Yablonka-Reuveni et al., 2015). Human ECs and vSMCs of aortic or umbilical origin in cultured singly or together in EC/vSMC co-culture spheroids strongly express
Fgfr1 mRNA and had lower expression of Fgfr2 or Fgfr4 (Antoine et al., 2005; Brogi et al., 1993).
222
In an extensive immunohistochemical study of adult human tissues, Hughes reported strong and
wide spread expression of Fgfr1-4 mRNAs in the endothelium and media of microvessels, veins
and arteries of epithelial and mesenchymal (S. E. Hughes, 1997).
The effect of isoform-specific FGF2 signaling on gene regulation has been investigated in
neovascularization processes. Findings suggest that FGF2 may exert an indirect control on
angiogenesis by promoting the expression of other angiogenic growth factors (Murakami &
Simons, 2008; Presta et al., 2005). This is proposed to occur in concert with its (FGF2) own direct
and potent modulation of angiogenesis. The strongest evidence for the indirect control of
angiogenesis by FGF2 is demonstrated by its effect on the VEGF system. FGF2-induced
angiogenesis in Matrigel implants coincided with increased expression of VEGF mRNA and
protein by infiltrating stromal cells and fibroblasts. Neutralizing VEGF antibodies suppressed the
FGF2-induced angiogenesis in this model (Claffey et al., 2001). Similarly, antibody blockade of
VEGFA and the VEGF receptor, Vegfr1 in murine brain ECs or HUVECs inhibited FGF2-
mediated capillary growth and morphogenesis (Kanda, Miyata, & Kanetake, 2004). In vivo, FGF2-
induced angiogenesis in the mouse cornea was associated with increased VEGF mRNA and
protein expression in endothelial cells of growing capillaries (Seghezzi et al., 1998). FGF2 gene
transfer to ischemic mouse hindlimbs targeted nonendothelial mesenchymal cells and induced
expression of angiogenic VEGF pathways (Fujii et al., 2006, 2008). Co-stimulation of cultured
endothelial cells with FGF2 and VEGF-A enhanced mural cell recruitment as well as endothelia-
mural interaction. Furthermore, neovessel formation required synergistic upregulation of Pdgfb nd
PDGF receptor β (Pdgfrβ) gene expression (Kano et al., 2005). FGF2 treatment with either PDGF-
A or PDGF-B synergistically promoted PDGFR and FGFR expression in ECs and vSMCs, respectively (Nissen et al., 2007). PDGF-A/FGF2 and PDGF-B/FGF2 combinations also
223 synergistically stimulated angiogenesis in mouse cornea and ischemic skeletal muscles (J. Zhang et al., 2009). PDGF, likewise plays a synergistic role in the FGF2-induced transcriptional and translation regulation of HGF expression in vitro in cultured cells and an in vivo ischemia model
(Onimaru et al., 2002). Overall, these studies suggest that FGF2 signaling lies upstream of enhanced growth factor signaling in ischemia-induced neovascularization. Interestingly, the
VEGF expression in ECs was similarly augmented in ECs transfected with LMW FGF2 or HMW
FGF2 cDNAs. However, when FGF2 neutralizing antibodies were administered, this upregulation is inhibited in LMW FGF2-expressing ECs but unchanged in HMW FGF2 cells (Seghezzi et al.,
1998). The intracrine effect of HMW FGF2 on VEGF expression follows the trend of FGFR- independent signaling by the HMW isoforms discussed previously.
The FGF2-isoform specific autocrine/intracrine signaling is proposed to underlie the selective transcriptional activation in response to overexpression of either only LMW FGF2
(18kDa) or HMW FGF2 (22, 22.5 and 24kDa) in NIH 3T3 cells. Quarto and colleagues performed a microarray analysis and reported that each class of FGF2 isoform differentially modulates a distinct profile of genes in stimulated cells (Quarto et al., 2005). In the HMW FGF2 -expressing cells, genes involved in growth arrest and tumor suppression were up-regulated while those associated with cell proliferation and tumor growth were down-regulated and unchanged in LMW
FGF2 cells. Using a proteome antibody array, the expression profile of angiogenesis-related proteins in HMW or LMW expressing skeletal muscle was investigated (Table 13). Protein isolated from sham or ischemic muscles after 7 days of ischemic was hybridized to an array spotted with antibodies to 53 different proteins. These proteins were classified based on their roles in angiogenic process into the following functional groups: growth factors, pro-angiogenic, angiostatic, ECM remodeling, chemokines and inflammatory mediators.
224
The array was spotted with several growth factors including FGFs, VEGFs, PDGFs, &
HGF which have been previously shown to be regulated by FGF2 (Murakami & Simons, 2008;
Presta et al., 2005). A novel role for FGF2 in the regulation of FGF7 protein expression was
observed. HMW FGF2 induced an increase in FGF7 expression during ischemia. The potential
role of FGF7 in ischemia-induced vascular growth is yet unknown as it has only been linked to the
proliferative phase of acute or chronic wound healing by promoting of keratinocyte proliferation
and migration (Maretzky et al., 2011; Niu et al., 2007; Yen et al., 2014). It is known that FGF2
autoregulates its own expression as described in the Chapter 1 of this dissertation as well as
modulating Fgf23 transcription and translation (Jimenez et al., 2004; Stachowiak et al., 2015; Xiao
et al., 2010). The presence of only the HMW isoforms up-regulated the expression of two IGF
binding proteins, IGFBP-3 and IGFBP-10, which modulate the activity of the IGF axis. IGFBP-3
is an angiostatic protein with inhibitory effects on tumor growth while IGFBP-10 promotes
angiogenesis by regulating angiogenic gene transcription in response to hypoxia (Kubota &
Takigawa, 2007; Malik et al., 2015; Yamada & Lee, 2009). Microarray studies detected increased
IGFBP3 gene expression within a day of hindlimb ischemia surgery and this change was
associated with regeneration of the ischemic muscle (Matsakas, Yadav, Lorca, Evans, & Narkar,
2012; Paoni et al., 2002). Two other genes in the insulin-like growth factor (IGF) axis, IGFBP-2
and IGF2 were both increased two-fold during 24kDa HMW FGF2-induced cell proliferation in
the AR4-2J cancer cell line (Hortala et al., 2005). Protein levels of CX3CL1 (fractalkine), a
chemokine was also elevated with expression of only HMW FGF2 and the presence of ischemia.
Comparison of the expression profile induced by selective HMW FGF2 expression in this
dissertation with the microarray analysis performed by Quarto and colleagues is not entirely
straightforward as the former analyzed expression of a specific group of proteins (angiogenesis-
225 related), while the later performed a whole genome expression survey (Quarto et al., 2005). The presence of selective FGF2 isoform expression occurred from birth in the muscle tissues analyzed while the fibroblasts were transfected with FGF2 cDNAs for only 48 hours. Nevertheless, it is important to note that like increased anti-angiogenic IGFBP-3, the microarray studies reported up- regulation of several genes with growth arrest and tumor inhibition activities in HMW FGF2 cells.
