Investigating the Role of Myh10 in the Epicardium: Insights from the EHC Mouse 2016 Liam A. Ridge

Investigating the Role of Myh10 in the Epicardium: Insights from the EHC Mouse

A thesis submitted to the University of Manchester for the degree of Doctor of Philosophy in the Faculty of Biology, Medicine and Health

2016

Liam A. Ridge

School of Biological Sciences

Division of Evolution & Genomic Sciences

1 List of Contents List of Contents ...... 2 List of Figures ...... 6 List of Tables ...... 8 List of Abbreviations ...... 9 Abstract ...... 15 Declaration ...... 16 Copyright Statement ...... 16 Acknowledgements ...... 17 Preface ...... 18 Chapter 1: Introduction ...... 19 1.1 Why Investigate Cardiac Development? – The Heart is the Problem ...... 19 1.2 Overview of Cardiogenesis in the Mouse Embryo ...... 21 1.2.1 Origins and Morphological Overview of Mammalian Heart Formation ...... 21 1.2.2 Cardiac Maturation and the Contribution of Extra-Cardiac Cell Populations .... 25 1.2.2.1 Outflow Tract Development ...... 25 1.2.2.2 Myocardial Proliferation and Ventricular Trabeculation...... 27 1.2.2.3 Development of the Coronary Vasculature ...... 30 1.3 Brief Overview of the Formation and Function of the Epicardium ...... 32 1.4 Epicardial Potential in Regenerative Medicine ...... 35 1.5 Myh10 is Essential for Correct Cardiogenesis ...... 38 1.5.1 Introduction to Non-Muscle Myosin IIB ...... 38 1.5.2 Role of NMIIB in Cell Migration ...... 41 1.5.3 Role of NMIIB in Cell Adhesion ...... 42 1.5.4 Role of NMIIB in Cell Polarity and Cell Division ...... 43 1.6 Expression of Myh10 in the Mouse ...... 45 1.7 Role of Myh10 in Human Disease ...... 47 1.8 The EHC Project ...... 50 1.9 Aims and Objectives ...... 55 Chapter 2: Materials and Methods ...... 56 2.1 Contribution Statement ...... 56 2.2 Animals ...... 57 2.2.1 Ethical Statement ...... 57 2.2.2 Mouse Lines ...... 57 2.3 Genotyping Animals ...... 57 2.3.1 DNA Isolation from Adult Mouse Tissue ...... 57 2.3.2 Genotyping PCR ...... 58 2.3.3 Agarose Gel Electrophoresis ...... 58 2.4 Collection and Processing of Embryonic Tissue ...... 58 2.4.1 Embryo Harvesting and Imaging ...... 58 2.4.2 Tissue Fixation ...... 59 2.4.3 Genotyping Embryos ...... 59 2.5 Sequencing ...... 59 2.5.1 PCR Product Purification ...... 59 2.5.2 Sequencing PCR Amplification and Purification ...... 59 2.6 Complementation Test ...... 60 2.7 X-gal Staining for β-galactosidase Expression ...... 60 2.8 Tamoxifen Injections ...... 60

2 2.9 Histology and Immunohistochemistry ...... 61 2.9.1 Paraffin Embedding and Microtome Sectioning ...... 61 2.9.2 OCT Embedding and Cryosectioning ...... 61 2.9.3 Haematoxylin Staining ...... 62 2.9.4 Whole Mount Platelet Endothelial Cell Adhesion Molecule 1 (PECAM-1) Staining ...... 62 2.9.5 Smooth Muscle α-Actin (SMαA) ...... 63 2.9.6 Wilms Tumour 1 (Wt1) ...... 64 2.9.7 Phospho-HistoneH3 (PHH3) ...... 65 2.9.8 Snail...... 65 2.9.9 Vimentin ...... 66 2.9.10 Non-Muscle Myosin II Isoforms A, B and C ...... 66 2.10 Cell Culture ...... 67 2.10.1 Culture of Epicardial Cells from Embryonic Heart Explants ...... 67 2.10.2 Immunocytochemistry ...... 67 2.10.3 Scratch-Wound Assay ...... 68 2.10.4 Culture of Lung Epithelial and Fibroblast Cells ...... 68 2.10.5 Culture of Cardiomyocytes and Cardiac Fibroblasts from Embryonic Hearts . 69 2.11 Image Analysis and Statistics ...... 70 2.11.1 Analysis of EPDC Migration In Vivo ...... 70 2.11.2 Analysis of Cell Proliferation In Vivo ...... 71 2.11.3 Analysis of Epicardial Cell Migration In Vitro ...... 71 2.12 Western Blot Analysis ...... 72 2.12.1. Generation of Protein Extracts ...... 72 2.12.1.1 Generation of Protein Extracts from Whole Embryonic Hearts ...... 72 2.12.1.2 Generation of Protein Extracts from Cultured Epicardial Cells and Explants ...... 72 2.12.2 SDS-PAGE ...... 73 2.12.3 Western Blotting Protocol ...... 73 2.12.4 Densitometry Analysis ...... 74 Chapter 3: Characterisation of EHC and Myh10∆ Mutant Embryos ...... 75 3.1 Introduction ...... 75 3.1.1 Background to Congenital Hydrocephalus in the NMIIB Ablated Mouse ...... 75 3.1.2 Coronary Vessel Formation ...... 77 3.1.2.1 Role of the Epicardium in Coronary Vessel Formation ...... 77 3.1.2.2 Vascular Endothelial Capillary Formation ...... 78 3.1.2.3 Vascular Smooth Muscle Cell Recruitment...... 79 3.1.3 Previously Reported Myh10 Knock Out Animals ...... 80 3.1.4 Complementation Testing ...... 82 3.2 Results ...... 83 3.2.1 Morphology of EHC Embryos ...... 83 3.2.2 Morphology of EHC Embryonic Hearts ...... 85 3.2.3 Complementation Assay ...... 89 3.2.3.1 Genotyping Progeny for the Flox Deletion and EHC Mutation ...... 89 3.2.3.2 EHC/Myh10∆ Animals are Embryonic Lethal ...... 89 3.2.3.3 Morphology of EHC/Myh10∆ Mutant Embryos ...... 90 3.2.3.4 Gross Morphology of EHC/Myh10∆ Mutant Hearts ...... 92 3.2.4 The Myh10∆ Allele is an Myh10 Null ...... 92 3.2.4.1 Generation of the Myh10∆ Mouse Line ...... 92 3.2.4.2 Myh10∆ Homozygous Mutants Are Embryonic Lethal ...... 93 3.2.4.3 Myh10∆ Homozygous Mutants Do Not Express Full Length NMHC IIB . 93 3.2.4.4 Morphology of Myh10∆ Homozygous Mutant Embryos ...... 96 3.2.4.5 Morphology of Myh10∆ Homozygous Mutant Hearts ...... 98

3 3.2.5 Characterisation of the Extent of Vascular Defects in the EHC Mouse ...... 100 3.2.5.1 Vascular Endothelial Cell Localisation ...... 100 3.2.5.2 Vascular Smooth Muscle Cell Localisation ...... 102 3.3 Discussion ...... 104 3.4 Conclusion ...... 112 Chapter 4: Analysis of Epicardial Cell Function ...... 113 4.1 Introduction ...... 113 4.1.1 NMII Expression and Function in the Embryonic Mouse Heart ...... 113 4.1.2 Epicardial Cell Function During Heart Development ...... 115 4.1.2.1 Molecular Regulation of Epicardial Function During Cardiogenesis ...... 115 4.1.2.2 Formation of the Epicardium ...... 116 4.1.2.3 Epicardial Proliferation and EMT ...... 117 4.1.2.4 EPDC Migration and Differentiation ...... 118 4.1.2.5 Epicardial Mediated Myocardial Proliferation ...... 121 4.2 Results ...... 123 4.2.1 NMHC IIB is the Predominant NMII Isoform in the Embryonic Epicardium ...... 123 4.2.1.1 Relative NMII Abundance in the Embryonic Heart ...... 123 4.2.1.2 NMII Sub-Cellular Localisation in Epicardial Cell Cultures ...... 126 4.2.2 Epicardial Cell Culture ...... 128 4.2.2.1 Cultured Epicardial Cell Morphology ...... 129 4.2.2.2 Myh10 Ablated Epicardial Cells Do Not Show Migration Defects in vitro . 131 4.2.3 EHC Mutants Show Defective EPDC Migration in vivo ...... 133 4.2.4 EHC EPDC Retain Their Differentiation Capacity ...... 135 4.2.5 Analysis of Epicardial Cell Proliferation ...... 138 4.2.6 EHC Mutants Show Defects in Epicardial EMT ...... 142 4.2.7 Investigating Mechanotransduction in Myh10 Ablated Epicardial Cells ...... 143 4.3 Discussion ...... 145 4.4 Conclusion ...... 152 Chapter 5: Generation of Myh10 Tissue-Specific Knock Out Animals ...... 153 5.1 Introduction ...... 153 5.1.1 Cre Recombinase/loxP Genomic Editing Technology ...... 153 5.1.2 Conditional Modulation of Cre Recombinase Expression ...... 156 5.1.3 Detection and Tracking Cre Recombinase Expression ...... 157 5.1.4 Epicardial-Specific Cre Drivers ...... 159 5.1.5 Myocardial-Specific Knock Out Mice ...... 159 5.1.6 The Myh10 Myocardial-Specific Knock Out Mouse ...... 160 5.2 Results ...... 161 5.2.1 Generation of Myocardial-Specific Myh10 Knock Out Animals ...... 161 5.2.2 α-MHC-Cre Breeding Scheme ...... 161 5.2.3 Confirmation that the α-MHC-Cre Mouse Expresses Cre Recombinase in the Ventricular Myocardium ...... 163 5.2.4 Confirmation that α-MHC-Cre; flox/flox Animals Show Reduced Cardiac NMIIB Protein ...... 163 5.2.5 Genotyping Animals to Identify Myocardial Myh10 Knock Out Embryos ...... 166 Figure 5.5: PCR Analysis of Myh10 Exon 2 Deletion in a panel of 'Second Generation' α-MHC-Cre/+; flox/+ Embryonic Tissues ...... 167 5.2.6 Validation of Myh10 Deletion in Cardiomyocytes ...... 168 5.2.7 Morphology of α-MHC-Cre; flox/flox Mutant Embryos ...... 170 5.2.8 Generation of Epicardial-Specific Myh10 Knock Out Animals ...... 173 5.2.9 The Wt1tm1(EGFP/cre)Wtp/J Mouse Line ...... 173 5.2.10 The Wt1tm2(cre/ERT2)Wtp Mouse Line ...... 175

4 5.2.11 Wt1-CreERT2 Breeding Strategy ...... 175 5.2.12 Heart Morphology of ‘First Generation’ Potential Epicardial-Specific Myh10 Knock Out Embryos ...... 177 5.2.13 Validation of Loss of Epicardial NMIIB ...... 180 5.2.14 Breeding Scheme for Colony Re-establishment at the University of Manchester ...... 182 5.2.15 Tamoxifen Dosing and Progeny Analysis ...... 184 5.2.16 Morphology of ‘Second Generation’ Potential Epicardial-Specific Myh10 Knock Out Embryos ...... 185 5.3 Discussion ...... 188 5.4 Conclusion ...... 194 Chapter 6: Final Discussion ...... 195 6.1 Phenotypic Analysis of the EHC Mouse ...... 195 6.2 Confirmation that a Mutation in Myh10 Causes the EHC Phenotype ...... 196 6.3 Defective Brain Development in Myh10 Knock Out Mice ...... 197 6.4 The EHC Mouse Shows Evidence of Epicardial Dysfunction ...... 199 6.5 Final Conclusion ...... 205 6.6 Limitations ...... 206 6.7 Future Directions ...... 208 6.7.1 Further Characterisation of the Myh10 Null Phenotype ...... 208 6.7.2 Exploring Epicardial Dysfunction ...... 209 6.7.3 Further Defining NMIIB Function in Cardiogenesis ...... 212 Appendices ...... 214 Bibliography ...... 226

Final Word Count (including Bibliography) = 74,322

5 List of Figures

Figure 1.1: Schematic Representation of Early Heart Development and the Contribution of the Primary and Secondary Heart Fields ...... 23 Figure 1.2: Schematic Representation of Outflow Tract Formation and Septation ...... 26 Figure 1.3: Schematic Representation of the Propagation of Cardiomyocyte Proliferation by the Epicardium ...... 29 Figure 1.4: Schematic Representation of the Formation of the Epicardium and Coronary Vasculature ...... 33 Figure 1.5: Schematic Representation of NMIIB Structure and Filament Assembly ...... 40 Figure 1.6: Schematic Representation of the Inv(11)8Brd Balancer Chromosome ...... 51 Figure 1.7: Schematic Representation of the EHC Mutagenesis Screen Breeding Scheme . 52 Figure 1.8: Schematic Representation of the Localisation of the EHC Point Mutation and the Predicted Effect on the NMHC IIB Protein...... 54 Figure 3.1: Schematic Representation of the Ventricular Segments During Embryonic Mouse Brain Development ...... 76 Figure 3.2: Morphology of the EHC Mutant Embryo ...... 84 Figure 3.3: Morphology of the EHC Mutant Heart...... 86 Figure 3.4: Genotyping and Morphology of EHC/Myh10∆ Mutants ...... 91 Figure 3.5: Western Blot Analysis of NMIIB Expression in Myh10∆ Animals ...... 95 Figure 3.6: Morphology of Myh10∆ Homozygous Mutant Embryos ...... 97 Figure 3.7: Morphology of Myh10∆ Homozygous Mutant Hearts ...... 99 Figure 3.8: Molecular Characterisation of the Coronary Vasculature in EHC Hearts...... 101 Figure 4.1: Schematic Representation of the Differentiation of EPDCs into Multiple Cardiac Cell Lineages ...... 119 Figure 4.2: Expression of NMII Isoforms in Control and Myh10∆ Mutant Embryonic Hearts ...... 124 Figure 4.3: NMIIA and NMIIB Expression in Cultured Epicardial Cells ...... 127 Figure 4.4: Morphology and Molecular Profile of Cultured Epicardial Cells ...... 130 Figure 4.5: Scratch-Wound Assay on Epicardial Cell Cultures ...... 132 Figure 4.6: Analysis of EPDC Migration in vivo by Wt1 Immunohistochemistry ...... 134 Figure 4.7: Analysis of Cardiac Fibroblast Localisation by Vimentin Immunohistochemistry ...... 137 Figure 4.8: Analysis of Cell Proliferation in Epicardial Cell Cultures ...... 139 Figure 4.9: Analysis of Epicardial Proliferation and EMT ...... 141 Figure 4.10: Analysis of MRTF-A Localisation in Cultured Epicardial Cells ...... 144 Figure 5.1: Schematic Representation of Cre-loxP Mediated Genomic Editing ...... 155 Figure 5.2: Schematic Representation of the ROSA26 Reporter Mouse Line ...... 158 Figure 5.3: Breeding Scheme for the Generation of Cardiomyocyte-Specific Myh10 Knock Out Animals - the α-MHC-Cre Mouse ...... 162 Figure 5.4: Analysis of 'First Generation' Cardiomyocyte-Specific Myh10 Knock Out Animals ...... 164 Figure 5.5: PCR Analysis of Myh10 Exon 2 Deletion in a panel of 'Second Generation' α- MHC-Cre/+; flox/+ Embryonic Tissues ...... 167 Figure 5.6: Immunocytochemical Analysis of NMIIB Localisation in Cultured Cardiomyocytes and Fibroblasts ...... 169 Figure 5.7: Morphological Analysis of 'Second Generation' Cardiomyocyte-Specific Myh10 Knock Out Embryos and Hearts ...... 172 Figure 5.8: Analysis of Cre Recombinase Expression in the Wt1-CreGFP Mouse Line ... 174 Figure 5.9: Breeding Strategy to Generate Epicardial-Specific Myh10 Knock Out Animals Using the Wt1-CreERT2 Mouse at the University of Oxford ...... 176

6 Figure 5.10: Morphological Analysis of 'First Generation' Potential Epicardial-Specific Myh10 Knock Out Embryonic Hearts ...... 178 Figure 5.11: Molecular Analysis of NMIIB and SMαA Localisation in ‘First Generation’ Potential Epicardial-Specific Myh10 Knock Out Embryonic Hearts ...... 181 Figure 5.12: Revised Breeding Strategy to Generate Epicardial-Specific Myh10 Knock Out Animals at the University of Manchester ...... 183 Figure 5.13: Morphological Analysis of 'Second Generation' Potential Epicardial-Specific Myh10 Knock Out Embryos and Hearts ...... 186 Figure 6.1: Revised Model Illustrating Defective Epicardial Function and Coronary Vessel Formation in the EHC Mouse ...... 201

7 List of Tables

Table 3.1: Compound Heterozygous EHC/Myh10∆ Animals Display Embryonic Lethality ...... 90 Table 3.2: Homozygous Myh10∆ Mutant Animals Display Embryonic Lethality ...... 93 Table 3.3: Phenotypic Summary of the Myh10 Ablated Mouse ...... 105

8 List of Abbreviations

α-MHC - α-myosin heavy chain Alk5 - TGFβ type 1 receptor kinase AMP – Adenosine monophosphate Angpt1 – Angiopoietin-1 aPKC – Atypical protein kinase C Apob – Apolipoprotein b APS – Ammonium persulphate APSC – Aorticopulmonary septation complex ATP – Adenosine triphosphate AU – Arbitrary units bp – Base pairs BHF – British heart foundation BMP – Bone morphogenetic protein BSA – Bovine serum albumin Bves – Blood vessel epicardial substance

CaCl2 – Calcium chloride Cas9 – CRISPR associated protein 9 Cdh1 – Cadherin 1 cDNA – Complimentary DNA CHD – Coronary heart disease CNCC – Cardiac neural crest cells

CO2 – Carbon dioxide CRISPR – Clustered regularly interspaced short palindromic repeats CRISPRi – CRISPR interference CSF – Cerebral spinal fluid Cy3 – Cyanine 3 Cy5 – Cyanine 5 DAB - 3,3'-diaminobenzidine tetrahydrochloride DAPI – 4’,6-diamidino-2-phenylindole dH2O – Distilled water ddH2O – Double distilled water dHAND - deciduum, heart, autonomic nervous system, neural crest derived DMEM – Dulbecco’s modified eagle medium

9 DORV – Double outlet of the right ventricle DPC – Days post-conception E – Embryonic day EBI - European bioinformatics institute eCFP – Enhanced cyan fluorescent protein ECM – Extracellular matrix EDTA – Ethylenediaminetetraacetic acid eGFP – Enhanced green fluorescent protein EGFR – Epidermal growth factor receptor EHC – Embryonic hydrocephalus and cardiac defects EMT – Epithelial-mesenchymal-transformation En-2 – Engrailed-2 ENU - N-ethyl-N-nitrosourea EPDC – Epicardial-derived cell Erbb – Erb-b2 receptor tyrosine kinase ERT2 – Estrogen receptor T2 EtOH – Ethanol Ets1 – E26 avain leukemia oncogene 1 eYFP – Enhanced yellow fluorescent protein FACS – Fluorescence-activated cell sorting FAK – Focal adhesion kinase FANTOM5 – Functional annotation of the mammalian genome 5 FBS – Featal bovine serum FGF - Fibroblast growth factor FGFR – Fibroblast growth factor receptor FITC – Fluorescein isothiocyanate Floxed – Flanked by loxP sites FMHS –Faculty of medical and human sciences FOG2 – Friend of GATA 2 G – Grams GAGs – Glycosaminoglycans Gata – GATA binding protein GTP – Guanosine-5’-triphosphate

H2O2 – Hydrogen peroxide HA – Hyaluronic acid Hand - Heart and neural crest derivatives

10 Has2 – Hyaluronic acid synthase 2 HRP – Horse radish peroxidase ICC – Immunocytochemistry Igf2 - Insulin-like growth factor 2 Igf1r - Insulin-like growth factor 1 receptor IHC – Immunohistochemistry Insr – Insulin receptor IP – Intraperitoneal Isl1 - Insulin gene enhancer protein Kb – Kilo base pairs KCl – Potassium chloride KDa – Kilo Dalton Kg – Kilo gram

KH2PO4 - Monopotassium phosphate loxP – Locus of cross-over M – Molar mA – Milli amp mm – Milli metre mM – Milli molar Mef2c – Myocyte enhancing factor-2c MeOH – Methanol mg – Milli gram

MgCl2 – Magnesium chloride Mkl – megakaryoblastic leukemia/myocardin-like mL – Milli litre µL – Micro litre MLC – Myosin light chain MLCK – Myosin light chain kinase MMRRC - Mutant mouse resource and research centers MRTF – Myocardin related transcription factor Mycn – Myelocytomatosis viral related oncogene, neuroblastoma derived Myh – Myosin heavy polypeptide n – Number

NaCl2 – Sodium chloride

Na2HPO4 – Disodium phosphate NaOAc – Sodium acetate

11 NIH – National institutes of health Nkx2.5 - NK2 transcription factor related, locus 5 nM – Nano molar NMII – Non-muscle myosin II NMHC II – Non-muscle myosin heavy chain II Nrg1 – Neuregulin-1 OCT – Optimal cutting temperature OFT – Outflow tract Otx-2 – Orthodenticle homolog 2 P – Progeny Par – Partitioning defective Pax3 – Paired box 3 PBS – Phosphate buffered saline PBS-T - Phosphate buffered saline + 0.1% (v/v) tween 20 PCR – Polymerase chain reaction PDGF – Platelet-derived growth factor PDGFR – Platelet-derived growth factor receptor PECAM-1 – Platelet endothelial cell adhesion marker 1 PEO – Proepicardial organ PHF – Primary heart field PHH3 – Phospho-histoneH3 PFA – Paraformaldehyde Prox1 - Prospero-related homeobox 1 PVDF - Polyvinylidene fluoride Raldh2 – Retinaldehyde-specific dehydrogenase type 2 Rbpj – Recombination signal binding protein for immunoglobulin kappa J region RCF – Relative centrifugal force RhoA – Ras homologue A RIPA – Radioimmunoprecipitation assay buffer RNA –Seq – RNA sequencing ROCK – Rho-associate kinase RPM – Revolutions per minute RT-PCR – Real-time polymerase chain reaction Rxra – Retinoid X receptor alpha Scx – Scleraxis SDS – Sodium dodecyl sulphate

12 SDS-PAGE – Sodium dodecyl sulphate polyacrylamide gel electrophoresis SEM – Standard error of the mean Sema3d – Semaphorin 3D SHF – Secondary heart field Shh – Sonic hedgehog Shox2 - Short stature homeobox-2 SLB – Sample loading buffer SMαA – Smooth muscle α-actin SM22 - Smooth muscle protein  Snai1 – Snail family zinc finger 1 Sox9 - Sex-determining region Y box 9 SRF – Serum response factor T0 – Time zero TAE – Tris-Acetate-EDTA TALLEN – Transcription activator-like effector nucleases Taz – Tafazzin TβR – Transforming growth factor receptor TBS – Tris buffered saline TBS-T – TRIS buffered saline + 0.05% (v/v) tween-20 Tbx – T-box transcription factor Tcf21 – Transcription factor 21 TEMED – Tetramethylethylenediamine TGF β – Transforming growth factor β Tris – Tris(hydroxymethyl)aminomethane Trp – Transformation related protein Twist - Twist basic loop-helix-loop transcription factor 1 V – Volts Vangl2 – Vang-like protein 2 VCAM1 – Vascular cell adhesion molecule 1 VEC – Vascular endothelial cell VEGF – Vascular endothelial growth factor VSD – Ventral septal defect vSMC – Vascular smooth muscle cells V/V – Volume per volume WB – Western blotting

13 Wnt – Wingless-type Wt1 – Wilms tumour 1 WT – Wild type W/V – Weight per volume X-gal – 5-bromo-4-chloro-3-indolyl--D-galactopyranoside Yap – Yes-associated protein 1 Zip – Zipper ZO-1 – Zonula occludens-1

14 Abstract

Aim Recent interest in cardiogenesis has focused on the epicardium, the outer epithelial layer that envelops the heart. Epicardial-derived cells (EPDCs) contribute vascular smooth muscle to developing coronary vessels and provide critical signalling cues to facilitate myocardial functionality. However, the precise molecular mechanisms that underpin epicardial biology remain unclear. Ablation of Myh10 in the EHC mouse results in embryonic lethal cardiac malformations, including epicardial and coronary defects. We sought to establish the role of Myh10 in epicardial cell function to further dissect the coronary vessel developmental pathway, a deeper understanding of which may inform the design of therapeutics to regenerate and repair the injured heart.

Methods Utilising multiple cell and developmental biology techniques, we generated a pathological evaluation of the EHC phenotype. EPDC migration was investigated in vivo with Wt1 immunohistochemistry, and in vitro by performing scratch wound assays on epicardial cell cultures. Congruently, we examined the ability of epicardial cells to undergo EMT in vivo by employing Snail and Phosphohistone-H3 immunohistochemistry.

Results Our studies reveal that EHC epicardial cells have a reduced capacity to invade the ventricular myocardium. Furthermore, we discovered increased proliferation and reduced Snail expression specifically within the EHC epicardium, consistent with EMT dysregulation. Interestingly, epicardial cell function did not appear to be disrupted in vitro.

Conclusion These results demonstrate a novel role for Myh10 in both EPDC migration and the promotion of epicardial EMT. Our finding that migration is unaffected in vitro suggests that the unique properties of the in vivo epicardial microenvironment dictate a requirement for Myh10 in order to elicit correct epicardial function. Together, this research enhances our understanding of the dysfunctional processes that contribute to abnormal cardiogenesis; these insights may aid our ability to determine the molecular regulators of coronary vessel development, and create therapeutics to regenerate vessel growth and repair injured cardiac tissue in cardiovascular disease.

15 Declaration No portion of the work referred to in the thesis has been submitted in support of an application for another degree or qualification of this or any other university or other institute of learning.

Copyright Statement i. The author of this thesis (including any appendices and/or schedules to this thesis) owns certain copyright or related rights in it (the “Copyright”) and s/he has given The University of Manchester certain rights to use such Copyright, including for administrative purposes. ii. Copies of this thesis, either in full or in extracts and whether in hard or electronic copy, may be made only in accordance with the Copyright, Designs and Patents Act 1988 (as amended) and regulations issued under it or, where appropriate, in accordance with licensing agreements which the University has from time to time. This page must form part of any such copies made. iii. The ownership of certain Copyright, patents, designs, trade marks and other intellectual property (the “Intellectual Property”) and any reproductions of copyright works in the thesis, for example graphs and tables (“Reproductions”), which may be described in this thesis, may not be owned by the author and may be owned by third parties. Such Intellectual Property and Reproductions cannot and must not be made available for use without the prior written permission of the owner(s) of the relevant Intellectual Property and/or Reproductions. iv. Further information on the conditions under which disclosure, publication and commercialisation of this thesis, the Copyright and any Intellectual Property University IP Policy (see http://documents.manchester.ac.uk/display.aspx?DocID=24420), in any relevant Thesis restriction declarations deposited in the University Library, The University Library’s regulations (see http://www.library.manchester.ac.uk/about/regulations/) and in The University’s policy on Presentation of Theses

16 Acknowledgements This thesis is dedicated to my wonderful wife, Rosy. Your love and understanding has got me to the end of this project through all the late nights and weekends spent in the lab; I will never be able to repay you for that. Thank you for reminding me that there is so much more to life than work. Although I study hearts for a living, I promise that you will always have mine.

To my Mum, Dad, and brother Gareth – thank you for being the best family. I know that what I do is very far removed from your everyday lives, but you have really helped me get through this. I extend that appreciation to my new family-in-law; Dave, Dianne and Caroline. I love you all.

I would especially like to thank my supervisor, Kathryn Hentges, for always having your door open, and never being too busy to help me. Thank you to the past and present members of the Hentges and Woolf groups, especially Gennadiy Tenin and Riccardo Coletta, for making the lab such a lovely place to work. Thank you to my PhD advisor Christoph Ballstrem, as well as Emma Barnes and Jayne Wright at Syngenta, for their valuable comments and advice. Thank you to the BBSRC and Syngenta for funding.

A special thank you to my friend, Sheona Drummond. You have always been there when I needed to talk about work or anything else. I will truly miss our daily chats. That extends to my pal Gareth Hughes for making me laugh every single day.

I would also like to say a special thank you to Tony Day for giving me my first chance in the research environment. As I move forward, I would like to thank Pete Scambler for giving me a fantastic opportunity to further my academic career at UCL. You have placed an enormous amount of trust in me and I promise I will not let you down.

Finally, I would like to thank my examiners, Nicola Smart and Bernard Keavney, for agreeing to read this thesis. Nicola, you have always been an approachable and friendly face at many-a-conference, and I value all the time you have taken to talk to me over the last few years. Bernard, you awarded me my first ever poster prize and gave me a great boost during my first year. I hope you both enjoy reading about the last 4 years of my life!

17 Preface I obtained by BSc Degree in Cell Biology (2:1, Hons) from Durham University in 2009. From October 2009 – September 2012, I was employed as a Research Technician in Prof. Tony Day’s laboratory at the University of Manchester, and co-authored 2 peer-reviewed research papers (Clark et al., 2013; Keenan et al., 2012) and a published abstract (Clark et al., 2011). During my PhD studies, I have co-authored a research paper (Ridge et al., Manuscript submitted, PLoS Genetics), and a review article (Clowes et al., 2014). I have also co-authored 3 published abstracts (Ridge et al., 2016, Ridge et al., 2015, and Barnes et al., 2014). I have presented my post-graduate research at both national and international conferences, and have been awarded a number of prizes for my presentation skills, including; AstraZeneca Runner up prize ‘Drug Discovery and Innovation’ (EuroTox, Porto 2015), Best Poster Prize (British Microcirculation Society Annual Meeting, Newcastle, 2016; The Heart – New Horizons, Manchester, 2016; MAHSC Symposium, Manchester, 2013), and Best Oral Presentation (PhD Conference, University of Manchester, 2015). I have also been the recipient of external funding grants, including: the Journal of Comparative Pathology Education Bursary, the British Society for Developmental Biology Travel Grant, and the Boehringer Ingelheim Fonds Travel Grant. I am currently a student member of the British Microcirculation Society (BMS), the British Society for Cardiovascular Research (BSCR) and the British Society for Developmental Biology (BSDB). Following the submission of this thesis, I will commence a 5-year BHF-funded post-doctoral Research Associate position in Prof. Scambler’s laboratory at the Institute of Child Health, University College London.

18 Chapter 1: Introduction

1.1 Why Investigate Cardiac Development? – The Heart is the Problem The importance of correct cardiac formation and function to an organism’s survival cannot be over emphasized. The heart is the first functional organ in the developing vertebrate embryo and has an essential and continued role in the delivery of oxygen and nutrients to its cells and tissues throughout life. However, the essentiality of the heart dictates that pathological processes that detrimentally affect its performance are often incompatible with life. A thorough and accurate appreciation of the root causes of these pathologies is the first step on a long road to establish effective treatments to improve the quality and length of life for patients.

Congenital malformations and acquired diseases of the heart pose an enormous health and economic burden throughout the world. Congenital heart defects have been reported to occur in approximately 1-5% of live births and represent the most frequent class of birth abnormalities (Clark et al., 2006). Although it is known that both genetic predisposition and exposure to certain environmental factors (e.g. teratogens) contribute to the development of these conditions, their precise aetiology remains elusive (Bruneau, 2008). Similarly, acquired cardiovascular disease is a major cause of morbidity and mortality, causing over 160,000 deaths every year in the UK alone (Townsend et al., 2014). Moreover, coronary heart disease (CHD), in which cessation of the coronary circulation leads to cardiac arrest and ischemic damage, represents the single largest cause of death in the UK (Townsend et al., 2014). The multitude of risk factors - both genetic and environmental - associated with CHD, are compounded by the interactions that these factors have with one another (Lusis, 2000). This intrinsic complexity makes it extremely difficult to unravel CHD aetiology, and therefore impedes our ability to understand how best to treat the disease. Current treatments for CHD patients are limited to invasive surgery and long-term medicinal routines (Nabel and Braunwald, 2012), which harbour their own inherent risks and financial outlay. In addition, the reduced instance of CHD in the last few decades due to the use of preventative medication has recently curtailed (Brown and O'Connor, 2010). Accompanied by the rising prevalence in obesity and an ageing population, it seems that our ongoing battle against CHD will persist for years to come. In combination, these factors demonstrate the growing demand for novel, therapeutic strategies to combat CHD incidence and associated morbidity. Investment in research to elucidate the processes underpinning the pathology of CHD and other conditions of cardiac dysfunction is required to accomplish this. 19

A thorough appreciation of cardiac development is an essential prerequisite to understanding cardiac dysfunction. Increasingly, investigators are adopting a developmental biology approach to assist the enhancement of our understanding and ability to treat cardiac disease. An increasingly recognised pattern within the field of regenerative medicine is that tissue repair is often facilitated by the reactivation of the embryonic conditions that initially created it (Riley and Smart, 2011; Masters and Riley, 2014; Foglia and Poss, 2016; Olivey and Svensson, 2010). By understanding the developmental program of cardiogenesis, we may be able to recapitulate embryonic conditions and induce the formation of new cardiac tissues and structures to repair the injured heart (Srivastava and Olson, 2000; Masters and Riley, 2014). From this, a deeper understanding of the cellular and molecular processes that underpin cardiogenesis will provide fundamental insights into the causes of cardiac dysfunction, and facilitate the design of novel therapies to more effectively treat both congenital and acquired cardiac disease (Tomanek, 2005).

To this end, the generation of gene knock out animal models, specifically the mouse, is integral to enhancing our understanding of mammalian cardiogenesis (Sung et al., 2012). Information gleaned from these studies is useful in both terms of regenerative medicine and toxicological studies to improve the safety of both pharmacological and agrochemical products. The impact and implications of such studies will be discussed in detail in later sections of this thesis. The adjoining sections of this introduction will briefly summarize the important morphological events and key molecular players known to be involved in mammalian cardiac formation.

20 1.2 Overview of Cardiogenesis in the Mouse Embryo The heart is the first organ to form and function in all vertebrates (Buckingham et al., 2005). Cardiac function is required in utero in order to supply tissues within the developing embryo - via the systemic circulatory system - with sufficient nutrients and oxygen to meet their increasing metabolic demands (Clowes et al., 2014). The delivery of blood to the cardiac tissue itself is provided by the coronary vasculature; a vessel network that envelops and permeates the heart once the myocardial walls become too thick for nutrient and oxygen transfer to occur via passive diffusion.

A significant proportion of our knowledge of mammalian cardiogenesis originates from studies on the mouse, so the description that follows relates to the formation of the mouse heart. Comparative embryological time points to the development of the human heart are provided where appropriate, and stated as days post conception (DPC) (Sylva et al., 2014). At this point, it is relevant to acknowledge that the mammalian species required by regulatory studies investigating the developmental toxicology of chemical compounds are usually the rat and rabbit (personal communication with Dr. Emma Barnes, Syngenta). However, as rodents and lagomorphs, these species are relatively comparable, and knowledge gained from either branch of research can be largely transferrable (Fischer et al., 2012; Danielsson et al., 2013). Equally, as mammals, mice and humans share similarities in physiology, developmental programmes and ultimately genetic composition, and as a result, many aspects of human development and disease processes are comparable to those in the mouse (Fougerousse et al., 2000; Krishnan et al., 2014).

1.2.1 Origins and Morphological Overview of Mammalian Heart Formation Correct cardiogenesis is reliant upon the precise orchestration of intricate signalling events and cellular processes to build a four-chambered, fully functioning muscular pump from early cardiac precursors. A brief summary of the morphological changes undertaken during this process is described below.

The heart begins to beat from approximately embryonic day (E) 8.25 (Wyszynski et al., 2010) (22-23 DPC) and is fully formed at E16.5 (56-60 DPC) (Schleich et al., 2013). The progenitor cells that form the future heart originate from the primitive streak following gastrulation. This region of the anterior mesoderm is induced by signals from the endoderm, such as BMP (bone morphogenetic protein). There are 5 key classes of

21 transcription factors involved in this early induction and the downstream determination of cardiac cell fate, which are highly conserved across vertebrate species, namely Nkx2.5, Mef2c, Gata-4, Tbx, and Hand (Olson, 2006). Cardiac precursor cells coalesce around the cephalic pole of the embryo to form the cardiac crescent (also referred to as the primary heart field, PHF) by E7.5 in the mouse (17-19 DPC) (Fig 1.1, A) (Dunwoodie, 2007; Schleich et al., 2013). The cardiac crescent subsequently fuses at the midline following anterior migration of cells in the PHF to form a linear heart tube at the ventral region of the embryo (Fig 1.1, B) (Dunwoodie, 2007). This primitive tube consists of two layers; an external myocardium, and an internal endothelium, separated by cardiac jelly (Schleich et al., 2013). The primitive heart tube incorporates additional cellular components from the secondary heart field (SHF) (also referred to in the literature as the anterior heart field). This region of the embryo, positioned dorsally and anteriorly to the heart tube, migrates medially and anteriorly as heart development progresses (Fig 1.1, A-B). Cardiac expansion during this stage is fuelled by myocardial proliferation.

At approximately E8.5 (22-23 DPC) the heart tube undergoes rightward looping (Fig 1.1, B, arrows) (Dunwoodie, 2007). Cardiac looping is the first right/left lateralisation event in the embryo (Schleich et al., 2013). Cardiac looping brings both the inflow and outflow portions of the heart into close proximity, in addition to aligning the future four chambers into position relative to one another (Buckingham et al., 2005; Schleich et al., 2013). Looping is induced due to uneven growth and remodelling of the primitive atria and ventricles (Xin et al., 2013). Alignment of the specified atrial and ventricular regions in the correct orientation is essential for the subsequent development and maturation of the four cardiac chambers.

Following looping, the heart experiences significant tissue remodelling (Dunwoodie, 2007). The SHF is now positioned close to the pharyngeal mesoderm (Fig 1.1, C). At this stage, cardiac growth is facilitated by both cardiomyocyte proliferation and the additional recruitment of cells to the expanding heart (Brade et al., 2013). Rapid cardiac expansion is accompanied by developmental processes encompassing the septation of the ventricles and atria, as well as the formation of the cardiac valves, which are complete by approximately E15.5. By E16.5, the embryonic heart has adopted a characteristic mature morphology.

Figure 1.1: Schematic Representation of Early Heart Development and the Contribution of the Primary and Secondary Heart Fields (A) Cardiac precursors emerging from the primitive streak coalesce at the embryonic midline to form the cardiac crescent. The secondary heart field resides dorsally to the cardiac crescent. (B) The cardiac crescent fuses at the midline to form the primitive heart tube at E8.0. The heart tube undergoes rightward looping at approximately E8.25. (C) Following looping, cells from the secondary heart field, now positioned dorsally and anteriorly, migrate and contribute to the expansion of the rudimentary heart (arrows). (D) By E10.5, the heart has adopted its characteristic four-chambered morphology. The primary heart field contributes predominantly to the left ventricle, with a modest contribution to both atria. The secondary heart field contributes to the right ventricle and outflow tract, and similarly contributes to both left and right atria. (E) By E14.5, the heart is fully septated, and the outflow tract is divided into the aorta and pulmonary trunk. Blue indicates the primary heart field. Yellow indicates the secondary heart field. Ao – aorta, IVS – interventricular septum, LA – left atria, LV – left ventricle, ML – midline, OFT – outflow tract, PhA – pharyngeal arches, PT – pulmonary trunk, RA – right atria, RV – right ventricle, SHF – secondary heart field, Tr – trabeculae. Adapted from Clowes, Boylen and Ridge et al., 2014.

23 The secondary heart field, marked by Isl1 (Cai et al., 2003) and Fgf8 (Buckingham et al., 2005) expression, contributes to the formation of the right ventricle and outflow tract, as well as both left and right atria (Fig 1.1, D) (Zaffran et al., 2004; Srivastava, 2006; Buckingham et al., 2005). In particular, anterior SHF cells contribute to the smooth muscle cell population within the region at the base of the aorta and pulmonary artery (Ward et al., 2005). In addition, cells towards the posterior SHF contribute cardiomyocytes to the atria walls and septum (Douglas et al., 2011). The PHF, marked by Nkx2.5 and Hand1 expression (Buckingham et al., 2005), contributes to the left ventricle, and both left and right atria (Fig 1.1, D). Additional cardiac progenitor cells are derived from the proepicardium and the cardiac neural crest; these sources will be discussed in detail in later sections.

The heart is divided into its classical four-chambered anatomy through the process of septation. For correct septation to occur, the heart aligns its distinct regions through the process of conversion, a sequential series of ‘morphological shifts’ (Schleich et al., 2013). As part of this convergence process, the outflow tract elongates via addition of myocardial cells from the SHF. Elongation of the outflow tract is essential for the correct positioning of the future aorta and pulmonary trunk. The outflow tract experiences a counter clockwise rotation that allows the aorta to be positioned behind the pulmonary trunk. By E14.5, (44- 48 DPC), the heart is connected to the pulmonary trunk and the aorta (Fig 1.1, E) (Buckingham et al., 2005; Brade et al., 2013). Disruption to this process results in the misalignment of the great vessels in relation to the ventricles, and a consequential failure in the fusion of the outflow tract septum with the primitive ventricular septum (as observed in Tetralogy of Fallot) (Shinebourne et al., 2006; Villafane et al., 2013).

The myocardium expands markedly following E9.5-10.5 (35-39 DPC), initiating the formation of increasing numbers of trabeculations (Zhang et al., 2013a). These structures vastly increase the surface area of the inner ventricular wall, and permit the efficient transfer of oxygen to the myocardial cells as late as E15.5 (Olivey and Svensson, 2010; Schleich et al., 2013). During early cardiogenesis, supply of oxygen and nutrients to the heart is achieved via passive diffusion across the myocardium. However, as cardiomyocytes proliferate and the cardiac walls thicken, these metabolic demands require vascularisation of the heart, leading to the eventual formation of the coronary vascular network (Riley and Smart, 2011). Coronary vessel formation and connection to the systemic circulation is a finely tuned process involving a series of vasculogenic, angiogenic, and remodelling events (Tomanek, 2005), and is essential for embryonic survival (Riley and Smart, 2011).

1.2.2 Cardiac Maturation and the Contribution of Extra-Cardiac Cell Populations As previously described, the heart is derived from cells originating in both the primary and secondary heart field during early embryogenesis. However, maturation of the primitive heart tube requires additional cellular contributions from extracardiac cell populations.