The proteome array identified epidermal growth factor (EGF) as elevated in the presence of LMW FGF2 (Table 13). Similarly to FGF7, EGF has been implicated in wound healing; particularly, in dermal wounds where it enhances proliferation and migration of fibroblasts and keratinocytes (Haase et al., 2003; Seeger & Paller, 2015). The pro-angiogenic factor angiopoietin- like 4 (Angptl4) was up-regulated in LMW FGF2 3T3 fibroblasts (Quarto et al., 2005). However, other members of the angiopoietin family (angiopoietins-1 and -3) analyzed on the array were not changed in the FGF2 LMW-only muscles. Angiopoetin-1 (Ang-1) is a potent angiogenic factor mainly expressed by perivascular cells of mature vessels (Eklund & Saharinen, 2013; Herbert &
Stainier, 2011). Ang-1-mediated activation of the EC-specific receptor Tie2 is important for the vessel stabilization and maturation (Eklund & Saharinen, 2013; Fagiani & Christofori, 2013).
Angiopoietin-3 also has positive angiogenic activity in human ECs and glioblastoma cells (H. J.
Lee et al., 2004; Morisada, Kubota, Urano, Suda, & Oike, 2006). This suggests that, in the context of chronic limb ischemia, there is an absence of LMW FGF2-induced expression of angiogenic factors.
The importance of FGF2-induced activation of angiogenic growth factor expression is further called into question when the expression profile of the angiogenic proteins is examined in
Fgf2-/- ischemic muscles. Like FGF2 LMW-only, expression of angiogenic factors like VEGF,
PDGF or HGF was not different from WT in Fgf2-/- mice. Two proteins of significant importance
226 during ECM remodeling were upregulated in Fgf2-/- muscles after 7 days of ischemia, MMP-9 and PAI-1. PAI-1 is the main inhibitor of the urokinase-type plasminogen activator (uPA) system which cleaves adhesion molecules, growth factors and their receptors as well as the ECM degradation required for endothelial and smooth muscle cell migration during angiogenesis
(Papetti & Herman, 2002; Van De Craen et al., 2012). The active enzyme of the uPA system is plasmin, a serine protease converted from the plasminogen proenzyme (Medcalf, 2007). In addition to the ECM, plasmin also converts latent pro-MMPs like MMP9 into active enzyme which in turn degrade the ECM (Arroyo & Iruela-Arispe, 2010; Q. Chen et al., 2013). The role of MMP-
9 is revascularization is as yet unclear since normal angiogenesis and arteriogenesis but increased necrosis was observed in MMP9-deficent ischemic limbs (Meisner et al., 2014). Pharmacologic inhibition of PAI-1 ameliorated tissue necrosis and enhanced angiogenesis in mice with hind limb ischemia (Tashiro et al., 2012). This improved angiogenesis was associated with increased immunolocalization of FGF2 in the PAI-1 inhibitor-treated muscles. This suggests that the presence of elevated PAI-1 and MMP-9 may underlie the reduced muscle viability observed in
Fgf2-/- mice (discussed in Chapter 1).
Surprisingly, no effect of selective FGF2 expression was observed in the expression of several inflammatory cytokines and chemokines. These chemokines/cytokines mediate the initial phase of wound healing/repair that occurs immediately following ischemic or other forms of tissue injury (Keeley et al., 2008; Silvestre et al., 2008). This is the inflammatory response that is critical for the eradication of infectious microbes, clearance of cellular waster or debris and eventually repair or resolution (Raman, Sobolik-Delmaire, & Richmond, 2011; Tidball, 2005, 2011).
Vascular growth and inflammation are closely linked and inter-dependent in a number of
(patho)physiological conditions including chronic ischemia (Keeley et al., 2008; Romagnani,
227
Lasagni, Annunziato, Serio, & Romagnani, 2004; Silvestre et al., 2008). The Presta group has
shown the necessity of inflammatory cytokines and chemokines for FGF2 induced-angiogenesis
in vitro and in vivo (Andrés et al., 2009; Presta et al., 2009). FGF2 stimulation of cultured
microvascular ECs up-regulated the expression of several pro-inflammatory genes including Ccl2,
Ccl7, Cxcl31, Cxcl1, Opn, IL-6. These cytokines and chemokines served as chemoattractants for
the recruitment of infiltrating leukocytes to vascularizing areas of FGF2-treated Matrigel plugs
(Andrés et al., 2009). Of these chemokines, CCL2 also known as MCP-1 is of particular
importance in FGF2-induced neovascularization. MCP-1, the most extensively studied of the CC
chemokine ligand family, regulates inflammation by recruiting leucocytes, such as monocytes, to
sites of tissue injury and infection (Capoccia, Gregory, & Link, 2008; Shireman, 2007). MCP-1
infusion potently induces angiogenesis and collateral growth in the ischemic rabbit hindlimb
(Arras et al., 1998; Deindl et al., 2003; Hoefer et al., 2001). Furthermore, MCP-1 mRNA and
protein is increased in response to FGF2 treatment in migrating BAECs and ischemic skeletal
muscle (Fujii et al., 2006; Jih et al., 2001; Onimaru et al., 2002; Wempe, Lindner, & Augustin,
1997). The time course of MCP-1 upregulation follows closely with the activation and trafficking
of monocytes/macrophages to the site of ischemic injury (Arras et al., 1998; Deindl et al., 2003).
The role of neutrophils, a subset of mononuclear leucocytes in FGF2-driven angiogenesis is not as
well understood.
Neutrophils predominate early in the inflammatory cell population that infiltrates injured/ischemic muscle where they may play a beneficial or negative role in the resolution of muscle injury (Tidball & Villalta, 2010; Tidball, 2011). The former is due to the phagocytic activity on necrotic debris and secretion of factors that attract and transactivate monocytes/macrophages while the latter is linked to the secondary damage from the generation of
228 free radicals (Smith et al., 2008; Tidball & Villalta, 2010; Tidball, 2005). Within 3-5 days of injury, they are replaced by monocytes/macrophages which is thought to signify the start of injury resolution (Smith et al., 2008; Tidball, 2005). Growth factors like FGF2 and other angiogenic cytokines are released by macrophages themselves or from activated ECs to mediate the angiogenesis process (Andrés et al., 2009; Presta et al., 2009; Shireman, 2007). The timing and extent of the inflammatory response in ischemic muscle is very critical to the regulation of ischemic revascularization by FGF2.
To account for differences due to selective FGF2 isoform expression, inflammatory cell
(neutrophil and monocyte) density was evaluated in non-ischemic skeletal muscle. Loss of LMW
FGF2 (FGF2 HMW-only muscles) did not significantly change neutrophil or monocyte density
(Figure 42A and 43A). Similarly, ablation of the LMW isoform had no effect on basal neutrophil or monocyte density (Figure 42B and 43B). This suggests that the form of FGF2 present in non- injured tissues does not affect the inflammatory cell population. In the presence of HMW FGF2, the expected increase in neutrophil recruitment was observed in the ischemic limbs (Figure 42B).
This was unlike the WT muscles where neutrophil density was unchanged from the contralateral
(sham-operated limb) after 3 days of ischemia. LMW FGF2-only mice also had no difference in neutrophil infiltration in response to ischemia.