1.2.2.1 Outflow Tract Development Proper development and septation of the outflow tract is essential for the correct separation of the pulmonary and systemic blood flow (Buckingham et al., 2005). It has been shown that the anterior most cells of the SHF make a major contribution to outflow tract development (Zaffran et al., 2004; Kelly et al., 2001). Lineage tracing has indicated that SHF cells expressing Tbx1, Fgf8 and Fgf10 contribute myocardial cells to muscularise the developing outflow tract (Buckingham et al., 2005). In addition, the outflow tract incorporates cardiac neural crest cells (CNCC), which are derived from the dorsal neural tube and migrate to the heart via the pharyngeal arches (Fig 1.2, A) (Xin et al., 2013). It is known that the induction of CNCC delamination and migration is facilitated by a number of classical signalling pathways, including but not limited to; BMP, FGF (fibroblast growth factor), TGF-β (transforming growth factor β), and retinoic acid (Brade et al., 2013). CNCCs migrate into the developing outflow tract and form a ‘U-shaped’ aorticopulmonary septation complex (APSC) at approximately E11.5 (Fig 1.2, B). By E14.5 (42-44 DPC), the condensing of the APSC with additional mesenchymal cells permits septation of the outflow tract into the aorta and pulmonary artery, which are patent with the ventricular chambers (Fig 1.2, C) (Brade et al., 2013). In addition to their contribution to the aorticopulmonary septum, CNCCs predominantly contribute to the formation of aortic smooth muscle cells that line the great arteries (Fig 1.2, D), and play an additional role in providing secreted signals to other cardiac cells (Brade et al., 2013; Kirby et al., 1983; Youn et al., 2003; Achilleos and Trainor, 2012).

Figure 1.2: Schematic Representation of Outflow Tract Formation and Septation (A) Cardiac neural crest cells (CNCCs; blue cells) migrate from the neural crest towards the developing heart via the pharyngeal arches at E8.5. (B) By E11.5, CNCCs populate the distal outflow tract and to facilitate elongation and form a ‘U-shaped’ aorticopulmonary septation complex. (C) Septation of the outflow tract is enabled by condensation of the aorticopulmonary septation complex. Simultaneously, the proximal outflow tract becomes populated by cardiomyocytes. This myocardialisation event enables the rotation of the outflow tract to properly align it with the ventricles (white block arrow). (D) By E14.5, the outflow tract is septated into the aorta and pulmonary trunk. The great vessels are lined by CNCC-derived smooth muscle cells. AA – aortic arch, Ao – aorta, APSC – aortopulmonary septation complex, CMs – cardiomyocytes, CNCC – cardiac neural crest cells, OFT – outflow tract, PhA – pharyngeal arches, PT – pulmonary trunk, SHF – secondary heart field, SMC – smooth muscle cells. Adapted from Brade et al., 2013.

Congenital heart defects relating to the formation and positioning of the great arteries, including double outlet of the right ventricle (DORV), are often associated with abnormal outflow tract shortening. The importance of the CNCC contribution to outflow tract development is highlighted by studies in which the neural crest is partially ablated, resulting in varied heart defects, including: tuncus arteriosis, Tetralogy of Fallot and DORV (Schleich et al., 2013; Achilleos and Trainor, 2012). Mutations in key genes expressed in the SHF have been shown to disrupt outflow tract development. In particular, Fgf8 mutant animals fail to survive due to primary outflow tract malformations (Abu-Issa et al., 2002). The Fgf8 mutant phenocopies the human 22q11 deletion syndrome (also known as DiGeorge syndrome), and is likely underpinned by disrupted CNCC migration (Frank et al., 2002; Buckingham et al., 2005). Additional genes associated with malformation of the outflow tract include: Isl1 (Buckingham et al., 2005), Mef2c (Lin et al., 1997), and Tbx20 (Takeuchi et al., 2005).

Furthermore, a major contributor to the development of congenital heart defects, including DORV, is a failure in myocardialisation, which results in incomplete convergence or wedging (Bartram et al., 2001). This can disrupt the rotation of the base of the great arteries, and cause aberrant alignment of the aorta and pulmonary truck in relation to the ventricles. This has been observed in the Gata4 mutant mouse, which demonstrates DORV with defects in outflow tract septation (Luttun and Carmeliet, 2003).

1.2.2.2 Myocardial Proliferation and Ventricular Trabeculation Embryonic cardiac growth is facilitated by cardiomyocyte proliferation (Ahuja et al., 2007). The majority of cardiomyocytes become binucleated following birth due to DNA replication in the absence of cytokinesis. Post-natal heart growth is achieved through the enlargement of cardiomyocytes via hypertrophy (Xin et al., 2013). However, the correct regulation of embryonic cardiomyocyte proliferation is key to the formation and development of the trabeculae in the maturation of the ventricular myocardial wall (Foglia and Poss, 2016). Early research suggested that the formation of the trabeculae is driven by molecular signalling events between the myocardium and the endothelial lining of the ventricular lumen, which induce the invagination of the proliferating myocardium to form ridged projections (Sedmera et al., 1997; Challice and Viragh, 1974). Endocardial Notch signalling has been shown to be crucial for correct trabeculae formation (Foglia and Poss, 2016) and myocardial maturation/compaction (D'Amato et al., 2016), with the former thought to induce the activity of both Bmp10 and Nrg1 (Foglia and Poss, 2016). Animals

27 with mutated Notch1, and Bmp10, as well as Nrg1 and its receptors Erbb2 and Erbb4 display impaired trabeculation and a thinned myocardial wall (Foglia and Poss, 2016). In addition, the formation of the trabeculae is impaired in mice deficient for Hand2 (also known as dHAND; deciduum, heart, autonomic nervous system, neural crest derived) (Srivastava et al., 1997), Angpt1 (Suri et al., 1996), and VEGF (vascular endothelial growth factor) (Carmeliet et al., 1996; Ferrara et al., 1996; Luttun and Carmeliet, 2003).

Maturation of the myocardial wall between E10.5-15.5 involves the progressive formation of a compact zone of myocardial tissue (Foglia and Poss, 2016). The development of the compact myocardium involves the proliferation of cone shaped myocardial clones, which are broadest at the outer surface of the myocardium, and taper towards the inner surface of the myocardial wall (Fig 1.3) (Mikawa et al., 1992). It is thought that this patterning of cardiomyocyte proliferation is initiated through the release of mitogenic factors from the epicardium, the hearts outer epithelial layer (discussed in detail in subsequent sections) (Chen et al., 2002; Perez-Pomares et al., 2002b). Disruption to erythropoietin and retinoic acid signalling, as well as mutations in Wt1, have been shown to result in a thinning of the myocardial wall (Wu et al., 1999; Stuckmann et al., 2003; Martinez-Estrada et al., 2010). In addition, retinoic acid/erythropoietin signalling is has been shown to regulate epicardial Igf2 expression, and disruption of both Igf2, and its receptors Insr and Igf1r has been shown to reduce ventricular myocardium proliferation. Other signalling cascades implicated in the control of myocardial development include: FGFs (Lavine and Ornitz, 2008), Wnt (wingless-type) (Merki et al., 2005; Zamora et al., 2007), and VEGF (Brade et al., 2013). Although the elucidation of the exact mitogenic molecules required for myocardial proliferation, and ultimately both trabeculae and compact zone formation, remain incomplete, it is clear that the epicardium plays an important role in these processes (Foglia and Poss, 2016).

Figure 1.3: Schematic Representation of the Propagation of Cardiomyocyte Proliferation by the Epicardium The epicardium secretes signalling molecules that induce cardiomyocyte proliferation. This facilitates the expansion and correct development of the myocardium. A sub-population of epicardial cells undergo EMT, which gives rise to epicardial-derived cells (yellow cells). These cells also contribute signalling molecules to induce cardiomyocyte proliferation. Key signalling molecules involved in this process are indicated. Please see main text for details. CM – cardiomyocytes, EMT – epithelial-mesenchymal transformation, EPDC – epicardial- derived cell. Adapted from Brade et al., 2013.

1.2.2.3 Development of the Coronary Vasculature The coronary vasculature is the network of blood vessels that perfuse the ventricular myocardium. These vessels deliver oxygen and nutrients to meet the working demands of the underlying cardiac muscle when passive diffusion becomes insufficient, in addition to permitting myocardial compaction (Dyer et al., 2014). Embryonic coronary vessel development encompasses two distinct stages; initial establishment of a primitive endothelial plexus, followed by plexus remodelling, maturation and connection to the systemic circulation via the aortic root (Dyer et al., 2014). These processes require the correct orchestration of distinct physiological events, including; (i) vasculogenesis: the de novo formation of primitive vessels, (ii) angiogenesis: the remodeling of, and formation of new vessels from, the initial vascular network, and (iii) arteriogenesis: the recruitment of vascular smooth muscle cells to coronary arteries and the establishment of hierarchy within the vascular tree.

Although the ultimate origin of these coronary endothelial cells is contentious (Dyer et al., 2014; Riley and Smart, 2011), recent data suggests that the endothelial plexus largely originates from the sinus venosus, with a modest contribution from both the ventricular endothelium and epicardium (Red-Horse et al., 2010; Tian et al., 2013b; Tian et al., 2015; Chen et al., 2014; Zhang et al., 2016; Katz et al., 2012). Initial observations from Wu and colleagues (2012) indicated that the majority of coronary endothelial cells were derived from the endocardium (Wu et al., 2012). However, recent studies that have refined the use of Cre recombinase based lineage tracing demonstrate that whilst the endocardium is the predominant source of coronary endothelial cells in the ventricular septum, there is a minimal endocardial contribution to the coronaries within the ventricular walls (Tian et al., 2015; Chen et al., 2014; Zhang et al., 2016).

At approximately E11.5, patches of endothelial cells branch from the sinus venosus and coalesce in the sub-epicardial space with minor endothelial populations derived from both the epi- and endocardium (Katz et al., 2012; Red-Horse et al., 2010; Tian et al., 2015). These primitive vessels continue to spread across the ventricular surface to envelope the heart and form a complete plexus at approximately E13.5. Blood-filled capillaries appear around the peritruncal region of the aorta at E12.5-13.0 (Gonzalez-Iriarte et al., 2003; Tian et al., 2013a) and are recruited to penetrate the wall of the aorta and establish the connection with the plexus via the aortic stems from approximately E15.5 (Tian et al., 2013a). This mechanism appears to be conserved in the human, with the formation of the

30 endothelial plexus beginning at 31-35 DPC, and connection to the aorta established by 44- 48 DPC (Hutchins et al., 1988).

Following complete plexus formation at E13.5, coronary capillaries undergo vascular remodelling and maturation to maintain vascular integrity in the presence of pressure generated by blood flow. This is accomplished through the recruitment of vascular smooth muscle cells and perivascular fibroblasts, which are known to derive from the epicardium (discussed in detail in the next section). The processes that initiate and regulate smooth muscle cell recruitment are poorly understood, but are thought to include mechanical stimulation from haemodynamic properties following the initiation of flow (Luttun and Carmeliet, 2003; Vrancken Peeters et al., 1999), and chemotaxis signalling gradients derived from the coronary endothelium (Dardik et al., 2005). These processes are discussed in greater detail in Sections 3.1.3.2-3.

1.3 Brief Overview of the Formation and Function of the Epicardium As alluded to in the preceding section, correct cardiogenesis is dependent upon a structure called the epicardium, the monolayer mesothelium of the heart. The epicardium is derived from a transient extracardiac source termed the proepicardial organ (PEO), which itself arises from the coelomic mesenchyme of the septum transversum at approximately E8.5 in the mouse (23-26 DPC) (Manner et al., 2001). However, the mechanisms that induce PEO formation are not currently known (Manner et al., 2001). The PEO is comprised of cyst- like cells that form a clustered structure posterior to the post-looped heart at the junction of the sinus venosus and inflow tract at E9.5 (Fig 1.4, A). Mammalian PEO cells traverse the pericardial space and adhere to the developing myocardium between E9.5-11.5 (26-30 DPC) to form an outer epithelial sheath that completely envelops the nascent heart by E12.5 (37-42 DPC) (Fig 1.4, B) (Komiyama et al., 1987). The PEO and epicardium have been described in a diverse range of embryonic vertebrate species, including chick (Manner, 1993), mouse (Viragh and Challice, 1981) and man (Hirakow, 1992) and therefore demonstrate a high level of evolutionary conservation. Continuity between cells within the epicardium is maintained by the extension of cellular processes (Viragh and Challice, 1981) and the formation of cell-cell gap junctions, which have been demonstrated to be essential for correct epicardial cell function (Li et al., 2002). In addition, the epicardium deposits extracellular matrix (ECM) into the sub-epicardial space, which separates it from the underlying myocardium (Fig 1.4, B). The sub-epicardial ECM plays an important role in molecular communication between these tissues, and is critical for other aspects of epicardial cell function (von Gise and Pu, 2012).

From approximately E12.5, epithelial-mesenchymal transformation (EMT) is activated in a sub-population of epicardial cells. These cells have been termed epicardial-derived cells (EPDCs) (Gittenberger-de Groot et al., 1998) and have acquired the capacity to delaminate from the epicardium and invade the underlying myocardium, where they differentiate and contribute to the structural scaffolding and coronary vasculature of the heart (Fig 1.4, B). It is not known if cells are committed to an EPDC fate in the PEO, but the transduction of localised signalling molecules has been implicated in the activation of EMT in epicardial cells (Smith et al., 2011; von Gise and Pu, 2012). These signaling events coordinate extensive alterations in epicardial gene expression, including the down regulation of adhesion and cell polarity molecules, cytoskeletal reorganization and enabling the ability to breakdown ECM (Perez-Pomares and de la Pompa, 2011).

Figure 1.4: Schematic Representation of the Formation of the Epicardium and Coronary Vasculature (A) The epicardium arises from an extra-cardiac structure called the proepicardial organ (PEO). The PEO forms posterior to the post-looped heart by approximately E9.5. (B) The cyst-like structures that make up the PEO dissociate and transverse the pericardial space, where they adhere to the developing myocardium. Following the formation of a uniform epithelium, a sub-population of epicardial cells undergo EMT to generate epicardial-derived cells (EPDCs). EPDCs invade the myocardium, where they differentiate into cardiac fibroblasts and vascular smooth muscle cells (vSMC). These vSMC are subsequently recruited to the endothelial coronary capillaries to form mature coronary vessels. (C) The epicardium is absolutely essential for the correct formation of the mature coronary vasculature. Ao – aorta, CV – coronary vessel, ECM – extracellular matrix, LA – left atrium, LAD – left anterior descending artery, LCA – left coronary artery, RA – right atrium, RCA – right coronary artery, PT – pulmonary trunk, VEC – vascular endothelial cell. Adapted from Clowes, Boylen and Ridge et al., 2014.

EPDCs retain a level of multipotency and contribute to multiple different cell populations within the heart (Kimura and Sadek, 2012). A number of pioneering studies have highlighted their capacity to differentiate into vascular smooth muscle cells, both perivascular and interstitial fibroblasts, and controversially, vascular endothelial cells (Mikawa and Fischman, 1992; Gittenberger-de Groot et al., 1998; Dettman et al., 1998; Perez-Pomares et al., 2002a). Fate mapping studies have also described the formation of functional cardiomyocytes from EPDCs (Cai et al., 2008; Zhou et al., 2008). Recent data suggests that the PEO is comprised of distinct sub-populations of cells; those expressing the classical epicardial markers Wt1/Tbx18, and those expressing Scx/Sema3d, and that these heterogeneous cell populations have distinct differentiation capacities (Katz et al., 2012). However, it is clear that the correct epicardial function is fundamental for the correct formation of the mature coronary vasculature by E16.5 (Fig 1.4, C).

Experimental evidence indicates that successful development of the coronary vasculature greatly depends on the correct migration and differentiation of EPDCs. Moreover, defects in myocardial development, as well as coronary abnormalities, strongly correlate to compromised EPDC function (Winter and Gittenberger-de Groot, 2007) indicating an additional epicardial role in the expansion and proliferation of the ventricular myocardium. Together, the potential of epicardial cells to differentiate into multiple cardiac cell lineages and regulate other aspects of cardiogenesis has driven research to uncover the molecular processes regulating epicardial cell function as a means of facilitating the design of novel therapies for cardiovascular diseases.

1.4 Epicardial Potential in Regenerative Medicine Understanding the molecular mechanisms that underpin embryonic epicardial function is critical to contemporary efforts to better treat cardiac disease. The formation and maintenance of the coronary circulation is essential for proper cardiac function, and disruption to these processes can cause significant mortality and morbidity in the form of congenital and acquired heart disease. Myocardial infarction causes the loss of billions of cardiomyocytes; unlike some species of lower invertebrate, the mammalian adult heart displays an extremely restricted ability to regenerate damaged cardiac muscle and coronary vasculature following tissue injury (reviewed in Masters and Riley, 2014). The mammalian adult epicardium is dormant, and classical embryonic epicardial marker genes are differentially expressed in the adult heart (van Wijk et al., 2012). Excitingly, the adult epicardium can “switch on” as least some elements of its embryonic gene expression profile in response to injury (Wagner et al., 2002; Smart et al., 2011), indicating that aspects of embryonic epicardial function may be similarly reactivated. Indeed, the potential of EPDCs to differentiate into functional cardiomyocytes following injury has been demonstrated by the reprogramming of adult progenitor cells (Smart et al., 2011). This seminal work highlights a potential application of EPDC redeployment in reparative and regenerative medicine; however, reactivation of epicardial proliferation and the secretion of factors to promote and modulate cardiac repair is critical to maintain function in the post- injured heart. A deeper knowledge of the factors that underpin epicardial function may be manipulated to harness more efficient cardiac muscle regeneration following injury.

Identification that the adult epicardium reactivates embryonic gene expression in response to injury has triggered therapeutic interest in epicardial biology. However, this does not inherently translate into the cell and tissue regeneration mechanisms necessary to repair the injured heart. The mammalian injury response centres on the restoration of the cardiac fibroblast population, which leads to extracellular matrix deposition at the site of injury and fibrotic scar formation (Kikuchi and Poss, 2012). Ultimately, mass fibrosis impedes contractility, leading to the pathological remodelling of cardiac tissue and heart failure due to the deterioration of function (Masters and Riley, 2014).

In contrast, it has been previously reported that fish and some amphibians can regenerate cardiac muscle following the resection of up to 20% of the ventricular mass (Poss et al., 2002). Interestingly, following the administration of a cryoinjury, which better recapitulates the effects of ischemia, initial scar formation is gradually replaced by functioning

35 myocardium in the zebrafish (Gonzalez-Rosa et al., 2011). It appears that the key difference underscoring increased heart regeneration capacity in the adult fish centres on the initiation of epicardial EMT and the subsequent secretion of paracrine factors that modify and enhance the myocardial injury response following ischemic reactivation of embryonic epicardial gene expression. The epicardium and sub-epicardial region are therefore proposed to act as a hypoxic niche for cardiac stem cells (Kimura and Sadek, 2012). Surprisingly, effective and efficient heart regeneration has been observed in the neonatal mouse heart (Porrello et al., 2011; Porrello et al., 2013; Mahmoud et al., 2013; Xin et al., 2013). However, this regeneration window appears not to extend beyond postnatal day 4, and the diminished production of trophic factors by the epicardium, accompanied by the decreased ability of the myocardium to respond to these cues, results in a default fibrotic response flowing injury in older animals (Porrello et al., 2011; Chen et al., 2002).

Although few key players in the regulation of this regenerative window have been identified, it is known that this decreased regeneration capacity is inversely correlated to the number of binucleated cardiomyocytes, which accumulate due to karyokinesis in the absence of cytokinesis in the perinatal mouse (Ikenishi et al., 2012). In contrast, adult zebrafish cardiomyocytes are mononuclear, and retain proliferation capacity indefinitely (Wills et al., 2008). Interestingly, the zebrafish epicardium plays a key role in cardiomyocyte homeostasis and turnover (Wills et al., 2008), whilst the involvement of the epicardium in mammalian cardiac homeostasis is not known. It may be that the continued receptive state of the zebrafish cardiomyocytes to epicardial signals better facilitates their response to paracrine factors released by the reactivated epicardium following injury.

This indicates that as yet unidentified extrinsic factors are essential to construct new heart tissue in mammals (Sturzu and Wu, 2011). Some of the outstanding questions within the field that need to be addressed in order to fully exploit the epicardium as a therapeutic target are as follows;  How are the myogenic properties of the epicardium regulated? (Kikuchi et al., 2011)  To what extent does the epicardium function as the birthplace of de novo cardiomyocytes in response to injury? (Russell et al., 2011)  What parameters/criteria need to be met to drive EPDC-mediated propagation of cardiomyocyte proliferation and coronary vessel regeneration in response to injury, as observed in the embryonic context?

The molecular mechanisms underpinning epicardial cell motility and differentiation are poorly understood (Trembley et al., 2015), but it is thought that cell shape, cytoskeletal tension and RhoA (ras homologue A) activity regulate cardiac stem cell lineage commitment in human mesenchymal stem cells (McBeath et al., 2004); similar mechanisms may apply in the epicardium. Investigating how the epicardium functions during development will shed light on the discrepancies in embryonic and adult epicardial function. Furthermore, if the novel properties of embryonic epicardial cells can be therapeutically recapitulated, it is possible that the reactivation of the cardiac developmental programme will repair the injured heart; reconciling the differential function of the embryonic and adult epicardium may facilitate the neovascularization and regeneration of functional myocardium following infarction. This exciting therapeutic potential substantiates further research into epicardial cell function during cardiogenesis.

A vital tool in the developmental biologists arsenal to delineate the mechanisms that underpin cardiogenesis is the generation of genetic knock out model organisms (Sung et al., 2012). Phenotypic analysis of such knock out animals can shed light on the function of particular genes during cardiogenesis, and guide the characterisation of the key molecular players and processes that underpin correct heart formation.

1.5 Myh10 is Essential for Correct Cardiogenesis One gene that has been associated with playing a key role in correct cardiogenesis is Myh10 (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003). This gene encodes the protein non-muscle myosin heavy chain IIB (NMHC IIB). Mouse model systems in which NMHC IIB has been globally disrupted or deleted display cardiac and brain abnormalities, resulting in embryonic/perinatal lethality (Tullio et al., 1997; Ma et al., 2007). Furthermore, mice harbouring a single point mutation in Myh10 have been shown to develop body wall closure defects, including omphalocele and cleft palate, highlighting the broad spectrum of developmental processes that are reliant upon NMHC IIB (Ma and Adelstein, 2014a). The cardiac phenotype encompasses: DORV, ventricular septal defects (VSDs), and defects in cardiomyocyte cytokinesis. Myocardial-specific conditional ablation of NMHC IIB has also been previously shown to cause cardiomyopathy in the adult heart, highlighting the extended requirement for this molecule beyond embryonic development (Ma et al., 2009). A summary of the phenotypic findings from previous studies on both the global and cardiomyocyte-specific knock out mouse is provided in Table 3.3. In the human, a point mutation in Myh10 has been shown to phenocopy aspects of the loss of function mutation presented by the mouse, suggesting a conserved interspecies functionality (Tuzovic et al., 2013). However, the precise role that Myh10 plays during cardiogenesis is not yet fully understood. In addition, the broad variety of defects presented in Myh10 disrupted mice suggests that Myh10 may function in a myriad of different molecular processes. This thesis will now discuss the biological importance of NMHC IIB in the context of its molecular structure and multifaceted function.

1.5.1 Introduction to Non-Muscle Myosin IIB The myosin superfamily represents a significant proportion of the force generating machinery within eukaryotic cells. The myosin II class is the most populous subfamily, and together with actin, make up the majority of contractile protein machinery in cardiac, skeletal and smooth muscle (Vicente-Manzanares et al., 2009). Despite their name, nonmuscle myosin II (NMII) molecules are found in both muscle and non-muscle cells, and are ubiquitously expressed in eukaryotic cells. As previously mentioned, Myh10 encodes non-muscle myosin heavy chain IIB, which is a component of non-muscle myosin IIB (NMIIB). The Myh10 gene is conserved in Bilateria (HomoloGene: 55941), and encodes a 1976 amino acid protein in both the mouse and human. NMIIB is an intracellular actin binding moiety that has the ability to generate contractile force on actin filaments, and is thus able to physically modify the organisation of the actin cytoskeleton (Conti and

Adelstein, 2008; Vicente-Manzanares et al., 2009). As such, NMIIB serves an essential function in cellular processes that require force generation. Mammals have 3 different NMII heavy chain isoforms – denoted as NMHC-IIA, -IIB and -IIC – encoded by the genes Myh9, Myh10 and Myh14 respectively. The heavy chain isoform dictates the isoform of the NMII complex, and these isoforms are referred to as NMIIA, NMIIB and NMIIC in this thesis. The NMII isoforms display a high level of sequence homology. Whilst they display some degree of functional overlap, they have distinct properties with regard to their ability to bind actin, including, duty ratios (Vicente-Manzanares et al., 2009) and force- dependent filamentous actin affinities (Kovacs et al., 2007), suggesting that some NMII dependent cellular functions are isoform specific.

NMIIB is a hexameric protein complex, consisting of two 230kDa heavy chains, and two pairs of myosin light chains, both regulatory (20kDa) and essential (17kDa) (Fig 1.5, A). The activation of NMIIB is reliant upon phosphorylation of the regulatory light chains by Rho kinase and myosin light chain kinase (MLCK), which enables NMIIB molecules to assemble into bipolar filaments through interactions between their rod domains (Fig 1.5, B) (Conti and Adelstein, 2008). The globular head domain of the heavy chain molecule can bind to actin, and exert contractile force via an ATPase mediated conformational change (Fig 1.5, C). NMIIB can act indirectly on integrins and signal transduction molecules via its binding to actin and as such, plays a central role in diverse cellular processes, including the establishment of cell polarity, cell adhesion and cell migration (Conti and Adelstein, 2008; Vicente-Manzanares et al., 2009). These are interdependent processes, and numerous NMIIB knock out cell lines and animal models have been used to piece together the complex role of NMIIB in cell behaviour. The reported NMIIB knock out studies discussed in the following section are particularly relevant to this thesis research project.

For clarity, this thesis will refer to the protein product of Myh10 as NMIIB, due to the total dependency of the NMIIB protein complex on correct heavy chain function.

Figure 1.5: Schematic Representation of NMIIB Structure and Filament Assembly (A) A single essential light chain (red) and regulatory light chain (orange) bind to the lever arms of each NMHC IIB molecule (purple). Homo-dimerisation is mediated by interactions between NMHC IIB rod domains. In the absence of phosphorylation, the NMIIB hexamer forms a compact molecule through head-tail interactions. This compact molecule is not able to associate into filaments. (B) Following phosphorylation of the regulatory light chain, NMIIB undergoes a conformational change and is now able to assemble into filaments via interactions mediated by the rod domains. (C) Assembly of bipolar NMIIB filaments facilitates the binding of the NMHC IIB globular head domain to the actin cytoskeleton, and allows the execution of NMIIB functionality. ELC – essential light chain, MLCK – myosin light chain kinase, NHT – non-helical tail, NMHC IIB – nonmuscle myosin heavy chain IIB, RLC – regulatory light chain. Adapted from Conti and Adelstein, 2008, and Vincente-Manzanares et al., 2009. 40

1.5.2 Role of NMIIB in Cell Migration Cell migration is facilitated by the actin-polymerisation dependent formation of a structure called the lamellipodium (Vincente-Manzanares et al., 2009). Cell migration, encompassing cell edge retraction and the formation of new adhesions, is initiated by NMII forces acting upon the actin network at the rear of the lamellipodium (Giannone et al., 2007). In addition, it has been shown that NMII phosphorylation facilitates cell migration in the absence of ECM proteolysis by generating contractile forces to deform collagen fibres (Sahai and Marshall, 2003). However, the three NMII isoforms show different spatial distribution in migrating cells, suggesting that the NMII isoforms serve distinct functions during migration (Vicente-Manzanares et al., 2009; Kovacs et al., 2007; Even-Ram et al., 2007).

NMIIB has been implicated in the regulation of cell migration; NMIIB depleted cells are significantly less well spread and demonstrate a reduced rate of migration in in vitro wound healing assays (Sandquist et al., 2006). Also, NMIIB ablated fibroblasts display a reduced ability to control the direction or speed of their migration, producing multiple, unstable protrusions, and highly disorganised traction forces (Lo et al., 2004). Furthermore, these cells were unable to respond to mechanical stimuli, such as changing substrate flexibility and compression forces (Lo et al., 2004). Similarly, depletion of NMIIB in human cancer cells has been shown to impair both migration and spreading of the lamellipodia (Betapudi et al., 2006). Expanding upon this, Thomas et al., (2015) have recently shown that NMIIB plays a central biophysical role in nuclear mechanics during 3D invasion in mammary gland carcinoma cells (Thomas et al., 2015). Here, the authors show that NMIIB functions to apply force to the nucleus during invasion, and facilitates nuclear translocation through tight spaces, possibly through coupling force generation to the nuclear membrane protein, nesprin-2 (Thomas et al., 2015).

Interestingly, inhibition of NMII with blebbistatin has revealed that NMII regulates the length of actin bundles and retrograde actin flow in neuronal growth cones, thus demonstrating involvement in the dynamic reorganization of the actin cytoskeleton in migrating cells (Medeiros et al., 2006). Together, these data indicate that NMIIB can mediate the cellular response to external mechanical stimuli and stabilizes cell migration.

41 1.5.3 Role of NMIIB in Cell Adhesion A link has also been established between NMIIB function and cell adhesion. Adhesion complexes play an essential role in signal transduction and molecular communication between adjacent cells and their environment. NMIIB is thought to play a central role in the formation, maturation and maintenance of these complexes by regulating the recruitment and localisation of their protein components (Conti and Adelstein, 2008).

NMIIB has been shown to actively recruit specific proteins to adhesion sites, and is essential for maintenance of non-periphery focal adhesions (Sandquist et al., 2006). Similarly, ablation of NMIIA leads to the loss of E-cadherin and β-catenin from adhesions (Conti et al., 2004). Similarly, pan-inhibition of NMII activity impairs E-cadherin recruitment to cell-cell contacts (Shewan et al., 2005). Concurrently, it has been shown that NMII becomes mislocalised at cell-cell junctions following the ablation of the β-catenin orthologue, armadillo, in Drosophila (Dawes-Hoang et al., 2005). In addition, NMII accumulation at cell-cell junctions has been shown to be dependent upon E-cadherin activity, as well as myosin light chain (MLC) phosphorylation (Shewan et al., 2005).

In migrating cells, NMIIB contractility induces the formation of new adhesion sites (Giannone et al., 2007), which act as traction points to propel the cell body in the direction of migration (Webb et al., 2005). Integrin accumulation occurs at these sites in response to mechanical stress induced by NMIIB, as well as NMIIA ((Cai et al., 2006) cited in (Puklin- Faucher and Sheetz, 2009)). NMIIB is known to accumulate at sub-cellular sites experiencing mechanical stress, and inhibition of NMIIB has been shown to affect the recruitment of other mechanosensitive proteins (Schiffhauer et al., 2016). Notably, NMIIB has been shown to recruit vinculin to E-cadherin mediated epithelial cell junctions in response to mechanical stress (Sumida et al., 2011). This is particularly relevant, as it suggests that NMIIB deficient cells are unable to properly maintain adhesions with adjacent cells due to an inability to respond to mechanical stimuli.

Moreover, the development of embryonic hydrocephalus in NMIIB null mice has been attributed to loss of cell-cell adhesion in the neuroepithelial cells lining the spinal canal (Ma et al., 2007). This study presents evidence suggesting that NMIIB plays a structural, rather than a locomotive role, at cell adhesion sites (Ma et al., 2007)

1.5.4 Role of NMIIB in Cell Polarity and Cell Division Establishment and maintenance of the apical-basal axis is an essential process in the control of cell division and a broad variety of other cellular processes (Conti and Adelstein, 2008). Recent work has shown that activation of NMII is a key mediator for establishment of cell polarity and mitosis in the fly (Lee et al., 2007). This study found that the AMP- kinase knock out Drosophila animals failed to phosphorylate NMII (encoded by zip), resulting in defects in cell shape, epithelial cell polarity establishment, and mitosis. In addition, subcellular NMII accumulation has been shown to play a crucial role in anterior- posterior patterning and the establishment of planar cell polarity in Drosophila during gastrulation, and may therefore influence polarized cell movement (Zallen and Wieschaus, 2004).

Moreover, failure to initiate NMIIB activation has been implicated in the defective polarization and migration of myocardial cells, resulting in DORV in the mouse (Phillips et al., 2005). These authors found that Vangl2 (Vang-like protein 2) - a component of the planar cell polarity pathway - deficient mice, displayed disruption to RhoA and ROCK1 (rho-associated kinase 1) expression in the developing outflow tract, suggesting a role for NMII activation in correct outflow tract formation (Phillips et al., 2005). This complements the finding that NMIIB null mice display DORV and other cardiac defects, which have been attributed to a failure to establish cell polarity (Tullio et al., 1997). Similarly, Myh10 ablation in the mouse has been found to result in cardiomyocyte cytokinesis defects, indicating a role for NMIIB in this process (Tullio et al, 1997; Takeda et al., 2003; Ma et al., 2009). Expanding upon this, recent work suggests that NMIIB plays an important role in myocardial karyokinesis through its organisation of microtubules (Ma et al., 2010; Takeda et al., 2003). NMIIB depleted mice present an increased accumulation of abnormally shaped and binucleated cardiomyocytes in the compact myocardium, attributed to an increase in microtubule stability in the absence of NMIIB and NMIIC (Ma et al., 2010). Compounding this, Yang and colleagues (2015) have reported that surprisingly, NMIIB plays a unique function in cytokinesis during meiotic cell division of the male, but not female, germline cells (Yang et al., 2012). This study reports that NMIIB is dispensable for mitotic cell division in male germ cells, but not for meiotic cell divisions, for which both NMIIA and NMIIC are not required (Yang et al., 2012). This highlights the deviation and non-redundancy in NMII function in different cell lineages and during specific divisions.

Finally, NMIIB has been show to play a role in the spatial segregation of specific integrin heterodimers; the translocation of α5β1 to the cell centre during fibrillogenesis, an additional process of cell polarization, is now known to be NMIIB-dependent (Puklin- Faucher and Sheetz, 2009). Similarly, NMIIB has been shown to play a crucial role in the elongation of dendritic branches during hippocampal neurone maturation (Ozkan et al., 2015). The authors speculate that the mechanism underpinning this process is likely to be the role that NMIIB plays in the spatial segregation of molecules/factors that selectively promote the elongation of particular types of dendritic branches over others (Ozkan et al., 2015).

The studies described above clearly demonstrate that NMII plays a pivotal function in a broad and diverse array of cellular functions. Furthermore, these studies have illustrated that the different NMII isoforms have both redundant and functionally distinct properties. The functions that NMIIB serves are often essential for embryonic survival, highlighted by the severity of phenotype presented by NMIIB deficient mouse models (Tullio et al., 1997; Takeda et al., 2003).

1.6 Expression of Myh10 in the Mouse Considering these diverse functions, it is likely that NMIIB plays a multifaceted role in cardiac development, which is reflected in its expression profile during embryonic development (Ma et al., 2010). NMHC IIB has been shown to undergo alternative splicing, which introduces additional exons into the ATP binding region (B1) and actin-binding (B2) region of the NMHC IIB globular head domain (Itoh and Adelstein, 1995); these are known to affect NMIIB motility properties (Kim et al., 2008). However, expression of these alternatively spliced isoforms is restricted to the nervous system (Itoh and Adelstein, 1995; Ma et al., 2010). The expression of NMIIB in the adult mouse and a range of both mouse and human cell lines has been characterised by Ma and colleagues (2010) by using mass spectrometry (Ma et al., 2010). They report that NMIIB is the predominant NMII isoform in the adult mouse cerebral cortex, cerebellum, and spinal cord (Ma et al., 2010). In addition, NMIIB is the predominant isoform in COS-7 cells, suggesting a conserved function in higher mammals. Whilst NMIIB makes a modest contribution to the total NMII content in other human cells lines analysed in this study, NMIIB is relatively abundant in the perinatal mouse heart (Ma et al., 2010).

Murine NMIIB expression appears to peak during embryonic development. NMIIB is expressed in mouse embryonic stem cells and continues to be operative until the death of the organism (Ma and Adelstein, 2014b). It composes approximately 20% of NMII content in mouse embryonic stem cells, and is strongly detectable in the brain and heart throughout development. NMIIB is abundant in embryonic cardiomyocytes from E7.5, and moreover, becomes the only detectable isoform in these cells between E8.5 and E13.5, following the loss of NMIIA expression, and before NMIIC expression is detectable (Ma et al., 2010). NMIIB is similarly expressed in all other cardiac cell types from E13.5, although analysis in other cardiac cell types before this stage has not been conducted (Ma et al., 2010). In addition, NMIIB expression is detectable in the both the paramesenchyme and epithelial cells of the E13.5 lung (Ma et al., 2010).

Similarly, the FANTOM5 project, an international research consortium which has undertaken a systematic analysis of gene expression in the majority of primary mammalian cell types, has identified high Myh10 expression in the heart throughout mid to late gestation in the mouse. Consistent with the findings of Ma and colleagues (Ma et al., 2010), they found that whilst Myh10 expression was detectable in the neonatal mouse, its expression decreased significantly in the juvenile and adult mouse heart. This suggests that NMIIB function in the heart is specific to embryogenesis. A summary of findings from the

45 FANTOM5 project and others for Myh10 expression can be found on the EBI expression atlas website (www.ebi.ac.uk, search term “Myh10”).

1.7 Role of Myh10 in Human Disease As the greater NMII family is known to function in a multitude of cellular processes, it is not surprising that mutations which either directly affect NMII, or indirectly affect the processes it regulates, have been found to contribute to a broad and diverse array of human diseases (reviewed extensively in (Newell-Litwa et al., 2015); (Ma and Adelstein, 2014b)).

NMII function has been implicated in neural tube closure in both Xenopus and chick models, where Rho mediated NMII contractility induces constriction and subsequent convergence of the apical neuroepithelium (Ma and Adelstein, 2014b). In addition, NMII has been shown to function in the migration of neurons (Ma et al., 2007), and is required for apical-basal translocation of the nucleus during neurogenesis and abscission to facilitate migration (Das and Storey, 2014). Synchronously, NMIIB is known to contribute to the pathology of a wide range of neurological conditions, encompassing neurodevelopmental (e.g. autism), neurodegeneration (e.g. Alzheimer’s), and neuronal migration disorders (Newell-Litwa et al., 2015). NMIIB is known to localise to growth cones, where it regulates process extension during neuronal repair (Rochlin et al., 1995). NMIIB is also the predominant NMII isoform at the synapse, where it functions to cycle synaptic vesicles and during synaptic maturation (Chandrasekar et al., 2013; Newell-Litwa et al., 2015). In light of this, it is not surprising that de novo mutations in MYH10 have been reported to contribute to the development of schizophrenia and autism in human patients, although the biochemical impact of these mutations on NMIIB protein function are yet to be investigated ((Li et al., 2016), cited in Newell-Litwa et al., 2015).

Altered NMII contractility has also been implicated in tumourigenesis; the tumour suppressor p53 has been shown to alter NMIIB expression and reduce tumour invasion (Yam et al., 2001). Concurrently, the essential role of NMII in cell division can also mediate cellular detachment, a key process in tumourigenesis (Newell-Litwa et al., 2015). NMII mediated contractility can also facilitate cancer cell invasion independent of ECM remodelling (Beadle et al., 2008). Expanding upon this, in combination with NMIIA, NMIIB is thought to play a key role in the regulation of hematopoietic stem cell fate determination through its differential expression in the differentiating cell and renewing stem cell (Shin et al., 2014). Increased NMII contractility has also been associated with the disruption of endothelial cell-cell junctions, leading to reduced blood vessel stability and increased permeability, which may contribute to conditions such as atherosclerosis (Newell-Litwa et al., 2015).

47 The Myh10 knock out mouse displays DORV and VSD - abnormalities frequently observed in patients with congenital heart disease (Ma and Adelstein, 2014b). The population of the proximal outflow tract by proliferating cardiomyocytes – a process known as myocardialisation – is essential for the correct positioning of the aorta in relation to the left ventricle (van den Hoff et al., 1999). This cardiomyocyte migration event is disrupted when NMIIB is either deleted (Tullio et al., 1997), or replaced with a motor impaired NMIIB isoform (Takeda et al., 2003; Ma and Adelstein, 2014a). It is thought that this impairment is underpinned by defective NMIIB mediated adhesion disassembly. In addition, this function in myocardialisation appears to be unique to NMIIB, as it cannot be rescued by replacement with NMIIA (Bao et al., 2007). In agreement with this, impaired planar cell polarity mediated RhoA/ROCK1 signalling has been found to contribute to disrupted outflow tract myocardialisation (Phillips et al., 2005). It has been suggested that NMIIB functions downstream of the planar cell polarity signalling axis to regulate myocardialisation (Ma and Adelstein, 2014b). Expanding upon this, ROCK2 copy number variants have also been shown to contribute to the development of human congenital heart defects (Fakhro et al., 2011), suggesting a role of ROCK-mediated NMII contractility in the development of these disorders, possibly through the aberrant regulation of TGF-β signalling (Zhang et al., 2009).

A genetic link between NMIIB and human congenital heart disease is yet to be explicitly made. Indeed, very little association has been made between MYH10 mutations and human disease. A de novo MYH10 mutation (E908X) has been found in a single patient displaying a broad range of conditions, including the neuropathies of developmental delay, cerebral and cerebellar atrophy, hydrocephalus and microcephaly (Tuzovic et al., 2013). This mutation is thought to disrupt NMIIB filament assembly. A second proband, carrying an independent de novo mutation (T1162M), has also been reported (Ma and Adelstein, 2014b). The unifying ailment shared by these patients was congenital diaphragmatic hernia; no heart defects were reported in either proband. That being said, Ma and Adelstein (2014a), generated a mouse model of a Myh10 point mutation that phenocopied the human condition, Pentalogy of Cantrell. Fascinatingly, this condition encompasses the congenital heart defects VSDs and DORV, in addition to ectopia cordis, where the heart is located either partially or completely outside of the thorax (described in (Mallula et al., 2013)). Evidently, additional studies are required to reveal unambiguous NMIIB dependent cardiac abnormalities in the human.

Understanding the role of NMIIB in human disease is an emerging field. It is clear that NMIIB is essential for correct mammalian heart formation, and it is hoped that advances in next generation sequencing will reveal mutations in MYH10 affiliated with congenital heart pathologies. The study of mouse models that phenocopy human conditions, such as that generated by Ma and Adelstein (2014a), will form an essential component of our continued efforts to understand these pathologies at both the cell and molecular level, and subsequently facilitate the design of novel therapies to better repair and regenerate the injured heart.

1.8 The EHC Project To improve our understanding of the molecules and processes required for correct mammalian coronary vessel development, our laboratory has employed a forward genetics approach to identify novel genes involved in cardiogenesis. Forward genetics is an unbiased technique which seeks to identify essential genes in a given biological process (Lawson and Wolfe, 2011). In this instance, the genetic basis of abnormal cardiogenesis was investigated by screening a population of animals with random gene altering mutations for a cardiac developmental defect phenotype (Kile et al., 2003).

The l11jus27 mouse was isolated from a balancer chromosome mutagenesis screen (Kile et al., 2003). This experiment used the potent mutagen N-ethyl-N-nitrosourea (ENU) to induce random point mutations into the male germline (Hentges et al., 2006). The balancer chromosome contains an inversion between Trp53 and Wnt3 in chromosome 11 (Fig 1.6), which represses recombination events within this genomic region and thus hastens mutation mapping to a defined chromosome (Hentges and Justice, 2004). In addition, the balancer chromosome contains a selection marker (K14-Agouti), which facilitates the identification of carriers by producing a yellow pigment in the tail and ears (Kile et al., 2003). Mouse chromosome 11 is gene rich and is highly conserved with human chromosome 17 (Fig 1.6) (Kile et al., 2003; Hentges and Justice, 2004). The inversion was created through manipulation of Cre-loxP site-specific recombination in mouse embryonic stem cells (Hentges and Justice, 2004). A schematic representation of the breeding scheme employed to generate homozygous mutants displaying interesting developmental phenotypes in this screen in shown in Fig 1.7. This phenotype driven technique permits the unbiased identification of essential genes, and facilitates the analysis of mammalian gene function. A meiotic mapping approach revealed that the l11Jus27 mouse line actually carried two independent embryonic lethal recessive mutations, one of which presented embryonic hydrocephalus and cardiac defects (EHC) (Mitchell, K., PhD Thesis, University of Manchester; Ridge et al., Manuscript submitted, PLoS Genetics). No mutations associated with cardiac defects had previously been mapped to mouse chromosome 11, suggesting the discovery of a novel gene required for cardiac development (Kile et al., 2003).

Figure 1.6: Schematic Representation of the Inv(11)8Brd Balancer Chromosome The distal two thirds of mouse chromosome 11 displays synteny with human chromosome 17 (grey region). The Inv(11)8Brd balancer chromosome contains a 24-cM sequence inversion between Trp53 and Wnt3 which represses recombination events in this chromosomal region (orange region). cM – centi Morgan. Taken from Kile et al., 2003.

Figure 1.7: Schematic Representation of the EHC Mutagenesis Screen Breeding Scheme ENU treated C57BL/6J male mice (dark grey mice) were crossed to females carrying the balancer chromosome (brown mice). The balancer chromosome carries the Agouti transgene under the expression of the K14 promoter, which gives carriers a light pigment in the tail, ears and ventrum. P1 balancer carriers which may harbour a new mutation (lightning bolt) were crossed to mice heterozygous for both the balancer and Rex, which marks the non-mutagenised chromosome by giving animals a curly coat (represented by dashed line on brown mice). Progeny from this cross carrying the balancer and mutation (light tailed mice with normal coat, P2) were intercrossed to produce homozygous offspring (double lightning bolt, dark tail – test cross, P3). Mice homozygous for the balancer chromosome are Wnt-3 deficient and die by mid-gestation. The genotype status of each animal is indicated by the chromosome schematics underneath each mouse (black – wild type, black with orange – balancer, black with lightning bolt – mutagenized, light grey dashed – Rex). P – progeny. Adapted from Kile et al., 2003; Hentges et al., 2006. 52

Candidate gene analysis prompted the sequencing of the homozygous EHC mutant (subsequently referred to as EHC, unless otherwise stated) Myh10 sequence, which revealed a G to T point mutation in the splice donor site following Myh10 exon 18, which corresponds to the globular head domain in the protein sequence (Fig 1.8, A). The wild type Myh10 transcript generates a 1976 amino acid NMHC IIB protein (Fig 1.8, B). In EHC embryos, Myh10 exon 18 is omitted from the Myh10 transcript, creating an abnormal fusion of exon 17 and exon 19. This novel fusion alters the translational reading frame, introducing a premature stop codon into the transcript. Translation of this transcript is predicted to generate an abnormal C-terminally truncated protein of 703 amino acids, in which the final three amino acids are divergent from the wild type sequence (Fig 1.8, C, red box). Bioinformatic predictive modeling of the molecular structure of this truncated protein revealed that mutant NMHC IIB would lack the wild type α-helical coiled-coil rod domain (Fig 1.8, D-E, blue region, arrowhead) (Mitchell, K., PhD Thesis, University of Manchester; Ridge et al., Manuscript submitted, PLoS Genetics).

Figure 1.8: Schematic Representation of the Localisation of the EHC Point Mutation and the Predicted Effect on the NMHC IIB Protein (A) The EHC mouse carries a G>T point mutation in the sequence corresponding to the globular head domain of NMHC IIB. (B) The wild type NMHC IIB protein is 1976 amino acids long. (C) The EHC mutation alters the translational reading frame, and is predicted to result in a protein truncated at 703 amino acids that lacks the C-terminal myosin rod domain. In addition, the final three amino acids are divergent from the wild type sequence (indicated by red box). (D) Computational model of the structure of the wild type NMHC IIB globular head domain (yellow) and N-terminal portion of the rod domain (blue, arrowhead). (E) Computational model of the predicted structure of EHC NMHC IIB, which lacks the rod domain. Images depicted in D and E courtesy of Prof. Simon Lovell (University of Manchester).

1.9 Aims and Objectives Reflecting the need to understand cardiac development, and our preliminary data concerning the EHC phenotype, the overall aim of this PhD project was to characterise the cardiovascular defects presented in the EHC mutant and to identify the disrupted process responsible for these abnormalities. The overarching hypothesis underpinning this thesis is that the EHC point mutation causes the global loss of function of NMIIB, which results in catastrophic deviation from the correct programme of cardiac development. This hypothesis was interrogated on three fronts. Firstly, we investigated whether or not the EHC mutation ablates NMIIB function. Secondly, we characterised the EHC cardiac phenotype to establish the nature of the processes detrimentally affected in the EHC line. Finally, we examined whether NMIIB was required in particular sub-populations of cardiac cells during cardiogenesis.

(1) Due to the nature of the ENU mutagenesis screen, it is not possible to state categorically that an identified point mutation in a given gene is the causative mutation of the observed phenotype. To overcome this, we performed a genetic complementation test with a known Myh10 null allele, denoted as Myh10∆. Homozygous Myh10∆ animals have previously been shown to display a Myh10 null phenotype (Ma et al., 2009; see Table 3.3 for a summary of phenotypic findings from previous studies of the Myh10 ablated mouse). We then analysed the phenotype of the resultant progeny for embryonic lethality and cardiac defects to determine whether or not allelic rescue between the lines had occurred.

(2) In order to identify the perturbed developmental processes that underpin the EHC phenotype, we employed immunohistochemistry to characterise the localisation of cellular components of the coronary vasculature. In addition, we examined the function of EHC epicardial cells by employing immunofluorescent microscopy and an in vitro migration assay to assess key aspects of epicardial behaviour including: migration, proliferation, and EMT.

(3) To establish the tissue-specific requirement for NMIIB during cardiogenesis, we utilised immunohistochemistry and western blotting to determine the normal expression profile of NMIIB in different cardiac tissues at different stages of development. Furthermore, we embarked upon a breeding strategy to generate both epicardial and myocardial-specific Myh10 knock out mice by using Cre-loxP genomic editing technology. We examined the embryo and heart morphology of the subsequent progeny to determine the effect of NMIIB loss specifically in these tissues.

Chapter 2: Materials and Methods

All protocols and procedures were undertaken at room temperature, unless otherwise stated.

2.1 Contribution Statement Day-to-day care of individual mouse strains and related animal husbandry (including ear punching of adult animals) was undertaken by the technical staff at the Biological Services Facility at the University of Manchester. The original Wt1-CreERT2 colony was maintained by Mark Evans and colleagues in the Biomedical Sciences Building at the University of Oxford.

Dr. Kathryn Hentges designed all primers and genotyping protocols. Sequencing of homozygous EHC Myh10 genomic DNA was performed by Drs. Karen Mitchell and Christopher Clowes (described at length in Ridge et al., Manuscript submitted, PLoS Genetics). I performed sequencing of the EHC/Myh10∆ embryos described in this thesis. Dr. Kathryn Hentges and myself devised the breeding schemes for the generation of tissue-specific Myh10 knockout animals. Dr. Kathryn Hentges performed the tamoxifen injections undertaken at the University of Manchester.

Dr. Kathryn Hentges and myself devised all experiments. I conducted all dissections and collected embryonic tissues for use in experiments. I similarly carried out all genotyping, sample processing, and optimisation of all experimental techniques detailed in this thesis. I carried out all immunohistochemistry and western blot experiments, except vimentin immunohistochemical staining of EHC mutant heart sections, which was undertaken by Irina-Elena Lupu.

I optimised the epicardial cell culture system in our laboratory based on helpful advice from Dr. Nicola Smart, University of Oxford. I performed the scratch wound migration assay with valuable advice from Dr. Patrick Caswell, University of Manchester. Dr. Caswell also provided useful comments for the cell tracking analysis. The optimisation of the lung epithelial cell culture system, cardiomyocyte culture system, and all immunocytochemistry experiments, was conducted by myself. The X-gal staining protocol was optimised by Dr.

Gennadiy Tenin. All images presented in this thesis were acquired by myself, unless otherwise stated. Similarly, I conducted all image and statistical analysis.

2.2 Animals

2.2.1 Ethical Statement All protocols were performed in accordance with UK Home Office Regulations under the personal license number 40/10715 (IEDB3DF8D), project license number 40/3406. Mouse colonies were maintained at the Biological Services Facility in the University of Manchester.

2.2.2 Mouse Lines The generation of the l11Jus27 mouse from which the EHC line was isolated has been described previously (Kile et al. 2003). The Wt1-CreEGFP (Wt1tm1(EGFP/cre)Wtp/J) line was purchased from the Jackson Laboratory (www.jax.org). The Wt1-CreERT2 (Wt1tm2(cre/ERT2)Wtp) founder males were provided by Professor Paul Riley at the University of Oxford as part of a collaboration funded by Boehringer Ingelheim Fonds. The α-MHC-Cre (B6.FVBTg(Myh6-cre)2182Mds/J) founder male was a generous gift from Dr. Elizabeth Cartwright at the University of Manchester. The Myh10flox/flox (Myh10tm7Rsad) mouse line was purchased from the MMRRC (www.mmrrc.org). To generate the Myh10∆ line, Myh10flox/flox mice were crossed to Tg(Nes-cre)Wme to generate global deletion of Myh10 exon 2.

2.3 Genotyping Animals

2.3.1 DNA Isolation from Adult Mouse Tissue Tissue was obtained by ear punching animals at weaning. Genomic DNA was prepared by adding 250µL of DNA lysis buffer (200mM NaCl2, 100mM Tris, 5mM EDTA, 0.2% (w/v) SDS, pH8.0) and Proteinase K (Roche Diagnostics Ltd.) to a working concentration of 80μg/mL. Samples were incubated at 55°C for a minimum of 4 hours. Following digestion, samples were centrifuged (Fisher Scientific, accuSpin Mirco) at 13,000 rpm to pellet cellular debris. The supernatant was transferred to a fresh 1.5 mL microcentrifuge tube and 300µL isopropanol was added. Samples were thoroughly mixed by inverting 7-8 times and

57 centrifuged again at 13,000 rpm for 20 minutes to pellet the precipitated DNA. The supernatant was carefully aspirated and 500µL 70% EtOH was added to each sample before briefly vortexing. Samples were centrifuged once again at 13,000 rpm for 10 minutes and the supernatant removed and disposed. The precipitated DNA pellet was then resuspended in an appropriate volume of ddH2O (typically 100-200µL depending on the size of the pellet).

2.3.2 Genotyping PCR Genotyping PCR samples were set up in 10µL reaction volumes, each containing; 5µL ddH2O, 3μL MyTaq Red Mix DNA polymerase (Bioline, BIO-25044), 1μL sample genomic DNA, and 1μL of the applicable forward and reverse primer mix (see Appendix 1). Reactions were placed in a thermocycler (peQlab, peQSTAR 2X Gradient) and appropriate programme run (see Appendix 2). Following the completion of the PCR cycle, reactions were stored at 4°C.

2.3.3 Agarose Gel Electrophoresis PCR products were separated by electrophoresis on 1-2% (w/v) agarose (Eurogentec, EP- 0010-05) gels. The gel mixture contained a 1:20,000 dilution of SafeView nucleic acid stain (NBS Biologicals) in TAE buffer (see Appendix 3). Gels were submerged in TAE buffer and run at 100V for 30-45 minutes, before imaging on a UV transilluminator (UVItec. BTS-20.M). HyperLadder 1Kb (Bioline, BIO-33053) was loaded at 5μL/lane as a DNA size standard.

2.4 Collection and Processing of Embryonic Tissue

2.4.1 Embryo Harvesting and Imaging Timed mating cages were set up and pregnancy determined by the presence of a vaginal plug. Pregnant female mice where sacrificed by cervical dislocation at the required stage of embryonic development. Embryos were removed in utero via cesarean section and placed in ice cold PBS (see Appendix 3) before being dissected out of the decidua and having extraembryonic membranes removed. Embryos were imaged using bright field Leica MZ6 dissecting microscope with Leica DFC420 digital camera. Required embryonic tissue was

58 dissected into fresh PBS using fine forceps and stored in fresh 1.5mL tubes for downstream analysis.

2.4.2 Tissue Fixation Embryos were fixed in 4% PFA (Sigma) in PBS, either at room temperature for 2 hours, or overnight at 4°C. Fixed tissue was then briefly washed in PBS, and dehydrated through washes in 50% MeOH:PBS, and 100% MeOH (2 x 5 minutes each). Dehydrated tissue was stored in fresh MeOH at -20°C until required.

2.4.3 Genotyping Embryos Tissue for genotyping was dissected from embryos and stored at -20°C. Yolk sac tissue was used for embryos ≤ E11.5, and tail clippings were used for older embryos. Genomic DNA was prepared as previously described (see Section 2.3.1). PCR reactions were similarly set up as previously described (see Section 2.3.2) with appropriate primers (see Appendix 1) and the appropriate PCR protocol was run (see Appendix 2). PCR products were then separated by electrophoresis and imaged as previously described (see Section 2.3.3).

2.5 Sequencing

2.5.1 PCR Product Purification For sequencing of the EHC mutation in EHC/∆ samples, appropriate bands were excised from the gel and centrifuged through a glass wool column at 5000 rpm for 5 minutes. An approximately equal volume of isopropanol, plus 10% of this volume of 3M NaOAc was added to each filtrate and mixed thoroughly. Samples were then stored overnight at -20°C. Samples were centrifuged at 13,000 rpm for 10 minutes, and the supernatant was carefully discarded before resuspending the DNA pellet in 20μL ddH2O. To ensure that the DNA had been purified, 3μL of each sample was run with 2μL Orange G loading dye (Sigma) on a 1% agarose gel and imaged as previously described (see Section 2.3.3).

2.5.2 Sequencing PCR Amplification and Purification Based on the intensity of the purified DNA band of each sample, an appropriate volume of purified DNA was added to 20μL cycle sequencing reactions (usually 5-8μL), each containing; 2μL BigDye dNTP mix (Applied Bioscience, BigDye Terminator V3.1), 3μL 59 BigDye sequencing buffer (Applied Bioscience, BigDye Terminator V3.1), 2μL EHC

Forward primer (see Appendix 1) and made up to 20μL with ddH2O. Samples were placed in a thermal cycler and the ABI sequencing programme was run (see Appendix 2). After completion, reaction volumes were transferred to fresh 1.5mL tubes and 16μL ddH2O plus 64μL 95% (v/v) EtOH was added to each before briefly vortexing. Samples were left at room temperature for 15 minutes to allow precipitation of extension products, and then centrifuged at 13,000 rpm for 20 minutes to pellet the precipitate. The supernatant was discarded and 250μL 70% EtOH was added to each sample, before briefly vortexing and centrifuging at 13,000 rpm for 10 minutes. Immediately after centrifugation, the supernatant was removed and pellets were allowed to air dry in a heat block at 95°C for 1 minute. Samples were sent to FMHS sequencing facility for analysis. Chromatographs were analysed using 4Peaks software (Version 1.7.2, available online at http://nucleobytes.com/index.php/4peaks).

2.6 Complementation Test Timed mating cages of Myh10∆/+ and EHC/+ male and female adult mice were set up and pregnancies were allowed to proceed to term. Litters were culled and tissue was obtained from tail clippings to genotype each sample for the Myh10∆ deletion and EHC mutation (as described in Section 2.2.1-3, and 2.4.1-2 respectively). The observed genotypes were compared to expected Mendelian ratios and data sets were analysed using a Chi squared test with 2 degrees of freedom (available online at http://graphpad.com/quickcalcs/chisquared1.cfm).

2.7 X-gal Staining for β-galactosidase Expression Freshly dissected embryonic hearts were incubated in X-gal staining solution (see Appendix 4) for 72 hours at room temperature on a bench top rocker (optimised incubation time, personal communication with Dr. Gennadiy Tenin). Samples were washed in PBS (2 x 5 minutes) and imaged using a Leica DFC420 camera mounted on a Leica MZ6 dissection microscope and Leica Firecam imaging software.

2.8 Tamoxifen Injections A 20mg/mL stock solution of tamoxifen (Sigma, T5648) was made by adding 300mg tamoxifen to 500µL 100% EtOH, briefly vortexing and then making up to 15mL with

60 sunflower oil (Sigma). This solution was incubated in a water bath at 50°C with occasional vortexing until all of the tamoxifen crystals had dissolved. The stock solution was aliquoted into 1.5mL tubes and stored at -20°C until required. For the injections, an aliquot was quickly thawed and administered via IP using a 29G insulin syringe (Terumo, MYJECTOR, U-100) at 100µL/25g body weight (80mg/Kg) to pregnant female mice (see Appendix 5 for injection schedule). Animals were observed daily to monitor for adverse side effects.

2.9 Histology and Immunohistochemistry Where a comparison between control and mutant is shown, sections were taken from an approximately equal tissue depth. A list of information on product codes and working dilutions of both primary antibodies and detection reagents is provided in Appendix 6 and Appendix 7 respectively. All primary antibodies were purchased commercially, and their specificity confirmed by quality control protocols performed by the manufacturer. Please see relevant antibody datasheets for details.

2.9.1 Paraffin Embedding and Microtome Sectioning Dehydrated tissue was washed in 100% EtOH (1 x 5 minutes), and then incubated in 50:50 Histoclear:EtOH (R.A. Lamb) for 30 minutes on an orbital shaker. Tissue samples were then incubated for 30 minutes at a time in 100% Histoclear, 50% Histoclear:paraffin (Paraplast®, Sigma), and then 100% paraffin at 65°C. Tissue in 100% paraffin was then transferred into individual plastic moulds (Dispomould, R.A. Lamb) and orientated using fine forceps and a dissection microscope (Leica MZ6). The paraffin was allowed to set overnight at 4°C before sample blocks were removed from the plastic moulds. Paraffin embedded samples were cut into 7μm sections using a microtome (Thermo Shandon Finesse). ‘Ribbons’ of sections were placed on the meniscus of a preheated 42°C water bath to facilitate specimen uncoiling. Sections were then adhered to glass microscope slides (Thermo Scientific, Polysine), and placed to dry on a heated slide rack at 40°C overnight. Slides were stored at room temperature until required.

2.9.2 OCT Embedding and Cryosectioning Embryonic tissue required for cryosectioning was dissected into ice cold PBS. Samples were then transferred to a thimble sized foil cup containing optimum cutting temperature (OCT) embedding media (R.A. Lamb) and snap frozen in liquid nitrogen. Samples were then stored at -80°C until required. Samples were cut into 14μm sections using a cryostat

61 microtome (Leica CM3050 S). Sections were adhered onto glass microscope slides (Thermo Scientific, Polysine) and stored at -80°C until required.

2.9.3 Haematoxylin Staining 7μm paraffin sections on microscope slides were de-waxed in xylene (2 x 10 minutes) and allowed to air dry for 15 minutes. Sections were then rehydrated by incubating in descending concentrations of EtOH in ddH2O (100%, 90%, 70%, 50% EtOH) for 30 seconds each, and then in 100% ddH2O for 2 x 1 minute. Sections were then incubated in freshly filtered acidified Harris’ haematoxylin (Thermo) for 2 minutes, before being washed in running tap water for a further 2 minutes. Sections were then dehydrated through an ascending EtOH concentration series (50%, 70%, 90%, 100% EtOH) for 30 seconds each. Sections were incubated in xylene (2 x 1 minute) before a coverslip was mounted to each slide using DEPEX mounting media (Fisher). Slides were imaged the following day using a Leica DFC420 camera mounted on a Leica DMLB2 brightfield microscope and Leica Application Suite (V2.8.1) imaging software.

2.9.4 Whole Mount Platelet Endothelial Cell Adhesion Molecule 1 (PECAM-1) Staining Whole mount immunohistochemistry was performed for PECAM-1 to permit the visualization of coronary endothelial cells on the surface of E16.5 embryonic heart samples. PECAM-1 is considered a classical endothelial cell marker, and has previously been used to highlight the endothelial components of the coronary vasculature (Liu and Shi, 2012). Embryonic hearts were removed from storage in 100% MeOH and transferred to 2mL tubes. The tissue was rehydrated through graded MeOH in PBS + 0.1% (v/v) Tween-20 (PBS-T) washes (75%, 50%, 25% MeOH to 100% PBS-T) for 1 x 5 minutes each. The endogenous peroxidase activity of the tissue was quenched by incubating in 3% hydrogen peroxide (H2O2) for 30 minutes. Tissue was then incubated in blocking buffer (10% goat serum (Vector) in PBS) to reduce non-specific antibody binding. Whole hearts were then incubated with rat anti-mouse PECAM-1 primary antibody (AbD Serotec, 1mg/ml stock) diluted 1:1000 in PBS-T + 1% (w/v) bovine serum albumin (BSA) overnight, with PBS-T + 1% BSA alone added to negative controls. The following day, samples were washed in PBS-T (3 x 5 minutes) before incubating with biotinylated goat anti-rat secondary antibody (Vector Labs, ABC Elite) diluted 1:500 in PBS-T, for 2 hours. Samples were washed with PBS-T (3 x 5 minutes) and incubated with avidin-biotin

62 horseradish peroxidase complex reagent (Vector Labs, ABC Elite kit, made up as per manufacturers protocol), for 30 minutes. Samples were again washed in PBS-T (3 x 5 minutes) and then incubated with DAB substrate reagent (Vector Labs, made up according to manufacturer’s protocol). Staining was allowed to develop to an appropriate level (typically 3-5 minutes) as determined by visual inspection using dissection microscope. To stop the staining reaction, sample tubes were flooded with tap water and left to wash for 30 minutes. Tap water was replenished and the staining allowed to resolve overnight. Hearts were imaged the next day using a Leica DFC420 camera mounted on a Leica MZ6 dissection microscope and Leica Firecam imaging software.

2.9.5 Smooth Muscle α-Actin (SMαA) Immunohistochemistry for SMA was performed on transverse sections of E16.5 embryonic heart samples to enable the visualization of the vascular smooth muscle cell components of the coronary vasculature. It has previously been shown that SMA is a marker of vascular smooth muscle cells (Skalli et al., 1989). 7μm paraffin sections on microscope slides were de-waxed in xylene (3 x 5 minutes) and allowed to air dry for 15 minutes. Sections were then rehydrated by incubating in descending concentrations of EtOH in PBS (100% EtOH, 50% EtOH:PBS, 100% PBS) for 1 x 5 minutes each. Rehydrated slides were then subjected to an antigen retrieval step by submerging in 10mM Sodium Citrate pH 6.0, and heating in a microwave (700 watt) for 20 minutes at 20% power. Whilst still submerged, slides were allowed to cool to ambient temperature before being briefly washed in dH2O. Individual tissue sections were isolated with a hydrophobic barrier pen (Daido Sangyo) and then incubated with 0.3% H2O2 for 30 minutes. Slides were then briefly washed with PBS and blocked in PBS + 10% Horse Serum (Vector) for 1 hour. Experimental sections were then incubated with mouse anti-mouse SMαA primary antibody (Sigma, A5228, ~2mg/ml stock), diluted 1:400 in PBS, for 48 hours at 4°C, whilst PBS alone was added to negative control sections. Following incubation with primary antibody, slides were washed in PBS (3 x 3 minutes) and all sections were incubated with biotinylated goat anti-mouse secondary antibody (Vector), diluted 1:1000 in PBS, for 2 hours. Slides were again washed in PBS (3 x 3 minutes) and then incubated with avidin- biotin horseradish peroxidase complex reagent (Vector, ABC Elite kit, made up according to manufacturer’s protocol) for 30 minutes. Slides were washed with PBS (3 x 3 minutes) and incubated with DAB substrate reagent (Vector, made up according to manufacturer’s protocol). Staining was allowed to develop to an appropriate level (typically 3-5 minutes) as determined by visual inspection using a dissection microscope. To stop the staining

63 reaction, slides were submerged in running tap water and left to resolve for 15 minutes. Slides were mounted with a coverslip using Immu-mount (Thermo Shandon), and sealed with nail varnish. Slides were imaged the following day using a Leica DFC420 camera mounted on a Leica DMLB2 brightfield microscope and Leica Application Suite (V2.8.1) imaging software. Images were processed using ImageJ software (Wayne Rasband, NIH, USA).

2.9.6 Wilms Tumour 1 (Wt1) Immunohistochemistry was performed on coronal sections of E14.5 embryonic heart samples to permit the detection and identification of both epicardial and epicardial-derived cells. It has previously been shown that Wt1 is one of a limited number of reliable of embryonic epicardial cell markers (Masters and Riley, 2014). As Wt1 is known to be strongly and continuously expressed in the developing mouse kidney from E10.5 (Rackley et al., 1993), staining of cardiac sections was performed in parallel to positive control staining of sectioned kidney tissue. 14μm cryosections on microscope slides were removed from storage and immediately fixed in 4% PFA for 15 minutes. Slides were washed in PBS (3 x 3 minutes) and then incubated with PBS + 0.1% (v/v) Triton X-100 for 15 minutes to enhance tissue permeabilisation. Slides were washed in PBS (3 x 3 minutes) and then blocked with blocking buffer (PBS + 1% (w/v) BSA, + 10% (v/v) normal goat serum (Vector)) for 1 hour. Experimental tissue sections were then incubated with rabbit polyclonal anti-mouse Wt-1 primary antibody (Calbiochem, CA1026) diluted 1:300 in blocking buffer, for 24 hours at 4°C. Blocking buffer alone was added to negative control sections. The next day, slides were washed in PBS (3 x 3 minutes) and all sections were then incubated with biotinylated goat anti-rabbit secondary antibody (Vector) diluted 1:500 in PBS + 0.1% (v/v) Triton X-100, for 2 hours. Slides were again washed in PBS (3 x 3 minutes) and incubated with Cy3 conjugated streptavidin (GE Healthcare, PA3001), diluted 1:3000 in PBS + 0.1% (v/v) Triton X-100, for 30 minutes, whilst protected from the light. Slides were then washed in PBS (3 x 3 minutes) before glass coverslips were mounted with Vectorshield + DAPI (Vector, H-1200) and sealed with nail varnish. Slides were stored in the dark at 4°C until imaging the following day. Images were collected on an Olympus BX51 upright microscope using 10x/ 0.30 UPlanFln and 20x/ 0.50 UPlanFLN objectives and captured using a Coolsnap ES camera (Photometrics) through MetaVue Software (Molecular Devices). Specific band pass filter sets for DAPI and Cy3 were used to prevent bleed through from one channel to the next.

2.9.7 Phospho-HistoneH3 (PHH3) Immunohistochemistry was performed on coronal sections of E14.5 embryonic heart samples to facilitate the detection of proliferating cells. PHH3 is a marker of mitosis and meiosis (Ribalta et al., 2004), and has previously been used to analyse proliferation in the embryonic heart (Iyer et al., 2016; Chakraborty and Yutzey, 2012). 14μm cryosections on microscope slides were removed from storage and treated as previously described (see Section 2.9.6), with some minor adjustments. Experimental tissue sections were incubated with rabbit anti-mouse PHH3 primary antibody (Merk Millipore, 06-570) diluted 1:300 in blocking buffer, overnight at 4°C. Following washing in PBS (3 x 3 minutes) all sections were then incubated with FITC conjugated goat anti-rabbit secondary antibody (Sigma, F9887) diluted 1:160 in PBS + 0.1% (v/v) Triton X-100, for 2 hours. Slides were again washed in PBS (3 x 3 minutes) and a glass coverslip was mounted to each with Vectorshield + DAPI. Slides were stored in the dark at 4°C until imaging the following day. Images were collected and captured as previously described (see Section 2.9.6). Specific band pass filter sets for DAPI and FITC were used to prevent bleed through from one channel to the next.

2.9.8 Snail Immunohistochemistry was performed on coronal sections of E14.5 embryonic heart samples to identify cells undergoing EMT. Snail is widely accepted as a universal marker of EMT activation (Vega et al., 2004). 14μm cryosections on microscope slides were removed from storage and treated as previously described (see Section 2.9.6), with some minor adjustments. Slides were blocked with blocking buffer (PBS + 10% (v/v) normal horse serum (Vector)) for 1 hour. Experimental tissue sections were then incubated with goat polyclonal anti-mouse Snail primary antibody (Abcam, ab53519) diluted 1:100 in PBS + 0.1% (v/v) Triton X-100, for 1 hour. Following washing in PBS (3 x 3 minutes) all sections were then incubated with biotinylated hoarse anti-goat secondary antibody (Vector) diluted 1:500 in PBS + 0.1% (v/v) Triton X-100, for 1 hour. Slides were again washed in PBS (3 x 3 minutes) and incubated with Cy3 conjugated streptavidin (GE Healthcare, PA3001), diluted 1:1000 in PBS + 0.1% (v/v) Triton X-100, for 15 minutes, whilst protected from the light. Slides were then washed in PBS (3 x 3 minutes) before glass coverslips were mounted as previously described (see Section 2.9.6). Slides were stored in the dark at 4°C until imaging the following day. Images were collected as previously described (see Section 2.9.6). Specific band pass filter sets for DAPI and Cy3 were used to prevent bleed through from one channel to the next.

2.9.9 Vimentin Immunohistochemistry was employed on coronal sections of E14.5 embryonic hearts to detect and identify cardiac fibroblasts. Vimentin is a marker of mesenchymal cells, and has previously been used to detect cardiac fibroblasts in the embryonic mouse heart (Camelliti et al., 2005; Zamora et al., 2007). 14μm cryosections on microscope slides were removed from storage and treated as previously described (see Section 2.9.6), with some slight adjustments. Slides were blocked with blocking buffer (PBS + 10% (v/v) normal goat serum) for 1 hour. Experimental sections were then incubated with rabbit anti-mouse vimentin primary antibody (Proteintech, 10366-1-AP), diluted 1:50 in PBS + 0.1% (v/v) Triton X-100 overnight at 4°C. Slides were washed with PBS (3 x 3 minutes) and then incubated with FITC conjugated goat anti-rabbit secondary antibody diluted 1:160 in PBS + 0.1% (v/v) Triton X-100 for 2 hours. Sections were then washed with PBS (3 x 3 minutes) and a coverslip was mounted to each with Vectorshield + DAPI. Images were collected and captured as previously described (see Section 2.9.6). Specific band pass filter sets for DAPI and FITC were used to prevent bleed through from one channel to the next.

2.9.10 Non-Muscle Myosin II Isoforms A, B and C Transverse 7μm sections of E14.5 embryos on microscope slides were de-waxed in xylene (2 x 10 minutes), and then rehydrated though descending EtOH concentrations in PBS (100/90/70/50%) for 30 seconds each. Sections were equilibrated in 100% PBS (2 minutes) and then incubated in PBS + 0.1% (v/v) Triton X-100 for 15 minutes. Tissue sections were blocked in PBS + 1% (w/v) BSA for 1.5 hours, and then incubated in rabbit anti-mouse NMIIA, B, or C primary antibodies (Biolegend, PRB-440P, -445P, and -444P respectively), diluted 1:500 in PBS + 0.1% (v/v) Triton X-100 for 1 hour. Slides were washed in PBS (3 x 3 minutes) and then incubated in biotinylated goat anti-rabbit secondary antibody, diluted 1:500 in PBS + 0.1% (v/v) Triton X-100 for 1 hour. Following an additional PBS wash (3 x 3 minutes), sections were incubated with AlexaFluor-488 conjugated streptavidin (Invitrogen, S11223), diluted 1:1000 in PBS + 0.1% (v/v) Triton X-100 for 30 minutes. Slides were then washed with PBS (3 x 3 minutes) and coverslips were mounted with Vectorshield + DAPI. Slides were stored at 4°C and protected from light until required for imaging. Images were collected and captured as previously described (see Section 2.9.6). Specific band pass filter sets for DAPI and FITC were used to prevent bleed through from one channel to the next.

2.10 Cell Culture

2.10.1 Culture of Epicardial Cells from Embryonic Heart Explants Embryos were harvested at E11.5 as previously described (Chen et al., 2002), with some modifications (see Section 2.4.1). Briefly, embryonic hearts were dissected into fresh PBS, and the atria and outflow tract were removed. Ventricular tissue was then carefully dissected into four pieces of comparable size, and transferred to a 0.5mL tube containing complete epicardial cell culture media (Dulbecco’s Modified Eagle Medium (DMEM) (Sigma, D5796) supplemented with 15% (v/v) heat inactivated foetal bovine serum (FBS) (Gibco, 10500064) and 1% (v/v) Penicillin/Streptomycin (Sigma, P0781)). Once all hearts had been dissected, the ventricular pieces were gently agitated to help to remove excess blood cells. Each piece of ventricular tissue was then transferred to a single well of a 24 well tissue culture plate (Corning) containing a 13mm diameter coverslip pre-coated with 0.1% (w/v) gelatin (Sigma, G2500) (1 hour incubation at 37°C). For the scratch-wound experiments (see Section 2.10.3), explants were directly placed in 0.1% (w/v) gelatin coated

24-well plates. Explants were incubated at 37°C with 5% CO2 for 48 hours, at which time a ‘halo’ of epicardial cells could be clearly observed surrounding the explant. The explant was gently removed with forceps, and care taken not to disturb the epicardial monolayer. Following removal of the explant, the epicardial cultures were washed with complete media (2 x 5 minutes) and returned to the incubator. After 24 hours, the area where the explant used to adhere had been completely covered by the epicardial monolayer. Media was replaced at least every 3 days until the cultures were required for experiments.

2.10.2 Immunocytochemistry After a total of 72 hours (following dissection), epicardial cell cultures were washed with tissue culture grade PBS + MgCl2 and CaCl2 (Sigma, D8662) (2 x 5 minutes) and then fixed in 4% PFA for 10 minutes on an orbital shaker. Cells were then washed with PBS (2 x 2 minutes) and cell monolayers were permeabilised by incubating in PBS + 0.1% (v/v) Triton X-100 for 15 minutes. Cultures were then blocked in either PBS + 10% (v/v) goat serum, or PBS + 1% (w/v) BSA, for at least 1 hour before addition of relevant primary antibody diluted in PBS + 0.1% (v/v) Triton X-100 for 1 hour (see Appendix 6 for antibody details and dilutions). Unbound antibody was removed by washing the cultures with PBS (2 x 2 minutes) before addition of appropriate biotinylated secondary antibody

67 (see Appendix 7) diluted 1:500 in PBS + 0.1% (v/v) Triton X-100 for 1 hour. Cultures were once again washed in PBS (2 x 2 minutes) and then incubated in Cy5 conjugated streptavidin (GE Healthcare, PA45001) diluted 1:500 in PBS + 0.1% (v/v) Triton X-100 for 30 minutes. Cells were washed with PBS (2 x 2 minutes) and then counter-stained with 100nM rhodamine-phalloidin (Cytoskeleton, PHDR1) where appropriate for 30 minutes as per manufacturer’s instructions to allow visualization of the actin cytoskeleton. Following a final wash with PBS (3 x 2 minutes), coverslips were mounted onto microscope slides using Vectorshield + DAPI mounting media (Vector, H-1200), and stored at 4°C in the dark until imaging. Images were collected and captured as previously described (see Section 2.9.6). Specific band pass filter sets for DAPI and Cy3 and Cy5 were used to prevent bleed through from one channel to the next.

Please note, for MRTF-A immunocytochemistry, 48 hour cultures were serum-starved for 24 hours by incubating in DMEM supplemented with 1% (v/v) Penicillin/Streptomycin only prior to the start of the staining protocol. Immunocytochemistry was carried out as described above.

2.10.3 Scratch-Wound Assay Epicardial cell cultures were generated and maintained as previously described (see Section 2.9.1). After a total of 72 hours in culture post-dissection, epicardial monolayers were scratched using a sterile P20 pipette tip, and immediately washed with complete media (2 x 5 minutes) to remove cellular debris. Fresh media was added, and the 24 well tissue culture plate was placed inside an AS MDW live cell imaging system (Leica). Phase contrast images were captured at 10-minute intervals for a period of 20 hours using a x20/0.5 Plan Fluotar objective with Image Pro 6.3 (Media Cybernetics LtD) imaging software. Point visiting was used to permit multiple fields of view to be imaged within the same time course. This was achieved by pre-saving field of view co-ordinates prior to initiating the imaging time course. Cells were maintained at 37°C and 5%CO2 for the duration of the experiment (20 hours).

2.10.4 Culture of Lung Epithelial and Fibroblast Cells Epithelial and fibroblast cells from the embryonic mouse lung were cultured as previously described (Lebeche et al., 1999; del Moral et al., 2006), with some modifications. Briefly, the distal lobe of the left E16.5 lung was dissected from the embryo, cut into 3-4 pieces and washed in PBS for 30 minutes to remove red blood cells. Tissue pieces were pelleted at

500RCF for 5 minutes at 4°C and then resuspended in 1% trypsin (Sigma, T3924). Large pieces of tissue were broken up by triturating through a 21G needle (Terumo). Cell suspensions were incubated at 37°C for 15 minutes with occasional agitation to homogenise the tissue pieces. Cell suspensions were pelleted at 500RCF for 5 minutes at 4°C and then washed with DMEM (Sigma, D6429) supplemented with 10% (v/v) FBS (2 x 5 minutes) to stop trypsin-mediated tissue degradation. Cell suspensions were pelleted again at 500G and then resuspended in DMEM (without serum) supplemented with 20 units/mL DNAse (Promega, M6101). Suspensions were incubated at 37°C for 10 minutes to reduces the suspension viscosity due to DNA contamination from damaged cells. Samples were pelleted as described above and resuspended in 1mL DMEM + 10% (v/v) FBS and 1% (v/v) Penicillin/Streptomycin. The cell suspension was filtered through pre- wetted 100, 70, 40µm cell strainers (Corning). After each filtration, 500µL media was added to ensure maximum recovery of cells through the strainer. The filtrate was made up to 3mL by adding complete media, and plated into uncoated 24 well tissue culture plates (500µL/well). Plates were incubated for 2 hours to allow the attachment of mesenchymal cells. Suspended cells were removed and re-plated into 0.1% (w/v) gelatin pre-coated wells (500µL/well). Fresh media was added to the mesenchymal cell population, and plates were incubated at 37°C with 5% CO2 for 24 hours. The next day, the media was replaced in all cell cultures to remove remaining red blood cells. Cultures were returned to the incubator for 48 hours. Subsequent immunocytochemistry was carried out as detailed above (see Section 2.10.2).

2.10.5 Culture of Cardiomyocytes and Cardiac Fibroblasts from Embryonic Hearts Both cardiomyocytes and cardiac fibroblasts were isolated and cultured from embryonic hearts as previously described (Louch et al., 2011; Sreejit et al., 2008), with some modifications. Briefly, embryonic hearts were dissected from E15.5 embryos into ice cold

PBS without MgCl2 and CaCl2 (Sigma, D8537). The atria and great vessels were excised, and hearts were incubated in DMEM (Sigma, D5796 supplemented 1% (v/v) Penicillin/Streptomycin) with 1mg/mL collagenase (Sigma, C1889) at 37°C for 45 minutes with occasional agitation to dissociated the heart tissue. Cell suspensions were pelleted at 500RCF for 10 minutes at 4°C, and each pellet was resuspended in 500µL DMEM (+ 15% (v/v) FBS, 1% (v/v) Penicillin/Streptomycin). Cell suspensions were plated onto 0.1% (w/v) gelatin pre-coated coverslips in 24 well tissue culture plates and incubated at 37°C with 5% CO2 for 2-3 hours to allow attachment of non-myocyte cells. Unattached cells were removed by aspirating the supernatant and transferring to fresh gelatin coated

69 coverslips. Fresh media was added to the fibroblast-enriched cell culture (500µL/well) and plates were returned to the incubator for 24 hours. The media was replaced to remove red blood cells, and plates were returned to the incubator for a further 5 days. Media was replaced every 2 days. Subsequent immunocytochemistry was carried out as described above (see Section 2.10.2).

2.11 Image Analysis and Statistics Following capture, all microscopy images were processed and analysed using ImageJ (http://rsb.info.nih.gov/ij) (Wayne Rasband, NIH, USA). Images were enhanced by adjusting the brightness and contrast settings relative to negative control images.

All statistical analysis was carried out using Prism 6 (GraphPad Software, www.graphpad.com/scientific-software/prism/). To ensure that the correct statistical test was used in each instance, the data was interrogated by the D’Aostino-Pearson normality test. If the data passed this test, a parametric test (un-paired, 2-tailed t-test) was employed. If the data failed the D’Aostino-Pearson normality test (i.e. data deviated significantly from Gaussian distribution), a non-parametric test (Mann-Whitney U test) was utilised. Graphical representation of these data is always shown +/- standard error of the mean (SEM).

2.11.1 Analysis of EPDC Migration In Vivo Images acquired from immunohistochemistry experiments (see Section 2.9.6) were analysed for the presence of Wt1 positive cells, determined by the co-localisation of Wt1 signal with the DNA marker, DAPI. As previously mentioned, Wt1 is known to be one of a limited number of reliable markers of both the embryonic epicardium, and epicardial- derived cells (Masters and Riley, 2014). A line was manually drawn to delineate the apical boundary of the epicardium on x20 microscope images. The distance between the centre of all Wt1 positive cell nuclei and this line was manually measured and calibrated from pixels to micrometres. One x20 image (field of view) was analysed per heart section. In total, 3 non-consecutive sections were used per sample, and three independent samples were used for both control (EHC/+) and mutant (EHC/EHC) genotypes. A combined total of 975 (control) and 1180 (mutant) migration measurements were analysed by Mann-Whitney U test (p<0.0001).