Several studies have shown increased neutrophil accumulation in WT skeletal muscle after
3 days of chronic injury (Contreras-Shannon et al., 2007; Martinez et al., 2010; Meisner et al.,
2014; Shireman et al., 2006). The divergence between the results presented here and these published reports may be attributed to the use of immunohistochemistry. The use of flow cytometry with multiple markers to detect and quantify neutrophil cells might prove to be more conclusive. This method has been applied in a number muscle injury studies to analyze resident
229
and circulating inflammatory cells and they show early, increased accumulation of neutrophils in
ischemic muscle (Capoccia et al., 2008; Contreras-Shannon et al., 2007; Martinez et al., 2010;
Tashiro et al., 2012). Secondary measures of histological staining using neutrophil markers like
myeloperoxidase (MPO), granulocyte-1 marker, or Ly-6 were also performed. Another reason for the discrepancy in the WT ischemic muscle may be timing of the tissue sampling (Oklu et al.,
2013). While some have suggested that neutrophils have a relatively short life span and are not detectable 1-2 days after induction of skeletal muscle injury (Meisner & Price, 2010; Tidball,
2011), others have observed neutrophil accumulation as late as 3 days in injured muscle
(Contreras-Shannon et al., 2007; Martinez et al., 2010; Uaesoontrachoon, Wasgewatte Wijesinghe,
Mackie, & Pagel, 2013). Furthermore, histological detection of robust neutrophil accumulation in ischemic muscle has been observed as late as 14 days after the ischemic surgery (Tashiro et al.,
2012). The lack of additional timepoints earlier than day 3 at 12h or 24h after ischemia is a limitation of this study. It is entirely possible that the peak of neutrophil density in the WT ischemic limbs occurs early and the day 3 results capture a moment when the inflammatory cell infiltrate switches over from a largely neutrophil population to monocytic one. This shift is very critical for the resolution of the inflammatory response and tissue healing as the persistence of neutrophils can further exacerbate the tissue damage by elevating oxidative stress and monocytes mediate the clearance of neutrophils.
Strong inflammatory cell infiltration has been observed during angiogenesis in chick embryonic CAM and subcutaneous Matrigel plug assays treated with recombinant LMW FGF2
(Andrés et al., 2009). An abundance of the cell infiltrate was identified to be positive for F4/80, a cell marker for monocytes. Similarly, Zittermann and Issekutz reported that FGF2 stimulated monocyte recruitment to dorsal skin sites as well synergistically enhanced the chemotactic
230
response to other proinflammatory cytokines (Zittermann & Issekutz, 2006a, 2006b). Consistent
with these studies, LMW FGF2-expressing limbs had elevated Mac-3-positive cell (monocytes)
densities in response to ischemia (Figure 43B). Monocyte recruitment was assessed in the ischemic
muscles at 3 or 7 days and increases were observed at both timepoints. HMW FGF2 expression
had no significant effect on monocyte infiltration until 7 days after the induction of ischemia. The
ability of LMW or HMW FGF2 to mediate chemotactic movement of neutrophils during ischemia
was similar and not changed from WT FGF2 expression. The pattern and timing of recruitment in
the WT tissues is consistent with several studies where neutrophil infiltration in ischemic muscle
was increased at 3 and 7 days (Contreras-Shannon et al., 2007; Martinez et al., 2010; Shireman et
al., 2007, 2006; Tashiro et al., 2012). Other studies reported declining monocytes levels starting at
7 days of ischemia (Capoccia et al., 2008; McClung et al., 2012). In cardiotoxin-induced muscle
injury, macrophage recruitment continued for 14 days after induction of the insult (Martinez et al.,
2010).
231
Figure 44: Schematic outlining the cellular pathways involved in the FGF2 isoform-mediated response to hindlimb ischemia.
Inhibit Activate Unknown interaction
232
Chapter 3: FGF2 isoforms in the regulation of skeletal muscle repair and regeneration
RESULTS III
Chronic ischemia induces an injurious phenotype in the vascular compartment of ischemic
limbs but also affects the viability of the muscles. Therefore, effective compensation for vascular
obstruction in PAD requires not only the promotion of vascular growth via angio- or arteriogenesis
but also the repair of ischemic-injured muscle (McClung et al., 2012; Scholz et al., 2002). Skeletal
muscle is capable of endogenous repair and regeneration in response to ischemic damage due to
the presence of a resident stem cell population known as satellite cells (Chang & Rudnicki, 2014;
Kang & Krauss, 2010; Le Grand & Rudnicki, 2007). These satellite cells, located between the
sarcolemma and basal lamina, are quiescent in resting adult muscle but become activated during
injury (Ceafalan et al., 2014). The myogenic repair program includes satellite cell activation,
proliferation, and ends with differentiation to myoblasts and fusion with existing tubes or to one
another to form new myofibers (Dumont, Wang, & Rudnicki, 2015; Rudnicki, Le Grand,
McKinnell, & Kuang, 2008; Yin, Price, & Rudnicki, 2013).
Satellite cell activation and subsequent differentiation to newly generated myofibers is
regulated by a family of transcription factors known as the myogenic regulatory factors (MRFs)
(Figure 45). These MRFs include the paired domain transcription factor, paired box 7 (Pax7) which
is expressed by all satellite cells, myogenic factor 5 (Myf5) and myoblast differentiation protein
(MyoD) which are highly expressed by activated satellite cells and some myoblasts, and myogenin
and muscle-specific regulatory factor 4 (MRF4) which are upregulated during differentiation of
myoblasts (Dumont et al., 2015; Karalaki, Fili, Philippou, & Koutsilieris, 2009; Yablonka-
Reuveni, Day, Vine, & Shefer, 2008; Yin et al., 2013). Traditional angiogenic factors like VEGF, angiopoietins, MMPs, or FGFs may potentially serve as responders to skeletal muscle injury by
233 modulating myogenesis (Doukas et al., 2002; McClung et al., 2015; Meisner et al., 2014; Rissanen et al., 2002; Yablonka-Reuveni et al., 2015).
234
Figure 45: Adult myogenesis is coordinated by a group of transcription factors (MRFs) that
regulate satellite cell activation, differentiation/self-renewal, and fusion. Satellite cells exit quiescence and become activated in response to muscle injury (exercise or ischemia). The satellite stem cells (Myf-) and satellite progenitor cells (Myf+) are characterized by their expression of Pax7. Proliferating satellite cells and their progeny are often referred to as myogenic precursor cells or adult myoblasts. Myoblasts express both MyoD and Myf5.
Following proliferation, myoblasts begin differentiation by downregulating Pax7. Terminal differentiation and fusion begins with the expression of Myogenin, which together with
MyoD activate muscle specific structural and contractile genes. MRF4 is further required for hypertrophy of the new fibers. During regeneration, activated satellite cells have the capability to return to quiescence to maintain the satellite cell pool. Reprinted with permission from Elsevier (Le Grand & Rudnicki, 2007).
235
Absence of LMW FGF2 influences Pax7 expression during ischemia
To determine the gene expression levels of the MRFs, RNA was isolated from non-
ischemic WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only skeletal tissues. First strand cDNA was amplified by qRT-PCR using primers for the MRFs (Pax7, Myf5, MyoD, Myogenin and
MRF4) and an internal control gene (18S). Pax7 and Myf5 mRNA expression were both unchanged
in Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only muscles relative to non-ischemic WTs
(Figures 46A and 47A).
To characterize the effect of FGF2 isoform and ischemia on Pax7 and Myf5 expression, mRNA expression was analyzed at 1, 3, 7 and 14 days the induction of ischemia. Transcript levels were presented as a ratio of sham muscles to address any changes related to the hindlimb ischemia surgery. After 3 days of ischemia, Pax7 expression in all groups was decreased from day 1 and increasing by day 7 (Figure 46B). Expression was not changed from WT in the absence of all
FGF2 (Fgf2-/-) and in the absence of the HMW isoforms (FGF2 LMW-only) at all timepoints of ischemia. In the absence of LMW FGF2 (FGF2 HMW-only), Pax7 expression was significantly increased after 14 days of ischemia (p<0.05 vs. WT; Figure 46B).
WT and Fgf2-/- ischemic muscles had a similar profile of Myf5 expression during ischemia where expression after 3 days was increased from day 1 and decreased at after 7 days and returned to normal levels at day 14 (Figure 47B). Ischemic skeletal muscle from FGF2 LMW-only and
FGF2 HMW-only also shared a similar pattern where day 3 expression was decreased from day 1 and increasing by day 7 and returned to normal after 14 days (Figure 47B). For all mouse groups, no significant changes in Myf5 expression were observed relative to WTs.
236
Pax7 expression (non-ischemic muscle) A. 2 WT Fgf2-/- FGF2 LMW-only 1.5 FGF2 HMW-only
1 (arbitrary units) (arbitrary
0.5 Relative mRNA expression expression mRNA Relative
0
Pax7 expression (ischemic muscle) B. 6
WT Fgf2-/- 5 FGF2 LMW-only FGF2 HMW-only # * 4
3
(arbitrary units) (arbitrary 2
1 mRNA Expression relative sham to relative Expression mRNA
† † 0 1 3 7 14 Days after ischemia surgery 237
Figure 46. Pax7 mRNA expression in non-ischemic, sham and ischemic skeletal muscles. (A)
Quantitative RT-PCR analysis for Pax7 gene expression in non-ischemic skeletal muscle relative to WT (white bar), Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar) or
FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes
relative to WT non-ischemic muscles (B) Time course of Pax7 mRNA expression. For each
timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA expression in ischemic
muscles are expressed relative to corresponding sham expression levels in WT (blue),
Fgf2-/- (red), FGF2 LMW-only (green), or FGF2 HMW-only (black) limbs. Data are
presented as mean ± SEM. n=4 for each mouse group at each timepoint. *p<0.05 vs. Day 1
cohort, #p< 0.05 vs. WT, †p<0.05 vs. FGF2 HMW-only.
238
A. Myf5 expression (non-ischemic muscle)
2 WT Fgf2-/- FGF2 LMW-only 1.5 FGF2 HMW-only
1 (arbitrary units) (arbitrary 0.5 Relative mRNA expression expression mRNA Relative
0
B. Myf5 expression (ischemic muscle) 12 WT Fgf2-/- FGF2 LMW-only 10 FGF2 HMW-only
8
6
4 (arbitrary units) (arbitrary
2 mRNA Expression relative sham to relative Expression mRNA
0 239 1 3 7 14
Days after ischemia surgery Figure 47. Myf5 mRNA expression in non-ischemic, sham and ischemic skeletal muscles. (A)
Quantitative RT-PCR analysis for Myf5 gene expression in non-ischemic skeletal muscle
relative to WT (white bar), Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar) or
FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes
relative to WT non-ischemic muscles (B) Time course of Myf5 mRNA expression. For each timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA expression in ischemic muscles are expressed relative to corresponding sham expression levels in WT (blue),
Fgf2-/- (red), FGF2 LMW-only (green), or FGF2 HMW-only (black) limbs. Data are presented as mean ± SEM. n=4 for each mouse group at each timepoint.
240
Loss of HMW FGF2 expression decreases basal levels of MyoD
MyoD and myogenin expression levels in non-ischemic WT, Fgf2-/-, FGF2 LMW-only and
FGF2 HMW-only skeletal muscles were also examined. MyoD mRNA expression was decreased
in both Fgf2-/- and FGF2 HMW-only mice (p<0.05 vs WT) while FGF2 LMW-only expression
was unchanged relative to non-ischemic WT (Figure 48A). Myogenin expression was unchanged
from WT in Fgf2-/-,FGF2 LMW-only or FGF2 HMW-only muscles (Figure 49A)
To describe the effect of specific FGF2 isoform expression and hindlimb ischemia on
MyoD and myogenin transcript levels, sham and ischemic muscles were isolated and analyzed for
MyoD and myogenin expression. In Fgf2-/-, FGF2 LMW-only, and FGF2 HMW-only limbs, no changes from WT expression levels of either MyoD (Figure 48B) or myogenin (Figure 49B) were detected at 1, 3, 7 or 14 days of ischemia.
A. MyoD expression (non-ischemic muscle)
2 WT Fgf2-/- FGF2 LMW-only 1.5 FGF2 HMW-only
1 # (arbitrary units) (arbitrary
0.5 # Relative mRNA expression expression mRNA Relative
241 0 B. MyoD expression (ischemic muscle) WT 9 Fgf2-/- FGF2 LMW-only 8 FGF2 HMW-only 7
6
5
4 (arbitrary units) (arbitrary 3
2 mRNA Expression relative sham to relative Expression mRNA 1
0 1 3 7 14 Days after ischemia surgery
Figure 48. MyoD mRNA expression in non-ischemic, sham and ischemic skeletal muscles.
(A) Quantitative RT-PCR analysis for MyoD gene expression in non-ischemic skeletal muscle
relative to WT (white bar), Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar) or
FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes
relative to WT non-ischemic muscles (B) Time course of MyoD mRNA expression. For each
timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA expression in ischemic
muscles are expressed relative to corresponding sham expression levels in WT (blue),
Fgf2-/- (red), FGF2 LMW-only (green), or FGF2 HMW-only (black) limbs. Data are
presented as mean ± SEM. n=4 for each mouse group at each timepoint. #p< 0.05 vs. WT.
242
Myogenin expression (non-ischemic muscle)
A. 2 WT Fgf2-/- 1.5 FGF2 LMW-only FGF2 HMW-only
1
(arbitrary units) (arbitrary 0.5 Relative mRNA expression expression mRNA Relative
0
B. Myogenin expression (ischemic muscle) 8 WT 7 Fgf2-/- FGF2 LMW-only 6 FGF2 HMW-only
5
4
3 (arbitrary units) (arbitrary
2
mRNA Expression relative sham to relative Expression mRNA 1
0 1 3 7 14 Days after ischemia surgery 243
Figure 49. Myogenin mRNA expression in non-ischemic, sham and ischemic skeletal muscles.
(A) Quantitative RT-PCR analysis for Myogenin gene expression in non-ischemic skeletal muscle relative to WT (white bar), Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar) or FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes
relative to WT non-ischemic muscles (B) Time course of Myogenin mRNA expression. For
each timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA expression in ischemic
muscles are expressed relative to corresponding sham expression levels in WT (blue),
Fgf2-/- (red), FGF2 LMW-only (green), or FGF2 HMW-only (black) limbs. Data are presented as mean ± SEM. n=4 for each mouse group at each timepoint.