2.11.2 Analysis of Cell Proliferation In Vivo Immunohistochemistry images (see Section 2.9.7) were analysed for positive PHH3 staining, as determined by broad co-localisation of PHH3 signal with DAPI. A line was manually drawn to delineate the boundary between the epicardium (cells on the immediate apical tissue surface) and the myocardium in x20 magnification images. The total number of cells in the epicardium (Xepi) vs the total number of PHH3 positive cells in the epicardium (Yepi) were counted and the proliferation rate (Pepi) was calculated (Pepi = (Yepi/

Xepi) x100). For the myocardium, a 100µm square box was superimposed onto the myocardial tissue in each image. Similarly, the total number of cells (Xmyo) and total number of PHH3 cells (Ymyo) inside the box were counted, and proliferation rate (Pmyo) calculated

(Pmyo = (Ymyo/ Xmyo) x100). One x20 image (field of view) was analysed per heart section. In total, 3 non-consecutive sections were used per sample, and three independent samples were used for both control (EHC/+) and mutant (EHC/EHC) genotypes. Cell proliferation rates were compared between the control (n=18) and mutant (n=18) epicardium, and similarly the control (n=18) and mutant (n=18) myocardium by unpaired t test (p<0.0001 and p=0.1684 respectively).

2.11.3 Analysis of Epicardial Cell Migration In Vitro Phase contrast image stacks generated from the scratch wound assay (see Section 2.10.3) were compiled for analysis in ImageJ. For each field of view, 10 cells at the leading edge of the scratch (also referred to as the denuded area) were identified in the first (time zero - T0) image. The migration of these cells was tracked through the image series until the cells had completely re-covered the denuded area using the MTrackJ plugin. The centre of the cell nucleus was used as a reference point in each image. In the event that a selected cell divided during the image series, tracking was terminated 1 hour (6 images) prior to cell division. All cells were tracked for a minimum of 6 hours. In total, 240 control (+/+ or Myh10∆/+) and 270 mutant (Myh10∆/Myh10∆) cells were tracked. Tracking measurements encompassed migration speed and directional persistence; control and mutant values were compared by Mann-Whitney U test (p=0.6717 and p= 0.2494, respectively).

2.12 Western Blot Analysis A full list of the western blot solutions used in these experiments is provided in Appendix 8.

2.12.1. Generation of Protein Extracts A table of volumes used to generate protein extracts and run in each experiment is provided in Appendix 9.

2.12.1.1 Generation of Protein Extracts from Whole Embryonic Hearts Embryonic hearts were harvested as previously described (see Section 2.4.1). Atria, aorta and pulmonary artery tissue was carefully removed using forceps, and ventricular tissue was stored in a 1.5mL tube at -80°C until required for experiments. Samples were rapidly thawed and 50µL RIPA buffer (see Appendix 8) was added to each. Samples were homogenized using a plastic pestle and left on wet ice with occasional vortexing for 1 hour. Insoluble material was pelleted by centrifuging at 14,000RCF at 4°C for 1 hour, after which, the supernatant was removed and transferred to a fresh 1.5mL tube. Samples were snap frozen on dry ice and stored at -80°C for future use.

2.12.1.2 Generation of Protein Extracts from Cultured Epicardial Cells and Explants Epicardial cell cultures were established as previously described (see Section 2.10.1). Following removal, heart explants were pooled according to genotype (control = +/+ and ∆/+, mutant = ∆/∆, explants from a total of six hearts/genotype), and 50µL RIPA buffer was added to each pool (see Appendix 9). Protein extracts were subsequently prepared as previously described (see Section 2.12.1.1).

After a total of 72 hours in culture, media was carefully aspirated and 250µL epicardial extraction buffer (see Appendix 8) was added to each epicardial culture. The epicardial monolayer was removed from the tissue culture plastic with a modified cell spreader (Falcon) and transferred to a 1.5mL tube. Cell suspensions were pelleted at 14,000RCF at 4°C for 5 minutes, and the supernatant was carefully removed. Samples were pooled according to genotype, and this process was repeated to harvest the remaining cultures. After addition of the final cell suspension, pooled samples were pelleted at 14,000RCF at

4°C for 10 minutes and the supernatant was removed. Visible pellets were resuspended in 50µL 4x sample loading buffer (SLB) (see Appendix 8), boiled at 95°C for 5 minutes, and then snap frozen on dry ice before being stored at -80°C until required for experiments. Prior to use in SDS-PAGE (see Section 2.12.2), epicardial protein samples were boiled at 95°C a further 2 times for 5 minutes.

2.12.2 SDS-PAGE An appropriate volume of 4 x SLB was added to each sample (see Appendix 9), and boiled at 95°C for 5 minutes. Tubes were centrifuged briefly to collect the mixture at the bottom of the tube and homogenized by pipetting up and down several times. The total volume was loaded on to a stacking gel and separated on a 12% polyacrylamide gel (see Appendix 8). A volume of 5µl/lane protein standard ladder (Biorad, Kaleidoscope Precision Plus) was loaded as a size marker. Gels were run vertically in running buffer (see Appendix 8) at 100V for approximately 1.5 hours, until the dye front reached the bottom of the gel.

2.12.3 Western Blotting Protocol Gels were briefly washed in transfer buffer (see Appendix 8) and proteins were then transferred to PVDF membrane (GE Healthcare, Hybond P) for 2 hours at 200mA in transfer buffer. Membranes were blocked in TBS-T (see Appendix 8) with 5% (w/v) powdered skimmed milk (Tesco) overnight at 4°C. The membrane was cut into two sections, just below the 75kDa marker, and the top portion was incubated with relevant primary antibody (see Appendix 6), with gentle agitation for 2 hours. The bottom section was incubated in HRP conjugated mouse anti- β-actin (Sigma, A3854) diluted 1:200,000 in TBS-T with gentle agitation for 4 hours as a loading control. The upper section was washed thoroughly with TBS-T (3 x 15 minutes) and then incubated with donkey anti- rabbit HRP conjugated secondary antibody (Santa Cruz, sc-2313) diluted 1:1000 in TBS-T for 1 hour at room temperature. Both sections of the membrane were then washed with TBS-T (3 x 15 minutes) and incubated in enhanced chemiluminescence reagent (Thermo Fisher, 34080) for 5 minutes according to the manufacturer’s protocol. Membranes were imaged simultaneously on a chemiluminescent imaging dock (Biorad) using an appropriate exposure time to maximise signal without reaching saturation.

73 2.12.4 Densitometry Analysis Western blot images were analysed using ImageLab software (Biorad, http://www.biorad.com/en-uk/product/image-lab-software). Background subtraction was set to 10mm, and the area under the peak curve was used to calculate total signal intensity of each band. Signal intensity values for the peak of interest were normalised to that of the β-actin loading control in each lane (see Appendix 10), to obtain a single relative abundance value for the peak of interest (detailed in Appendix 11).

Chapter 3: Characterisation of EHC and Myh10∆ Mutant Embryos

3.1 Introduction

3.1.1 Background to Congenital Hydrocephalus in the NMIIB Ablated Mouse Hydrocephalus is characterised by the dilation of the brain ventricles due to an excess accumulation of cerebral spinal fluid (CSF) (Vogel et al., 2012). This fluid accumulation can lead to the damage and destruction of the brain tissue. The flow of CSF originates from the lateral ventricle, passing to the third and then fourth ventricle via narrow channels before being drained into the lymphatics in the subarachnoid space (Fig 3.1) (Fliegauf et al., 2007). Congenital hydrocephalus can arise through multiple causes; impaired CSF flow, excess CSF production, or reduced CSF absorption (Crews et al., 2004). There are two major classes of hydrocephalus; when the interruption of CSF flow occurs due to blockage of the interconnecting channels between ventricles (non-communicating/obstructive), and when adsorption of CSF in the subarachnoid space is impaired following exit from the ventricles (communicating) (Vogel et al., 2012). The most common cause of congenital hydrocephalus is aqueductal stenosis, where the connection between the third and fourth ventricle (the aqueduct of Sylvius) becomes obstructed (Hydrocephalus Association, www.hydroassoc.org). It is estimated that congenital hydrocephalus affects 1-3 out of every 1000 live births, and represents the most common pediatric neurological condition requiring surgical intervention (Casey et al., 1997). Interestingly, the presence of non- communicating hydrocephalus has been associated with an increased prevalence of cardiovascular disease in adults (Eide and Pripp, 2016)

It is thought that both genetic and environmental factors, including exposure to certain teratogenic compounds, contribute to the development of congenital hydrocephalus (Gilbert-Barness, 2010; Vogel et al., 2012). However, the delineation of the causative genetic mutations associated with this condition is hampered by the polygenic nature of hydrocephalus and background specific modifying elements (Vogel et al., 2012). This later component is likely to be responsible for the varying severity of the hydrocephalus phenotype displayed in mutated mouse strains that harbour the same genetic mutation, but are derived from different genetic backgrounds (Homanics et al., 1995; Vogel et al., 2012).

Figure 3.1: Schematic Representation of the Ventricular Segments During Embryonic Mouse Brain Development (A) Illustration of the three primary brain ventricles in the E11.5 mouse embryo; telencephalic (red region), mesencephalic (blue region), and rhombencephalic (yellow region). (B) These primary ventricles develop into the two lateral ventricles (red region), as well as the third (blue region), and fourth (yellow region) ventricles. The production of cerebral spinal fluid predominantly originates from the lateral ventricles, and flows through the ventricular system until being drained into the subarachnoid space via the fourth ventricle. The cerebral aqueduct (Aqueduct of Sylvius), which connects the third and fourth ventricles (not shown), forms later in embryonic development. III – third ventricle, IV –fourth ventricle, LV – lateral ventricles, Mes – mesencephalic ventricle, Rho – rhombencephalic ventricle, SAS – subarachnoid space, Tel – telencephalic ventricle. Adpated from Fliegauf et al., 2007.

Although few genes have been directly associated with this pathology, a broad variety of genetically engineered mice have been shown to present with autosomal recessive congenital hydrocephalus (Vogel et al., 2012). Indeed, previous investigators have reported that the global NMIIB knock out mouse displays congenital hydrocephalus, accompanied by cardiac abnormalities (discussed at length in Section 3.1.3) (Tullio et al., 2001; Bridgman et al., 2001; Ma et al., 2007; Ma et al., 2009). Whilst the precise nature and extent of NMIIB function in the brain is yet to be determined, it is clear that NMIIB plays a key role during normal brain development that is functionally distinct to its role during cardiogenesis.

3.1.2 Coronary Vessel Formation During early cardiogenesis, the metabolic demands of the working myocardium are met by the passive diffusion of oxygen and nutrients across the myocardial wall. However, as the ventricles expand and thicken due to rapid growth and myocardial proliferation, diffusion becomes inadequate, and delivery of these components is replaced by infiltration of the heart by a vascular network. The development of this coronary vasculature and its connection to the systemic embryonic circulation is essential for continued embryonic development (Reese et al., 2002; Olivey and Svensson, 2010; Riley and Smart, 2011). The formation of the mature coronary vasculature is wholly reliant upon the correct orchestration and tight regulation of a wide variety of processes, which encompass vasculogenic, angiogenic, and arteriogenic events (Tomanek, 2005; Olivey and Svensson, 2010; Riley and Smart, 2011).

3.1.2.1 Role of the Epicardium in Coronary Vessel Formation It is well established that the epicardium contributes to the formation of the coronary vasculature. The initiation of EMT in a subset of epicardial cells gives rise to the EPDC population, which delaminate from the epicardial epithelium and invade the underlying myocardium. The EPDC population is multipotent, and it is widely accepted that the majority of perivascular fibroblasts and the vascular smooth muscle cell (vSMC) components of the coronary vessels derive from these progenitor cells (Gittenberger-de Groot et al., 1999; Dettman et al., 1998; Vrancken Peeters et al., 1999). Differentiation of EPDCs into cardiomyocytes has also been reported (Zhou et al., 2008; Cai et al., 2008), although this has recently been challenged (Christoffels et al., 2009; Kispert, 2012). Additionally, the generation of vascular endothelial cells from EPDCs is the subject of continued debate; whilst Katz and colleagues (2012) have described the formation of

77 coronary endothelial cells from distinct sub-populations of the epicardium (Katz et al., 2012) it appears that the majority of the coronary endothelial cell lineage is not epicardial- derived (Singh and Epstein, 2013; Riley and Smart, 2011; Tian et al., 2015).

3.1.2.2 Vascular Endothelial Capillary Formation It has been shown that at approximately E11.5, mesenchymal cells in the sub-epicardial space coalesce to form the primitive endothelial coronary plexus (Reese et al., 2002). These primitive vessels spread over the ventricular surface by angiogenesis until approximately E13.5, and precede the formation of mature vascular structures that adorn the fetal heart (Sylva et al., 2014). Many studies have utilised either chimera experiments or Cre-loxP genomic editing technology to determine the cellular origins of the endothelial capillary plexus (Riley and Smart, 2011). The origin of these initial vascular precursors is unclear, with a number of conflicting studies providing evidence of contributions from the epicardium (Katz et al., 2012), endocardium (Wu et al., 2012), and sinus venosus (Red- Horse et al., 2010; Tian et al., 2013b). However, recent fate mapping studies have clarified this contention, and indicate that the coronary endothelium is likely to derive predominantly from the sinus venosus, with a modest contribution from the ventricular endothelium, specifically to vessels within the interventricular septum (Red-Horse et al., 2010; Tian et al., 2015; Zhang et al., 2016; Chen et al., 2014). Concurrently, it is suggested that the initial primitive plexus arises from the endothelial outgrowth of the sinus venosus, which is subsequently consolidated by ‘blood islands’, which are derived from endocardial budding of the ventricular endothelium (Morabito et al., 2002; Riley and Smart, 2011; Red- Horse et al., 2010). It is also suspected that the endothelium makes a continued contribution to the coronaries following the establishment of the endothelial plexus, and in the neonatal heart (Riley and Smart, 2011; Tian et al., 2015). However, before drawing definitive conclusions on the cellular origins of the coronaries, it is important to consider the technical limitations of the molecular tools employed, as well as inter-species variation of the animal models used in these studies. The close proximity of the sinus venosus to the proepicardial organ makes it difficult to accurately quantify the contribution of either tissue to the building of the endothelial plexus architecture (Riley and Smart, 2011; Zhang et al., 2013b).

3.1.2.3 Vascular Smooth Muscle Cell Recruitment Following the establishment of a rudimentary coronary capillary plexus at E13.5, an intricate remodelling process is initiated which prompts the differentiation and maturation of the venous and arterial coronaries (Brade et al., 2013). The molecular mechanisms that determine endothelial cell fate to either a venous or arterial identity remain largely unclear (Riley and Smart, 2011). However, it is widely acknowledged that following the establishment of patency between the coronary plexus and the aortic root, blood flow/pressure stimuli and other haemodynamic properties contribute at least in part to the induction of vSMC recruitment to capillaries and the establishment of arterial identity (Luttun and Carmeliet, 2003; Vrancken Peeters et al., 1999). Vascular SMC recruitment is required to provide mechanical support to nascent vessels bearing a blow flow load. This model of vSMC recruitment dictates a requirement for reciprocal signalling between the capillary vascular endothelium and smooth muscle cells to guide their migration. In this regard, the PDGF-B (platelet-derived growth factor B)/PDGFR-β (platelet-derived growth factor receptor β) signalling axis has been implicated in vSMC recruitment in the wider embryo, as endothelial expression of PDGF-B propagates the proliferation and recruitment of PDGFR-β positive vSMCs (Hellstrom et al., 1999; Tomanek et al., 2008). Similarly, mice deficient for all VEGF isoforms except VEGF120, display reduced SMC recruitment to coronary arteries (Carmeliet et al., 1999; van den Akker et al., 2007), although it is unclear whether loss of VEGF impacts directly on vSMC migration, or via reducing other recruitment factors (Luttun and Carmeliet, 2003).

In addition, TGF-β1 signalling is required for both the recruitment and differentiation of vSMC, as mutations in both TGF-β1 and its receptors TβRII (TGF beta receptor 2) and Alk5 (TGFβ type 1 receptor kinase) cause a reduction of smooth muscle in the vascular wall (Fuster et al., 2005). Expanding upon this, epicardial-specific ablation of Rbpj revealed that Notch actively cooperates with TGFβ signalling to promote vSMC differentiation in EPDCs (Grieskamp et al., 2011). Furthermore, the differentiation of EPDCs into vSMC is known to involve the regulation of RhoA-Rho kinase signalling by serum response factor (SRF) (Lu et al., 2001). Mice deficient for β-catenin similarly show impairment of vSMC differentiation and recruitment, suggesting a role for canonical Wnt-β-catenin signalling in this process (Zamora et al., 2007).

79 3.1.3 Previously Reported Myh10 Knock Out Animals Previous studies have established that the Myh10 knock out mouse displays embryonic defects in the development of the brain and heart (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003). Global deletion of Myh10 results in embryonic hydrocephalus and homozygous mutant embryos do not survive after perinatal day 0 (Tullio et al., 1997). Global ablation of Myh10 causes defects in the development of the outflow tract, resulting in either DORV, or overriding aorta accompanied by muscular obstruction of the right ventricle (Tullio et al., 1997). Loss of Myh10 in the heart has also been shown to result in cardiomyocyte hypertrophy (Tullio et al., 1997; Ma et al., 2009) and multinucleated cardiomyocytes arising from cytokinesis dysfunction (Takeda et al., 2003; Ma et al., 2010). Mice heterozygous for the Myh10 null allele do not display haploinsufficiency and develop normally (Tullio et al., 1997). Indeed, animals expressing as little as 12% of wild type levels of Myh10 in the heart do not display cardiac defects at birth (Uren et al., 2000). Interestingly, genetic replacement of NMIIB with a targeted NMIIA knock in at the Myh10 locus rescues brain, but not cardiac defects (Bao et al., 2007). The majority of homozygous mutants harbouring this modification display embryonic lethality, likely due to cardiac defects, including ventral septal defects, and either the presence of DORV, or overriding aorta (Bao et al., 2007). This highlights the specific requirement for NMIIB during cardiogenesis. A summary of the phenotypic findings from studies on the Myh10 ablated mouse discussed here is provided in Table 3.3 in the Section 3.3.

The underpinning cause of the hydrocephaly phenotype of Myh10 null mice is yet to be definitively determined; NMIIB has been shown to play a role in the outgrowth of neuronal growth cones (Tullio et al., 2001; Bridgman et al., 2001), as well as playing a more structural function in the neuroepithelium to prevent blockage of the spinal canal by epithelial cells (Ma et al., 2007). In addition, it appears that the hydrocephalus phenotype in Myh10 disrupted mice demonstrates a gene dosage dependency, which dictates the severity and onset of brain abnormalities (Uren et al., 2000). Previous reports have suggested that the hydrocephalus phenotype is caused by a loss of the structural integrity of the spinal canal neuroepithelium, resulting in erroneous occlusion of the spinal canal cavity (Tullio et al., 2001; Ma et al., 2007). The disruption to the neuroepithelium has been directly attributed to loss of NMIIB, where it has been proposed to function as a scaffolding protein component of an apical mesh-like structure, also containing β-catenin and N- cadherin (Ma et al., 2007). The authors of this study propose that loss of NMIIB leads to the collapse of this mesh structure, allowing the invasion and subsequent obstruction of the spinal canal (Ma et al., 2007). Interestingly, the structural integrity of the mesh can be

80 reestablished by the expression of a motor-impaired NMIIB isoform, suggesting that NMIIB motor function is not required in the brain. In addition, these authors failed to detect NMIIB phosphorylation (a read out for NMIIB motor activity) in these mice, augmenting the proposal that NMIIB plays a structural, rather than a motor role during brain development (Ma et al., 2007).

Intriguingly, tissue-specific ablation of Myh10 via Cre recombinase-loxP technology in either the brain (driven by Nestin promoter activity) or heart (driven by α-MHC promoter activity) circumvents the embryonic lethality phenotype of the global knock out (Ma et al., 2009). Whilst deletion of Myh10 in the brain causes lethality between postnatal days 12-22 due to severe hydrocephalus, cardiomyocyte-specific Myh10 ablation does not cause lethality, and animals survive into adulthood (Ma et al., 2009). Surprisingly, these animals do not exhibit defects in outflow tract development, such as the DORV phenotype reported for the global Myh10 null mouse, and similarly, show a reduced instance of VSDs (Ma et al., 2009). However, these authors report that deletion of Myh10 from cardiomyocytes does result in cardiomyocyte hypertrophy and cytokinesis defects at birth, and adults develop cardiomyopathy at approximately 10 months (Ma et al., 2009). Ma and colleague (2009) speculate that the absence of structural defects associated with defective cardiomyocyte functionality (i.e. DORV), is a direct consequence of the timing of Myh10 deletion (Ma et al., 2009). However, the progressive development of cardiomyopathy in these mutants must be interpreted with caution; recent findings relating to the toxicological impact of continuous Cre recombinase expression in the mouse may offer an alternative explanation as to the development of this phenotype (Schmidt-Supprian and Rajewsky, 2007; Doetschman and Azhar, 2012). Further studies are required before the consequences of cardiomyocyte specific Myh10 ablation are definitively established.

Additionally, ablation of Myh9, the gene encoding NMIIA, causes early embryonic lethality due to defects in visceral endoderm formation, associated with cell adhesion defects (Conti et al., 2004). In contrast, ablation of Myh14, and the subsequent loss of NMIIC, has no aberrant phenotype in the mouse (Ma et al., 2010). Interestingly, the genetic replacement of NMIIB with NMIIA can rescue brain, but not heart, developmental defects, suggesting that NMIIB serves distinct functions during brain and cardiac development (Bao et al., 2007). Furthermore, diversity of phenotypes observed in studies of NMII knock out mice suggests that these proteins are at least in part, functionally distinct.

81 3.1.4 Complementation Testing The EHC mouse was isolated from a mutagenesis screen that employed the chemical mutagen ENU to induce random point mutations throughout the mouse genome (Kile et al., 2003). Previous work in the Hentges laboratory revealed a point mutation in the Myh10 gene in EHC mutants (Ridge et al., Manuscript submitted, PLoS Genetics). However, due to the nature of the mutagenesis screen and the likely introduction of multiple base changes throughout the genome, one is never certain that a point mutation in a given gene is the causative mutation of the mutant phenotype. In light of this, it would be necessary to perform a complementation test to establish whether or not the candidate EHC point mutation in Myh10 was the causative mutation of the EHC phenotype.

In its simplest form, a complementation test determines whether or not two recessive mutations reside in the same gene (Alberts, 2002). Mice heterozygous for the inherited EHC linked region develop normally (Ridge et al., Manuscript submitted, PLoS Genetics). Similarly, previous investigators have shown that the null allele of the EHC candidate gene, Myh10, has a recessive phenotype, such that only homozygous Myh10 null animals display developmental defects (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003). A complementation test can therefore be employed to determine if the causative mutations of both the EHC and Myh10 null phenotype are alleles of the same gene, i.e. Myh10. If the mutations are in the same gene, compound heterozygous animals will display the mutant phenotype, as they will carry no wild type copies of the Myh10 gene. In contrast, if the mutations are in different genes, i.e. the EHC causative mutation is not in Myh10, the compound heterozygous animals will display a normal phenotype, as they obtain one normal copy and one mutant copy of each gene (Alberts, 2002). In the instance of the latter, the mutations are therefore said to have complemented one another and rescued the mutant phenotype.

3.2 Results

3.2.1 Morphology of EHC Embryos Our laboratory has previously found that the EHC mutation causes embryonic lethality, as EHC homozygous mutants (subsequently referred to as EHC) are not present in the litters of intercrossed heterozygous EHC animals (Ridge et al., Manuscript submitted, PLoS Genetics). In light of this, we examined the morphology of EHC mutant embryos during embryonic development. As embryos were clearly alive at E11.5, but rarely viable post- E16.5 (Ridge et al., Manuscript submitted, PLoS Genetics), we investigated mutant embryo morphology from this mid- developmental time point to late gestation. Importantly, this window of embryonic development encompasses key morphological changes in cardiac development. As cardiac abnormalities have previously been described for NMHC IIB ablated animals (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003) it is appropriate to examine the EHC mutant phenotype at these stages.

At E11.5, EHC mutant embryos are approximately comparable in size to littermate controls (EHC/+). However, mutants display an abnormal head morphology, specifically in relation to the structure of the brain ventricles (Fig 3.2, A, B, arrowhead, n>10, 100% penetrance). At E12.5, the dilated brain ventricle phenotype has become more pronounced, and resembles embryonic hydrocephalus (Fig 3.2, C, D, arrowhead, n>10, 100% penetrance). The hydrocephalus phenotype persists through to E14.5, although at this later time point, excess fluid accumulation appears to be restricted to the mesencephalic vesicle (Fig 3.2, E-F, arrowhead, n>10, 100% penetrance). At E14.5, oedema in the cervical spinal cord region becomes apparent (Fig 3.2, E, F, arrow, n>30, 100% penetrance). In addition, EHC mutant embryos also appear to have an elongated body structure (Fig 3.2, E, F). By E16.5, the excessive bulging of the mesencephalic vesicle subsides to give rise to a characteristic ‘domed’ head morphology (Fig 3.2, G, H, arrowhead, n>30, 100% penetrance). This morphological abnormality is consistent with embryonic hydrocephalus (McAllister, 2012). It is interesting to note that we do not detect any other gross morphological abnormalities in the general anatomical structure of the EHC mutant embryo throughout this developmental time frame. In addition, general vascularisation of the extra-cardiac tissue does not appear to be disrupted, as mutant vasculature is comparable to littermate controls. We do observe diffuse areas of vascular haemorrhaging (Fig 3.2, F), however, these abnormalities are less apparent at E16.5.

Figure 3.2: Morphology of the EHC Mutant Embryo (A, C, E, G) Comparison of control (EHC/+) and (B, D, F, H) mutant (EHC/EHC) embryos at different developmental time-points. (B) Mutants display irregular head morphology from E11.5 (arrowhead). (D) By E12.5, the distended head morphology has become more pronounced, and resembles embryonic hydrocephalus (arrowhead). (F) At E14.5, oedema in the spinal cord region is apparent (arrow). The hydrocephalus appears to be restricted to the mesencephalic vesicle (arrowhead). Mutant embryos also appear to display an elongated body structure. (H) The excessive bulging of the mesencephalic vesicle subsides to give a characteristic ‘domed’ head morphology (arrowhead). Scale bars = 1mm (A-F), 2mm (G, H).

3.2.2 Morphology of EHC Embryonic Hearts The embryonic hydrocephalus phenotype in isolation is not thought to be sufficient to cause embryonic lethality. Indeed, many disease states have been recorded that show a manifestation of hydrocephalus at birth in both the mouse and human, clearly showing that hydrocephalus is not systematically incompatible with life (Zhang et al., 2006). Similarly, it is unlikely that the localised regions of vascular haemorrhaging are responsible for the EHC embryonic lethality phenotype, as these abnormalities appear to subside as embryonic development progresses to E16.5. As defects in cardiogenesis have been previously reported for NMHC IIB disrupted mice, we next examined cardiac development in the EHC mutant embryos.

At E12.5, EHC mutant hearts display a largely undisrupted developmental programme when compared to heterozygous littermates (Fig 3.3, A, B). Cardiac looping appears to have occurred correctly, and the initial formation of a four-chambered heart is evident (Fig 3.3, A, B). However, mutant hearts do appear to show unusually distended atria when compared to stage matched controls (Fig 3.3, A, B, n>10, 100% penetrance). In addition, EHC mutants show a slightly flattened ventricular morphology (Fig 3.3, A, B, n>10, 100% penetrance). Histological examination of the ventricular myocardium reveals the presence of many binucleated cardiomyocytes, which are not observed in controls (Fig 3.3, C, D, black arrows).

At E14.5, EHC mutant hearts show more obvious cardiac developmental abnormalities. Firstly, the EHC heart shows a reduction in overall size when compared to control littermates (Fig 3.3, E, F). In addition, mutants show the manifestation of DORV, a condition characterised by the aorta erroneously arising from the right ventricle (Fig 3.3, F, dashed line, n>30, 100% penetrance). Furthermore, the ventricular surface of the mutant heart appears to be decorated by blood-filled nodules (Fig 3.3, F, arrowheads, n>30, 100% penetrance). In contrast, blood-filled coronary vessels become apparent on the ventricular surface of control hearts (Fig 3.3, E, arrow). Moreover, the EHC ventricles have adopted a more pronounced rotund appearance, and lack the formation of the ventricular apex (Fig 3.3, F). Additional EHC cardiac abnormalities are apparent upon histological evaluation of transverse heart sections (Fig 3.3, G, H). Mutant hearts show a reduction in ventricular myocardial trabeculation (Fig 3.3, H, asterisk, n>10, 100% penetrance). In addition, there is evidence to suggest that mutant hearts show defects in the formation of the ventricular septum (Fig 3.3, H, arrow).

Figure 3.3: Morphology of the EHC Mutant Heart

Figure 3.3: Morphology of the EHC Mutant Heart (A) Heterozygous control (EHC/+) and (B) mutant (EHC/EHC) hearts at E12.5. Mutants display a relatively comparable morphology to controls, although distended atria are frequently observed. (C) H and E staining of the ventricular myocardium of control and (D) mutant embryos. Mutants display many binucleated cells (arrows), which are not seen in controls. (E) Control and (F) EHC mutant hearts at E14.5. Nascent blood-filled coronary vessels are observed in control hearts (E, arrow). Mutant hearts are reduced in size when compared to controls, and display DORV (F, dashed line). In addition, the mutant ventricular surface is decorated by blood filled nodules (F, arrowheads). (G) H and E staining of transverse sections of control and (H) mutant hearts at E14.5. Mutants show a reduction in trabeculation (H, asterisk) and a thinned interventricular septum (H, arrow). (I) Control and (J) EHC mutant hearts at E16.5. Blood-filled coronary vessels that traverse the surface of the ventricle are evident in control hearts (I, arrow). In contrast, mutant hearts display the aforementioned ventricular nodules (J, arrowheads). In addition, the DORV abnormality is more prominent at this stage (J, dashed line). (K) Haematoxylin staining of transverse sections of both control and (L) mutant hearts at E16.5. The reduced thickness of the compact myocardium and interventricular septum is increasing apparent at this later developmental stage. (M) Histological evaluation reveals the presence of mature coronary vessels in the myocardial wall of control hearts. (N) These vascular structures are absent from the myocardial wall in the EHC mutants. In addition, the profile view of the ventricular nodules reveals that they are filled with blood (white block arrow), and protrude from the ventricle whilst being covered by the epicardium (arrowhead). (O) Transmission electron microscopy shows that the control epicardium forms a contiguous epithelial layer (arrowhead). In addition, epicardial cells have a flattened appearance. (P) In contrast, EHC epicardial cells have an abnormal rounded morphology, and display discontinuous contacts with adjacent cells (arrow). Image pairs (C and D, E and F) were acquired at the same magnification. Scale bars = 200µm (G, H), 1mm (I, J) and 5µm (O, P). Ao – aorta, DORV – double outlet of the right ventricle, IVS – interventricular septum, LA – left atria, LV – left ventricle, OFT – outflow tract, PT – pulmonary trunk, RA – right atria, RV – right ventricle. Panels C, D, G, H, O and P taken from Ridge et al. (manuscript in preparation). Images in C, D, G, and H courtesy of Karen Mitchell. Transmission electron micrographs (O, P) courtesy of Yinhui Lu and Karl Kadler (University of Manchester).

At comparable tissue depths, control hearts show a complete ventricular septum, which longitudinally dissects the ventricles and connects the apex and base of the heart; separating the lumen of the right ventricle from the lumen of the left ventricle. In EHC hearts, the ventricular septum is thinner in the fibrous region, and appears vulnerable to tearing.

At E16.5, EHC mutant hearts show gross morphological abnormalities. The DORV phenotype persists (Fig 3.3, J, dashed lines, n>50, 100% penetrance), and a number of other structural defects have become apparent (Fig 3.3, I-P). The ventricles have adopted a highly rotund appearance, and mutant hearts do not show the formation of the ventricular apex (Fig 3.3, I, J, n>50, 100% penetrance). In addition, whilst the left and right atria are present, they are positioned aberrantly in relation to the ventricles, and have a distended appearance (Fig 3.3, I, J, n>50, 100% penetrance). Histological evaluations show that the E16.5 EHC ventricular myocardium shows a reduction in the thickness of the myocardial wall and ventricular septum (Fig 3.3, K, L, n>10, 100% penetrance). The myocardium also exhibits decreased trabeculation, and the reduced thickness of the compact myocardium is also evident (Fig 3.3, M, N, n>10, 100% penetrance).

Our most striking observation however, concerns the formation of the coronary vasculature. At this developmental time point, the heart should be infiltrated by a network of coronary vessels, which supply the working myocardium with oxygen and nutrients required to meet its metabolic demands. In control hearts, blood filled coronary vessels that spread across the surface of the ventricles are clearly present (Fig 3.3, I). However, these structures are completely absent from both the surface (Fig 3.3, J, n>50, 100% penetrance), and myocardial wall of EHC hearts (Fig 3.3, N, n>10, 100% penetrance). Instead, the mutant ventricular surface remains decorated by blood filled nodules (Fig3.3, J, N, arrowhead, n>50, 100% penetrance).

The aforementioned defects in coronary vessel formation, accompanied by unusual myocardial development, hint that the EHC cardiac phenotype is rooted in disrupted epicardial cell function. Indeed, transmission electron microscopy reveals that EHC epicardial cells display an irregular lobe-shaped morphology (Fig 3.3, O, P, n>3, 100% penetrance). In control hearts, epicardial cells are flattened, and form a contiguous epithelial layer (Fig 3.3, O, arrowhead). In contrast, mutant epicardial cells display discontinuous connections with adjacent epicardial cells (Fig 3.3, P, arrow, n>3 100% penetrance). In addition, epicardial cells show irregular nuclear architecture (Fig 3.3, P).

Although many of the aspects of the EHC cardiac phenotype closely resemble those presented in previous studies of the NMHC IIB ablated mouse, the abnormalities relating to the coronary vasculature have not yet been reported. The EHC mouse therefore represents a novel model system in which to further examine the function of NMHC IIB in cardiogenesis, providing that the EHC mutation has been correctly mapped to Myh10, and causes a subsequent loss of function to NMHC IIB. To confirm the utility of the EHC mouse for studying Myh10, it is necessary to perform a genetic complementation test with a known Myh10 null allele.

3.2.3 Complementation Assay In order to perform the complementation test, we generated a global Myh10 null mouse line, denoted as Myh10∆. The generation and characterisation of this line is extensively described in Section 3.2.4.1.

3.2.3.1 Genotyping Progeny for the Flox Deletion and EHC Mutation Timed matings were set up between Myh10∆ heterozygous (∆/+) and EHC heterozygous (EHC/+) animals. Pregnancies were allowed to proceed to birth, and the resultant progeny were genotyped for the presence of both the Myh10∆ deletion and EHC mutation (see Section 2.3 and 2.5 respectively). To identify animals with the EHC mutation, Myh10 genomic DNA encompassing exon 18 and intron 18 was amplified by PCR. Amplified DNA was purified and subjected to sequencing PCR. Precipitated sequencing products were sequenced by technical staff at the FMHS sequencing facility (University of Manchester), and chromatographs were analysed using 4Peaks software.

3.2.3.2 EHC/Myh10∆ Animals are Embryonic Lethal From the above cross, numbers of wild type (+/+), heterozygous (EHC/+ or ∆/+) and mutant (EHC/∆) pups were collated from multiple independent litters and the observed frequencies were compared to expected Mendelian ratios (Table 3.1). This analysis revealed that no EHC/∆ pups were present at birth in the litters of EHC/+ and ∆/+ intercrossed animals, and the deviation from expected Mendelian ratios was statistically significant (Chi Squared = 11.091 with 2 degrees of freedom, 2-tailed p=0.0039). From this analysis, we conclude that animals with the EHC/∆ genotype are embryonic lethal, and that the Myh10∆ allele fails to rescue the embryonic lethality phenotype of the EHC mutants.

Genotype +/+ EHC/+ or ∆/+ EHC/∆

Expected 5 11 5 Frequency

Observed 8 13 0 Frequency

Table 3.1: Compound Heterozygous EHC/Myh10∆ Animals Display Embryonic Lethality Table comparing observed frequencies of wild type (+/+), heterozygous (∆/+ or EHC/+) and compound heterozygous (EHC/∆) animals in neonatal litters to expected Mendelian frequencies. Chi Squared = 11.091 with 2 degrees of freedom, 2-tailed p=0.0039.

3.2.3.3 Morphology of EHC/Myh10∆ Mutant Embryos In light of the complementation test findings, we examined the morphology of EHC/∆ embryos during development. As expected, compound heterozygous embryos (EHC/∆) were heterozygous for the Myh10∆ deletion (Fig 3.4, A) (explained in detail in Section 3.2.4.1) and the EHC point mutation (Fig 3.4, B, arrow, G>T at splice donor site of intron 18). The morphology of mutant embryos (EHC/∆) was compared to that of stage matched heterozygous controls (either EHC/+ or ∆/+) (Fig 3.4, C-F). Interestingly, we found that mutants displayed brain developmental defects, as observed in the EHC homozygous embryos. However, EHC/∆ embryos displayed a more severe brain malformation, evident from E11.5 (Fig3.4, C, D, arrowhead, n>5, 100% penetrance). The brain ventricles were clearly distended compared to controls and more pronounced than previously observed in EHC homozygous mutant embryos (Fig 3.4, C, D, arrowhead). At E16.5, EHC/∆ mutants displayed a pronounced mushroom-shaped protrusion of brain tissue from the cranium, resulting in the exposure of brain tissue to amniotic fluid in the yolk sac (Fig 3.4, E, F, arrowhead, n>10, 100% penetrance). This brain malformation is consistent with the phenotype of embryonic exencephaly (Copp et al., 2003). In addition, EHC/∆ embryos displayed an unusual body arrangement, with an apparent widening of the relative position of the fore and hind limbs (Fig 3.4, E, F, n>10, 100% penetrance). Although EHC/∆ embryos display exencephaly, rather than the hydrocephalus phenotype present in EHC homozygous mutants, the manifestation of brain abnormalities is consistent with previous reports of Myh10 ablated mice (Tullio et al., 1997, Uren et al., 2000). We suspect that the development of exencephaly results from the genetic background of the Myh10∆ line, which differs from that of the EHC strain, and is discussed in greater detail in Section 3.3.

Figure 3.4: Genotyping and Morphology of EHC/Myh10∆ Mutants (A) Amplification of the Myh10 exon 2 sequence in mutant animals (EHC/∆) produces both the wild type (~1.1Kb) and deleted (~600bp) band. (B) Sequencing of EHC/∆ Myh10 reveals a point mutation in the splice donor site immediately following Myh10 exon 18. Note, as these animals are heterozygous for the EHC point mutation, two peaks are detected in the sequencing data (arrow). These peaks correspond to the presence of both G and T bases. (C) Dissection images of heterozygous Myh10∆ control (∆/+) and (D) mutant (EHC/∆) embryos at E11.5. Mutant embryos display an irregular head morphology that resembles exencephaly (arrowhead). (E) Control and (F) mutant embryos at E16.5. Brain tissue is observed protruding from the cranium in mutant embryos (arrowhead). (G) Images at dissection of control and (H) mutant hearts at E16.5. Coronary vessels are clearly detected on the surface of the control heart (G, arrow). In addition, controls display the correct positioning of the great arteries (G, dashed line). In contrast, the mutant heart displays DORV (H, dashed line), and blood-filled ventricular surface nodules (H, arrowheads). Scale bars = 1mm (C, D, G, H), 2mm (E, F). Ao – aorta, LA – left atria, PT – pulmonary trunk, RA – right atria.

3.2.3.4 Gross Morphology of EHC/Myh10∆ Mutant Hearts We next examined the morphology of EHC/∆ mutant hearts at E16.5. Mutants displayed striking phenotypic similarity to EHC homozygous mutants (Fig 3.4, G, H, n>10, 100% penetrance). The mutant hearts display a rounded ventricular morphology, in addition to distended and irregularly positioned atria (Fig 3.4, H). Furthermore, mutants show abnormal positioning of the great arteries, including DORV. (Fig 3.4, H, dashed lines). Finally, whilst blood filled coronary vessels are observed on heterozygous control littermates (Fig 3.4, G, arrow), we fail to observe these in mutant hearts, which display ventricular nodules (Fig 3.4, H, arrowheads, n>10, 100% penetrance). These cardiac defects phenocopy those displayed by EHC mutants, and as previously mentioned, the absence of coronaries has not previously been reported in studies of Myh10 ablated hearts.

3.2.4 The Myh10∆ Allele is an Myh10 Null

3.2.4.1 Generation of the Myh10∆ Mouse Line The global Myh10 null mouse has been previously investigated (please refer to Table 3.3 in Section 3.3 for a summary of the findings from these reports). However, a comprehensive examination of the cardiac phenotype in these animals, specifically in relation to the coronary vasculature, is lacking. In order to generate animals harboring a Myh10 null allele in our laboratory, we utilised Cre recombinase-loxP site-specific recombination technology (discussed in detail in Section 5.1.1). The Myh10tm7Rsad mouse line (subsequently referred to as Myh10flox/flox) contains loxP sites that flank exon 2 of the Myh10 gene on both chromosomes. These animals were crossed to the Tg(Nes-cre)Wme mouse, which expresses Cre recombinase in the oocyte. To confirm the deletion of the Myh10 exon 2 genomic DNA sequence, genotyping was performed with primers flanking Myh10 exon 2. We have previously described the sequencing of PCR amplification products to confirm specificity of this primer pair to Myh10 (as described in (Ridge et al., Manuscript submitted, PLoS Genetics). Briefly, performing PCR with primers that flank Myh10 exon 2 generated two distinct PCR products. The wild type (WT) Myh10 allele generated an amplicon of approximately 1.1-Kb (Appendix 12, +/+ lane). The exon 2 deleted (Myh10∆) allele generated an amplicon of approximately 600-bp (Appendix 12, ∆/∆ lane). This reduction in amplicon size is due to the deletion of the Myh10 exon 2 DNA sequence. Note that for Myh10∆ heterozygotes, we observed an additional band at approximately 1.3-Kb (Appendix 12, ∆/+ lane, arrow). This represents the non-deleted floxed Myh10 allele. The

size difference between the WT and non-deleted floxed alleles is attributed to the integration of both the loxP sequences and partial viral plasmid sequence into the Myh10 locus. The subsequent Myh10 null allele is denoted as Myh10∆, or ∆ in all following genotype annotations. A schematic representation of the Myh10 wild type, floxed non- deleted, and exon 2 deleted loci and the relative primer binding sites is provided in Appendix 12.