244
HMW FGF2 upregulates MRF4 expression in response to ischemia
MRF4 expression in non-ischemic WT, Fgf2-/-, FGF2 LMW-only and FGF2 HMW-only skeletal muscles was also examined. For all mouse groups, MRF4 mRNA expression was not different from WT (Figure 50A).
To determine the role of FGF2 isoform expression on MRF4 gene expression in response to ischemia, transcript levels were measured at 1, 3, 7 or 14 days of ischemia. Ischemic WT,
Fgf2-/-, and FGF2 LMW-only limbs had a similar pattern of expression in which MRF4 levels were significantly decreased by 3 days, increased at day 7 and returned to normal by day day14
(Figure 50B). Expression at 3 days of ischemia was significantly elevated in FGF2 HMW-only muscles compared to WT (p<0.05, Figure 50B). Furthermore, the pattern of expression over time in FGF2 HMW-only muscles differed from the other groups with highest expression of MRF4 occurring on 3 days of ischemia, decreasing levels by 7 days and a return to normal expression at
14 days.
Overall, the MRF expression data suggest that HMW FGF2 is important for maintaining a population in ischemic skeletal muscle. It also enhances the progression of the activated satellite cells to differentiated myoblasts and incorporation into repairing/regenerating muscle.
245
A. MRF4 expression (non-ischemic muscle) 2
WT Fgf2-/- 1.5 FGF2 LMW-only FGF2 HMW-only
1 (arbitrary units) (arbitrary Relative mRNA expression expression mRNA Relative 0.5
0 MRF4 expression (ischemic muscle) B. 5 WT Fgf2-/- FGF2 LMW-only 4 FGF2 HMW-only
3 # ‡
2 (arbitrary units) (arbitrary
1 mRNA Expression relative sham to relative Expression mRNA *
0 1 3 7 14 246 Days after ischemia surgery Figure 50. MRF4 mRNA expression in non-ischemic, sham and ischemic skeletal muscles.
(A) Quantitative RT-PCR analysis for MRF4 gene expression in non-ischemic skeletal muscle relative to WT (white bar), Fgf2-/- (light gray bar), FGF2 LMW-only (dark gray bar) or FGF2 HMW-only (black bar) transcript expression levels are represented as fold-changes relative to WT non-ischemic muscles (B) Time course of MRF4 mRNA expression. For each timepoint of ischemia (1, 3, 7, or 14 days), fold changes of mRNA expression in ischemic muscles are expressed relative to corresponding sham expression levels in WT (blue),
Fgf2-/- (red), FGF2 LMW-only (green), or FGF2 HMW-only (black) limbs. Data are presented as mean ± SEM. n=4 for each mouse group at each timepoint. *p<0.05 vs. Day 7 cohorts, #p<0.05 vs. WT, ‡p<0.05 vs. FGF2 LMW-only.
247
DISCUSSION III
The results presented in this chapter of the dissertation demonstrate that the protection of skeletal muscle from ischemia-induced damage by the endogenous HMW FGF2 isoforms is possibly mediated by muscle regeneration pathways (Figure 51).
Satellite cell activation, expansion and subsequent differentiation during embryogenesis or myogenic repair is tightly regulated by the Pax genes and the myogenic regulatory factors (MRFs)
(Buckingham & Relaix, 2007; Kang & Krauss, 2010; P. Zhao & Hoffman, 2004). To determine the role of the FGF2 isoforms in the regeneration potential of skeletal muscle during ischemia, the expression of the myogenic markers was determined in normal or ischemic muscles. The Pax genes, Pax3 and Pax7 are expressed by satellite cells (Chang & Rudnicki, 2014). Pax3 expression was not detected in any of the muscles analyzed. Pax3 is critical for embryonic myogenesis but is down-regulated in muscle before or at birth (Buckingham & Relaix, 2007; Rudnicki et al., 2008).
Pax3-/- mice die in utero (Buckingham & Relaix, 2007). Pax7 is dispensable for embryonic myogenesis but is essential after birth and is expressed by all quiescent satellite cells. Pax7-/- mice have complete loss of satellite cell pool, smaller-sized muscles and impaired skeletal muscle regeneration (Sambasivan et al., 2011; von Maltzahn, Jones, Parks, & Rudnicki, 2013). Pax7 was detected in FGF2 LMW-only and FGF2 HMW-only muscles and its expression was unchanged from WT in non-ischemic muscles (Figure 46A). This confirms that selective FGF2 isoform expression does not affect the satellite cell population. In response to ischemia, Pax7 expression in WT or LMW FGF2 muscles decreased from day 1 to undetectable levels at day 3 and was returned to normal level at day 14 (Figure 46B). In the presence of only HMW FGF2 however, the expression of Pax7 was elevated over four-fold. The population of Pax7-expressing cells at 14 days of ischemia represents satellite cell numbers in recovering muscle after cycles of self-renewal
248
and myogenic differentiation. This is possibly the true myogenic potential of the HMW FGF2
muscles which was enhanced by the presence of ischemia and muscle injury. Downstream of the
Pax transcription factors are the MRFs which are required for the commitment, progression and
differentiation of proliferating satellite cells and their progeny (myoblasts) into mature myofibers
(Kang & Krauss, 2010; Rudnicki et al., 2008; Yin et al., 2013). Pax3/Pax7 induce the expression
of two MRFS, Myf5 and MyoD by direct interaction with distal enhancer elements and proximal promoters of both genes.
Myf5 is another early myogenic marker expressed by satellite progenitor cells which also express Pax7 (Ceafalan et al., 2014; Yin et al., 2013). Expression of only LMW FGF2 or only
HMW FGF2 did not affect normal Myf5 expression (Figure 47A). After induction of ischemia,
Myf5 expression follows the pattern of Pax7 expression (Figure 47B). The increased myogenic potential of the HMW FGF2 muscles is evident as Myf5 expression is elevated and at a maximum by day 14. The addition of Myf5 expression marks early commitment of the activated satellite cells to the myogenic lineage; these cells leave the basal lamina niche and will eventually undergo differentiation (Cooper et al., 1999; Gayraud-Morel et al., 2007). Myf5 regulates the proliferation rate and homeostasis of satellite cells and is expressed by activated, proliferating myoblasts
(Gayraud-Morel et al., 2007; Rudnicki et al., 2008).
Similar to Myf5, MyoD expression is up-regulated in activated cells and expressed by proliferating myoblasts. Both genes have been termed myogenic determination factors (Cooper et al., 1999; Yin et al., 2013). HMW FGF2 lowered MyoD expression in non-ischemic muscle while
LMW FGF2 had no effect (Figure 48A). During ischemia, the MyoD is required for the differentiation potential of satellite cells. MyoD-/- mice have reduced muscle mass, myoblasts which continually self-renew, are differentiation-defective and do not fuse into myotubes
249
(Cornelison, Olwin, Rudnicki, & Wold, 2000; Le Grand & Rudnicki, 2007). The reduced MyoD
expression in normal muscle may be of limited importance as the absolute expression of MyoD in
myoblasts is less relevant than the Pax7-to-MyoD ratio. A high ratio is observed in non-activated
satellite cells and keep them in their quiescent state and cells with a low Pax7-to-MyoD ratio begin
to differentiate (Montarras, L’honoré, & Buckingham, 2013; Yin et al., 2013). This is evident in the ischemic expression of MyoD which is highest at Day 3 and coincides with the timepoint of lowest Pax7 expression (Figure 48B). The Pax7-to-MyoD ratio has shown to decrease further after activation of Myogenin (Chang & Rudnicki, 2014).