3.2.4.2 Myh10∆ Homozygous Mutants Are Embryonic Lethal Heterozygous Myh10∆ animals were intercrossed and resulting pups were genotyped for the inheritance of the Myh10∆ allele. The numbers of wild type (+/+), heterozygous (∆/+) and mutant (∆/∆) pups were collated from several independent litters, and the observed frequency of each genotype was compared to expected Mendelian ratios (Table 3.2). This analysis revealed that no Myh10∆ mutants were recovered at birth, and that this deviation from the expected Mendelian frequency was statistically significant (Chi Squared = 14.029 with 2 degrees of freedom, 2-tailed p=0.0009). We can therefore conclude that the Myh10∆ homozygous mutants are embryonic lethal. This finding is consistent with previous reports in which targeted deletion of Myh10 exon 2 results in embryonic lethality (Tullio et al., 1997; Takeda et al., 2003).

Genotype +/+ ∆/+ ∆/∆

Expected 9 17 8 Frequency

Observed 14 20 0 Frequency

Table 3.2: Homozygous Myh10∆ Mutant Animals Display Embryonic Lethality Table comparing observed frequencies of wild type (+/+), heterozygous (∆/+) and homozygous (∆/∆) Myh10∆ animals in neonatal litters to expected Mendelian frequencies. Chi Squared = 14.029 with 2 degrees of freedom, 2-tailed p=0.0009.

3.2.4.3 Myh10∆ Homozygous Mutants Do Not Express Full Length NMHC IIB To examine whether the Myh10∆ deletion affects expression of NMHC IIB, we performed western blotting analysis of embryonic protein extracts. As it has been previously reported that NMHC IIB is enriched in the embryonic heart compared to other tissues (Ma et al.,

93 2010), we probed protein extract from E14.5 embryonic hearts using a C-terminal NMHC IIB antibody (Fig 3.5). Samples were also probed with an antibody against the house keeping gene β-actin, as a loading control and proxy measurement of total protein loaded onto the gel. All samples show a detectable band at ~42kDa, which corresponds to β-actin (Fig 3.5, lower band). This indicates that protein is present in each sample. A clear, distinct band is detectable at approximately 250kDa in both wild type (Fig 3.5, lanes 2 and 4) and ∆/+ (Fig 3.5, lanes 3 and 5) samples. This band corresponds to the reported molecular mass of NMIIB (Conti and Adelstein, 2008; Vicente-Manzanares et al., 2009). However, no signal was detected for NMIIB in ∆/∆ protein extracts (Fig 3.5, lanes 7-9, n=3). This result indicates that full length NMHC IIB is not synthesized in the hearts of Myh10∆ mutant embryos. As the Myh10∆ deletion is ubiquitous throughout the entire embryo, we extrapolate this result to conclude that full length NMHC IIB is not synthesized in the ∆/∆ mouse.

Figure 3.5: Western Blot Analysis of NMIIB Expression in Myh10∆ Animals Western blotting on E14.5 embryonic heart protein extracts using a C-terminal anti-NMIIB antibody. NMIIB was detected in wild type and heterozygous Myh10∆ animals, but not in homozygous Myh10∆ animals. The lower band at approximately 42kDa corresponds to β- actin, which is used as a loading control. Representative blot image (total of six mutant samples from two independent experiments).

3.2.4.4 Morphology of Myh10∆ Homozygous Mutant Embryos As Myh10∆ homozygous mutants were not present at birth, we analysed the morphology of mutant embryos during development. The morphology of ∆/∆ mutant embryos was compared to control (+/+ or ∆/+) littermates at the same developmental stages as described previously (see Section 3.2.1). As early as E9.5, mutant embryos display morphological abnormalities that are not present in heterozygous controls (Fig 3.6, A, B). Although mutants appear of comparable size to controls, they display an irregular head morphology (Fig 3.6, B, arrowhead, n>10, 100% penetrance). At E10.5, this irregular head morphology manifests as a severe dilation of the brain ventricles (Fig 3.6, C, D, arrowhead, n>10, 100% penetrance). Similarly, at E11.5 the brain ventricles are evidently distended and apparently filled with fluid, and are strikingly similar to the EHC/∆ brain phenotype (Fig 3.6, E, F, arrowhead, n>50, 100% penetrance). At E12.5, the severity of the head defects increases; the fluid filled membrane covering the embryo head is no longer apparent, and brain tissue is exposed through the cranium (Fig 3.6, G, H, arrowhead, n>10, 100% penetrance). By E14.5, the brain tissue protrudes from the cranium in a ‘mushroom’ structure (Fig 3.6, I, J, arrowhead, n>50, 100% penetrance). This results in the brain tissue being exposed to the amniotic fluid inside the yolk sac. This phenotype fulfills the characteristics of exencephaly, and phenocopies that of the EHC/∆ mutant. Similarly, ∆/∆ embryos display an unusual body arrangement in relation to the positioning of the limbs (Fig 3.6, J, n>50, 100% penetrance). These abnormalities and exencephaly remain at E16.5 (Fig 3.6, K, L, arrowhead) and although embryos are frequently alive at this stage at the point of dissection (as determined by the observation of beating hearts), it is very rare to recover mutant embryos post-E16.5.

Figure 3.6: Morphology of Myh10∆ Homozygous Mutant Embryos (A-L) Comparison of control (∆/+) and mutant (∆/∆) embryos at different developmental time points. Please see figure labels for genotype and stage details. Arrowheads indicate irregular head morphology in mutant embryos. Scale bars = 1mm (A-J), 2mm (K-N). Representative images from multiple independent litters.

3.2.4.5 Morphology of Myh10∆ Homozygous Mutant Hearts As expected, Myh10∆ mutants generated in our laboratory showed similar cardiac abnormalities to those previously reported for the Myh10 ablated mouse (Tullio et al., 1997). At E11.5, homozygous Myh10∆ mutant hearts display a very similar morphology to heterozygous controls (Fig 3.7, A, B, n>50, 100% penetrance). The process of looping appears to have occurred correctly, and the four cardiac chambers are evident (Fig 3.7, B). However, at E12.5, the morphology of the mutant heart begins to deviate from controls (Fig 3.7, C, D). The mutant ventricles have adopted a flattened appearance, and there is evidence to suggest that the outflow tract is misplaced (Fig 3.7, D, n>10, 100% penetrance). In addition, whilst the formation of primitive vessels can be observed on the ventricular surface of heterozygous hearts (Fig 3.7, C, arrow), these structures are not present in mutants, which display blood-filled nodules (Fig 3.7, D, arrowhead, n>10, 100% penetrance). By E14.5, the mutant heart displays gross morphological abnormalities. The atria appear distended and misplaced compared to controls (Fig 3.7, E-F). In addition, whilst the control heart shows the correct development of the ventricular apex, the mutant ventricular display an abnormal rounded morphology (Fig 3.7, E-F). Furthermore, blood from the right ventricle can be seen to enter both the pulmonary vessel and the aorta, indicating the presence of DORV and/or a ventricular septal defect (Fig 3.7, F, dashed line). In addition, the ventricular surface of the mutant heart is decorated by blood-filled nodules (Fig 3.7, F, arrowheads). All of the aforementioned defects were present at 100% penetrant (n>50). Similarly, at E16.5, the rounded ventricular morphology and presence of DORV persist (Fig 3.7, G-H, dashed lines). Moreover, mutants do not display evidence of blood-filled coronary vessels on the surface of the ventricles, in contrast to control hearts (Fig 3.7, G, arrow). Once again, the surface of the mutant heart is decorated by blood-filled nodules (Fig 3.7, H, arrowheads). Similarly, these defects were present at 100% penetrance (n>50). It is clear that the cardiac abnormalities presented in ∆/∆ embryos phenocopy both those observed in the homozygous EHC and EHC/∆ mutant hearts. Importantly, the phenotypic similarities of the ∆/∆ mutant hearts to those of the published Myh10 knock out signify that the Myh10∆ allele generated by our lab represents a true Myh10 null allele.

Figure 3.7: Morphology of Myh10∆ Homozygous Mutant Hearts (A) Dissection images of control (∆/+) and (B) mutant (∆/∆) hearts at E11.5. (C) Control hearts begin to show evidence of coronary vessel formation at E12.5 (arrow). (D) In contrast, the mutant ventricular surface is covered in blood-filled nodules (arrowhead). (E) The control heart shows correct alignment of the aorta and pulmonary trunk in relation to the left and right ventricle at E14.5 (dashed line). (F) At E14.5, mutant hearts show misalignment of the great vessels, which resembles DORV (dashed line). Blood can be seen entering both the aorta and pulmonary trunk from the right ventricle. The presence of ventricular nodules is apparent (arrowheads). (G) By E16.5, coronary vessels that transverse the length of the ventricular surface are evident in controls (arrows). (H) These vessel structures are absent from the surface of the mutant heart, which is covered in blood-filled surface nodules (arrowheads). In addition, the DORV phenotype persists (dashed lines). Scale bars = 0.5mm. Ao – aorta, LA – left atria, OFT – outflow tract, PT – pulmonary trunk, RA – right atria.

3.2.5 Characterisation of the Extent of Vascular Defects in the EHC Mouse Our observations at the point of dissection revealed that EHC mutant hearts showed an absence of blood filled coronary vessels on their ventricular surface (Fig 3.3, J) and within the myocardial wall (Fig 3.3, N). In order to determine the extent of these coronary abnormalities, we employed an immunohistochemical approach to characterise the localisation of key cellular components of the coronary vessel architecture.

3.2.5.1 Vascular Endothelial Cell Localisation Vascular endothelial cells line the lumen of the coronary vasculature. These endothelial cells are predominantly thought to arise from the sinus venosus, and begin to colonise the sub-epicardial myocardium from approximately E10.5 (Red-Horse et al., 2010; Chen et al., 2014; Tian et al., 2015). The endothelial cells are organised into a primitive capillary network, or plexus, by angiogenesis during a distinct window of embryonic development between E10.5 – E13.5. As the vascular endothelial cell population precedes the localisation of other vascular cell components to the ventricular surface, they are an ideal marker to investigate fundamental defects in vessel formation. We performed whole mount immunohistochemistry on E16.5 hearts for the pan-endothelial cell marker, PECAM-1 (platelet endothelial cell adhesion molecule 1, also known as CD31). Although it is known to be expressed by all types of endothelial cells, including non-vascular residing endothelial cells, PECAM-1 has been previously used to highlight endothelial components of the coronary vasculature in a number of seminal studies, and is considered the classical endothelial cell marker (Liu and Shi, 2012).

Immunohistochemical staining for PECAM-1 shows that the vascular endothelial components in heterozygous control hearts are organised into large coronary network that traverses the entire ventricular surface (Fig 3.8, A, arrows). The major large diameter vessels are clearly identifiable. These vessels branch at many points into the coronary vascular tree (Fig 3.8, C). In contrast, the vascular endothelial cells in EHC mutant hearts appear to be restricted to a primitive and rudimentary capillary network. The major, large diameter vessels are not present, and the capillaries have vastly limited branching points when compared to controls (Fig 3.8, B, arrows). In addition, we observe PECAM-1 immunoreactivity in the aforementioned ventricular nodules, which decorate the ventricular surface (Fig 3.8, D). These defects were present at 100% penetrance (n=5).

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Figure 3.8: Molecular Characterisation of the Coronary Vasculature in EHC Hearts (A) Whole mount PECAM-1 staining of control (EHC/+) and (B) mutant (EHC/EHC) E16.5 hearts. Vascular endothelial cells are organised into large coronary vessels in control hearts (A, arrows). In contrast, PECAM-1 immunoreactivity is restricted to capillary vessels and ventricular surface nodules in mutant hearts (B, block arrows). (C) Enlarged image of PECAM-1 staining from panel A. (D) Enlarged image of PECAM-1 staining from panel B. (E) SMα A staining of transverse sections of control and (F) mutant E16.5 hearts. Positive SMα A staining indicates vascular smooth muscle cell coalescence around vessel structures in the myocardial wall of control hearts (E, arrow). These vessel structures are absent in the mutant heart. Images in A-B, and C-D, acquired at the same magnification. Scale bars = 50µm (E, F). Representative images (n=5).

The presence of vascular endothelial cells within these structures suggests that the endothelial network is incorrectly patterned, and that vascular endothelial components are not appropriately remodeled into the vascular tree. The limited expanse of the coronary endothelial network in E16.5 EHC mutant hearts resembles the rudimentary capillary plexus present in wild type embryos earlier in cardiogenesis (personnel communication, Dr. Kathryn Hentges). Unfortunately, as it is extremely rare to recover EHC mutant embryos post E16.5, it has not been possible to assess whether the endothelial phenotype displayed in the mutants is a developmental delay in the coronary vessel formation pathway, or whether this plexus is expanded and rectified later in development. However, considering that embryonic lethality occurs at a point very soon after E16.5, it is highly unlikely that the endothelial network we observe is significantly altered during this time.

3.2.5.2 Vascular Smooth Muscle Cell Localisation We next analysed the localisation of vSMCs within the coronary network of mutant hearts. The vSMC components of the coronary vasculature are thought to derive exclusively from the epicardium during development (Riley and Smart, 2011). Epicardial-derived smooth muscle begins to coalesce around nascent coronary capillaries from approximately E14.5, providing mechanical support to the vessels as they begin to bear a blood flow load. It is thought that the initiation of blood flow induces the expression of signalling molecules by the vascular endothelium. These signals act in a non-cell autonomous fashion to stimulate vSMC migration and recruitment. It has previously been shown that smooth muscle α- actin (SMαA) is a marker of vascular smooth muscle cells (Skalli et al., 1989). We employed immunohistochemistry to stain sections of E16.5 hearts to profile the localisation of vSMCs within the myocardial wall. This developmental time point was selected for analysis as at this point, the coronary vasculature is undergoing maturation, and vSMC localisation to the vessels should be observable.

Our analysis revealed that smooth muscle cells are clearly localised around the periphery of vessel structures in the ventricular myocardial wall of heterozygous control hearts (Fig 3.8, E, arrow). We observe a low level of diffuse SMαA immunoreactivity throughout the myocardial tissue section, but this staining is enriched around vessels. As SMαA, in addition to other vSMC markers, such as SM22 (smooth muscle protein ), is known to be expressed in cardiomyocytes between approximately E12.5-E15.5 (personal communication, Dr. Nicola Smart), we propose that this level of background staining is a

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feature of non-vSMC specific SMαA expression, and not due to the ubiquitous presence of vSMC throughout the tissue.

In the EHC mutant heart, we do not observe this SMαA staining profile. Vascular smooth muscle cells are detectable, as determined by positive SMαA immunoreactivity in the myocardium (Fig 3.8, F). However, in contrast to control hearts, we do not observe localisation of vSMCs to vessel structures. Indeed, these structures are entirely absent from the mutant myocardium, and the resulting smooth muscle localisation appears to be more randomly dispersed throughout the ventricular wall (Fig 3.8, F). This disrupted vSMC localisation profile was present at 100% penetrance (n=5). From this evidence, we conclude that vascular smooth muscle cells are not appropriately recruited to the coronary capillaries. However, we are not able to categorically determine whether the vessel structures are absent from the EHC myocardium due to a failure to recruit smooth muscle cells, or whether smooth muscle cell localisation is disrupted due to a lack of vessel structures.

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3.3 Discussion To briefly summarise our findings of the EHC phenotype, we have shown that the EHC mouse harbors a recessive, embryonic lethal mutation in Myh10. Homozygous EHC embryos exhibit embryonic hydrocephalus, accompanied by extensive defects in the formation of the heart. EHC mutant hearts display gross anatomical abnormalities including DORV, thinning of the ventricular myocardium and ventricular septum, consistent with previous reports of Myh10 ablated mice (summarised in Table 3.3). Surprisingly, EHC mutants also show aberrant non-epithelial epicardial cells with discontinuous cell connections. Moreover, the EHC heart shows defects in the formation of the coronary vasculature, compounded by a failure to recruit vascular smooth muscle cells to the developing endothelial capillaries. These profound defects in the formation of the coronary vasculature have not previously been reported in the Myh10 knock out mouse (Table 3.3).

To address the first aim of the project and validate that the EHC point mutation disrupts Myh10 and causes complete loss of functional NMHC IIB, we performed a complementation assay with the Myh10∆ mouse. It has been previously shown that targeted deletion of the Myh10 exon 2 sequence ablates NMHC IIB protein production; animals therefore display a Myh10 null phenotype (Tullio et al., 1997; Uren et al., 2000; Ma et al., 2009). In agreement with this, we do not detect full length NMIIB in homozygous Myh10∆ mutants, as determined by western blotting with a C-terminal anti-NMIIB antibody. As expected, homozygous Myh10∆ mutants display embryonic lethality and cardiac defects, consistent with previous reports of the Myh10 null phenotype (Tullio et al., 1997; Uren et al., 2000).

When the EHC mutation and Myh10∆ deletion are combined in trans, EHC/∆ embryos recapitulate major aspects of both the EHC and Myh10∆ homozygous mutant phenotypes, presenting late gestation embryonic lethality and cardiac defects. The finding that the Myh10∆ allele does not rescue either the embryonic lethality or morphological defects presented in the EHC phenotype, indicates that the Myh10 null allele fails to genetically complement the EHC allele. The nature of the complementation test thereby confirms that the EHC mutation has been correctly mapped to the Myh10 locus. Furthermore, as the Myh10∆ allele is a known Myh10 null allele, and does not synthesise functional NMHC IIB protein, we can therefore conclude that the EHC point mutation causes the same phenotype as the Myh10 null allele. Further experimentation with an antibody against the

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Phenotypic Description Details of Myh10 Genetic Myh10 KO Study Coronary Vessel Ventricular Septal Outflow Tract Other Cardiac Mutation Background Onset of Lethality Brain Abnormalities Defects Defects Defects Abnormalities CM hypertrophy evident Global disruption Overriding Aorta or from E12.5, increased CM of Myh10 exon 2 DORV accompanied Tullio et al., 1997 129/Sv 65% embryonic Membranous VSD in binucleation, 70% reduction Hydrocephalus with insertion of Not described by pulmonary Takeda et al., 2003 C57BL/6J lethal, 35% P0 6/7 (86%) newborns in CM population at E14.5, (100%) Neomyocin stenosis in 6/7 associated with CM cassette (86%) of newborns cytokinesis defects Increased CM binucleation, Global 92% of F2 progeny 70% reduction in CM No evidence of replacement of embryonic lethal, Membranous VSD in 129/Sv DORV in 5/7 (71%) population, dilated hydrocephalus, but migration Bao et al., 2007 Myh10 exon 2 with 100% of F3 Not described 7/7 (100%) embryos C57BL/6 embryos analysed cardiomyopathy in surviving defects in facial and pontine NMIIA-GFP progeny embryonic analysed adults associated with CM neurons cDNA lethal hypertrophy CM hypertrophy and CM-specific cytokinesis defects (P0), 129S6/SvEvTac Membranous VSD in Ma et al., 2009 deletion of Myh10 Animals are viable Not described Not present defects in structural integrity Not present B6.FVB 2/9 (22%) animals exon 2 of intercalated discs and cardiomyopathy (10 months) Global G>T point Multinucleated Absence of mature Thinning of

105 mutation in splice cardiomyocytes from E12.5, 100% lethality vessels associated membranous donor site Overriding Aorta or reduced trabeculation and Present Study C57BL/6J between E14.5- with vSMCs, ventricular septum, Hydrocephalus (100%) following Myh10 DORV (100%) reduced ventricular E17.5 abnormal epicardial clear breaks occasionally exon 18 (EHC myocardial wall thickness morphology (100%) observed line) from E14.5 Multinucleated Absence of mature Thinning of cardiomyocytes from E12.5, Global deletion of 100% lethality vessels associated membranous 129S6/SvEvTac Overriding Aorta or reduced trabeculation and Exencephaly Present Study Myh10 exon 2 between E14.5- with vSMCs, ventricular septum, C57BL/6 DORV (100%) reduced ventricular (100%) (Myh10∆ line) E17.5 abnormal epicardial clear breaks occasionally myocardial wall thickness morphology (100%) observed from E14.5 Table 3.3: Phenotypic Summary of the Myh10 Ablated Mouse Table summarising the phenotypic observations made from studies of global and tissue-specific Myh10 null mouse models. Coronary defects described in the present study have not previously been reported in the literature. CM – cardiomyocyte, vSMCs – vascular smooth muscle cells.

N-terminal region of NMHC IIB, would help to establish whether or not an aberrant, C- terminally truncated protein is synthesised in the EHC mutants, or if the erroneous Myh10 transcript is ubiquitously degraded before translation. However, the phenotypic similarity between the EHC/∆ embryos, and both the homozygous EHC and homozygous Myh10∆ mutants augments our hypothesis that the EHC allele represents a Myh10 null allele. We have therefore established that the point mutation in Myh10 is the causative mutation of the EHC phenotype.

The EHC mutants develop an embryonic hydrocephalus phenotype, which is strikingly similar to that previously reported in Myh10 deficient embryos (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003; Ma et al., 2004). The development of embryonic hydrocephalus in the Myh10 ablated mouse has been attributed to the erroneous obstruction of the spinal canal by neuroepithelial cells, leading to the aberrant dilation of the brain ventricles and aqueduct of Sylvius from E11.5 (Ma et al., 2004; Ma et al., 2007). We similarly observe dilation of the brain ventricles from approximately E11.5 in the EHC mutant embryos. Ma and colleagues (2007) attribute abnormal invasion of the spinal canal by neuroepithelial cells to the destruction of a NMIIB mediated mesh-like scaffold (Ma et al., 2007). This mesh structure is reinstated and hydrocephalus relieved following re- expression of NMIIB to wild type levels (Ma et al., 2004). This suggests that NMIIB plays a crucial role in maintaining the integrity of cell adhesions within the neuroepithelium. Whilst we have not investigated the hydrocephalus aspect of the EHC phenotype further, we have confirmed that NMIIB is the only detectable NMII isoform in the neuroepithelium of the spinal canal in Myh10 ablated mice (Appendix 13), in agreement with the findings of others (Ma et al., 2007). This finding supports the conclusion of others that NMIIB plays an important role in the maintenance of correct neuroepithelial function. However, it is interesting to note that replacement of NMIIB with NMIIA can rescue the hydrocephalus phenotype (Bao et al., 2007), suggesting that NMIIA can compensate for NMIIB function in the neuroepithelium, and that NMII isoforms show a level of functional redundancy with regard to adhesion maintenance in the developing brain.

Intriguingly, whilst both EHC/∆ and ∆/∆ embryos phenocopy the EHC cardiac abnormalities, they display different brain defects. Both EHC/∆ and ∆/∆ embryos develop exencephaly at a similar developmental stage to the first observation of hydrocephalus in the EHC embryo. The finding that brain but not cardiac defects displayed by the Myh10 null can be rescued by replacement of NMIIB with either a motor-impaired NMIIB isoform (Ma et al., 2007) or by NMIIA (Bao et al., 2007) provides evidence that NMIIB is

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a pleiotropic gene with a multitude of functions in different developmental processes. In light of this, it should not be surprising that differences in brain defects arise in Myh10 ablated mice from different genetic backgrounds (please refer to Table 3.3). Indeed, the influence of genetic background has been speculated as a cause of the variation in severity of neural tube defects in Apob modified mice, which manifest as either hydrocephalus or exencephaly (Homanics et al., 1995). The development of exencephaly in Myh10 null animals derived from the Myh10∆ line could be further explored through analysis of neuronal specificity markers in the developing brain using immunohistochemistry and in situ hybridization approaches. Previous work in the Hentges laboratory has demonstrated that the EHC mutant embryo does not show significant differences in the localisation of these markers compared to controls (Mitchell, K., PhD Thesis, University of Manchester). A similar experimental approach in Myh10∆ homozygous mutants would help to ascertain whether cells in the developing brain are specified in a correct spatial and temporal manner, and provide a potential explanation as to why EHC and Myh10∆ homozygous mutants develop differing brain defects.

In agreement with previous studies of the Myh10 ablated mouse, the EHC mutant heart displays myocardial developmental defects, including the presence of DORV, VSDs, thinning of the myocardial wall and cardiomyocyte hypertrophy (Tullio et al., 1997; Takeda et al., 2003; summarised in Table 3.3). Previous studies have associated the development of these cardiac defects with a failure to correctly establish planar cell polarity in the myocardium (Phillips et al., 2005; Phillips et al., 2008). Indeed, these authors speculate that these defects are underpinned by loss of NMIIB activation, indicating that the cardiac abnormalities observed in the EHC mouse may be caused by a polarity defect. This supports previous work that illustrates a prominent role for NMIIB in the correct establishment of cell polarity (reviewed in Vincente-Manzanares et al., 2009; Conti and Adelstein, 2008).

As previously discussed, myocardial proliferation is essential for the correct development of the ventricular myocardial wall, including the formation of the trabeculae and compact myocardial zone (Brade et al., 2013). NMIIB is known to play a key role in the regulation of cell proliferation (Gutzman et al., 2015). Indeed, it has been previously shown that Myh10 null hearts show an approximate 70% reduction in the number of cardiomyocytes at E14.5, attributed to defects in NMIIB-mediated cytokinesis that generate multinucleated cardiomyocytes (Takeda et al., 2003). Consistent with these findings, the EHC myocardium shows an increased prevalence of multinucleated cardiomyocytes, suggesting that NMIIB

107 functions during cytokinesis in embryonic cardiomyocytes. It is possible that the defects we observe in the EHC myocardium are at least in part underpinned by a reduction of cardiomyocyte cytokinesis, leading to a reduction in the number of cardiomyocytes and the overall reduced size of the EHC heart.

In addition, reduced cardiomyocyte proliferation, which results in defects in the thickness and trabeculation of the myocardial wall, has been observed following the disruption of Mycn, Bmp10 and Nrg1 (Harmelink et al., 2013). As a number of molecular signalling pathways are known to converge on NMIIB (Vincente-Manzanares et al., 2009), it is possible that disrupted Mycn-Bmp10-Nrg1 signalling underpins the myocardial proliferation defects observed in Myh10 mutants.

Similarly, cardiomyocyte proliferation is also thought to play a key role in the development of the outflow tract (Brade et al., 2013), and proliferation defects may underpin abnormalities surrounding the abnormal placement of the great arteries observed in the EHC heart. The development of the outflow tract depends upon the integration of cardiomyocytes into this structure (myocardialisation), which permits the muscularisation of the outlet septum and simultaneous rotation of the outflow tract wall to properly align the great vessels with the left and right ventricles (Rana et al., 2007; Bajolle et al., 2006). The cardiomyocyte cytokinesis defects observed in the EHC heart may therefore reduce the number of myocytes available for this myocardialisation event, leading to the development of DORV in the EHC phenotype due to reduced outflow tract rotation (Neeb et al., 2013). Indeed, myocardial ablation of Gata4 and Tbx5 has been shown to reduce cardiomyocyte proliferation, resulting in outflow tract abnormalities (Misra et al., 2014).

Interestingly, replacement of NMIIB with NMIIA cannot rescue either the DORV and VSD abnormalities, or the presence of multinucleated cardiomyocytes (Bao et al., 2007). It has been shown previously that NMIIA is not expressed in cardiomyocytes (Takeda et al., 2003), suggesting that NMIIB plays a novel function in cardiomyocytes that cannot be compensated by NMIIA. However, whilst deletion of Myh10 specifically from cardiomyocytes results in ventricular septum thinning and cardiomyocyte hypertrophy, these animals show a reduced incidence of VSD, and moreover, do not display DORV (Ma et al., 2009). This result suggests that loss of Myh10 from another cell population primarily contributes to the formation of DORV in Myh10 ablated animals.

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To this end, it is important to consider the important contribution of CNCC to the developing outflow tract. These cells are known to contribute to outflow tract elongation, and as previously mentioned, outflow tract malformations are frequently associated with shortening of the outflow tract (Brade et al., 2013; Neeb et al., 2013). A recent study has shown that Ets-1 deficient mice develop outflow abnormalities due to impaired CNCC migration (Gao et al., 2010). It is well documented that NMIIB plays a crucial role in cell migration (Vincente-Manzanares et al., 2009; Conti and Adelstein, 2008). It is therefore possible that NMIIB functions to facilitate the migration of CNCCs to the developing outflow tract, and loss of NMIIB leads to insufficient CNCC migration and integration into the developing outflow tract. This hypothesis would explain the disparity between the development of DORV in the global Myh10 knock out, but not in cardiomyocyte-specific Myh10 mutants. The recruitment of CNCCs is also important for outflow tract septation (Brade et al., 2013). Whilst we have not assessed outflow tract length or CNCC migration directly, septation of the outflow tract does appear to occur in the Myh10 ablated mouse. Further studies are required to examine whether CNCC migration is perturbed in order to fully determine the pathogenic processes underpinning DORV manifestation in the EHC and Myh10∆ mutant hearts.

Intriguingly, we have found that NMIIB is the predominant NMII isoform in the wall of the descending aorta (see Appendix 13). CNCCs are known to contribute to the smooth muscle coating that ensheathes the outflow tract (Li et al., 2005), and NMIIB has previously been used as a marker of embryonic smooth muscle cells (El-Mounayri et al., 2013). This suggests that NMIIB plays a crucial role in maintaining the structural integrity of the arterial wall, possibly in relation to the maintenance of force and vascular tone (Brozovich et al., 2016). It would be extremely interesting to investigate whether these NMIIB containing smooth muscle cells derive from CNCCs, and establish whether or not loss of CNCC derived smooth muscle in the outflow tract impedes normal outflow tract development.

Surprisingly, we observe abnormalities in the formation of the epicardium in the EHC heart; mutant epicardial cells display an abnormal rounded morphology and disrupted contacts with adjacent cells. Epicardial defects have not previously been reported for Myh10 ablated mice (refer to Table 3.3). It has previously been shown that NMIIB plays a key role in cell adhesion, and ultimately the regulation of cell shape (Conti and Adelstein, 2008; Vicente-Manzanares et al., 2009; Gutzman et al., 2015). Ablation of the adhesion proteins β-catenin and connexin-43 has previously been shown to disrupt epicardial

109 formation (Zamora et al., 2007; Rhee et al., 2009; Li et al., 2002), indicating that NMIIB may function in the maintenance of epicardial adhesions. Expanding upon this, ablation of NMIIA has been shown to reduce E-cadherin and β-catenin localisation at cell-cell adhesions, leading to defective cell adhesion and disordered tissue organisation (Conti et al., 2004). This implies that a reduction in the NMII protein content within the EHC epicardium following the loss of NMIIB may detrimentally impact upon the adhesion properties of these cells; this could possibly underpin the disrupted epicardial cell contacts and the disorganised myocardium that characterise the EHC heart. In addition, as a key component of the actomyosin cytoskeleton, NMIIB is highly likely to influence the structural integrity of cellular compartments such as the nucleus (Conti and Adelstein, 2008; Vicente-Manzanares et al., 2009). NMIIB has been shown to exert tension forces on the cell nucleus (Thomas et al., 2015), which may explain why EHC epicardial cells display a disrupted nuclear architecture. Interestingly, alterations in the forces acting upon chromatin structure have been shown to directly affect transcription regulation (Tajik et al., 2016). This implies that loss of NMIIB may influence epicardial gene expression - and ultimately function - through changes in chromatin stretching.

Our characterisation of the cellular components of the coronary vasculature suggests that EHC hearts fail to form mature coronary vessels. Defects in the formation of the coronary vasculature have not previously been reported for the Myh10 knock out mouse (refer to Table 3.3). However, the detection of an endothelial plexus that traverses the ventricular surface of the EHC heart suggests that NMIIB is not required for plexus formation. In addition, this would indicate that the observed defects in the epicardium do not impede plexus formation, supporting the findings of others that the vascular endothelial cells do not derive from the epicardium (Red-Horse et al., 2010; Wu et al., 2012). In addition, we were able to detect PECAM-1 positive cells in abnormal surface nodules that decorate the surface of the EHC heart, suggesting that coronary vasculogenesis may be disrupted. These structures have previously been observed following the ablation of thymosin β4 signalling (Smart et al., 2007), and disruption to epicardial polarity establishment (Rhee et al., 2009). Myocardial thymosin β4 signalling is required for correct epicardial development, suggesting that NMIIB may be required for the transduction of extracellular signals to establish polarity in the epicardium.

The recruitment of vSMC to the endothelial coronary plexus is impeded in the EHC heart. As previously mentioned, altered haemodynamic properties following the initiation of blood flow through these rudimentary vessels is thought to drive the recruitment of vSMC,

110 thus facilitating vessel maturation and the establishment of venous/arterial identity (Tomanek, 2005; Luttun and Carmeliet, 2003; Riley and Smart, 2011). Recruitment of vSMC is essential for correct coronary vessel formation (Winter and Gittenberger-de Groot 2007). Whilst the resolution of individual SMαA positive cells via the peroxidase detection system employed here is poor compared to immunofluorescent methods, this technical limitation does not detract from our finding that vSMC recruitment does not correctly occur in the EHC heart. The resulting failure to mechanically support developing vessels may lead to vessel rupture and subsequent cardiac haemorrhage. This cardiac insufficiency may underpin the sudden late gestation embryonic lethality of the EHC phenotype.

Defective vSMC recruitment is observed in the β-catenin knock out mouse, suggesting a prominent requirement for the correct regulation of epicardial cell adhesions (Zamora et al., 2007). In contrast, this study also found that the differentiation vSMC was drastically impaired. We are able to detect vSMC in the EHC heart, indicating that the processes underpinning impaired vSMC differentiation in the β-catenin knock out mouse are not disrupted in the EHC heart. This suggests that whilst NMIIB may function in the maintenance of epicardial cell adhesions, it does not play a part in wider β-catenin/Wnt signalling processes. Abnormalities in the migration of vSMC have been reported for Notch3 deficient mice, and it is thought that this process requires spatial regulation of αvβ3 integrin activity (Scheppke et al., 2012). Interestingly, NMIIB has been implicated in the spatial distribution of integrins (Puklin-Faucher and Sheetz, 2009), indicating a possible molecular process that is dysregulated following the loss of NMIIB.

In addition, the integral and well-documented role of NMIIB in cell migration may provide the simplest explanation for insufficient vSMC recruitment; vSMC may be fundamentally unable to navigate through the myocardium to reach the endothelial capillaries in the EHC heart. Conversely, it may be that the recruitment factors that drive vSMC migration are insufficiently expressed in the EHC mutant. It will be extremely informative to analyse the expression of components of the PDGF (platelet-derived growth factor) signalling pathway in the context of the EHC mutants, to investigate whether this signalling axis is disrupted following the loss of NMIIB (Hellstrom et al., 1999; Smith et al., 2011).

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3.4 Conclusion Together, these data suggest that coronary vessel formation in the EHC mouse is severely impaired and implies that whilst mutant hearts are capable of forming primitive endothelial coronaries, they lack mature, functional vessels. These observations suggest that the process of vessel remodelling and maturation is defective in EHC hearts. These coronary defects are strikingly similar to those observed in studies that have either physically ablated, or genetically disrupted the epicardium, leading to a primary defect in epicardial cell function (Zamora et al., 2007; Mellgren et al., 2008; Hellstrom et al., 1999; Moore et al., 1999; Martinez-Estrada et al., 2010). Although epicardial and coronary abnormalities have not been reported in previous studies of Myh10 knock out mice, it is possible that background specific modifying genes affecting the expression of the Myh10 locus alter the phenotypic effect of Myh10 ablation (Barbaric et al., 2007; Wong et al., 2005). This thesis has previously discussed the importance of the epicardium to the formation of the coronary vasculature. In light of this, and considering the abnormalities we observe with regard to the disruption of epicardial cell morphology and abnormal coronary vessel formation in the EHC mouse, we next sought to investigate whether or not epicardial function was compromised following loss of Myh10.

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Chapter 4: Analysis of Epicardial Cell Function

4.1 Introduction Although the three mammalian NMII isoforms are widely expressed in various tissues, they exhibit largely distinct expression profiles at the cellular level (Ma and Adelstein, 2012). In the mouse, NMIIB is enriched in both neuronal (cerebral cortex, cerebellum, spinal cord) and cardiac tissue (Ma et al, 2010). Although NMIIB has previously been shown to play a crucial role in cardiac formation and function (Ma and Adelstein, 2012), the precise function that NMIIB plays is yet to be fully elucidated. Intriguingly, epicardial and coronary defects have not been previously reported for the NMIIB ablated mouse. In light of this, we sought to investigate the relationship between NMIIB ablation and epicardial function in the EHC mouse.

4.1.1 NMII Expression and Function in the Embryonic Mouse Heart All three NMII isoforms are expressed in both the mouse and human heart (Ma and Adelstein, 2012). In the adult mouse brain, NMIIB represents the predominant NMII isoform (Ma et al., 2010). Similarly, NMIIB represents approximately 37% of the total NMII protein content in the perinatal mouse heart (Ma et al., 2010). Both NMIIA and NMIIB are expressed early in mouse development, and are found in mouse embryonic stem cells, whilst NMIIC expression is first detected at approximately E13.5 (Ma and Adelstein, 2012). Furthermore, the IIA and IIB isoforms have been shown to be expressed in the non-myocyte cardiac cell populations (including epicardial, fibroblast and endocardial cells) (Ma and Adelstein, 2012). The enrichment of NMIIB in the mouse brain and heart suggests a key function in the development and function of these tissues (Tullio et al., 1997; Takeda et al., 2003; Ma et al., 2010). Concurrently, the relative abundance of NMIIB decreases significantly in the adult mouse heart and is accompanied by a dramatic sub-cellular redistribution of NMIIB protein immediately following birth (Ma et al., 2010). This suggests that NMIIB functions predominantly during embryonic development, and that this function is distinct from that in the adult heart.

Immunofluorescence microscopy data from the Adelstein group has previously shown that NMIIB is detected in both ventricular myocytes and non-myocytes in the E13.5 embryonic heart (Ma et al., 2010). In contrast, from E7.5, expression of NMIIA in the ventricles is limited to the non-myocyte populations, whilst NMIIC is found exclusively in ventricular

113 myocytes (Ma et al., 2010). The importance of NMIIB to cardiomyocyte function has been previously reported; ablation of cardiac NMIIB causes defects in cardiomyocyte cytokinesis, resulting in the premature accumulation of binucleated cardiomyocytes and a reduction in the total myocytes population (Takeda et al., 2003; Ma et al., 2010). It is thought that NMIIB plays a critical role in formation and constriction of the contractile actin ring, which is essential for correct cell division (Ma and Adelstein, 2012). Interestingly, replacement of NMIIB with NMIIA does not rescue cardiomyocyte cytokinesis defects (Bao et al., 2007). In addition, whilst loss of NMIIC does not cause myocardial defects, double ablation of NMIIB and NMIIC greatly impairs myocytes cell division, reducing the number of cardiomyocytes and simultaneously causes abnormalities in the morphology of cardiomyocyte nuclei (Ma et al., 2010). The presence of multi-lobed cardiomyocyte nuclei in NMIIB/NMIIC double knock out animals suggests a role of NMII in karyokinesis, thought to manifest due to dysregulation of microtubule dynamics during mitotic spindle formation (Ma et al., 2010). It is thought that the cardiomyocyte defects associated with loss of NMIIB may contribute to the development of DORV through a failure to support the myocardialisation of the distal outflow tract (Ma and Adelstein, 2012). This is supported by a previous study, which attributes the presence of DORV in Vangl2 mutant mice to the inhibition of NMII activation (Phillips et al., 2005).

Interestingly, the findings outlined above show that the three NMII isoforms display distinct functions, and loss of NMIIB function in cardiomyocytes cannot be fully compensated by either NMIIA or NMIIC. In addition, the ability of NMIIB to compensate for the loss of NMIIC, indicates that NMIIB plays a distinct yet multifaceted role during cardiogenesis, and demonstrates the importance of correct NMIIB function to the developing heart. That being said, the relative expression of the NMII isoforms in specific cell populations within the heart, such as the epicardium, has not been fully explored and is yet to be resolved. This characterisation will facilitate the elucidation of NMIIB function in different cell types within the developing heart.

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4.1.2 Epicardial Cell Function During Heart Development As previously discussed, it is well established that the epicardium plays an integral role in the formation of the coronary vasculature and cardiogenesis as a whole. Considering the abnormal morphology of epicardial cells and coronary defects displayed by the EHC mouse, it is important to evaluate the relationship between the loss of Myh10 and plausible dysfunction to epicardial cell biology.

4.1.2.1 Molecular Regulation of Epicardial Function During Cardiogenesis Normal epicardial behaviour is dependent on the accomplishment of multiple cellular events, and failure to undergo these correctly results in the cardiovascular defects. As previously mentioned, epicardial cells are derived from the proepicardial organ, which forms posterior to the post-looped heart at approximately E8.5 in the mouse. After being released from the PEO at approximately E9.5, epicardial progenitor cells migrate across the pericardial space and adhere to the developing myocardium, completely enveloping the nascent heart by E11.0 (Master and Riley, 2014).

Following the formation of a uniform epithelial layer, epicardial cells enter into a signalling dialogue with the myocardium, the orchestration of which is absolutely essential to the correct execution of epicardial function and the completion of cardiogenesis (Olivey and Svensson, 2010; von Gise and Pu, 2012; Masters and Riley, 2014). The epicardium releases trophic factors that facilitate the proliferation and maturation of the ventricular myocardial wall. In addition, a sub-set of epicardial cells undergo EMT, generating the EPDC population that actively contributes to both the formation of the coronary vasculature, and the structural development of the heart. The epicardium therefore represents a unique and critical modulator of cardiac development. The molecular regulation of each of these duel aspects of epicardial function is discussed in the subsequent sections.

In contrast to its embryonic counterpart, the adult epicardium demonstrates a dramatic and rapid decrease in activity following the first perinatal week (Xin et al., 2013; Master and Riley, 2014). The adult epicardium is said to enter a quiescent state and is thought not to contribute to cardiac homeostasis following perinatal day 4 in the mouse. Interestingly, epicardial cells can withdraw from this dormant state following myocardial infarction/ischemia, which sees the concomitant reactivation of epicardial embryonic gene

115 expression and secretion of paracrine signalling molecules to mediate the myocardial response to injury (van Wijk et al., 2012; Zhou et al., 2011). Unfortunately, whilst this injury response is sufficient to prevent exsanguination, it cannot rehabilitate muscle function following cardiomyocyte loss, culminating in congestive heart failure. Epicardial reactivation following infarction predominantly promotes the proliferation of cardiac fibroblasts rather than cardiomyocytes, consequently inducing mass fibrosis and scar formation (Zhou and Pu, 2012; Kispert, 2012). Whilst recent efforts have reported increased cardiomyocyte production following infarction by priming with thymosin β4, which acts to promote the differentiation of Wt1-marked EPDCs to a cardiomyocyte fate (Smart et al., 2011), the number of regenerated cardiomyocytes needs to increase enormously to compensate for the loss of billions of cardiomyocytes following infarction. Accordingly, there is currently a concerted research focus on understanding the sequential mechanisms that underpin embryonic epicardial cell function, in order to better harness epicardial potential in heart regeneration and repair.