After limited rounds of proliferation, myoblasts withdraw from the cell cycle and terminal
differentiation is initiated by the expression of Myogenin and MRF4 which is prompted by the
transcriptional activity of MyoD (Kang & Krauss, 2010; Yin et al., 2013). Neither Myogenin nor
MRF4 was affected by expression of only HMW FGF2 or only LMW FGF2 in non-ischemic
muscle (Figures 47A and 48A). Myogenin and MRF4 drive the expression of several muscle-
specific structural and contractile proteins including sarcomeric myosin. The differentiating
myoblasts fuse to form elongated multinucleated myotubes and nascent myofibers. MRF4 also
appears to be necessary for hypertrophy of the new fibers. It is the only MRF that is expressed to a
high degree in mature myofibers and its increased expression is proposed to be associated with
downregulation of Myogenin (Gayraud-Morel et al., 2007; Yablonka-Reuveni, 2011). During
ischemia, Myogenin expression peaked at day 7 with a non-significant elevation at day 3 in the
HMW FGF2-expressing cells (Figure 49B). Ischemic expression of MRF4 followed a similar
pattern as Myogenin expression except for HMW FGF2 muscles whose expression peaked after 3
days and continued to decline (Figure 50B). These results suggest that the rate of myoblast
differentiation is much faster and more pronounced in the presence of HMW FGF2. This increased
250 myogenic potential of HMW FGF2 may underlie the complete protection from toe/limb necrosis observed in the FGF2 HMW-only mice as well as the rapid recovery from limb use impairment
(as discussed in Chapter 1). Another FGF, FGF6 plays an important in muscle repair and regeneration. Like FGF2, FGF6 is capable of stimulating rat satellite cell proliferation in vitro
(Sheehan & Allen, 1999). Fgf6-/- mice display impaired muscle regeneration linked to reduced numbers MyoD and Myogenin-expressing myoblasts and decreased migratory ability of the existing myoblasts (Floss et al., 1997; Neuhaus et al., 2003). The degree of migratory capability was further reduced in satellite cells-derived myoblasts isolated from Fgf2/Fgf6 double knockout mice (Neuhaus et al., 2003).
An important limitation to note is that the analysis of MRF gene expression was performed in whole muscle. This analysis while extremely useful has the potential to dilute the results and make subtle differences in expression of the transcriptions factors difficult to detect (Danoviz &
Yablonka-Reuveni, 2012; Yablonka-Reuveni et al., 2015). Satellite cells isolated from muscle tissue would be a useful alternative and may further highlight the effects of the HMW FGF2 isoforms on ischemia-induced muscle regeneration.
251
Figure 51. Schematic representation of the muscle regeneration pathway that may mediate the HMW FGF2 protection of ischemic muscle.
Inhibit
Activate Unknown interaction
252
CONCLUSIONS AND CLINICAL RELEVANCE
This dissertation provides significant advances in the understanding of the FGF2’s biological functions in vascular biology, in particular and for the first time, the in vivo actions of the HMW protein isoforms of FGF2. The results presented here unequivocally demonstrate that the HMW FGF2 isoforms are necessary and sufficient for inducing post-ischemic vascular growth, protecting skeletal muscle from damage and preserving functional capacity. The LMW isoform, conversely, was shown to have no effect on tissue survival during chronic hindlimb ischemia.
Furthermore, HMW FGF2 expression alone induced growth of capillaries (angiogenesis), small and medium-sized collateral vessels (arteriogenesis). The pathways which via which HMW FGF2 enhanced salvage of ischemic muscle were independent of FGF receptor activation and were associated with regulation of growth factor expression including FGF2, and other ischemia- triggered signals such as inflammation and myogenesis. Taken together, the findings presented here suggest a confluence of vascular remodeling and tissue repair/regeneration in HMW FGF2- induced recovery from chronic ischemia (Figure 52).
Since the discovery of FGF2 in 1984 as an angiogenic growth factor, only a handful of studies have examined the biological roles of the FGF2 isoforms as well as treated the two classes of FGF2 proteins as separate entities. The genesis of the isoform-specific knockout mouse models prompted studies that have started to show that there are differences between the FGF2 proteins
(Azhar et al., 2009). Three groups, in particular, have focused on researching the in vivo roles of the endogenous HMW isoforms distinct from LMW FGF2 in bone development and homeostasis
(Xiao et al., 2009, 2014, 2013), neuronal differentiation and degeneration (Claus, Bruns, & Grothe,
2004; Grothe, Schulze, et al., 2000; Haastert et al., 2006; Timmer et al., 2004) and cardiovascular pathophysiology (Liao et al., 2007, 2009, 2010; Manning et al., 2013). Previous studies conducted
253
by our laboratory have determined that the LMW isoform and the HMW isoforms have different
roles in acute ischemia-reperfusion injury and cardioprotection (Liao et al., 2007, 2009, 2010).
This dissertation offers, for the first time, important insights into the in vivo “angiogenic” functions
of the endogenous FGF2 isoforms, particularly the HMW isoforms which were previously
considered to act in a similar if not redundant fashion as LMW FGF2. The mitogenic effects of
HMW FGF2 in vascular biology have largely focused on the use of recombinant exogenous HMW
FGF2 in transformed and/or immortalized cells. While the later can provide valuable information
on the function of proteins, in vitro cell activity does not always depict the entire picture of the
protein's functions. This is the first study to identify the vascular effects of HMW FGF2 in vivo
and to demonstrate that the HMW isoforms are sufficient for ischemia-induced vascular growth,
both angiogenesis and arteriogenesis.
There is a large body of pre-clinical and clinical studies focusing on the therapeutic
potential of exogenous LMW FGF2 as described in the Introductory section (Tables 4-6). In summary, the multi-species animal studies provided adequate proof-of-concept data which were not translated into beneficial results in several randomized controlled trials. In this dissertation, the response to chronic ischemia included induction of angiogenesis (capillary growth) in WT,
Fgf2-/- and FGF2 HMW-only muscles as well as FGF2 LMW-only muscle. The extent of capillary growth was greater in the HMW FGF2-expressing limbs while Fgf2-/- mice were similar to WT.
FGF2 LMW-only ischemic muscles had no change in capillary number after 42 days of ischemia.
These results reveal antithetical functions of HMW and LMW FGF2 in vascular growth because ablation of both classes of FGF2 isoforms (LMW and HMW FGF2) in the Fgf2-/- mice has a milder phenotypic effect than is present in tissues expressing only LMW or only HMW FGF2.
Additionally, the absence of an angiogenic response in LMW FGF2-only mice suggests that this
254 endogenous isoform of FGF2 may exert an inhibitory effect on the angiogenic potential of HMW
FGF2 (in FGF2 LMW-only and WT muscles). This calls into question or, at the very least, warrants a re-examination of the previously identified role for endogenous LMW FGF2 as a vascular growth factor. Most important, however, is the clinical potential for the HMW isoforms of FGF2 to serve as robust inducers of revascularization in response chronic ischemia.