4.1.2.2 Formation of the Epicardium The processes that initiate and guide epicardial cell migration from the PEO to the myocardial surface remain unclear. However, it is thought that migration may be facilitated by the formation of an extracellular matrix bridge that spans the coelomic cavity in the chick embryo (Nahirney et al., 2003). The conservation of this structure in the mouse is yet to be confirmed or rebutted.

Studies in the mouse have indicated that adhesive interactions between vascular adhesion molecule 1 (VCAM-1) and α4β1 integrin play a crucial role in the process of spreading epicardial cells across the surface of the myocardium (Kwee et al., 1995; Yang et al., 1995). Reciprocal expression of α4 integrin and fibronectin in the epicardium, and VCAM-1 in the sub-epicardial myocardium facilitate the adhesion of epicardial cells, and deletion of either α4 integrin or VCAM-1 results in the loss of the epicardium, failure to form coronary vessels and cardiac haemorrhage (Kwee et al., 1995; Yang et al., 1995). It has also been suggested that α4 integrin mediated interactions are essential for the migration of PEO vesicles across the pericardial cavity (Sengbusch et al., 2002) and in the regulation of EMT activation in epicardial cells (Dettman et al., 2003; Winter and Gittenberger-de Groot, 2007). This highlights the importance of correct modulation of cell adhesion in the formation and function of the epicardium, and that disruption to this adhesion mechanism results in cardiac defects.

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The correct establishment of cell polarity has similarly been demonstrated to be critically important for the formation of the epicardium (Hirose et al., 2006). Mice lacking Par3 show abnormalities in the formation of epicardial precursor cyst structures and do not develop an epicardium, resulting in mid-gestation embryonic lethality (Hirose et al., 2006). Epicardial precursors show disrupted localisation of polarity markers including, aPKC (atypical protein kinase C) and PAR6β (partitioning defective 6-β), suggesting that epicardial precursors are required to sense and respond to polarity establishing cues from their extracellular environment in order to generate PEO cysts.

4.1.2.3 Epicardial Proliferation and EMT Correct epicardial cell function has been implicated in the formation and maturation of the coronary vasculature. Coronary vessel formation is dependent upon the cellular contribution of epicardial-derived cells, which are generated by the activation of EMT in a subset of epicardial cells. The initiation of EMT endows EPDCs with the ability to delaminate from the epicardium, and invade the underlying myocardium, where they differentiate into the vascular smooth muscle and perivascular fibroblast components of the coronary vasculature. Additionally, it is suspected that EPDCs exert a regulatory influence on the development of the coronary vessels and other cardiac tissue through their expression of paracrine signaling molecules (Stuckmann et al., 2003; Lavine et al., 2006). For example, epicardial-derived FGF signaling regulates the activation Hedgehog, which in turn, stimulates VEGF and angiopoietin expression to remodel the nascent vasculature (Lavine et al., 2006). The epicardium must receive and correctly respond to EMT initiating factors in order to generate migrating EPDCs. This process encompasses a multitude of cellular changes that must be correctly orchestrated.

A number of signalling cascades have been implicated in the promotion of epicardial EMT. It has previously been shown that the transcription factor Wt1 is essential to initiate epicardial EMT through direct transcriptional control of Snai1 and Cdh1, which encode Snail and E-cadherin respectively (Moore et al., 1999; Martinez-Estrada et al., 2010). Snail transcriptionally represses E-cadherin expression in the epicardium, promoting the adoption of a mesenchymal phenotype by epicardial cells (Martinez-Estrada et al., 2010). The importance of Wt1 to epicardial EMT is highlighted by the finding that Wt1 deficient epicardial cells fail to undergo EMT, resulting in a lack of coronary vessels and a failure of ventricular compact myocardium expansion (Moore et al., 1999; Martinez-Estrada et al., 2010). It has been shown that Wt1 is expressed specifically in the PEO, epicardium and

117 EPDCs, but decreases as EPDCs terminally differentiate (Perez-Pomares et al., 2002b). It has been hypothesised that Wt1 expression maintains EPDCs in a condition to activate myocardial proliferation, prior to terminal EPDC differentiation (Perez-Pomares et al., 2002b). In addition, activation of epicardial EMT by platelet derived growth factor (PDGF) signalling has been shown to be essential for the development and recruitment of epicardial derived coronary smooth muscle cells to developing vessels (Hellstrom et al., 1999; Smith et al., 2011).

It has also been shown that the plane of epicardial cell division dictates the fate of daughter epicardial cells and thus the generation of EPDCs (Wu et al., 2010). Cells remain in the epicardial layer if they align their mitotic spindle parallel to the plane of the basement membrane, and migrate into the myocardium to generate EPDCs if spindle orientation is perpendicular to the basement membrane (Wu et al., 2010). Loss of β-catenin causes disrupted epicardial cell adherence and the randomisation of mitotic spindle orientation. The subsequent dysregulation of EMT indicates the importance of cell adhesion in epicardial function and EPDC generation.

4.1.2.4 EPDC Migration and Differentiation Epicardial derived cells are distinguished by their expression of Wt1, Tbx18, Raldh2 and Tcf21 (Masters and Riley, 2014). It is well established that EPDC differentiate into both cardiac fibroblast and vascular smooth muscle cells in both the mouse and chick (Cai et al., 2008; Zhou et al., 2008; Mikawa and Fischman, 1992; Dettman et al., 1998; Gittenberger- de Groot et al., 1998) (Fig 4.1). The extent of the contribution of EPDC to the cardiomyocyte and vascular endothelial linages is subject of considerable debate (Gittenberger-de Groot et al., 2012). Fate-mapping studies have reported a minor contribution of EPDC to the cardiomyocyte population (Fig 4.1) (Cai et al., 2008; Zhou et al., 2008), which can be recapitulated during the adult response to injury in a thymosin β4 dependent manner (Smart et al., 2011). However, an epicardial origin for embryonic cardiomyocytes has recently been challenged (Christoffels et al., 2009; Kispert, 2012). Similarly, it has been suggested that EPDCs can differentiate into vascular endothelial cells, and that this process involves the Shh (sonic hedgehog)-VEGF-Ang2 signalling axis (Fig 4.1) (Olivey and Svensson, 2010; Smart and Riley, 2012). However, Red-Horse and colleagues (2010) have reported that the coronary endothelial plexus originates from the sinus venosus (Red-Horse et al., 2010). It has also been suggested that the vascular endothelium arises from the endocardium (Wu et al., 2012).

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Figure 4.1: Schematic Representation of the Differentiation of EPDCs into Multiple Cardiac Cell Lineages A sub-set of epicardial cells undergo EMT to generate epicardial-derived cells (EPDCs). These cells have been shown to differentiate into numerous cell-fates within the embryonic heart. Key molecular signalling molecules involved in each differentiation process are indicated. Please see full text for details. CM – cardiomyocyte, ECM – extracellular matrix, VEC – vascular endothelial cell, VSMC – vascular smooth muscle cell.

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However, it has been demonstrated that the epicardial progenitor cells are not homogeneous, and that the PEO contains distinct populations of either Wt1 and Tbx18, or Sema3D and Scx expressing cells (Katz et al., 2012). Here the authors report that whilst vSMCs and fibroblasts were derived from both Wt1/Tbx18 and Sema3D/Scx expressing progenitors, only Sema3D/Scx expressing cells contributed to the coronary endothelium lineage (Katz et al., 2012). The contention surrounding these somewhat contradicting findings is likely rooted in a combination of interspecies variation, and the limitations of the genetic fate-mapping tools employed. Endogenous expression of EPDC markers in both cardiomyocytes and endothelial cells compromises the accurate interpretation of such lineage tracing experiments (Christoffels et al., 2009; Rudat and Kispert, 2012; Zhou and Pu, 2012).

The differentiation of EPDC into vSMCs is essential for coronary artery development and is dependent upon Notch signalling (del Monte et al., 2011; Grieskamp et al., 2011). Smooth muscle cells are required to provide mechanical support to the nascent vessel, and the absence of these cells causes vessel rupture and cardiac hemorrhage. It has been suggested that the molecular signals driving differentiation to this cell fate include PDGF, Wnt/β-catenin and TGFβ (Olivey and Svensson, 2010). Similarly, ablation of β-catenin specifically in the PEO severely impedes coronary artery, but not vein, formation (Zamora et al., 2007). This study reported that whilst β-catenin deficient embryos display correct epicardial formation, migration of EPDC and subsequent differentiation into vSMC is drastically impaired (Zamora et al., 2007). Similarly, it has also been shown that migration and differentiation of EPDCs into vascular lineages is dependent upon myocardial thymosin β4 expression (Smart et al., 2007). In addition to cellular contributions to developing heart structures, EPDCs provide paracrine signals to mediate cardiomyocyte proliferation. Myocardial-specific deletion of thymosin β4 causes defects in epicardial formation and development of the compact myocardium. Interestingly, mutants also displayed ventricular blood filled nodules, as observed in the EHC mouse (Smart et al., 2007). Treatment of epicardial explant cultures with a combination of thymosin β4, VEGF and FGF7 facilitated vasculogenic outgrowth (Smart et al., 2007). In addition, the FOG2 (friend of GATA 2) mutant phenotype is rescued by recovering myocardial FOG2 expression (Tevosian et al., 2000). This highlights the important of paracrine signalling from the myocardium for multiple aspects of epicardial function.

Similarly, FGF signalling has been further implicated in the migration of EPDCs, in addition to the differentiation of cardiac fibroblasts (Vega-Hernandez et al., 2011). These 120

authors propose that cardiac fibroblasts are required to support myocardial proliferation, and report that disruption to the FGF10/FGFR2b signalling axis disrupts the migration of EPDC derived fibroblasts into the myocardium. Loss of myocardial FGF10 signalling, concurrent with a loss of FGF responsiveness in the epicardium, disrupts epicardial and myocardial development simultaneously. Furthermore, FGF signalling is intricately associated with epicardial EMT and coronary neovascularization during the injury response in the zebrafish (Lepilina et al., 2006). This again emphasizes the critical importance of molecular communication between the epicardium and myocardium in both heart formation and repair. The ability of each of these tissues to interpret and correctly respond to reciprocal signals is imperative to these processes.

4.1.2.5 Epicardial Mediated Myocardial Proliferation The epicardium has been consistently associated with the correct development and maturation of the ventricular myocardial wall through secretion of paracrine signals (Watt et al., 2004; Perez-Pomares et al., 2002b; Moore et al., 1999; Zamora et al., 2007; Niederreither et al., 2001; Lavine et al., 2006). Defects in myocardial development are observed when the epicardium is physically excised from the developing embryo (Stuckmann et al., 2003). The differentiation of EPDC into fibroblasts coincides with the proliferation of embryonic cardiomyocytes and the respective expansion of the myocardium (Ieda et al., 2009). Co-culture experiments have indicated that stimulation of cardiomyocyte proliferation involves the secretion of the extracellular matrix components fibronectin and collagen by fibroblasts, and that the interpretation of these signals is mediated by myocardial β1-intgrin (Ieda et al., 2009).

In addition, it is thought that retinoic acid signalling is critically important for correct myocardial proliferation. Mice deficient for the retinoic acid receptor, Rxra, specifically in the epicardium, display a detached epicardium and hypomorphic myocardium (Sucov et al., 1994). Congruently, blocked retinoic acid synthesis in mice lacking Raldh2 (retinaldehyde dehydrogenase-2), causes gross heart defects and midgestation embryonic lethality (Niederreither et al., 1999). The epicardium responds to retinoic acid by expressing trophic factors, including FGF2 (Merki et al., 2005). FGF2 deficient mice display cardiac proliferation defects (Virag et al., 2007). This is supported by findings that heart slices incorporating the epicardium can initiate myocardial proliferation in response to retinoic acid, but that this response is not observed in slices lacking the epicardium (Stuckman et al., 2003). Furthermore, cardiomyocytes express FGF receptors, and genetic ablation of

121 FGF9-FGFR1/FGFR2 signalling results in defective formation of the compact myocardium (Lavine et al., 2005). In addition, expression of hepatic erythropoietin in response to retinoic acid induces epicardial Igf2, which in turn induces myocardial proliferation (Brade et al., 2011). However, it appears that both the ability of the epicardium to produce trophic factors, and the ability of the myocardium to respond to these factors, diminishes and is rapidly lost in the first few perinatal days (Porrello et al., 2011; Chen et al., 2002).

In consideration of both the studies outlined above, and the evidence presented in the previous chapter suggestive of epicardial defects in EHC hearts, we embarked upon a series of experiments to determine whether or not the EHC mutant heart displayed compromised epicardial cell function; these experiments are central to determine the elements of the coronary vessel developmental programme that are disrupted in EHC mutants. We utilised a combination of immunofluorescent microscopy and western blotting techniques to profile both the spatial and temporal expression of NMIIB and a number of key molecules associated with proper epicardial cell function. These experiments will help to identify the defective processes that may underpin abnormal coronary vessel formation, and directly address the following questions: (a) is NMIIB expressed in the epicardium? (b) what is the consequence of NMIIB deletion on the migration of EPDCs? and (c) do EHC epicardial cells display defects in EMT?

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4.2 Results

4.2.1 NMHC IIB is the Predominant NMII Isoform in the Embryonic Epicardium The three NMII isoforms are encoded by three independent genes, located on different chromosomes. Although a degree of overlap between their expression profiles can be found in some tissues, they exhibit largely distinct temporal and spatial expression throughout the developing embryo (Vincente-Manzanares et al., 2009; Conti and Adelstein, 2008). It has shown by a number of different techniques that NMIIB is enriched in the heart and brain of the embryonic mouse (Ma et al., 2010). The developmental defects observed in these organs in the Myh10 ablated mouse have therefore been attributed to the predominant expression of NMHC IIB in these tissues. However, whilst the relative expression profiles of the NMII isoforms has been examined in a number of different adult cell lineages within the heart (Ma et al., 2007; Ma et al., 2009; Ma et al., 2010), this has not been extended to a detailed analysis of the embryonic epicardium. In light of this, the relative abundance of the individual NMII isoforms in epicardial cells is yet to be resolved. To address this knowledge gap, we undertook a combination of both immunohistochemistry and western blotting experiments to profile the localisation and relative abundance of NMII in control and NMIIB ablated samples.

4.2.1.1 Relative NMII Abundance in the Embryonic Heart We performed western blotting on protein extracts from E11.5 whole embryonic hearts with antibodies against the three NMHC II isoforms. To allow for comparisons between samples, protein extracts were also probed for β-actin. As a universally considered ‘house keeping’ gene, β-actin expression can be assumed to be relatively uniform between samples and can therefore be used as a normalisation measurement of the total level of protein in a given sample. Densitometry measurements are provided in Appendix 11.

From this analysis, we were able to determine that both NMIIA and NMIIB are abundant in the E11.5 control (∆/+) embryonic heart (Fig 4.2, A, B). We were unable to detect NMIIC in these protein extract samples (Fig 4.2, C). All samples contained a relatively similar total protein content, as determined by the similar immunoreactive intensities of the β-actin band (at approximately 42kDa) (Fig 4.2, A-C).

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Figure 4.2: Expression of NMII Isoforms in Control and Myh10∆ Mutant Embryonic Hearts (A) Western blots were performed on pooled protein extracts from control (∆/+, n=9) and mutant (∆/∆, n=3) E11.5 hearts for NMIIA, (B) NMIIB and (C) NMIIC. Bands at approximately 250kDa correspond to the NMII isoform. The bands at approximately 42kDa correspond to β-actin, which was used as a loading control to compare protein content across samples. (D) Immunohistochemistry on paraformaldehyde-fixed cardiac sections from control (∆/+, n=3) E14.5 hearts for NMIIA, (E) NMIIB and (F) NMIIC. NMIIB is the predominant NMII isoform expressed in the epicardium (E, arrowheads). (G-I) As a positive control, immunohistochemistry for the three NMII isoforms was performed on paraformaldehyde-fixed E14.5 mouse lung tissue (n=3). NMIIA expression is found in lung epithelial cells (G, arrowheads). NMIIB is expressed throughout the E14.5 lung, but is predominantly localised to the paramesenchyme tissue (H, asterisk). NMIIC is found exclusively in lung epithelial cells (I, arrowhead).

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As expected, we did not detect any NMIIB immunoreactivity in NMIIB ablated heart extracts. Interestingly, we observed a small reduction in the intensity of the NMIIA band in mutant samples, as compared to controls (Fig 4.2, A). We were also unable to detect NMIIC in mutant samples (Fig 4.2, C). These data suggest that in the early mouse heart, NMIIA and NMIIB are the predominant NMII isoforms. Additionally, we were able to detect all NMII isoforms in protein extract from adult mouse lung tissue, which was used as a positive control as all three isoforms are known to be expressed in this tissue (Ma et al., 2010).

In order to delineate the spatial expression profile of NMII isoforms, we complimented our western blot data with immunohistochemical analysis at E14.5 (n=3). This later developmental time point was selected to further resolve our understanding of the temporal expression of the NMII isoforms. In addition, at E14.5, the embryonic epicardium is highly involved in the formation of the coronary vasculature, and influences myocardial development and functionality.

Our immunohistochemical observations showed once again that NMIIB is highly abundant in the control (∆/+) embryonic heart (Fig 4.2, D-F). We found strong NMIIB immunoreactivity throughout the tissue sections, indicating that NMIIB is localised throughout the ventricular myocardial wall (Fig 4.2, E). This immunoreactivity for NMIIB was noticeably more intense than signals for NMIIA and NMIIC in serial sections of the same hearts (Fig 4.2, D-F). By comparing immunoreactive signal in NMII stained sections, we concluded that NMIIB is the predominant NMII isoform in the heart at E14.5. Somewhat surprisingly, we found that NMIIB staining was extremely prominent within the epicardium itself (Fig 4.2, E, arrowheads). Moreover, our analysis indicates that NMIIB is the predominant NMII isoform in this tissue (Fig 4.2, D-F). This study expands upon pre- existing data for NMII levels within the heart, and further resolves the spatial and temporal expression of NMII during cardiac development.

Furthermore, we were able to readily detect all NMII isoforms within the embryonic lung of the same sections used for the heart analysis (Fig 4.2, G-I, n=3). As previously reported, NMIIA was detectable predominantly in the lung epithelial cells (Fig 4.2, G, arrowhead), NMIIB predominantly in the paramesenchyme (Fig 4.2, H, asterisk), and NMIIC exclusively in the lung epithelium (Fig 4.2, I, arrowhead) (Ma et al., 2010). The use of this positive control tissue indicates that our immunohistochemistry experimental data from the heart can be reliably interpreted.

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4.2.1.2 NMII Sub-Cellular Localisation in Epicardial Cell Cultures Our finding that NMIIB is enriched in the epicardium of control E14.5 hearts prompted us to further examine NMII protein expression in cultured epicardial cells. As we were unable to detect NMIIC in E11.5 embryonic hearts, we performed NMIIA and NMIIB immunocytochemistry on cultured epicardial monolayers from E11.5 heart explants (see Section 4.2.2 for details of culture generation and epicardial validation criteria). We were able to detect positive staining for both NMII isoforms in control (∆/+) cells (Fig 4.3, A- D). All antibodies and detection reagents were used at the same dilutions in each protocol. Although one cannot ascertain quantitative values of expression via immuno-staining, it did appear that NMIIB signal was more intense than that obtained for NMIIA (Fig 4.3, A, C). Interestingly, we found that NMIIA and NMIIB occupy distinct sub-cellular localisation in cultured epicardial cells. NMIIA signal was mainly restricted to the cell periphery, with very little staining in the proximal cell body. NMIIA staining was always found to co-localise with rhodamine-phalloidin staining of the filamentous actin cytoskeleton, particularly at the leading edge of migrating cells (Fig 4.3, A, B, arrowheads). In contrast, we found that NMIIB demonstrated a more ubiquitous expression profile in cultured epicardial cells (Fig 4.3, C, D). We found diffuse staining throughout the cell body, often independent of actin staining (Fig 4.3, D, asterisks). In addition, we also observed that NMIIB frequently associated with actin stress fibres that traverse the cell body (Fig 4.3, C, D, arrowheads). In stark contrast with our findings for NMIIA, NMIIB appeared to be completely absent from the leading edge of migrating cells, but was enriched at the trailing edge (Fig 4.3, C, D). Although we have not directly performed the experiment to show localisation of NMIIA and NMIIB in the same cells, our data suggests that both isoforms occupy largely distinct sub-cellular localisation, with a small degree of co-localisation at the cell periphery.

As expected, we were unable to detect NMIIB immunoreactivity in NMIIB ablated cells (Fig 4.3, F). Interestingly, we did not see a change in either the relative abundance or sub- cellular localisation of NMIIA in mutant epicardial cells (Fig 4.3, E). This experiment compliments our western blot data by suggesting that the expression of other NMII isoforms is not up-regulated in response to the loss of NMIIB in embryonic heart tissue. For completeness, we attempted to profile the relative expression of the three NMII isoforms in cultured epicardial cells via western blot (data not shown, see Appendix 11 for densitometry measurements). A total of 18 epicardial explant cultures (from 6 embryonic hearts) were pooled for both control (+/+ and ∆/+) and mutant (∆/∆) samples.

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Figure 4.3: NMIIA and NMIIB Expression in Cultured Epicardial Cells (A-B) NMIIA localisation in cultured epicardial cells derived from control (∆/+, n=6) E11.5 hearts. NMIIA predominantly co-localizes with actin filaments at the cell periphery (A, B, arrowheads). (C-D) NMIIB localisation in cultured epicardial cells derived from control (∆/+, n=6) E11.5 hearts. NMIIB staining is more pronounced and present throughout the cell body (D, asterisk). NMIIB often co-localizes with actin stress fibres that transverse the cell body (C, D, arrowheads). (E) NMIIA localisation in cultured epicardial cells derived from mutant (∆/∆, n=4) E11.5 hearts. The relative abundance and sub-cellular localization of NMIIA is not noticeably altered in mutant epicardial cells. (F) As expected, NMIIB is not detected in mutant epicardial cells. Scale bars = 100µm (A, C, E, F), 50µm (B, D).

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Epicardial cells were harvested via centrifugation and resuspended in western blot sample loading buffer. Unfortunately, the quality of the resultant western blots was poor due to the limited number of cells, making interpretation of the blots unreliable. However, although the immunoreactive band was extremely weak, NMIIB was the only detectable isoform in control epicardial cells. No band was observed for either of the three NMII isoforms in mutant cultures (data not shown).

Together, these western blotting and immune-staining experiments have shown that NMIIB is abundant in both the whole embryonic heart and cultured epicardial cells at different developmental time points (E11.5 and E14.5). These embryonic stages correlate to a time window during cardiogenesis when the epicardium is highly active. The finding that NMIIB is enriched in the epicardium and appears to be the predominant form of NMII in this tissue at the time points analysed, suggests that NMIIB expression is required for embryonic epicardial cell function.

4.2.2 Epicardial Cell Culture To further examine the functionality of mutant epicardial cells, we employed an epicardial cell culture system (Chen et al., 2002). Isolating epicardial cells from embryonic hearts and maintaining them in vitro for future experimentation allows for the control of a number of parameters that are not possible in the in vivo context. As our laboratory experienced technical difficulties in isolating epicardial cells from whole embryonic hearts by using flow cytometry (i.e. FACS), we employed an epicardial cell culture model whereby we enriched epicardial cells from ventricular explants of E11.5 embryonic hearts. This developmental time point was selected based on previous optimization of the culture protocol; epicardial cultures from E11.5 ventricular explants express embryonic epicardial markers, do not express markers of cardiomyocytes or the endocardium, and display a highly migratory phenotype that permits maximal epicardial migration form the explant to the culture substrate (Chen et al., 2002). We utilised the Myh10∆ line for use in these cell culture experiments, due to the simplified genotyping protocol required; the one-step PCR protocol permitted the simultaneous generation of cultures and genotyping results, whereas the EHC mutant mice must be genotyped by a longer protocol requiring Sanger sequencing.

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4.2.2.1 Cultured Epicardial Cell Morphology Ventricular tissue from E11.5 hearts was dissected into 3-4 pieces and plated on gelatin coated tissue culture plates. After a period of 24 hours, a ‘halo’ of cells that have migrated from the explant can be observed adhered to the plate. These cells can be maintained in standard culture conditions (37°C, 5% CO2) for approximately 3-4 days. We were able to confirm that the cells isolated from this method displayed a number of epicardial characteristics. Cultured cells displayed a typical ‘cobblestone-like’ staining for the epithelial cell markers ZO-1 (zonula occludens-1), which delineates tight junctions (Fig 4.4, A, B) and filamentous actin, which highlights the actin cytoskeleton (Fig 4.4, C, D). These cells also displayed positive staining for the epicardial cell and EPDC marker, Wt1 (Fig 4.4, E, F). From this analysis, and based on these well-defined epicardial molecular characteristics, we estimated that the purity of cultured epicardial monolayers derived from this explant model was in excess of 90%.

Crucially, we found that epicardial cells cultured from both control (+/+ or ∆/+, n=6) and mutant (∆/∆, n=3) hearts explants displayed a very similar staining profile for these epicardial cell markers (Fig 4.4). That being said, we did notice that mutant epicardial cells occasionally displayed gaps in the epithelial sheet (Fig 4.4, G, H, asterisks, approximately 25% of cultures). This subtle morphological aberration mimics that seen in the in vivo epicardium.

After 5 days in culture, we found that cells began to lose their epithelial morphology, and adopted a mesenchymal phenotype as determined by altered filamentous actin staining and loss of ZO-1 staining at the cell periphery (data not shown). This indicates that cultured epicardial cells show a propensity to spontaneously differentiate after prolonged incubation in culture. For this reason, experiments were carried out no later than after a total of 4 days in culture. Whilst we cannot rule out contamination of our culture model by other cardiac cell populations, we are confident that these cultures are enriched for epicardial cells, and represent a good model to examine epicardial cell function in our in vitro experiments

For completeness, we also examined the morphology of cells derived from EHC mutant heart explants. Cells displayed an extremely similar phenotype to that shown by ∆/∆ mutant epicardial cells, and stained positive for Wt1 and ZO-1 (data not shown, n=3). We therefore concluded that ∆/∆ mutant cultures were a suitable model to study the effects of the loss of NMIIB in epicardial cells.

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Figure 4.4: Morphology and Molecular Profile of Cultured Epicardial Cells (A) Both control (∆/+, n=4) and (B) mutant (∆/∆, n=3) epicardial cells display positive staining for the cell junction protein and epithelial marker, ZO-1. (C) Control and (D) mutant epicardial cells display similar ‘cobblestone-like’ filamentous actin localization, as determined by rhodamine-phalloidin staining. (E) Positive staining in control and (F) mutant epicardial cell cultures for the epicardial marker, Wt1. (G-H) An estimated 25% of mutant cultures displayed gaps in the cell monolayer, suggestive of defects in cell-cell adhesions (G, H, asterisks). All images acquired at the same magnification.

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4.2.2.2 Myh10 Ablated Epicardial Cells Do Not Show Migration Defects in vitro As previously discussed, NMIIB is enriched in the epicardium during a period of embryonic development in which the epicardium is functionally active. This suggests that NMIIB plays a key role in normal epicardial functionality and ultimately, correct cardiogenesis. NMIIB is known to play a crucial role in a broad variety of cellular processes, including cell migration (Vincente-Manzanares et al., 2009; Lo et al., 2004). Similarly, the correct function of epicardial cells in vivo is reliant upon migratory events, such as the migration of cells from the proepicardial organ across the pericardial space, and the invasion of EPDCs into the ventricular myocardium. In light of this, we sought to establish whether or not epicardial cells in which NMIIB has been functionally ablated exhibit cell migration defects.

To study epicardial cell migration, we performed a scratch-wound assay on day 3 epicardial monolayers. This experiment required the removal of an area of cells from the monolayer by scratching with a sterile pipette tip. Time-lapse microscopy was then used to image the repopulation of the denuded a-cellular area over a 20-hour time period (Fig 4.5, A-F). We could clearly observe mutant cells (Fig 4.5, D-F) migrating into the denuded area at a similar rate as seen in the control cultures (Fig 4.5, A-C). In order to investigate specific aspects of cell migration, we selected 10 cells per field of view that were situated on the leading edge of the scratched area at time zero (T0). We tracked the location of these cells in sequential images of the image series to generate tracked data of individual cell migration in space and time (Fig 4.5, C, F, multicoloured lines). We then analysed this track data and compared the average migration speed and directional persistence between control (n=240) and mutant (n=270) cells (Fig 4.5, G, H). This analysis revealed that there is no significant difference in either the migration speed (p=0.6717, Mann Whitney U test) or directional persistence (p=0.2494, Mann Whitney U test) between control and mutant epicardial cells in vitro (Fig 4.5, G, H). Please refer to the supplemental video files for full time-lapse microscopy tracking data (Appendix 14, Attached CD).

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Figure 4.5: Scratch-Wound Assay on Epicardial Cell Cultures (A-C) Time-lapse microscopy imaging of wound healing in control (∆/+) and (D-F) mutant (∆/∆) epicardial cell cultures. A ‘denuded’ area was created by scratching day 3 epicardial cell culture monolayers with a P20 pipette tip (as indicated by dashed line in A and D). Point visiting was used to image the same field of view at 10-minute intervals for a period of 20 hours. Ten cells per field of view were tracked using the MTrackJ plugin in ImageJ (as indicated by multicoloured lines in C and F). (G) Track data was used to compare the average migration velocity in control and mutant cell cultures over the duration of the assay. Average migration speed of control cells = 0.3099µm/min (n=240), and mutant cells = 0.3098µm/min (n=270) (p=0.6717, Mann Whitney U test). (H) Track data was used to compare the average migration directional persistence in control and mutant cell cultures over the duration of the assay. Average directional persistence of control cells = 0.7342 (n=240), and mutant cells = 0.7411 (n=270) (p=0.2494, Mann Whitney U test). Control refers to scratch-wound assays performed on epicardial cell cultures derived from both wild type (+/+) and heterozygous Myh10∆ (∆/+) hearts. All images were acquired at the same magnification. Scale bars = 250µm (A, D).

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4.2.3 EHC Mutants Show Defective EPDC Migration in vivo Our in vitro migration data indicates that NMIIB mutant cells do not display gross migratory defects. However, as epicardial cell function during cardiogenesis is reliant upon the correct orchestration of multiple migratory events, we sought to establish whether these processes occur correctly in the context of the developing embryo.

In the EHC mutant, it is evident that cells from the proepicardial organ retain the capacity to migrate across the pericardial space, as we are able to identify epicardial cells on the surface of the ventricular myocardium. In addition, it appears that after making initial contact with the myocardium, the epicardial cells are able to migrate on the surface of the nascent heart. Although the mutant epicardium shows abnormal gaps between individual cells, it does extend to surround the entire myocardium. As a consequence, we decided to investigate in vivo epicardial cell migration in relation to the invasion of EPDCs into the myocardial wall.

Epicardial cells acquire the ability to leave the epicardial epithelium via the process of EMT. This process is initiated in a sub-set of epicardial cells at approximately E11.5-E12.5 (Perez-Pomares and de la Pompa, 2011), and subsequently gives rise to the EPDC population. EPDCs migrate into the myocardium, where they differentiate into a number of different cell lineages. The transcription factor Wt1 (encodes Wilms tumour protein 1, Wt1) is expressed in both the epicardium and EPDCs. Therefore, Wt1 is considered one of the most reliable markers of epicardial cells and EPDCs during embryonic development in the mouse.

We utilised Wt1 immunohistochemistry to identify the localisation of EPDCs in the ventricular myocardium of E14.5 hearts (Fig 4.6, A-F). Cells showing Wt1 positive immunoreactivity are indicated by yellow crosshairs (Fig 4.6, C, D). We then manually measured the distance between the centre of the Wt1 positive cell nuclei and the apical boundary of the epicardium, and compared the mean migration distance of control (EHC/+, n=975 from >3 individual hearts) and mutant (EHC/EHC, n=1180 from >3 individual hearts) Wt1 positive cells. This analysis revealed that the average distance between Wt1 positive cells and the apical epicardial boundary was significantly reduced in the EHC mutant heart (Fig 4.6, G, p<0.0001, Mann Whitney U test).

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Figure 4.6: Analysis of EPDC Migration in vivo by Wt1 Immunohistochemistry (A) Representative images of coronal sections of E14.5 control (EHC/+) and (B) mutant (EHC/EHC) hearts stained for Wt1. Boxed area indicates area of magnification shown in panels C and D. (C, D) Wt1 positive cells (red) were identified by co-staining with the nuclear marker DAPI (blue), and are indicated by yellow crosshairs. (E) Wt1 staining identified clusters of EPDCs in mutant hearts in the sub-epicardial region (arrowheads). (F) In addition, mutant EPDCs were found to surround the blood-filled ventricular nodule structures (arrowheads). (G) The distance of Wt1 positive EPDCs from the apical epicardial boundary was manually measured using ImageJ. Average migration distances were compared between genotypes. Mean migration distance of control cells = 33.37µm (+/- 1.197µm, n=975), and mutant cells = 20.14µm (+/- 0.8136µm, n=1180) (p<0.0001, Mann Whitney U-test). (F) Histogram comparing the proportions of Wt1 positive cells at different myocardial depths. A larger proportion of total mutant cells (87.37%) resides <50µm from the apical epicardial boundary than in controls (68.4%). Scale bars = 250µm (A-B), 100µm (C-D), 50µm (E-F).

Concurrently, when we examined the relative proportion of Wt1 cells at different depths within the myocardial wall, we found that a higher proportion of mutant Wt1 cells (87.37%) were located within a sub-epicardial region of approximately 30-50µm when compared to controls (68.4%) (Fig 4.6, E, H). In addition to forming sub-epicardial ‘clusters’ (Fig 4.6, E, arrowheads), EHC Wt1 positive cells appear to surround the blood filled ventricular nodules (Fig 4.6, F, arrowheads). These data show that the ability of mutant EPDCs to invade the myocardium is significantly reduced compared to controls, and as a consequence, the localisation of the vast majority of these cells is restricted to the sub-epicardial zone.

This result contradicts our in vitro findings from the epicardial cell culture model. However, one can therefore interpret that the ability of NMIIB ablated epicardial cells to migrate is therefore highly dependent upon context of the epicardial microenvironment. The constitution of the cell microenvironment in vitro permits epicardial cell migration in the absence of NMIIB, whereas the in vivo microenvironment dictates a requirement for NMIIB to facilitate correct migration.

4.2.4 EHC EPDC Retain Their Differentiation Capacity After establishing that EHC EPDCs show defects in cell migration in vivo, we sought to investigate whether the differentiation capacity of these cells was similarly disrupted. Following invasion of the myocardium, it is known that EPDCs differentiate into a number of cell lineages. As previously mentioned, the vast majority of the vSMCs components of the coronary vessels are derived from these EPDCs. In addition, EPDCs have been shown to contribute to the cardiac fibroblast lineage, and differentiate into both interstitial and perivascular fibroblasts. It is clear that the correct differentiation of EPDCs following migration into the myocardium is fundamental for correct cardiogenesis, and particularly indispensible for coronary vessel formation.

Our previous immunohistochemistry experiments examining the localisation of vSMC in the EHC mutant heart shows positive SMαA staining throughout the myocardium, although this staining profile is aberrant when compared to controls (Fig 3.8, E, F). This suggests that whilst migration is disrupted, mutant EPDCs retain the ability to differentiate into the vSMC lineage.

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To expand upon this finding, we performed vimentin immunohistochemistry to detect the presence of cardiac fibroblasts in the mutant heart. Vimentin is a marker of mesenchymal cells, and has previously been shown to identify cardiac fibroblasts in the embryonic murine heart (Camelliti et al., 2005; Zamora et al., 2007) In control samples, vimentin staining is evident throughout the ventricular wall, indicating the presence of fibroblast cells throughout this tissue (Fig 4.7, A, B, n=3). At higher magnification, the filamentous characteristics of vimentin staining become apparent (Fig 4.7, E, F, arrowheads), mimicking the spindle-like morphology of fibroblasts. Similarly, fibroblasts are detectable throughout the myocardium of EHC mutants (Fig 4.7, C, D, n=3). Once again, higher magnification images show that filamentous vimentin staining highlights the fibroblast morphology (Fig 4.7, G, H, arrowheads). It is interesting to note that the mutant myocardium shows a reduction in cell density, which we have attributed primarily to defects in cardiomyocyte cytokinesis (Fig 4.7, H). The observation that vimentin immunoreactivity is detectable in the EHC heart suggests that mutant EPDCs retain the capacity to differentiate into the fibroblast cell lineage. Together, the observation that both vSMCs and cardiac fibroblast are present in the EHC mutant myocardium, clearly demonstrates that mutant EPDCs retain a level of multipotency.

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Figure 4.7: Analysis of Cardiac Fibroblast Localisation by Vimentin Immunohistochemistry (A-B) Vimentin staining of cardiac fibroblasts in control (EHC/+, n=3) and (C-D) mutant (EHC/EHC, n=3) E14.5 hearts. (E-F) Higher magnification images of control heart depicted in A and B. (G-H) Higher magnification images of mutant heart shown in C and D. Spindle-like vimentin staining indicates the presence of cardiac fibroblast throughout the tissue section in both control (E-F) and (G-H) mutant hearts (arrowheads). Scale bars = 200µm (A-D), and 100µm (E-H). Images courtesy of Irina-Elena Lupu.

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4.2.5 Analysis of Epicardial Cell Proliferation Although our analysis of the Wt1 immunohistochemistry experiments shows a clear reduction in the migration distance of EHC mutant EPDCs, it is important to eliminate other defects that may influence this analysis and lead to inaccurate interpretation of our results. For instance, defects in cell proliferation may lead to the generation of fewer EPDCs, and skew our migration measurement analysis. In addition, reactivation of epicardial cell proliferation is a key component of the cardiac injury response.

We performed immunohistochemistry for the proliferation marker phospho-Histone H3 (PHH3). The phosphorylation of histone H3 correlates with the condensation of genetic material during mitosis and meiosis (Ribalta et al., 2004). PHH3 has previously been used in isolation to analyse proliferation in the embryonic heart (Iyer et al., 2016; Chakraborty and Yutzey, 2012). PHH3 is universally accepted as a marker of cell proliferation. Initially, we independently examined the expression of PHH3 in epicardial cell cultures derived from control (+/+ or ∆/+) and mutant (∆/∆) hearts. We calculated the number of PHH3 positive cells as a proportion of total cells in 3 fields of view of each culture monolayer derived from a total of seven control and four mutant hearts. Control epicardial cells displayed positive PHH3 staining (Fig 4.8, A, B) at an average rate of 18.68% +/- 1.296% SEM. Comparably, NMIIB abolished epicardial cells displayed positive PHH3 staining (Fig 4.8, C, D) at an average rate of 16.83% +/- 0.6785% SEM. In addition, cells actively undergoing division were detectable in cultures from each genotype (Fig 4.8, E, F, arrowheads). This analysis revealed that there was no significant difference in the proliferation rates between control and mutant epicardial cells in vitro (Fig 4.8, G, p=0.3103, unpaired 2-tailed t-test).

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A ∆/+ B ∆/+

PHH3 PHH3 actin DAPI C ∆/∆ D ∆/∆

PHH3 PHH3 actin DAPI E ∆/+ G Comparing Average Proportion of PHH3 +ve Cells 25 ns

n 20

merge l 15

F ∆/∆ l

a 10

T 5

0 ∆Het/+ ∆∆/∆∆ merge Genotype

Figure 4.8: Analysis of Cell Proliferation in Epicardial Cell Cultures (A-B) Phospho-Histone H3 (PHH3) staining of day 3 control (∆/+) and (C-D) mutant (∆/∆) epicardial cell cultures. (E) Cells actively undergoing cell division are identifiable in both control and (F) mutant cultures (arrowheads). (G) Graph comparing the average proportion of PHH3 positive cells in control and mutant cultures. Average proliferation rate for controls = 18.68% (+/- 1.296%, n=7), and mutants = 16.83% (+/- 0.6785%, n=4) (p=0.3103, unpaired 2-tailed t-test). Scale bars = 100µm.

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We expanded this study by next investigating the expression of PHH3 in E14.5 control (EHC/+, n=3) and mutant (EHC/EHC, n=3) hearts (Fig 4.9, A-D). We calculated the number of PHH3 positive cells, identified by punctate staining (Fig 4.9, A-D), as a proportion of total cells present in the field of view for both the epicardium and myocardium. To simplify our analysis, and to compensate for the reduced cell density in the EHC myocardium, we analysed a 100µm square area of the myocardium where the cell density was comparable between control and mutant sections. We analysed 2 fields of view in each of 3 non-sequential cardiac sections per heart. A total of three hearts were used for each genotype, giving a total of 18 fields of view per genotype. This analysis showed that there was no significant difference in proliferation rate in the ventricular myocardium of control and mutant hearts (Fig 4.9, E, p=0.1684, unpaired 2-tailed t-test). However, we were surprised to discover that the rate of cell proliferation in the epicardium was significantly higher in the EHC mutant compared to controls (Fig 4.9, A –D, arrowheads, E, p=<0.0001, unpaired 2-tailed t-test). The control epicardium shows a proliferation rate of 4.953% +/- 0.9674% SEM, which is considerably lower than proliferation in the myocardium of 16.70% +/- 1.348% SEM. The EHC epicardium has a proliferation rate of 14.17% +/- 1.356% SEM, which is similar to that displayed by the EHC myocardium of 14.09% +/- 1.275% SEM. This indicates that the mutant epicardial cells fail to reduce their rate of proliferation in accordance with the reduction shown in control epicardial cells.

Once again, these in vivo observations contrast with our in vitro data. The proliferation of cultured epicardial cells, regardless of genotype, is roughly comparable to that of the embryonic myocardium during development. This suggests that additional factors restrict epicardial cell proliferation in vivo, which EHC epicardial cells do not appropriately respond to. The contrasting results between our in vivo and in vitro studies lead us to conclude that the behaviour of NMIIB mutant epicardial cells is highly dependent upon the substrate microenvironment on which the epicardial cells reside.

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Figure 4.9: Analysis of Epicardial Proliferation and EMT (A-B) PHH3 staining in coronal cryosections of control (EHC/+) and (C-D) mutant (EHC/EHC) E14.5 hearts. (E) Graph comparing average number of PHH3 positive cells as a proportion of total cells in both the epicardium and myocardium of control and mutant hearts. Average proliferation rate in the control epicardium = 4.953% (+/- 0.9674%, n=18), and mutant epicardium = 14.17% (+/- 1.356%, n=18) (p<0.0001, unpaired 2-tailed t-test). Average proliferation rate in the control myocardium = 16.70% (+/- 1.348%, n=18) and mutant myocardium = 14.17% (+/- 1.356%, n=18) (p= 1.684, unpaired 2-tailed t-test). (F-G) Localisation of the EMT marker Snail in the control and (H- I) mutant E14.5 heart. Snail positive epicardial cells are indicated by arrowheads. Scale bars = 100µm (A-D), 50µm (F-I).

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4.2.6 EHC Mutants Show Defects in Epicardial EMT Our finding that EHC mutant epicardial cells fail to reduce their proliferation rates is suggestive that another key aspect of epicardial behaviour may be compromised due to loss of NMIIB. Previous investigators have reported that cells exhibiting an increased rate of proliferation may consequently display dysfunctional EMT. Vega and colleagues (2004) have shown that various epithelial cell lineages reduce proliferation in response to EMT activation, and that an inverse correlation exists between proliferation and the expression of the EMT marker Snai1 (encodes Snail) during mouse embryonic development (Vega et al., 2004). Snai1 encodes Snail, a transcription factor expressed in the nucleus of cells in which EMT has been initiated and is considered a universal marker of EMT. Snai1 expression precedes the activation of transcriptional changes and cytoskeletal reorganization associated with the transition from an epithelial to mesenchymal morphology.

As previously discussed, the initiation of EMT in a subset of epicardial cells is an essential prerequisite to the generation of EPDCs. Failure to undergo EMT results in defects in cardiogenesis, with particular abnormalities in the formation of the coronary vasculature. In order to ascertain whether the increased proliferation rates observed in the EHC mutant epicardium correlates to EMT discrepancies, we assessed epicardial expression of Snail in E14.5 cardiac sections by performing immunohistochemistry for the Snail protein.

We were able to identify Snail positive epicardial cells in all control (EHC/+, n=7) hearts analysed (Fig 4.9, F, G). Moreover, where Snail positive epicardial staining was observed, we frequently saw staining of multiple, contiguous epicardial cells (Fig 4.9, F, G, arrowheads), giving the impression of ‘patches’ of the epicardium which simultaneously expressed Snail. In contrast, in the vast majority (85%) of the mutant (EHC/EHC, n=7) hearts analysed (6/7), we saw a dramatic reduction in the number of Snail positive epicardial cells (Fig 4.9, H, I). In addition, Snail staining appeared to be confined to single cells in the EHC epicardium, and we did not observe ‘patches’ of Snail positive epicardium (Fig 4.9, H, I, arrowheads).

This data indicates that the EHC epicardium shows evidence of EMT dysregulation. Reduced Snail staining, accompanied by a failure to reduce proliferation rate (as indicated by PHH3), lends to the conclusion that the EHC mutant epicardium shows a reduced incidence of EMT. Congruently, this suggests that NMIIB plays a key role in the activation

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of EMT in the epicardium. Failure to correctly activate EMT in a sufficient number of epicardial cells may underpin the abnormalities we observe downstream in the coronary vessel developmental process in the EHC mutant heart. Excitingly, a role for NMIIB in the initiation or execution of EMT has not previously been reported.

4.2.7 Investigating Mechanotransduction in Myh10 Ablated Epicardial Cells In light of the discovery that Myh10 ablated epicardial cells show defects in EMT activation, we sought to examine the mechanism underpinning this abnormality. Whilst some epicardial EMT activation factors have been described, the functional mediators of this process are poorly understood (Trembley et al., 2015). It has previously been shown that the myocardin-related transcription factors, MRTF-A and MRTF-B (encoded by Mkl1 and Mkl2 respectively) translocate to the epicardial nucleus in response to TGFβ signalling to activate motile gene expression during EMT and thus mediate the motility of EPDCs (Trembley et al., 2015). In addition, mice deficient for MRTF transcription factors display cardiac developmental abnormalities (Mokalled et al., 2015). Substrate mechanical tension plays a crucial role in the activation of EMT, and similarly has been shown to dictate the sub-cellular localisation of MRTF-A (O'Connor et al., 2015). Nuclear accumulation of MRTF-A is thought to be dependent upon the activation of the Rho-actin signalling pathway (Miralles et al., 2003). NMIIB is known to play a role in the transduction of mechano-signals from the cellular microenvironment (Vincente-Manzanares, et al., 2009; Conti and Adelstein, 2008). In light of this, we sought to establish whether loss of NMIIB affected the sub-cellular localisation, and thus function, of MRTF-A.

We performed immunohistochemistry for MRTF-A in day 3 epicardial culture monolayers. Unfortunately, we observed radical variance in MRTF-A sub-cellular localisation in epicardial cells, which did not correlate to genotype (Fig 4.10, A-D). Moreover, we frequently observed either exclusively nuclear (Fig 4.10, A, B), exclusively cytoplasmic/peri-nuclear (Fig 4.10, C, D), or a combination of each in epicardial cell cultures derived from the same embryonic heart sample (data not shown, n>10). Since conducting this experiment, it has subsequently transpired that other investigators have experienced similar technical difficulty with this antibody. Consequently, we have not been able to successfully interpret these data, as it appears that the anti-MRTF-A antibody demonstrates a high degree of non-specific antigen binding.

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Figure 4.10: Analysis of MRTF-A Localisation in Cultured Epicardial Cells (A-D) Day 3 epicardial cell cultures which had been serum starved for 24-hours were stained for MRTF-A. (A-B) Predominantly nuclear sub-cellular MRTF-A localisation in wild type (+/+) epicardial cell cultures. (C-D) Predominantly cytoplasmic and peri-nuclear sub-cellular MRTF-A localisation in different cell cultures derived from the same embryonic heart as depicted in panels A and B. This diversity in MRTF-A localisation was observed in cultures obtained from both heterozygous (∆/+) and homozygous (∆/∆) embryonic hearts. The MRTF-A staining profile did not correlate to either genotype, or treatment type (with/without serum starvation) (data not shown). Scale bars = 100µm.

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4.3 Discussion We have shown, through a combination of immunofluorescence microscopy and western blotting, that NMIIB is abundant throughout the embryonic heart, and is the predominant NMII isoform in the epicardium. Moreover, NMIIB occupies a distinct sub-cellular localisation in cultured epicardial cells. In Myh10 ablated hearts, the expression of the other NMII isoforms does not increase, and their sub-cellular localisation in epicardial cells does not change. We have established that Myh10 knock out hearts show a reduction in EPDC migration, but do not show any significant motility defects in vitro. Similarly, whilst cultured Myh10 null epicardial cells do not show proliferation abnormalities, the EHC epicardium has significantly higher proliferation rates compared to controls in vivo. Accordingly, the EHC epicardium shows a reduction in Snail protein expression, suggesting dysregulation of epicardial EMT. Whilst the potential of mutant EPDC to differentiate into vSMC and cardiac fibroblasts does not appear to be compromised, the aforementioned dysregulation of other aspects of epicardial biology indicate that the gross morphological defects observed in the EHC heart phenotype are underpinned by epicardial dysfunction.

Epicardial dysfunction has not previously been described for the Myh10 ablated mouse. The indication that the EHC phenotype may be underpinned by potential epicardial defects prompted us to examine the expression of NMIIB in the embryonic heart. Here, we show for the first time that NMIIB represents the predominant NMII isoform in the epicardium during a developmental window that corresponds to peak epicardial activity. Together with our observation that NMIIB occupies a distinct sub-cellular localisation to NMIIA in cultured epicardial cells, these findings suggest that NMIIB plays a crucial and novel role in epicardial function, which is not compensated by the other NMII isoforms in Myh10 null epicardial cells. Indeed, NMIIA has been found to actively suppress epithelial cell invasion of 3D substrates, suggesting that NMIIA would be unable to rescue defective EHC EPDC migration (Babbin et al., 2009). This indicates that it is not total NMII protein content of the epicardium, but the specific NMII isoform that is responsible for correct epicardial function.

Moreover, we have established that in response to the loss of NMIIB in Myh10 ablated hearts, the relative expression or localisation profile of the other NMII isoforms does not increase. Indeed, we observed a slight reduction in immunoreactivity for NMIIA in Myh10 null hearts via western blotting (see Appendix 11). It has recently been shown that force- mediated chromatin stretching can modulate transcription (Tajik et al., 2016). The

145 prominent role of NMIIB in the generation of tension forces suggests possible alterations to transcription regulation in the EHC embryo.

As correct epicardial cell function is critically required for the formation of the coronary network, we sought to examine which aspects of epicardial cell function are defective in EHC mutants to provide a possible explanation as to when the programme of coronary vessel formation is disturbed.

The correct migration of EPDC is crucial for the development of the coronary vasculature (Olivey and Svensson, 2010, Perez-Pomares and de la Pompa, 2011). These cells express Wt1, one of a limited number of molecular markers for the epicardium and EPDCs, and which is essential for epicardial EMT activation (von Gise et al., 2011; von Gise and Pu, 2012). We reveal here for the first time that Wt1 positive cells display a disrupted localisation profile in EHC mutant hearts. We identified abnormal ‘clusters’ of Wt1 positive cells in close proximity to the subepicardial space. Diminished EPDC migration has previously been shown in studies following disruption to the epicardium (Vega- Hernandez et al., 2011) and strongly correlates to defective coronary vessel development (Zamora et al., 2007; Rhee et al., 2009). Abnormally restricted EPDC migration is therefore highly likely to underpin abnormal vessel development in the EHC heart.

In addition, failure to infiltrate the myocardium to sufficient depths and in sufficient numbers may contribute to the myocardial defects observed in the EHC phenotype. EPDC are known to secrete paracrine signals to nurture myocardial proliferation (Olivey and Svensson, 2010; Perez-Pomares and de la Pompa, 2011). Correct myocardial responses to these signals are likely to be compromised by inadequate EPDC migration, as speculated by Mahtab et al., (2008). Interestingly, we found that cardiac fibroblasts are present in the EHC heart. This suggests that NMIIB is not required for the differentiation of EPDCs into fibroblasts; as this process is known to require FGF signalling (Vega-Hernandez et al., 2011), we tentatively suggest that NMIIB is not involved in the mediation of FGF signalling. Conversely, it has been shown that the cardiac fibroblast population derives from multiple tissue sources during development (Zeisberg and Kalluri, 2010); it is possible that the cardiac fibroblasts observed in the EHC heart are derived from a non-epicardial source, and that loss of NMIIB has a detrimental impact on the differentiation of EDPCs into the fibroblast lineage. We did not conduct analysis to determine if the number of cardiac fibroblasts was reduced in the EHC heart. Further studies are required to clarify

146 whether or not the proportion of cardiac fibroblasts is altered in the EHC mutant, and whether or not these cells do indeed derive from the epicardium.

The loss of NMIIB function has been demonstrated to affect cell adhesion, migration, and the ability of cells to respond to external stimuli (Vincente-Manzanares et al., 2009; Conti and Adelstein, 2008; Lo et al., 2004; Phillips et al., 2005; Ma et al., 2009). In light of these observations, the finding that EHC EPDCs exhibit migration defects is understandable, and perhaps even expected. However, our finding that migration is not completely abolished in vivo, suggests that NMIIB may function in an additional capacity to facilitate EPDC migration. Indeed, it has been shown that the establishment of cell polarity and the correct regulation of cell adhesions are crucial mediators of EPDC invasion (Rhee et al., 2009; Zamora et al., 2007). As NMIIB is known to play a prominent role in these cellular processes, further examination of the expression and localisation of polarity and adhesion molecules is required to delineate the role of NMIIB in EPDC invasion.

The basis for this speculation is augmented by our findings that Myh10 null epicardial cells do not display migration defects in vitro, indicating that additional as yet unknown factors influence defective EHC EPDC migration in vivo. As an aside, our studies of NMIIB ablated cell migration in vitro conflict with the findings of others (Lo et al., 2004; Betapudi et al., 2006), which may suggest that NMIIB function in migrating cells is highly cell- specific. Interestingly, defective EPDC migration has been previously described in studies in which non-migration aspects of epicardial cell function are directly compromised (Moore et al., 1999; Mellgren et al., 2008; Tevosian et al., 2000). These observations led us to hypothesise that the reduced EPDC migration phenotype in EHC hearts results from upstream epicardial cell dysfunction.

By examining epicardial processes upstream of EPDC migration, we have been able to establish additional discrepancies in the EHC heart. Mutant epicardial cells display elevated proliferation rates, which are associated with a reduction in Snail staining; this indicates a reduced incidence of epicardial EMT activation. Snail expression has been proposed to be directly regulated by Wt1 (Martinez-Estrada et al., 2010), and is known to promote EMT through repression of the adhesion molecule E-cadherin, permitting cellular delamination from the epithelium and the induction of dramatic phenotypic alterations (Thiery et al., 2009). In addition, Vega and colleagues (2004) have shown that Snail impairs cell cycle progression in various mouse epithelial cells, indicating an inverse correlation between cell proliferation and the activation of EMT morphological changes (Vega et al., 2004). In

147 conjunction with this, we have shown that the EHC epicardium shows diminished EMT activation, implying that NMIIB plays a critical role in the processes required for EMT signalling in these cells.

Defects in EMT activation are surprising; as an important component of the cytoskeletal machinery, NMIIB would be expected to act downstream of EMT during actin-myosin mediated cell motility (Conti and Adelstein, 2008; Vincente-Manzanares et al., 2009). Similarly, it is remarkable that EMT activation appears to be affected exclusively in the epicardium, as other EMT dependent processes (e.g. gastrulation, craniofacial development) occur correctly in the EHC embryos. Additionally, formation of the atrioventricular valves, a process dependent upon endothelial cushion EMT (von Gise and Pu, 2012) is apparent in EHC mutant hearts (data not shown). This suggests that NMIIB plays a novel function in the epicardium that predisposes epicardial cells to defects following Myh10 ablation.

It is plausible that this disruption is related to the role of NMIIB in signal transduction. Previous work has established that expression of Bves (blood vessel epicardial substance) is regulated by EGFR (epidermal growth factor receptor) signalling (Lin et al., 2007). Bves has been shown to regulate migration and EMT in epicardial cells via the modulation of fibronectin recycling (Benesh et al., 2013). Interestingly, NMIIB has been shown to function in the internalisation of EGFR and the subsequent activation of downstream signalling events (Kim et al., 2012). It is therefore possible that NMIIB mediates epicardial cell migration and ultimately function via the EGFR/Bves signalling pathway. Expanding upon this, it has previously been shown that PDGF signalling is essential for epicardial EMT and the development of vSMC from EPDCs (Smith et al., 2011; Rudat et al., 2013); it is therefore possible that NMIIB functions during PDGF signalling in the epicardium. In addition, recent data has shown that Yap and Taz, mediators of the Hippo signalling pathway, are required specifically within the epicardium to mediate coronary vessel formation (Singh et al., 2016). Disruption to these genes results in the dysregulation of epicardial EMT, and impairs formation of the coronary vasculature, as observed in the EHC mutant heart. NMIIB may therefore function in Hippo signalling transduction. Interestingly, canonical Wnt, Hedgehog and FGF signalling in the epicardium has been recently shown to be dispensable for correct epicardial and EPDC formation (Rudat et al., 2013); this implies that NMIIB does not function in these signalling pathways.

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It is interesting to note that epicardial EMT has been reported to occur independent of Snail expression (Casanova et al., 2013). The study utilised Wt1- and Tbx18-Cre recombinase mouse lines to drive the deletion of Snai1 in epicardial cells, and observed no detrimental phenotype in resultant mice (Casanova et al., 2013). However, there are serious technical limitations associated with these Cre drivers that may restrict interpretations of this finding. In addition, we do not feel that this report detracts from our finding that NMIIB contributes to epicardial EMT. The identification of an albeit reduced population of Snail positive epicardial cells, accompanied by the presence of a limited number of EPDC deep within the EHC myocardium, indicates that a reduced level of EMT does occur in the mutant epicardium. As the epicardium is known to be a heterogeneous cell population (Katz et al., 2012), it may be that a sub-set of epicardial cells do not require Snai1 to initiate EMT. Additionally, we cannot rule out that Snail paralogues may compensate for loss of Snail to induce EMT. Further experimentation is required to definitively address these possibilities. Based on our current data, we suggest that initiation of epicardial EMT requires a finely balanced response to a threshold of pro-EMT signals. NMIIB functions in an important, but not essential, role in initiating EMT in response to these signals.

Our finding that cultured Myh10 ablated epicardial cells do not show proliferation or migration abnormalities - as demonstrated by their EHC in vivo counterparts - is particularly interesting. In addition, epicardial cells from embryonic heart explants have been previously shown to express Snai1 (Takeichi et al., 2013), suggesting that epicardial cell migration in vitro involves EMT activation. These conflicting findings suggest that the behaviour of Myh10 null epicardial cells is dependent upon their in situ extracellular environment; the in vitro microenvironment recapitulates many aspects of normal epicardial cell behaviour that are not permitted in the context of the EHC embryo.

A pivotal difference between the microenvironment in these two systems is the constitution of the extracellular matrix. The sub-epicardial matrix is known to play a key role in normal epicardial function (Riley and Smart, 2011; Olivey and Svensson, 2010; von Gise and Pu, 2012; Perez-Pomares and de la Pompa, 2011). Several studies have directly shown that disruption to the sub-epicardial matrix results in cardiac defects and disrupted heart regeneration in response to injury (Missinato et al., 2015; Lockhart et al., 2011). NMIIB has been shown to play a role in the secretion of extracellular matrix components, including collagen (Kalson et al., 2013) and fibronectin (Lin et al., 2007; Benesh et al., 2013). Accordingly, the EHC heart shows disruption in the thickness and composition of

149 the sub-epicardial extracellular matrix (data not shown, discussed in Chapter 6). This suggests a direct role for NMIIB in the synthesis of this biologically important substrate; disruption of which may lead to abnormal epicardial cell function.

Furthermore, the sub-epicardial matrix generates substrate tension, which has previously been implicated in the regulation of epicardial EMT activation (O’Connor et al., 2015; Trembley et al., 2015). It has been shown that cells respond to their mechanical environment through the regulation of the three NMII isoforms (Raab et al., 2012; Vicente-Manzanares et al., 2009). Additionally, NMIIB is known to accumulate at regions of the cell where mechanical stress is exogenously applied (Schiffhauer et al., 2016). In addition, chemical inhibition of myosin II phosphorylation and therefore motor/actin binding activity has been shown to reduce the accumulation of other mechano-sensitive elements (Schiffhauer et al., 2016). This suggests that the mechano-accumulation response in at least in part orchestrated by a myosin II dependent pathway. Moreover, NMIIB has been shown to exhibit a highly cell-type and cell-cycle-specific responsive behaviour to mechanical stresses, which are not mirrored by NMIIA and NMIIC (Schiffhauer et al., 2016). The correct maintenance of mechano-tension is essential for the correct establishment of cell polarity, migration and division, and the established role of NMIIB in force generation may underpin the compromised ability of EHC epicardial cells to activate EMT signalling (Norstrom et al., 2010).

Speculatively, it has previously been shown that mechanical forces can be translocated into biochemical signals associated with EMT (cell differentiation and ECM remodelling) through the phosphorylation of focal adhesion kinase (FAK) (Lee and Nelson, 2012). Integrin mediated binding to the ECM induces FAK phosphorylation, which in turn activates Rho kinase mediated actomyosin contractility (Lee and Nelson, 2012). Activated Rho kinase is required for epicardial EMT and the generation of EPDCs (Artamonov et al., 2015). Moreover, cell adhesion to fibronectin has been shown to activate both PDGF and FAK activity in an α5β1 integrin-dependent manner to regulate mesenchymal cell migration (Veevers-Lowe et al., 2011). As both PDGF and α5β1 are strongly implicated in correct epicardial cell functionality, NMIIB may therefore function to link alterations in epicardial cell contractility with Snail expression and EMT activation; conversely, loss of NMIIB may affect ECM deposition, and therefore directly affect the mechanism of FAK phosphorylation and the induction of downstream signalling cascades to promote EMT gene expression.

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Although we were unable to interpret our data from the MRTF-A localisation experiments, it would be extremely interesting to repeat this immunocytochemical analysis with a reliable antibody. The translocation of MRTF-A to the nucleus is known to be involved in the initiation of epicardial EMT signalling (Trembley et al., 2015). In addition, cardiac defects have been reported following the targeted deletion of MRTFs (Mokalled et al., 2015), suggesting a crucial role for these transcription factors in normal cardiogenesis. Interestingly, the sub-cellular localisation of MRTF-A is dictated by substrate tension, and increased matrix rigidity promotes TGFβ induced EMT (O’Connor et al., 2015). Interestingly, in vitro studies that restrict cell shape have been found to limit the expression of EMT genes associated with TGFβ-MRTF signalling (O'Connor and Gomez, 2013). This may offer an explanation as to decreased instance of EMT in the EHC embryo, and suggests that epicardial substrate in vivo may detrimentally affect cell shape and EMT initiation. Establishing whether or not MRTF-A localisation is disrupted in NMIIB ablated epicardial cells would clarify the role of NMIIB in tension generation and mechano- signalling in the epicardium, and offer an exciting insight into the link between matrix adhesion, cell morphology and EMT regulation in the epicardium.

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4.4 Conclusion To conclude, we have shown that NMIIB plays a crucial role in both the initiation of epicardial EMT, and the subsequent migration of EPDC. These data indicate that the EHC coronary vessel defects are underpinned by compromised epicardial cell function. Epicardial NMIIB function is not rescued by the other NMII isoforms, consistent with the findings of others that suggest NMIIB plays distinct and varying functions in different tissues (Bao et al., 2007; Ma et al., 2007; Ma et al., 2009). Ma and Adelstein (2012) have proposed that cellular processes dependent upon the cross-linking function of NMII are more amenable to isoform substitution (Ma and Adelstein, 2012), indicating that the functionality of NMIIB in the epicardium may be more dependent upon the activity of the NMIIB motor domain. Our data suggests that loss of NMIIB causes disruption to the upstream signalling events required for EMT activation, which may be underpinned by the pleiotropic functions of NMIIB in extracellular matrix secretion (Kalson et al., 2013), tension modulation (Norstrom et al., 2010; Schiffauer et al., 2016) and receptor internalisation (Kim et al., 2012). Identification of the mechanisms responsible for epicardial dysfunction in the EHC mouse may inform therapeutic strategies to reactivate embryonic epicardial cell function in the diseased heart, and facilitate the regeneration and repair of injured cardiac tissue.

However, the complex task of delineating the precise function(s) of NMIIB during cardiogenesis is compounded by the ubiquitous expression of NMIIB throughout the developing heart, accompanied by the cumulative defects in both the epicardium and myocardium in global Myh10 knock out animals. This dictates a requirement to generate tissue-specific Myh10 knock out animals to elucidate the overriding cause of the EHC phenotype and the novel function fulfilled by NMIIB during mammalian heart formation.

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Chapter 5: Generation of Myh10 Tissue-Specific Knock Out Animals

5.1 Introduction A prominent problem with classical gene knock out studies is the generation of embryonic lethal mutants, which prevents the analysis of gene function at late gestation, post-natal and adult stages (Sung et al., 2012). The adoption of strategies to generate conditional knock out animals that can control gene deletion in a spatial and/or temporal manner offers a means of circumventing these challenges (Matthaei, 2007). In addition, conditional knock out approaches to generate mouse models of human diseases that affect specific tissues may provide a more realistic phenotype that better recapitulates the human condition. Therefore, generation of conditional gene knock out animals and the analysis of subsequent phenotypic changes is central to enable our full appreciation of gene function in vivo, and thus further our understanding of human disease (Sung et al., 2012).

A varied range of genomic editing technologies exist, including; zinc-finger nucleases, TALEN (transcription activator-like effector nucleases) and CRISPR (clustered regularly interspaced short palindromic repeats)/Cas9 (CRISPR associated protein 9), with the latter seemingly superseding all other technologies in recent years. However, this thesis research project has utilised the Cre-loxP system, and this introduction will therefore focus on this technology.

5.1.1 Cre Recombinase/loxP Genomic Editing Technology Genomic editing technologies have evolved into a crucial tool for the developmental biologist to determine the tissue-specific requirements and functions of genes during embryogenesis. Site-specific recombinase technology permits sequence deletion, insertion, translocation and inversion at defined sites within the genome. Currently, the most widely used site-specific DNA recombinase system is Cre-recombinase (causes recombination)/loxP (locus of cross-over (x) in P1) (subsequently referred to as Cre-loxP) (van der Weyden et al., 2002). The Cre-loxP system is composed of a single enzyme and short recombination sequences derived from bacteriophage P1 (Hoess and Abremski, 1984). The Cre recombinase enzyme, a member of the DNA recombinase family, consists of a 130 residue N-terminal domain and a 211 residue C-terminal domain (Van Duyne, 2015). Cre recombinase catalyses site-specific recombination by recognising and binding to

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34-bp loxP sequences, which can be integrated into the target host DNA sequence. The loxP site is comprised of two symmetrical recombinase-binding elements of 13-bp, separated by an 8-bp core region (Fig 5.1, A) (Van Duyne, 2015). Two Cre-recombinase monomers bind cooperatively to each loxP site (one per recombinase-binding element), resulting in a Cre-dimer at each recombination site (Van Duyne, 2015). Two Cre-bound loxP sites then associate to form an anti-parallel synaptic complex, in which strand cleavage is catalysed by the Tyr324 residue of Cre. Exchange of strands between the two loxP sites forms a Holliday junction intermediate, which initiates the second cleavage event of the second DNA strand. Subsequent strand exchange completes the recombination reaction (Van Duyne, 2015).

In essence, the Cre-loxP system cleaves DNA at a distinct target sequence and ligates it to a second identical site to generate a contiguous strand via high efficiency homologous recombination (van der Weyden et al., 2002). Cre-loxP recombination can therefore be manipulated to induce the deletion of sequences from the genome. The 6-bp core region of the loxP site dictate directionality of the reaction; cis loxP sites orientated in the same direction leads to the excision of the intervening DNA sequence (Fig 5.1, B), whilst inverted cis loxP sites results in the inversion of the intervening DNA sequence (Fig 5.1, C) (van der Weyden et al., 2002). In light of this, one can inactivate gene expression (loss-of function) by inserting directionally identical loxP sites to flank a coding DNA sequence (usually one or several exons) of interest (Soriano, 1999). The allele created is termed “floxed” (flanked by loxP). The floxed allele is inserted into the correct location by homologous recombination into mouse embryonic stem cells, injected into mouse blastocysts and animals that carry the floxed allele of interest subsequently generated (Matthaei, 2007). The floxed sequence is fully functional until it is excised following activation of Cre expression. This occurs in animals derived from crossing the floxed line to a transgenic Cre line, in which Cre expression is usually driven by a tissue-specific promoter (Matthaei, 2007).

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A 8-bp core

5’-ATAACTTCGTATA-ATGTATGC-TATACGAAGTTAT-3’

13-bp RBE 13-bp RBE

loxP loxP + Cre

loxP C

loxP loxP + Cre

loxP loxP Figure 5.1: Schematic Representation of Cre-loxP Mediated Genomic Editing (A) The loxP recombination sequence consists of two symmetrical 13-bp recombinase binding elements (black arrows). These sites are separated by an 8-bp core sequence (block arrow). The core region dictates the directionality of the loxP sequence. (B) When two loxP sites (grey arrowheads) are integrated into the genome in the same orientation, Cre- mediated recombination results in the excision of the intermittent sequence (indicated by the yellow and red region). (C) When loxP sites are orientated in opposite directions, Cre- mediated recombination results in the inversion of the intermittent sequence (indicated by the inversion of the yellow and red regions). RBE – recombinase binding elements. Adapted from van der Weyden et al., 2002.

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5.1.2 Conditional Modulation of Cre Recombinase Expression Controlling Cre recombinase expression by placing it under the control of a tissue-specific promoter allows for the modulation of gene deletion in a temporal/spatial manner (Matthaei, 2007). Since this strategy was first reported by Gu and colleagues (1994), a plethora of tissue-specific Cre expressing transgenic mouse lines have been generated (Gu et al., 1994). Further control of Cre recombinase expression has been permitted by the introduction of inducible promoters, which remain inactive (i.e. no Cre expression) until being “switched on” following the administration of an exogenous chemical inducer (Matthaei, 2007). One such example of this adapted system is the Cre-ERT2 allele. In this instance, Cre recombinase is fused to a modified human estrogen receptor (ERT2), which recognises the ligand tamoxifen and its 4-hydroxytamoxifen derivative (van der Weyden, 2002). Administration of tamoxifen induces the translocation of the Cre-ERT2 fusion protein from the cytoplasm to the nucleus, upon which excision of the floxed sequence is initiated (Indra et al., 1999; Li et al., 2000). Whilst this method permits greater restriction of Cre activity in space and time, complications relating to the administration of these chemical inducers have been reported, including embryo abortion and behavioral changes (Doetschman and Azhar, 2012). These caveats must be considered in the interpretation of such studies (Matthaei, 2007).

The simplicity and broad utility of the Cre-loxP system has established it as the preferred method of genetic editing technique in the mouse over recent years; it can function on a variety of DNA substrates and over mega-base distances. In addition, the short loxP recombination sequence does not interfere with gene expression when inserted into chromosomal DNA. Furthermore, Cre can mediate transmissible genetic modifications by functioning in mammalian germ line cell (van der Weyden, 2002). For these reasons, a vast number of Cre expressing mouse lines have been generated, and they have served a vital role in the elucidation of mammalian gene function.

However, increased use of the Cre-loxP system has unveiled potential caveats to the interpretation of mouse models of human disease generated by this technique. High levels of Cre expression have been associated with chromosomal aberrations due to cryptic loxP sites within the mammalian genome (van der Weyden, 2002; Doetschman and Azhar, 2012). Moreover, Cre expression has been shown to impair cell proliferation and is implicated in arrest of the cell cycle (Adams and van der Weyden, 2001; Naiche and Papaioannou, 2007; Loonstra et al., 2001). In addition, a major complication with Cre-loxP

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arises from ‘leaky’ Cre expression, where Cre is non-specifically expressed in other cells and tissues (Sung et al., 2012). Alternatively, Cre may not be expressed sufficiently under a given tissue-specific promoter, therefore diminishing the efficiency of recombination of the floxed allele (Matthaei, 2007; Sung et al., 2012). These problems are especially pertinent in transgenic lines where the expression of Cre does not recapitulate that of the driver gene due to the exclusion of important elements of the endogenous promoter region (e.g. enhancers). This issue can be overcome in part by the use of knock in rather than transgenic approaches (Jin et al., 2000).

5.1.3 Detection and Tracking Cre Recombinase Expression Expression of Cre recombinase at the cellular level can be identified and tracked by crossing Cre expressing animals to so-called reporter lines. Monitoring is necessary to ensure correct temporal and spatial restriction of Cre expression. One such commonly used reporter line is the ROSA26 reporter (R26R) (Soriano, 1999). The ROSA26 locus is situated on mouse chromosome 6 (Srinivas et al., 2001) is ubiquitously expressed (Scearce- Levie et al., 2001) and constitutively active during mouse embryonic development (Friedrich and Soriano, 1991). The R26R line contains a lacZ knock in at the ROSA26 locus, which encodes the bacterial enzyme β-galactosidase. A floxed stop codon/neomycin-resistance cassette is inserted upstream of the LacZ gene (Fig 5.2, A). In cells that express Cre, the stop codon is excised by homologous recombination, permitting lacZ transcription and the production of β-galactosidase exclusively in these cells (Fig 5.2, B). One can subsequently detect β-galactosidase expression by X-gal staining, which is hydrolysed by β-galactosidase to generate a blue coloured staining product (Soriano, 1999).

Similarly, a broad range of ROSA26 reporter strains have now been developed which utilise the visualization properties of bioluminescent and biofluorescent molecules, such as eYFP (enhanced yellow fluorescent protein) and eCFP (enhanced cyan fluorescent protein) (Srinivas et al., 2001). A comparison of the ROSA26 reporter strains commercially available can be found on the Jackson Laboratory website (available at https://www.jax.org/research-and-faculty/tools/cre-repository/comparison-of-cre- reporters). These reporter lines have been employed by numerous studies in conjunction with cell-specific Cre drivers to fate map the cardiac progenitor cell populations in the mammalian heart (Chong et al., 2014). A broad variety of cardiac cell specific Cre driver lines are available (reviewed in Doetschman and Azhar, 2012).

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Figure 5.2: Schematic Representation of the ROSA26 Reporter Mouse Line (A) The ROSA26R mouse carries the lacZ gene (blue region), which encodes the bacterial enzyme β-galactosidase, under the control of the ROSA26 promoter (green box). Two loxP sites (grey arrowheads) orientated in the same direction flank a STOP sequence (red octagon) upstream of the lacZ insertion, which prevents lacZ transcription in normal conditions. (B) In cells that express Cre recombinase, recombination excises the stop sequence, allowing the transcription of lacZ. Detection of lacZ expression in cells and tissues is achieved by X-gal staining (see main text for details).

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5.1.4 Epicardial-Specific Cre Drivers As previously mentioned, embryonic epicardial cells are identified by expression of Wt1, Tbx18, Raldh2 and Tcf21 (Masters and Riley, 2014). A number of Cre driver mouse lines are available that have been shown to express Cre in the epicardium and EPDCs (Doetschman and Azhar, 2012). These include; Gata5-Cre (Merki et al., 2005), Tbx18-Cre (Cai et al., 2008), Wt1-GFP/Cre (Zhou et al., 2008). However, due to the technical limitations of these lines with regard to ‘leaky’ Cre expression, and recent claims that other cardiac cell populations may transiently express these markers, extreme caution must be excised when interpreting data generated from these lines. A further consideration to appreciate is that the recombination efficiency and effects of genetic background may strongly influence the findings of experiments utilising these lines (Zhang et al., 2013b). In addition, the identification of heterogenicity in the epicardial cell population (Katz et al., 2012) compounds these complications. In light of this, it has been argued that the Wt1-CreERT2 and Tcf21-CreERT2 driver lines are the only well characterised, and therefore, most reliable lines to regulate Cre expression in a spatial and temporal manner in the epicardium at the present time (Doetschman and Azhar, 2012).

5.1.5 Myocardial-Specific Knock Out Mice The α-MHC (α-myosin heavy chain) promoter is the most commonly selected promoter to induce myocardium-specific gene deletions and has been implemented to control myocardial deletions in both a spatial and temporal manner (Agah et al., 1997; Heine et al., 2005; Doetschman and Azhar, 2012). The B6.FVBTg(Myh6-cre)2182Mds/J mouse line (subsequently referred to as α-MHC-Cre) has been used previously to drive Cre recombinase mediated recombination in cardiomyocytes from E9.5, with high levels of Cre expression detected at E11.5 (Gaussin et al., 2002; McFadden et al., 2005; Agah et al., 1997). This line carries a Cre recombinase knock in at the α-MHC, also referred to as Myh6, locus. This promoter induces greater than 90% recombination in cardiomyocytes (Agah et al., 1997) and is considered the standard Cre line to generate cardiomyocyte specific knock out mouse strains (Heine et al., 2005). However, it has also been reported that the α-MHC-Cre promoter becomes more broadly transcriptionally active in the adult mouse (Agah et al., 1997). In addition, recent findings have suggested that prolonged Cre exposure has cytotoxic implications (Schmidt-Supprian and Rajewsky, 2007), specifically in the 8-12 month adult heart (Doetschman and Azhar, 2012). This indicates that studies evaluating α- MHC-Cre mediated gene deletions in the adult mouse must be cautious when interpreting their results, and ensure appropriate controls are established.

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5.1.6 The Myh10 Myocardial-Specific Knock Out Mouse As previously discussed, Ma and colleagues have reported the generation of a cardiomyocyte-specific Myh10 knock out mouse line by crossing the α-MHC-Cre line to a Myh10 exon 2 floxed allele (Fig 5.3, B) (Ma et al., 2009). This study reports that loss of NMIIB exclusively from cardiomyocytes circumvents the embryonic lethality defect presented by global Myh10 knock out animals (Tullio et al., 1997; Takeda et al., 2003). However, mutants are born with enlarged, multinucleated cardiac myocytes and thin ventricular septum, recapitulating elements of the global Myh10 ablated phenotype. Moreover, cardiomyocyte Myh10 deficient animals developed a cardiomyopathy between 6- 10 months, with interstitial fibrosis and inflammatory cell invasion of the myocardium (Ma et al., 2009). The authors report the loss of NMIIB from cardiomyocytes at E13.5 by immunohistochemistry and western blotting (Ma et al., 2009). Interestingly, myocardial- specific NMIIB hearts do not display DORV, and have a greatly reduced incidence of VSD. In addition, the survival of these animals into adulthood suggests that coronary vessel formation must not be severely compromised. These findings are particularly pertinent to this thesis research, as it indicates that the severe cardiac defects displayed by the EHC mouse, and other globally ablated Myh10 mutants, is underpinned by loss of NMIIB from a non-cardiomyocyte cell population.

As Ma and colleagues did not report an analysis of the epicardium in their study, we sought to generate cardiomyocyte-specific Myh10 knock out mice on our own colony background to investigate coronary vessel development, as background strain differences have been shown to affect α-MHC-Cre expression (Agah et al., 1997).

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5.2 Results

5.2.1 Generation of Myocardial-Specific Myh10 Knock Out Animals In order to generate myocardial-specific Myh10 knock out animals, we adopted a Cre recombinase/loxP genomic editing approach to delete the Myh10 exon 2 sequences in cardiomyocytes. We employed the α-MHC-Cre mouse (available from the Jackson Laboratory under the name B6.FVBTg(Myh6-cre)2182Mds/J) (Fig 5.3, A), which expresses Cre exclusively in cardiomyocytes from approximately E9.5 in embryonic development, together with the Myh10flox/flox line, in which exon 2 of Myh10 is floxed (Fig 5.3, B).

5.2.2 α-MHC-Cre Breeding Scheme Our α-MHC-Cre founder male was a kind gift from Dr. Elizabeth Cartwright (University of Manchester). The α-MHC-Cre male was crossed to Myh10flox/flox female mice and the resultant progeny were weaned and genotyped (Fig 5.3, C). Male progeny carrying the Cre recombinase transgene and heterozygous for the floxed Myh10 allele (α-MHC-Cre; flox/+) were selected for subsequent breeding to Myh10flox/flox female mice to generate α-MHC- Cre/+; flox/flox embryos (Fig 5.3, C, boxed P2 progeny). Theoretically, these embryos should carry a homozygous Myh10 deletion in cardiomyocytes from E9.5 onwards. Progeny from this P2 generation were genotyped and subsequently analysed for a cardiovascular phenotype.

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Figure 5.3: Breeding Scheme for the Generation of Cardiomyocyte-Specific Myh10 Knock Out Animals - the α-MHC-Cre Mouse

(A) The α-MHC-Cre mouse contains a Cre recombinase knock in (light blue region) under the activity of the α-MHC promoter (orange region). The α-MHC-Cre mouse is heterozygous for this allele. (B) The Myh10flox/flox mouse carries two loxP sites (grey arrowheads), orientated in the same direction, that flank the Myh10 exon 2 sequence (purple region). The Myh10 exon 2 sequence is deleted in cells which express Cre recombinase. The Myh10flox/flox mouse is homozygous for this floxed allele. (C) Breeding scheme used to generate myocardial-specific Myh10 knock out mice (boxed progeny in P2). α-MHC – alpha myosin heavy chain, P – progeny.

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5.2.3 Confirmation that the α-MHC-Cre Mouse Expresses Cre Recombinase in the Ventricular Myocardium Although the α-MHC-Cre mouse is considered a reliable cardiomyocyte specific Cre line, we sought to confirm that Cre expression was detectable and limited to the Myh6 expressing myocardium in the embryonic heart. In order to achieve this, we bred the α- MHC-Cre founder male to ROSA26 reporter females. From the α-MHC-Cre to R26R cross, we isolated E14.5 embryonic hearts and performed X-gal staining (Fig 5.4, A-C). The left ventricle was severed to allow visualization of the internal ventricular tissue (Fig 5.4, B). The generation of a blue staining product was clearly observable in the ventricular myocardium of Cre positive hearts (as determined by genotyping) (Fig 5.4, C, n=4). Staining also appeared to be present in the left atria (Fig 5.4, C). Importantly, it appeared that staining was absent from the ventricular surface and great vessels, suggesting that Cre expression was limited to the cardiomyocyte population. We observed no X-gal staining in Cre negative samples (data not shown, n=9). Unfortunately, staining development required a 72-hour incubation period at room temperature. As a consequence, we observed loss of heart tissue integrity as the samples began to degrade. However, this experiment crucially shows that the expression of Cre recombinase in the α-MHC-Cre mouse is not only detectable, but seemingly restricted to the myocardial tissue in the embryonic heart.

5.2.4 Confirmation that α-MHC-Cre; flox/flox Animals Show Reduced Cardiac NMIIB Protein In order to confirm that ventricular Cre recombinase expression resulted in a reduction in NMIIB protein, we performed western blotting and immunohistochemistry experiments to compare the relative abundance of NMIIB in control (+/+; flox/flox, n=3) and myocardial knock out (α-MHC-Cre/+; flox/flox, n=3) hearts (Fig 5.4, D-J). Probing for NMIIB in adult heart protein extracts appeared to show a reduction in NMIIB signal intensity in α-MHC-Cre/+; flox/flox samples (Fig 5.4, D, n=3). Analysis of peak intensity using Image Lab Software (Bio-Rad) revealed that whilst all samples showed approximately equal signal for the β-actin loading control (Fig 5.4, E), α-MHC-Cre/+; flox/flox samples showed a reduction in NMIIB peak intensity (Fig 5.4, F, arrows). In addition, immunohistochemical staining for NMIIB in the interventricular septum of E14.5 hearts revealed a reduction in NMIIB staining in α-MHC-Cre/+; flox/flox samples (Fig 5.4, G-J). Together, this analysis indicates that expression of Cre in α-MHC-Cre/+; flox/flox samples causes a detectable decrease in NMIIB protein.

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Figure 5.4: Analysis of 'First Generation' Cardiomyocyte-Specific Myh10 Knock Out Animals

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Figure 5.4: Analysis of 'First Generation' Cardiomyocyte-Specific Myh10 Knock Out Animals (A) The α-MHC-Cre founder male was crossed to a ROSA26 reporter female, and embryos were harvested at E14.5. Embryos positive for the Cre knock in (as determined by genotyping) displayed a normal cardiac phenotype, with correct positioning of the aorta and pulmonary trunk. The left ventricle was excised to permit infiltration of the X-gal staining solution (straight dashed line). (B) Schematic showing the rotation of the heart pieces for orientation in panel C. (C) Hearts were submerged in X-gal staining solution for 72 hours. The production of a blue stain product was visible in the ventricular myocardium, and left atria (n=4). No staining was observed in Cre negative hearts (data not shown, n=9). (D) Western blot analysis of NMIIB expression in protein extracts from control (+/+; flox/flox, n=3) and myocardial-specific Myh10 knock out (α-MHC/+; flox/flox, n=3) E14.5 hearts. Blots were also probed for β-actin as a loading control. (E) Peak intensity analysis of western blot image shown in D with ImageLab imaging software. The peak intensity for β-actin is relatively equal across all samples. (F) Peak intensities for NMIIB in myocardial-specific Myh10 knock out hearts is decreased compared to controls (arrows). (G-H) Immunohistochemistry for NMIIB on paraformaldehyde-fixed cardiac sections from control (n=3) and (I-J) myocardial-specific Myh10 knock out (n=3) E14.5 hearts. Images show a portion of the interventricular septum. Scale bars = 1mm (A, C), 100µm (G-J). Ao – aorta, LA – left atria, LV – left ventricle, PT – pulmonary trunk, RA – right atria, RV – right ventricle.