The FGF2 HMW-only muscles were protected from the toe/foot necrosis that was observed in WT, Fgf2-/-, and FGF2 LMW-only mice to an equal degree. Limb use impairment was also ameliorated in FGF2 HMW-only limbs and the recovery of normal function was faster than WT limbs. Recovery was present in LMW FGF2-expressing mice but occurred at a significantly slower rate (3x) than WT and FGF2 HMW-only. Fgf2-/- limbs did not recover full use of ischemic limbs.
Although both classes of FGF2 isoforms are capable of protecting skeletal muscle from ischemic injury, HMW FGF2 appears to be more efficient at this preserving limb viability. This is important because the salvage of limbs in animal models of chronic ischemia is used as a surrogate for a predictor of clinical success in humans (healing of lower extremity ulceration) (Cooke & Losordo,
2015; Dragneva et al., 2013; Waters et al., 2004).
In line with the bulk of the in vivo FGF2 studies, characterization of the downstream signaling mediated in response to FGF2 have focused on the cell surface activation of FGFRs by secreted or ECM-bound/released FGF2. Evidence from our laboratory and others has clearly shown that while LMW FGF2 is secreted during cellular stress, HMW FGF2 is largely confined to the nucleus (Chlebova et al., 2009; Jiang et al., 2009; Liao et al., 2009, 2010; Yu et al., 2007).
This localization of HMW FGF2 is critical for the cellular responses elicited by the protein isoform as evidenced when exogenously administered HMW FGF2 produces similar effects as LMW
FGF2 in cultured cells but transgenes expressing only the HMW isoforms produce differential
255
effects from LMW FGF2 expressing cells (Jiang et al., 2007; Kardami et al., 2007; Liao et al.,
2010). The absence of changes in FGFR activation in HMW FGF2-induced revascularization of
ischemic muscles suggests that the effects of this endogenous FGF2 isoform are intracrine in
nature. The nuclear localization of HMW FGF2, which is important for its protective effect in
chronic ischemia, adds a layer of complexity when considering the development of a HMW FGF2
drug. The HMW FGF2 therapeutic would have to actively get across two barriers, the plasma
membrane and nuclear envelop. Cell penetrating peptides like synthetic polyarginine or HIV-1 Tat
DNA binding domain have been successfully utilized to transport molecules into cells by an unknown mechanism (Biswas & Torchilin, 2014; Sakhrani & Padh, 2013). The inherent nuclear localization signals present on HMW FGF2 could be enhanced to ensure targeted delivery to the nucleus (Liao et al., 2009). Small peptides from viruses also that confer nuclear localization, such as the KKKRKV peptide from SV40 large T antigen may also be utilized to deliver the drug to the nucleus (Biswas & Torchilin, 2014).
Decades after the introduction of therapeutic revascularization as a potential treatment for coronary and peripheral artery disease, this intervention has yet to be successfully translated to the clinic and the need for alternative treatments persists. The research presented in this dissertation identifies a prospective new player in the field of therapeutic revascularization that has the
potential to be beneficial for patients suffering from critical limb and cardiac ischemia.
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Figure 52. Schematic representing the hypothetical pathways activated and/or inhibited by
HMW FGF2 during adaptive revascularization and protection from ischemic muscle injury
Inhibit Activate
Unknown interaction
257
FUTURE DIRECTIONS
The studies presented herein suggest distinct roles for HMW and LMW FGF2 in regulating postnatal vessel growth/remodeling during ischemia. There remains, however, a number of important questions that still need to be addressed to better determine the roles of HMW and LMW
FGF2 in vascular growth and ischemic injury recovery. These studies may prove useful for the understanding of the molecular mechanisms by which the FGF2 isoforms regulates vascular development and promotes tissue repair in ischemic muscle as well as the potential development of HMW FGF2 as a new application for the treatment of ischemic vascular diseases. They include:
1) Role of FGF2 isoforms in muscle functional recovery: This dissertation showed that the
HMW isoforms accelerated complete functional recovery of ischemic limbs. This was
evaluated via an index of muscle function; ischemic hindlimb use scoring (Chalothorn et al.,
2007; Faber et al., 2011; Helisch et al., 2006; Peng et al., 2011). A different index of muscle
functional capacity, exercise performance measured with a treadmill endurance test, more
closely recapitulates the desired primary outcomes of PAD and CAD clinical trials (Dragneva
et al., 2013; Haas et al., 2012; Lavu, Gundewar, & Lefer, 2011). A preliminary study of this
endpoint in ischemic mice was reported in Chapter 1 and trends towards decreased functional
capacity were observed in the absence of all FGF2 expression (Fgf2-/-). In order to
conclusively determine the effect of selective FGF2 isoform expression on post-ischemic
exercise tolerance, increased sample sizes and the inclusion of additional timepoints before
and after 14 days of ischemia are necessary. A more robust examination of exercise
performance would be important for confirmation of these preliminary results and as a
secondary measure of muscle function recovery.
258
2) Role of vascular cell proliferation, and migration in the FGF2 isoform-specific response
to chronic ischemia: This dissertation showed that while HMW FGF2 augments ischemia-
induced capillary growth, LMW FGF2 inhibited angiogenesis in ischemic muscles.
Angiogenesis and arteriogenesis during embryonic development or in response to ischemia
requires the activation, proliferation, and migration of endothelial cells (ECs) and/or vascular
smooth muscle cells (vSMCs) (Carmeliet, 2000; Fischer et al., 2006). Differences in the
angiogenic capacity of HMW or LMW FGF2-expressing vascular cells may contribute to the
observed differences in FGF2 isoform stimulation of vessel growth To address this, ECs or
vSMCs isolated from FGF2 isoform knockout mice could be evaluated for proliferation and
migration using nanofiber peptide scaffold assays (an in vitro model of vascularity) (Cho et
al., 2012; Narmoneva et al., 2005). Additionally, in vivo assessment of microvascular cell
proliferation could be performed with immunohistochemical detection of proliferation markers
along with and EC or vSMC-specific markers can be evaluated.
3) Role of FGF2 isoforms in ischemia-induced myocyte death: An interesting question raised
by this dissertation research is whether the improvement in skeletal muscle functional recovery
and prevention of limb necrosis observed when only HMW FGF2 is expressed is related to the
degree of myocyte survival and/or regeneration. In ischemic muscle injury, myocyte death (via
necrosis or apoptosis) is followed by inflammatory cell accumulation and regeneration (Tidball
& Villalta, 2010; Tidball, 2011). The role of the FGF2 isoforms in the determining of the
degree myocyte survival and necrosis during chronic ischemia is still unknown.
4) Role of FGF2 isoforms in ischemia-induced myocyte regeneration: Preliminary results
presented in Chapter 2 of this dissertation suggest that the HMW isoforms have a greater
capacity for myocyte regeneration. Specifically, HMW FGF2 increased expression of satellite
259
cell activation and myoblast differentiation markers. These results were obtained from whole
muscle gene expression which is a limitation. Determination of myogenic gene expression in
satellite cells and myoblasts isolated from isoform knockout muscle tissue would confirm these
initial results and may further highlight the effects of the HMW FGF2 isoforms on ischemia-
induced muscle regeneration (Fukada et al., 2013; Sheehan & Allen, 1999; Yablonka-Reuveni
et al., 2015). In murine muscle, the large lobular nuclei of regenerating myofibers are centrally
located while healthy mature fibers have small peripheral nuclei (Paoni et al., 2002; Scharner
& Zammit, 2011). Histological analysis of this morphological marker of ongoing muscle
regeneration over time would allow for determination of the timing and extent of ischemic
muscle repair regulated by the FGF2 isoforms.
5) Role of inflammatory cell infiltration: Neutrophils and macrophages have been shown to
infiltrate ischemic muscle in hindlimb ischemia models (G. Brevetti, Giugliano, Brevetti, &
Hiatt, 2010; Silvestre et al., 2008; Tidball, 2005) and modulate the angiogenic/arteriogenic
response (Fung & Helisch, 2012; Naldini & Carraro, 2005; Silvestre et al., 2008). Preliminary
results presented here suggest that the total ablation of Fgf2 gene does have some effect on
neutrophil and macrophages infiltration after ischemia. Less clear, however, is the precise role
of the inflammatory microenvironment in FGF2 isoform-specific revascularization. The
absence of data on inflammatory cell infiltration between 0 and 3 days after ischemia is a
limitation. Neutrophil infiltration begins within a few hours of ischemic muscle injury and is
typically cleared by 3 days when monocyte infiltration starts to increase (Nguyen & Tidball,
2003; Tidball, 2011). Determination of the degree of inflammation as early as hours after the
induction of ischemia, the timing of resolution of this inflammation, and the subset of
inflammatory cells involved will provide useful information on their contribution to FGF2
260
isoform-induced vascular growth. As in Chapter 2, ischemic muscle would be evaluated for
immunohistochemical detection of myeloperoxidase (MPO) and Mac-3, markers for
neutrophils and macrophages, respectively. Additionally, the role of inflammation in ischemic-
induced revascularization could be examined by antibody depletion, genetic or
pharmacological ablation of specific inflammatory cell populations.
6) Role of FGFR signaling: Results presented in Chapter 2 of this dissertation suggest that FGFR
signaling is not associated with HMW FGF2-induced revascularization. To confirm the
relationship between the HMW isoforms and FGFR signaling in ischemia-induced
revascularization, the hindlimb ischemia studies (as described in Chapter 1) should be repeated
in the presence of FGFR inhibitors (Liao et al., 2010). Furthermore, it would be important to
determine the role of nuclear FGFR signaling as distinct from cell surface FGFR signaling in
HMW FGF2-mediated response to chronic ischemia.
7) Involvement of HSPG signaling in FGF2 isoform-induced vascular growth: In addition to
their roles as co-receptors for FGF-FGFR binding and cell signaling, heparan sulfate
proteoglycans (HSPGs) are also capable of FGF ligand binding and signaling independent of
FGFR interactions (Murakami et al., 2008; Sarrazin et al., 2011). Syndecans are trans-
membrane HSPGs that regulate cell adhesion, proliferation, and migration in inflammation,
wound healing and angiogenesis (Couchman, Gopal, Lim, Nørgaard, & Multhaupt, 2015;
Murakami et al., 2008). The most widely studied syndecan, syndecan-4 is ubiquitously
expressed in many cell types including myofibroblasts, epithelial, endothelial, and smooth
muscle cells. Syndecan-4 null mice while viable, have defects in inflammatory responses to
wound healing and lipopolysaccharide injection. The role of syndecan-4 and other HSPGs in
FGF2 isoforms-mediated angiogenesis is not yet understood.
261
8) Potential function of HMW FGF2 as a transcription factor: As mentioned above,
preliminary results suggest that HMW FGF2-induced revascularization does not correlate with
activation of FGFR signaling. This fact coupled with the exclusive nuclear localization of
HMW FGF2 is suggestive of a role for the HMW isoforms as a direct modulator or co-factor
of gene transcription. Identification of DNA binding sites and other motifs present on the FGF2
isoforms may serve as protein-DNA or protein-protein interactions (interactome) and could
highlight putative targets of HMW FGF2 that mediate the vascular response to ischemia. These
interactomes can be detected by chromatin immunoprecipitation followed by sequencing
(ChIP–seq) assays. Confirmed partners of HMW FGF2 in other cell systems include the
survival of motor neurons protein (SMN), antiapoptotic protein 5 (Api5), and the ribosomal
protein L6/TAXREB 107 (Chlebova et al., 2009; Förthmann et al., 2015). In neuronal cells,
HMW FGF2 has been shown to interact with SMN and Api5 both anti-apoptotic proteins, to
potentiate cell survival and differentiation (Berghe et al., 2000; Claus, Bruns, et al., 2004;
Woodbury & Ikezu, 2013). It would be important to test if these interactions persist in and if
they promote HMW-FGF2 induced protection of myocytes and/or vascular cells.
9) Role of HMW FGF2 in other models of tissue injury: The chronic ischemia model described
in this dissertation has shown that HMW FGF2 is a promoter of functional recovery of
ischemic tissues. It would be interesting to explore if this protection extends to other chronic
ischemia models in the heart or brain. Studies are currently underway to determine the role of
HMW FGF2 isoforms in revascularization and protection from cardiac dysfunction in hearts
subjected to permanent LAD ligation to induce chronic myocardial ischemia.
10) Role of HMW FGF2 in models of ischemia with underlying compromised vascular
biology: Patients with CAD or PAD also have other risk factors including advanced age,
262
diabetes, endothelial dysfunction or stress, and hypercholesteremia that abrogate physiological
vascular growth (Dragneva et al., 2013; Haas et al., 2012). Models with forms of these human
risk factors would better mimic human ischemic disease and further characterize the
“angiogenic” efficacy of the HMW FGF2 isoforms. This can be done by crossing the FGF2
HMW-only mice with diet-induced or genetic models of diabetes or hyperlipidemia.
Additionally, the ischemia experiments could be performed in older HMW FGF2-expressing
mice.
11) Role of HMW FGF2 in tumor growth: In addition to their functions in vascular remodeling,
the FGF2 isoforms are implicated in tumor progression and metastasis. This could potentially
affect the use of the HMW isoforms as a revascularization therapeutic agent (Thomas-Mudge
et al., 2004). HMW FGF2 overexpression induces radioresistance in NIH3T3 fibroblasts and
HeLa cells (Cohen-Jonathan et al., 1997; Delrieu et al., 1998). The NIH3T3 cells had a
transformed morphology and induced tumor formation in nude mice (Quarto et al., 1991).
Conversely, bovine aortic ECs (BAECs) overexpressing HMW FGF2 had no effect on tumor
growth in nude mice but tumorigenesis did occur when all the FGF2 isoforms (LMW and
HMW) were overexpressed or when only the LMW isoform was overexpressed. Overall, these
studies indicate a potential role for the FGF2 isoforms in cancer development. These actions
of the FGF2 isoforms need to be taken into consideration if HMW FGF2 is to be used clinically
as a revascularization agent. Clinical trials in patients of PAD and CAD that received
recombinant LMW FGF2 did not report any malignancies that correlated with receipt of the
drug (Khurana & Simons, 2003; Lederman et al., 2002; Simons et al., 2002).
263
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