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5.2.5 Genotyping Animals to Identify Myocardial Myh10 Knock Out Embryos In order to determine whether Cre recombinase mediated homologous recombination had correctly occurred in α-MHC-Cre/+; flox/flox hearts, we performed PCR amplification of the Myh10 exon2 sequence to identify whether or not the floxed exon 2 sequence had been deleted in cardiac tissue. The genotyping protocol has been previously described for the Myh10∆ line (see Section 3.2.4.1). In order to establish that Cre mediated deletion of Myh10 exon 2 was specific to the cardiac cell population, we genotyped a panel of embryonic tissues, including heart, tail, brain and liver of E14.5 animals.

Unfortunately, whilst we were able to identify that that the Myh10 exon 2 deletion occurred within cardiac tissue, it appeared that the deletion was not limited to the heart, and we also observed the deleted PCR product when genotyping adult ear punches and other embryonic tissues (see Appendix 15, n>3). The finding that we could identify the deletion of Myh10 in tissues other than the heart immediately indicated that we were not able to continue our analysis of progeny derived from the initial α-MHC-Cre founder male.

To reestablish the α-MHC-Cre colony, we repeated the breeding scheme outlined above (see Section 5.2.2) with a new α-MHC-Cre founder male (courtesy of Dr. Elizabeth Cartwright). Genotyping was performed on the resultant progeny as described above to identify α-MHC-Cre/+; flox/+ embryos. We then performed PCR genotyping for the Myh10 exon 2 deletion on a panel of embryonic tissues including heart, tail, brain and liver, at both E14.5 and E16.5, as these time points would be used for subsequent phenotypic analysis (Fig 5.5). From this experiment, we were able to detect the exon 2 deleted PCR product in the hearts of α-MHC-Cre/+; flox/+ embryos (Fig 5.5, A, n=9). We did not observe this deleted product in either α-MHC-Cre/+; +/+ or +/+; flox/+ control littermate hearts (data not shown, n>3). Reassuringly, we were similarly unable to detect the deleted PCR product in either tail (Fig 5.5, B), brain (Fig 5.5, C), or liver (Fig 5.5, D) samples from α-MHC-Cre/+; flox/+ embryos. This shows that progeny descended from the second α-MHC-Cre founder male express Cre recombinase exclusively in the heart, and that this expression is sufficient to induce the excision of Myh10 exon 2. Expanding on this, we were similarly able to detect the deleted PCR product in ventricular tissue of α- MHC-Cre/+; flox/flox embryos (see Appendix 16, n=3). This additional control experiment has provided a sound basis for our future interpretations of the α-MHC- Cre/+; flox/flox phenotype.

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Figure 5.5: PCR Analysis of Myh10 Exon 2 Deletion in a panel of 'Second Generation' α-MHC-Cre/+; flox/+ Embryonic Tissues

(A) Genotyping results on ventricular tissue of α-MHC-Cre/+; flox/+ embryos. The deletion of Myh10 exon 2 from the floxed allele generates a smaller PCR product (600bp). (B) This smaller PCR amplicon is not detected in tail, (C) brain, or (D) liver tissue from the same embryos. Data shown is from E16.5 embryos. Experiment was repeated on E14.5 embryos and the same results were observed (total heterozygous embryos analysed = 9). The right-hand lane (∆/+) was used as a positive control to compare band sizes. For clarification, a schematic representation of the PCR amplicons shown above is provided in Appendix 12.

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5.2.6 Validation of Myh10 Deletion in Cardiomyocytes Once we had established that Myh10 was specifically deleted in the heart of α-MHC- Cre/+; flox/flox animals, it was necessary to confirm that this deletion resulted in a reduction in NMIIB protein specifically in cardiomyocytes. Due to our lack of antibodies against cardiomyocyte specific markers, we employed a modified cardiomyocyte isolation protocol (Louch et al., 2011; Sreejit et al., 2008) to enrich embryonic cardiomyocytes, and performed immunocytochemistry for NMIIB protein in both control (+/+; flox/flox, n=3) and myocardial-specific Myh10 knock out (α-MHC-Cre/+; flox/flox, n=3) samples (Fig 5.6, A-L). This protocol allowed the isolation of both cardiomyocyte and fibroblast cell populations by employing the pre-plating method, which removes the majority of cardiac fibroblasts from the collagenase digested heart cell suspension (Louch et al., 2011; Sreejit et al., 2008).

By using this method, we were able to detect NMIIB positive staining in both control fibroblasts and cardiomyocytes (Fig 5.6, A-F). Similarly, we found that α-MHC-Cre/+; flox/flox fibroblasts showed positive NMIIB staining (Fig 5.6, G-I). Interestingly, we found that cells in the cardiomyocyte-enriched population did not display NMIIB staining (Fig 5.6, J-L), suggesting that NMIIB is lost in cardiomyocytes, but not cardiac fibroblasts. However, before drawing a definitive conclusion, it is vital to confirm that these NMIIB- ablated cells are indeed cardiac myocytes by performing dual-staining with a cardiomyocyte specific marker, such as the intermediate filament protein, desmin (Ma et al., 2009), or the cardiomyocyte specific transcription factors, Nkx2.5 and Gata-4 (Kodama et al., 2005).

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Figure 5.6: Immunocytochemical Analysis of NMIIB Localisation in Cultured Cardiomyocytes and Fibroblasts (A-C) Immunofluorescence microscopy images showing the localisation of NMIIB in cardiac fibroblasts cultured from control (+/+; flox/flox, n=3) E15.5 hearts. (D-F) NMIIB expression in cardiomyocyte enriched cell cultures from control hearts. (G-I) Similarly, NMIIB is found in cardiac fibroblasts cultured from myocardial-specific Myh10 deleted (α-MHC-Cre/+; flox/flox, n=3) hearts. (J-L) In contrast, NMIIB staining was not detected in cells from the cardiomyocyte enriched population derived from myocardial- specific Myh10 deleted hearts. Scale bars = 50µm. CM – cardiomyocyte, Fibro - fibroblast.

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5.2.7 Morphology of α-MHC-Cre; flox/flox Mutant Embryos As previously discussed, we have found that global loss of Myh10 results in severe brain and cardiac developmental defects in the EHC mouse, which consequently lead to embryonic lethality in this strain. Although myocardial-specific Myh10 knock out animals are viable, we next sought to investigate whether α-MHC-Cre/+; flox/flox animals displayed any phenotypic similarity to the EHC mouse during embryonic development. This analysis was required as deletion of Myh10 from a single cardiac cell population, as opposed to the global deletion in the EHC line, may manifest as a subtle, non-lethal cardiac phenotype. Additionally, the background of our α-MHC-Cre line deviates from that of the previously reported myocardial-specific Myh10 knock out (Ma et al., 2009). As genetic background is known to influence α-MHC-Cre expression (Agah et al., 1997), this may therefore influence the severity of the myocardial-specific Myh10 deletion in our colony.

Embryos from the cross of the ‘second generation’ α-MHC-Cre/+; flox/+ males to Myh10flox/flox females (Fig 5.3, C, P2) were genotyped, and analysed at E16.5. This developmental time point was selected as both the brain and cardiac abnormalities displayed in the EHC mutants are extremely prominent at this stage. From our observations at dissection, we found that α-MHC-Cre/+; flox/flox embryos did not display any gross morphological defects at E16.5, and appeared phenotypically identical to control littermates (Fig 5.7, A, B, n>10). As expected, this demonstrates that myocardial ablation of Myh10 does not lead to developmental hydrocephalus, and is in agreement with previously reported studies of the cardiomyocyte-specific Myh10 knock out (Ma et al., 2009; Ma et al., 2010).

Closer examination of the α-MHC-Cre/+; flox/flox embryonic heart similarly revealed no phenotypic similarity to the EHC mutant (Fig 5.7, C-F). Mutant hearts displayed an identical phenotype to control hearts, and moreover, did not show any evidence of DORV or other gross morphological defect associated with the EHC heart at the same developmental stages (Fig 5.7, C-F, n>10). Interestingly, we were able to identify blood filled coronary vessels on the ventricular surface of myocardial Myh10 knock out hearts (Fig 5.7, D, F, arrowheads). Furthermore, we have confirmed the presence of coronary vessels at the molecular level in α-MHC-Cre/+; flox/flox embryonic hearts by utilising PECAM-1 immunohistochemistry in a limited number of samples (see Appendix 17, n=2). In addition, we observed no evidence of the ventricular nodules that decorate the surface of the global Myh10 deleted heart. Moreover, control (+/+; flox/flox) littermates displayed

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a wild type phenotype, illustrating that the floxed Myh10 allele does not disrupt cardiogenesis. This morphological evaluation demonstrates that both α-MHC-Cre/+; flox/flox, and Myh10flox/flox embryos do not show developmental defects at the gross anatomical level. However, a more extensive examination of vascular components (i.e. vECs and vSMCs localisation) is required (as described in Section 3.2.5) before it can be definitely established that α-MHC-Cre/+; flox/flox embryos do not display coronary vessel defects related to the global Myh10 ablated heart.

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Figure 5.7: Morphological Analysis of 'Second Generation' Cardiomyocyte-Specific Myh10 Knock Out Embryos and Hearts

(A) Dissection image of control (+/+; flox/flox) and (B) cardiomyocyte-specific Myh10 knock out (α-MHC-Cre/+; flox/flox) E16.5 embryos. (C) Ventral view of control and (D) myocardial knock out hearts at E18.5. Both hearts show correct positioning of the aorta and pulmonary trunk (dashed line). In addition, blood-filled coronary vessels can be observed on the ventricular surface (arrow heads). (E) Dorsal view of control and (F) mutant hearts at E18.5. Similarly, coronary vessels are clearly evident (arrowheads). Scale bars = 2mm (A-B), 1mm (C-F). Ao – aorta, LA – left atria, LV- left ventricle, PT – pulmonary trunk, RA –right atria, RV – right ventricle.

5.2.8 Generation of Epicardial-Specific Myh10 Knock Out Animals Our finding that the myocardial-specific Myh10 knock out mice did not recapitulate any aspect of the EHC phenotype led us to form the hypothesis that the cardiac abnormalities associated with the EHC line were caused by a loss of NMIIB from a different cardiac cell population. In light of our previously described findings, which strongly suggest that the EHC mouse shows epicardial dysfunction, we hypothesize that loss of NMIIB from the epicardium is responsible for these defects. To test this hypothesis, we sought to establish an epicardial-specific Myh10 knock out line and analyse the phenotype of mutant embryos.

5.2.9 The Wt1tm1(EGFP/cre)Wtp/J Mouse Line Our first attempt to generate epicardial-specific Myh10 knock out embryos utilised the Wt1tm1(EGFP/cre)Wtp/J transgenic mouse line (subsequently referred to as Wt1-CreGFP), which expresses a Cre recombinase/eGFP (enhanced green fluorescent protein) fusion protein under the control of the Wt1 promoter (Fig 5.8, A). As previously mentioned, Wt1 is considered a strong marker of epicardial cells during development. We sought to delete Myh10 specifically from the epicardium by utilising Cre recombinase mediated recombination to excise Myh10 exon 2 in Wt1 expressing cells. We then hoped to utilise the properties of the eGFP expression in these cells to isolate them by flow cytometry (FACS) for further experimentation.

To ensure adequate quality control, we first crossed the Wt1-CreGFP founder male (purchased from the Jackson Laboratory) to the aforementioned ROSA26 reporter line to monitor Cre recombinase expression in the Wt1-CreGFP line. Embryos were genotyped for the presence of Cre and X-gal staining was performed on E14.5 hearts (Fig 5.8, B-E). Surprisingly, we were only able to detect positive X-gal staining in 2 of 12 Cre positive hearts (Fig 5.8 B-E). In addition, in samples in which positive X-gal staining was observed, β-galactosidase activity (and therefore Cre expression) appeared to be limited to either weak staining in very few cells (Fig 5.8, C, D, arrowheads), or tightly restricted to distinct regions of the ventricular surface (Fig 5.8, E, arrow).

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Figure 5.8: Analysis of Cre Recombinase Expression in the Wt1-CreGFP Mouse Line (A) The Wt1-CreGFP mouse line caries a Cre-eGFP fusion sequence (blue and green region) knock in which is under the activity of the Wt1 promoter (grey region). (B-E) Wt1- CreGFP male mice were crossed to the ROSA26 reporter line. Embryos were harvested at E14.5 and submerged in X-gal staining for 72 hours. From a total of 12 Cre positive hearts, only two showed positive X-gal staining (C, D, arrowheads and E, arrow). No staining was observed in Cre negative hearts (data not shown). The heart shown in D and E was accidentally damaged during dissection, removing the atria. Scale bars – 1mm. LA – left atria, LV – left ventricle, RA – right atria, RV – right ventricle.

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The unreliable expression of Cre in the Wt1-CreGFP line was confirmed by performing RT-PCR on adult heart tissue (data not shown). It has previously been reported that Wt1 is expressed in adult epicardial cells at the atrioventricular sulcus and at the ventricular apex, but not throughout the epicardium covering the ventricular wall (van Wijk et al., 2012). We were only able to detect Cre expression in one of four Cre positive hearts by RT-PCR (data not shown).

Due to the inconsistencies observed in both the level and localisation of Cre expression in the Wt1-CreGFP line, we concluded that this line could not be used to reliably induce the deletion of Myh10 in the epicardium, and was therefore not an appropriate tool to pursue the generation of epicardial-specific Myh10 knock out animals.

5.2.10 The Wt1tm2(cre/ERT2)Wtp Mouse Line To circumvent the issues we encountered with the Wt1-CreGFP mouse line, we next adopted an approach to utilise the Wt1tm2(cre/ERT2)Wtp inducible conditional knock out mouse (subsequently referred to as Wt1-CreERT2 for simplicity). This line similarly contains a heterozygous Cre recombinase knock in at the Wt1 locus, but in this instance, the protein produced by the activity of the Wt1 promoter is a Cre-ERT2 fusion protein (Fig 5.9, A). The Cre-ERT2 fusion is restricted to the cytoplasm unless exposed to the synthetic estrogen receptor ligand tamoxifen, upon which Cre-ERT2 translocates to the nucleus to initiate loxP site directed recombination. This system permits the generation of conditional Myh10 knock out animals in which Myh10 deletion is restricted not only spatially (to the epicardium), but also temporally. This tool theoretically allows the user to further assess the requirement of NMIIB in the epicardium at different developmental stages.

5.2.11 Wt1-CreERT2 Breeding Strategy We crossed Myh10flox/flox female mice to a number of Wt1-CreERT2 males in Professor Paul Riley’s colony at the University of Oxford to generate Wt1-CreERT2/+: flox/+ male mice (Fig 5.9, B). These animals were subsequently crossed to additional Myh10flox/flox females to generate Wt1-CreERT2/+; flox/flox embryos (Fig 5.9, B, boxed P2 progeny). Cre mediated recombination was induced by administering 4-hydroxy-tamoxifen via intraperitoneal injection (as described in Zhou et al., 2008) to pregnant females at either E9.5 and E11.5 (‘early’), or E10.5 and E12.5 (‘late’) (Fig 5.9, B). This injection schedule was selected on the advice of the Riley laboratory. Embryos were then collected at E14.5 for phenotypic analysis.

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Figure 5.9: Breeding Strategy to Generate Epicardial-Specific Myh10 Knock Out Animals Using the Wt1-CreERT2 Mouse at the University of Oxford (A) The Wt1-CreERT2 mouse harbours a Cre-ERT2 fusion sequence under the activity of the Wt1 promoter. Cre-mediated recombination is only induced following the administration of tamoxifen, which promotes the nuclear translocation of the Cre-ERT2 fusion protein. (B) Breeding scheme to generate epicardial-specific Myh10 knock out animals (boxed progeny, P2). Please see main text for full details. This work was carried out in collaboration with Prof. Paul Riley’s laboratory at the University of Oxford.

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5.2.12 Heart Morphology of ‘First Generation’ Potential Epicardial- Specific Myh10 Knock Out Embryos Following genotyping, it became apparent that Wt1-CreERT2/+; flox/flox embryos did not display gross morphological defect associated with the EHC phenotype (data not shown). Similarly, Wt1-CreERT2/+; flox/flox hearts displayed a highly comparable morphology to control (+/+; flox/flox) hearts, with correct positioning of the aorta and pulmonary arteries (Fig 5.10, A-F, dashed lines, n=6). In addition, we frequently observed blood filled vessel structures on the ventricular surface (Fig 5.10, A-F, arrowheads). However, in the cohort of hearts from pregnant females injected at the ‘early’ time points (E9.5 and E11.5), it appeared that there were areas of the ventricular coronary vasculature that exhibited micro-haemorrhaging (Fig 5.10, B, C, arrows, n=3). This observation indicated that the development of the coronary vasculature was subtly disrupted.

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Figure 5.10: Morphological Analysis of 'First Generation' Potential Epicardial-Specific Myh10 Knock Out Embryonic Hearts

Figure 5.10: Morphological Analysis of 'First Generation' Potential Epicardial- Specific Myh10 Knock Out Embryonic Hearts (A) Dissection images of control (+/+; flox/flox) and (B-C) potential epicardial-specific Myh10 knock out (Wt1-CreERT2/+; flox/flox) E14.5 hearts injected with tamoxifen at E9.5 and E11.5 (early schedule). All hearts show the correct orientation of the great arteries (dashed lines), and blood-filled coronary vessels are apparent (A, B, arrowhead). Areas of micro-haemorrhaging are evident on the ventricular surface of potential knock out hearts (B, C, arrows, n=3). (D) Dissection images of control and (E-F) potential knock out E14.5 hearts injected at E10.5 and E12.5 (late schedule). Correct positioning of the great arteries (D-F, dashed lines), as well as the presence of blood-filled coronary vessels are evident (D- F, arrowheads). All images were acquired at the same magnification. Ao – aorta, LA – left atria, PT – pulmonary trunk, RA – right atria. In the panel label, Wt1-CreERT2 has been shortened to Wt1-Cre due to space restrictions. This work was carried out in collaboration with Prof. Paul Riley’s laboratory at the University of Oxford.

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5.2.13 Validation of Loss of Epicardial NMIIB Before drawing any significant conclusions, it was imperative to establish that Myh10 had been deleted from the epicardium of Wt1-CreERT2/+; flox/flox hearts. Due to the difficulty in isolating epicardial cells from the whole heart sample, we were unable to conduct a PCR based genotyping strategy to detect the genomic Myh10 exon 2 deletion. Therefore, we performed immunohistochemistry for NMIIB on these Wt1-CreERT2/+; flox/flox hearts to determine whether or not NMIIB protein had been ablated in epicardial cells. We performed dual staining with SMαA to detect whether these ‘micro- haemorrhaging vessels showed disruption to the localisation of vascular smooth muscle cells (Fig 5.11, A-I). Prominent NMIIB staining was detected in the epicardium of control (+/+; flox/flox, n=3) hearts (Fig 5.11, A). In addition, we found clear coalescence of smooth muscle cells around vessel structures in control hearts (Fig 5.11, B, C, arrows). Unfortunately, we detected prominent NMIIB immunoreactivity in the epicardium of mutant (Wt1-CreERT2/+; flox/flox, n=3) hearts at comparable levels to control littermates (Fig 5.11, D, G). However, the recruitment of vascular smooth muscle cells did appear to be subtly disrupted in mutants, with an abundance of SMαA staining in the sub- epicardial region that failed to completely delineate the vessel structure (Fig 5.11, E, arrowheads, n=2). Interestingly, mutant hearts demonstrated large areas of epicardial detachment (Fig 5.11, G, I, asterisks, n=2). Whilst this may contribute to the micro- haemorrhage phenotype, the root cause of this defect was shown not to be due to loss of epicardial NMIIB expression. Instead, it is likely that tamoxifen toxicity was responsible for this epicardial detachment, and the subtle disruption to coronary vessel development.

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Figure 5.11: Molecular Analysis of NMIIB and SMαA Localisation in ‘First Generation’ Potential Epicardial-Specific Myh10 Knock Out Embryonic Hearts

(A-C) Immunohistochemical analysis of NMIIB and SMαA localisation in control (+/+; flox/flox) and (D-I) potential epicardial-specific Myh10 knock out (Wt1-CreERT2/+; flox/flox) paraformaldehyde-fixed E14.5 cardiac sections. NMIIB is present in the epicardium (A). Positive SMαA staining indicates the localisation of vascular smooth muscle cells around vessel structures in the myocardial wall (B, C, arrows). (D) Surprisingly, NMIIB is detectable in the epicardium of potential knock out hearts. The migration of vascular smooth muscle cells appears to be disrupted, as indicated by abundant SMαA staining in the sub-epicardial region (E, F, arrowheads). In addition, there is incomplete SMαA staining around some vessel structures (D, E). Potential knock out hearts also display extensive regions of epicardial detachment (G, I, asterisks). Scale bars = 25µm. In the figure labels, Wt1-CreERT2 has been shortened to Wt1-Cre due to space restrictions.

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5.2.14 Breeding Scheme for Colony Re-establishment at the University of Manchester In order for us to optimise our breeding strategy and tamoxifen dosing regime, we imported the P1 Wt1-CreERT2/+; flox/+ males (Fig 5.9, B) from Paul Riley’s laboratory and reestablished the Wt1-CreERT2 colony at the University of Manchester. We implemented a revised breeding strategy, shown in Fig 5.12. By breeding the Wt1- CreERT2/+; flox/+ founder males to the Myh10∆ line (Fig 5.12, A), we generated Wt1- CreERT2/+; flox/∆ males (Fig 5.12, A, boxed progeny). Simultaneously, we intercrossed +/+; flox/+ and +/+; ∆/+ heterozygous animals to generate +/+; flox/∆ females (Fig 5.12, B, boxed progeny). Subsequently, we intercrossed Wt1-CreERT2/+; flox/∆ males and +/+; flox/∆ females (Fig 5.12, C) and induced Cre mediated recombination by tamoxifen injection. Using this breeding strategy, we were able to generate; Wt1- CreERT2/+; flox/flox, Wt1-CreERT2/+; flox/∆, Wt1-CreERT2/+; ∆/∆ and +/+; ∆/∆ embryos (Fig 5.12, C, boxed P2 progeny). This strategy is beneficial on two fronts; firstly, the generation of both Wt1-CreERT2/+; ∆/∆ and +/+; ∆/∆ embryos acts as an internal control for the global Myh10 knock out phenotype, which is informative to compare to epicardial-specific conditional Myh10 knock out embryos. Secondly, the presence of Wt1- CreERT2/+; flox/∆ embryos requires the recombination of just one floxed allele to generate an epicardial-specific Myh10 knock out. This reduces the required efficiency of the Cre recombinase if activity is not sufficient to delete both alleles in Wt1-CreERT2/+; flox/flox embryos, and increases the probability of generating homozygous Myh10 null epicardial cells. Due to the heterozygous nature of the Wt1 Cre in the males, 50% of the progeny generated from this cross will be negative for the Cre. This serves as an ideal negative control, to which any tamoxifen toxicity can be finely assessed in each genetic combination.

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Figure 5.12: Revised Breeding Strategy to Generate Epicardial-Specific Myh10 Knock Out Animals at the University of Manchester (A) In the first cross, Wt1-CreERT2/+; flox/+ males imported from the University of Oxford were crossed to heterozygous Myh10∆ (∆/+) females to generate Wt1- CreERT2/+; flox/∆ animals (boxed progeny, P1). (B) In the second cross, heterozygous Myh10flox/+ males (+/+; flox/+) were crossed to heterozygous Myh10∆ females to generate +/+; flox/∆ animals (boxed progeny, P1). (C) In the final cross, Wt1-CreERT2/+; flox/∆ males were bred to +/+; flox/∆ females (the desired progeny from the previous crosses), to generate an array of both epicardial-specific, and global Myh10 deleted progeny (boxed progeny, P2) following tamoxifen administration. This array of genotypes is useful for comparison of the epicardial-specific Myh10 null phenotype to both the global Myh10 null, and Cre negative controls, in addition to increasing the frequency of epicardial knock out animals. Please see text for full details. P = progeny.

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5.2.15 Tamoxifen Dosing and Progeny Analysis As outlined in the breeding scheme (Fig 5.12, C), we crossed Wt1-CreERT2/+; flox/∆ males to +/+; flox/∆ female mice. Tamoxifen was administered via intraperitoneal injection (40mg/Kg) at either E10.5 and E11.5, or E10.5 and E12.5 (see Appendix 5) to optimise the dosing regime. These administration time points were selected based on personal communication with Dr. Nicola Smart (Oxford University). Pregnant females were sacrificed at E16.5 for embryo phenotypic analysis. We selected this late gestation time point primarily to analyse the cardiac morphology of the embryos and compare this to the EHC phenotype, as the maturing coronary vasculature should be readily observable at this time. As an appropriate control to test for Cre toxicity or the presence of a phenotype in the absence of gene deletion, we crossed Wt1-CreERT2/+; flox/∆ males to +/+; flox/+ female mice. The resultant control progeny (Wt1-CreERT2/+; flox/+ or Wt1- CreERT2/+; +/∆) were phenotypically identical to no Cre controls (data not shown).

By consulting our compiled list of tamoxifen-injected females (see Appendix 5), one can appreciate that we experienced great difficulty in generating live litters at E16.5. From a total of 18 injected females (plus two non-injected females), we obtained live embryos in just four litters (see Appendix 5). It is worth noting that these live litters were generated from females at the earlier stages of this experiment. We did not identify a trend between the injection schedule and the presence of live litters, as each injection regime generated two live litters (see Appendix 5).

There are a number of protocols described for the IP administration of tamoxifen. To establish whether the type of solvent we used for dissolving the tamoxifen was influencing the survival of the litters, we made up a batch to the same concentration in sesame oil (‘flox∆ 157’), and injected at a single time point (E11.5) (see Appendix 5). Similarly, the litter from this female was dead at dissection.

We did not include the administration of sham injections in this protocol. To establish whether the injection procedure (or tamoxifen exposure itself) caused the high levels of attrition in the litters, we sacrificed 2 plugged females at E16.5 without subjecting them to injections. In one of these females (‘Wt1 301’), embryos were present but at a much earlier embryonic stage than suggested by the recorded plug date (approx. E6.5 vs. E16.5) (see Appendix 5). The other non-injected female presented with a distended fluid-filled uterus.

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Whilst the number of females in this non-injected cohort is limited, this result indicates that the health of our litters was disrupted without tamoxifen exposure.

5.2.16 Morphology of ‘Second Generation’ Potential Epicardial-Specific Myh10 Knock Out Embryos Unfortunately, the difficulties described above have severely limited the number of potential epicardial-specific Myh10 knock out embryos at our disposal for analysis. At the time of writing, we have accrued a total of six Wt1-CreERT2/+; flox/∆ or Wt1- CreERT2/+; flox/flox embryos from 4 separate litters, which have the potential to have Myh10 deleted specifically in the epicardium. At the point of dissection, these E16.5 embryos did not display gross morphological defects associated with the EHC phenotype and appear phenotypically similar to control littermates (Fig 5.13, A, C, E, G). Similarly, the majority of potential epicardial-specific Myh10 knock out embryos did not display cardiac defects, and presented a highly similar morphology to controls (Fig 5.13, B, D, F). However, it did appear that the heart of one Wt1-CreERT2/+; flox/∆ embryo, which had been injected at E10.5 and E12.5, displayed a coronary vessel abnormality (Fig 5.13, H). Whilst some vessels were clearly visible on the ventricular surface (Fig 5.13, H, arrows), there was evidence of micro-haemorrhaging and ventricular blisters (Fig 5.13, H, arrowheads). In addition, the positioning of the great arteries (pulmonary trunk and aorta) appeared to abnormally deviate from that in control hearts (Fig 5.13, H, dashed lines). However, before suggesting that epicardial Myh10 ablation contributes to these abnormalities, it is essential to validate the deletion of NMIIB from the embryonic epicardium. In addition, our ability to interpret this result in the event that NMIIB is absent from the epicardium is overwhelmingly hindered by the presence of this defect in just a single embryo.

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Figure 5.13: Morphological Analysis of 'Second Generation' Potential Epicardial- Specific Myh10 Knock Out Embryos and Hearts

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Figure 5.13: Morphological Analysis of 'Second Generation' Potential Epicardial- Specific Myh10 Knock Out Embryos and Hearts (A-B) Dissection images of control (+/+; flox/flox) and (C-D) potential epicardial-specific Myh10 knock out (Wt1-CreERT2/+; flox/flox, n=1) E16.5 embryos and hearts injected at E10.5 and E11.5. Both animals show an identical wild type phenotype. (E-F) Dissection images of control (+/+; flox/∆) and (G-H) potential epicardial-specific Myh10 knock out (Wt1-CreERT2/+; flox/∆, n=2) E16.5 embryos and hearts injected at E10.5 and E12.5. Potential knock out embryonic heart shows abnormalities in the positioning of the aorta and pulmonary trunk, when compared to controls (F, H, dashed lines, n=1 50% penetrance). Whilst coronary vessels are present on the ventricular surface (H, arrows), potential knock out hearts show evidence of ventricular nodules and micro-haemorrhaging (H, arrowheads, n=1 50% penetrance). Scale bars = 1mm (A, C, E, G), 0.5 mm (B, D, F, H). Ao - aorta, LA – left atria, PT, pulmonary trunk, RA – right atria. In the figure labels, Wt1-CreERT2 has been shortened to Wt1-Cre due to space restrictions.

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5.3 Discussion To briefly summarise, after several unsuccessful attempts, we have preliminary data to suggest that we have generated a myocardial-specific Myh10 ablated mouse derived from the α-MHC-Cre line. This mouse does not display gross anatomical embryonic cardiac defects, consistent with previously reported studies (Ma et al., 2009; Ma et al., 2010). In addition, we have generated a limited number of inducible epicardial-specific Myh10 knock out embryos by utilising the Wt1-CreERT2 line. One of these embryos appeared to display multiple cardiac defects associated with the EHC phenotype. However, we have yet to confirm that the genotyped status of this cohort of animals translates to the deletion of epicardial NMIIB, and so are unable to determine whether this experiment has been successful at the time of writing.

The generation of cardiac-specific Myh10 knock out animals is essential to facilitate the delineation of NMIIB functions during cardiogenesis. Our findings from the EHC mouse indicate that loss of NMIIB causes defects in both the epicardium and myocardium. It is widely accepted that intricate molecular communication between these two tissues is fundamentally important for correct heart formation. This thesis has described severe defects in epicardial cell function in the Myh10 ablated heart. Such dysfunction has previously been reported to underpin subsequent defects in myocardial development. In light of this, we sought to test the hypothesis that NMIIB is essential for correct epicardial cell function, and that loss of NMIIB in the EHC mouse causes epicardial dysfunction and downstream myocardial defects. To achieve this, we aimed to generate both epicardial and myocardial-specific Myh10 knock out animals by utilising Cre-loxP genomic editing technology, and phenotypically evaluate these for cardiac defects.

We utilised the ROSA26 reporter mouse to determine the spatial expression pattern of Cre expression in both the epicardial-specific Wt1-CreGFP and myocardial-specific α-MHC- Cre lines. Through this quality control, we were able to establish inconsistencies in Cre expression in the Wt1-CreGFP mouse. Expression was detected via X-gal staining in just two of the 11 Wt1-Cre positive samples analysed. Moreover, Cre expression was either tightly restricted to distinct regions of the ventricular surface, or limited to weak expression in very few cells. This inconsistency was compounded by our failure to detect Cre expression by RT-PCR in three of four Wt1-Cre positive adult hearts (data not shown). It has previously been shown that Wt1 is expressed in epicardial cells at the atrioventricular sulcus and ventricular apex of the adult heart (van Wijk et al., 2012). Although Wt1 is not

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expressed in the majority of the epicardium that enshrouds the ventricular wall, we did detect Cre expression in one Wt1-Cre adult heart, indicating that Wt1-driven Cre expression in adult epicardial cells is detectable. Concern about the reliability of the Wt1- CreGFP line has previously been reported (Doetschman and Azhar, 2012). The highly unreliable Cre expression observed in the Wt1-CreGFP mouse led us to abandon this line as a vehicle to generate epicardial-specific Myh10 knock out animals.

Our experiments with the ROSA26R mouse to identify the Cre expression domain in α- MHC-Cre mice confirmed that Cre is expressed in the ventricular myocardium. Similarly, by utilising a combination of western blotting and immunohistochemistry on adult and E14.5 hearts respectively, we found that α-MHC-Cre/+; flox/flox samples showed a reduction in NMIIB signal. This suggested that cardiomyocytes-specific Myh10 deletion had occurred earlier in development, consistent with the findings of previous studies (Agah et al., 1997; Gaussin et al., 2002; McFadden et al., 2005; Ma et al., 2009. Indeed, by genotyping ventricular tissue of E14.5 α-MHC-Cre/+; flox/flox embryos, we established that the Cre-mediated deletion of the Myh10 exon 2 sequence had occurred (data not shown). Unfortunately, we identified that this deletion event had also occurred in the tail, brain and liver of α-MHC-Cre/+; flox/flox embryos, suggesting erroneous Cre expression in these tissues. Interestingly, whilst it has been previously shown that the α-MHC-Cre transgene can become more broadly and non-specifically expressed in the adult mouse (Agah et al., 1997), the non-specific Cre mediated deletion that we observe in a panel of embryonic tissues has not been previously described. We remain unsure whether or not this ‘leaky’ Cre is an aberrant feature unique to our initial α-MHC-Cre founder male, or if this animal was incorrectly labelled as possessing the α-MHC-Cre transgene. Needless to say, these unusual results compromised our analysis of animals derived from this founder male. However, this quality control analysis was crucially important; we identified erroneous Cre expression before incorrectly interpreting the myocardial-specific Myh10 null phenotype, and additionally established a more robust control protocol to test for the future generation of α-MHC-Cre mediated Myh10 knock out animals.

Following colony re-establishment with a new α-MHC-Cre founder male, we were able to confirm that Cre-mediated Myh10 exon 2 deletion was restricted to ventricular cardiac tissue. We then sought to establish that this genomic deletion resulted in the loss of NMIIB protein specifically in cardiomyocytes, and not other cardiac cell populations. Due to our lack of antibodies against cardiomyocyte specific markers, we undertook the

189 pragmatic approach to isolate and culture cardiomyocytes to generate enriched cardiomyocyte cell populations from embryonic hearts by employing a modified cardiomyocyte isolation protocol (Louch et al., 2011; Sreejit et al., 2008). Whilst we were able to identify positive NMIIB staining in fibroblasts derived from α-MHC-Cre/+; flox/flox hearts, we found that cells in the cardiomyocyte population did not display NMIIB immunoreactivity. It has been shown that cells isolated by means of this protocol display Gata-4 and Nkx2.5 positive staining (Sreejit et al., 2008). Both Gata-4 and Nkx2.5 are cardiomyocyte-specific transcription factors (Kodama et al., 2005). Combined with our data, this suggests that Myh10 is ablated in cardiomyocytes derived from α-MHC-Cre/+; flox/flox hearts. However, due to time and reagent limitations, we have not been able to explicitly confirm that NMIIB null cells generated in our hands are cardiomyocytes. Performing dual-staining for NMIIB and a cardiomyocyte marker, such as Gata-4/Nkx2.5, or alternatively, desmin (Ma et al., 2009; Ma et al., 2010), or α-sarcomeric actin (Rodgers et al., 2009), would quickly and robustly establish whether or not we have generated a true cardiomyocyte specific Myh10 null mouse line. Our confidence that the loss of NMIIB is exclusive to the cardiomyocyte population in α-MHC-Cre/+; flox/flox hearts could be further increased at the genetic level by performing the Myh10 exon 2 deletion PCR experiment on both cardiomyocytes and fibroblasts isolated in this culture protocol.

Analysis of α-MHC-Cre/+; flox/flox embryos derived from this second α-MHC-Cre founder male do not display any phenotypic similarity to the EHC mouse. Under gross morphological examination, we found that presumptive myocardial-Myh10 knock out animals are phenotypically normal and identical to control littermates. Moreover, we found that α-MHC-Cre/+; flox/flox animals are viable and survive to adulthood. The absence of EHC associated cardiac defects in these mutant mice compliments the findings of previously reported myocardial-specific Myh10 knock out animals (Ma et al., 2009; Ma et al., 2010). Here, we report that α-MHC-Cre/+; flox/flox animals clearly display blood filled coronary vessels on their ventricular surface, indicating that epicardial function is not disrupted in these hearts. We have shown preliminary data in these hearts confirming the presence of coronary vessels at the molecular level with PECAM1 staining (see Appendix 17). However, we have not closely examined the structure of these vessels, which could be achieved by performing SMαA staining on embryonic hearts to characterise the vasculature, as described in Sections 3.2.5.2. Our findings expand upon the work of Ma and colleagues (2009), who do not report on epicardial functionality in their studies (Ma et al., 2009). However, as the original Myh10 global knock out mouse described by this group

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(Tullio et al., 1997) was not reported to display coronary vessel abnormalities, it may be that genetic background modifiers influence the Myh10 null phenotype in a strain specific manner.

It has previously been reported that myocardial-specific Myh10 knock out mice develop a cardiac myopathy at around 10 months of age (Ma et al., 2009). Similarly, the Adelstein group reports that these mice are born with enlarged, multinucleated cardiomyocytes (Ma et al., 2009). We have not performed histology on α-MHC-Cre/+; flox/flox hearts to determine whether these two aspects are recapitulated in the myocardial-specific Myh10 null mice generated in our hands. Interestingly, recent research has associated prolonged Cre activity with cytotoxic affects, particularly in the adult mouse heart (Doetschman and Azhar, 2012; Schmidt-Supprian and Rajewsky, 2007). Combined with the increased non- specific expression of Cre in the adult α-MHC-Cre mouse (Agah et al., 1997), we would suggest that the interpretation of the adult α-MHC-Cre/+; flox/flox phenotype must be very carefully considered. Similarly, the observation that NMIIB expression is both rapidly decreased and dramatically re-dispersed throughout cardiac cells following birth (Ma et al., 2010), suggests that NMIIB serves different functions in embryonic and mature cardiomyocytes. Findings in the adult can therefore not reliably be extrapolated to infer embryonic NMIIB function.

From the above description, it appears that myocardial-Myh10 expression is dispensable for correct cardiogenesis, and indeed embryonic survival. Whilst we have not examined α- MHC-Cre/+; flox/flox hearts for defects in cardiomyocyte cytokinesis, we strongly suspect that these will be apparent as reported previously in the EHC heart, and by others (Ma et al., 2009; Ma et al., 2010). Our observation that α-MHC-Cre/+; flox/flox hearts develop normal coronary vessels suggests that epicardial function is not compromised. This indicates that the epicardial dysfunction observed in the EHC heart is not underpinned by loss of NMIIB function in the cardiomyocyte cell population, and implies that NMIIB is not required to generate myocardial derived signalling cues to the epicardium. This reinforces our hypothesis that NMIIB serves different and distinct cell- specific functions during cardiogenesis. In addition, our finding that α-MHC-Cre/+; flox/flox hearts do not display DORV, suggests that potential defects in cardiomyocyte cytokinesis are not sufficient to cause this EHC associated phenotype. This again strengthens our proposal that NMIIB is required in additional cardiac cells types, and may contribute to correct positioning of the great arteries by propagating the population of the

191 developing outflow tract by; orchestrating signalling events between the epicardium and myocardium required for correct myocardial proliferation (von Gise and Pu, 2012; Olivey and Svensson, 2010), or facilitating CNCC migration to the outflow tract (Achilleos and Trainor, 2012; Brade et al., 2013). Generating additional cardiac cell-specific Myh10 ablated animals will help to establish the tissue specific requirement for NMIIB during cardiogenesis, and advance our understanding of the precise role that NMIIB serves in this process.

In relation to this, we embarked upon further attempts to delete Myh10 specifically in the embryonic epicardium by utilising the Wt1-CreERT2 mouse line. Unfortunately, we encountered numerous problems in the generation of potential knock out embryos that we have attributed to tamoxifen toxicity. Whilst we found that our experiments in collaboration with Professor Paul Riley’s group at the University of Oxford generated potential knock out embryos with evidence of perturbed coronary vessels development (micro-haemorrhaging, epicardial detachment), we failed to delete Myh10 in the epicardium, as determined by immunofluorescent microscopy. These abnormalities are therefore not attributable to loss of epicardial NMIIB.

We attempted to overcome this recombination failure by revising our breeding scheme to introduce the Myh10∆ allele when re-establishing the Wt1-CreERT2 colony at the University of Manchester. By generating Wt1-CreERT2/+; flox/∆ embryos, only one floxed allele needs to be recombined to generate epicardial-Myh10 null embryos, therefore reducing the required Cre efficiency. However, our efforts to optimise this breeding strategy and tamoxifen-dosing regime were impeded by a high level of litter abortion. This issue was not overcome by varying the timing of tamoxifen administration (based on personal communication with Drs. Nicola Smart and Catherine Roberts), or changing the solvent used to dissolve the tamoxifen. Tamoxifen administration has previously been associated with high rates of litter reabsorption (Doetschman and Azhar, 2012) and, at the cellular level, initiation of the DNA damage response (Bersell et al., 2013). Further experimentation is required to determine if fetal survival can be enhanced by either changing the method of tamoxifen administration (i.e. gavage), or using the tamoxifen active metabolite, 4-hydroxytamoxifen, which is known to bind to estrogen receptors with a greater affinity (Asp et al., 2013). Additionally, dosing dams with progesterone may help to counteract tamoxifen toxicity effects (Jackson Laboratory, www.jax.org). This optimisation is essential to expand upon our existing study and generate sufficient embryos for analysis.

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That being said, we have been able to generate an, albeit limited, cohort of potential epicardial-specific Myh10 knock out embryos from our re-establishment of the Wt1- CreERT2 colony at the University of Manchester. The majority of these samples (five of six) do not display gross morphological cardiac defects. However, one embryo appears to show abnormal placement of the great arteries, in addition to areas of ventricular micro- haemorrhage and ventricular surface blisters. Furthermore, this heart shows irregularly placed atria and an abnormal ventricular morphology. These defects are highly comparable to those displayed in EHC mutants at the same developmental stage. It is interesting to note that the genotype of this embryo (Wt1-CreERT2/+; flox/∆) requires just a single floxed allele to be recombined to generate the epicardial-Myh10 knock out status. This phenotype was not present in Wt1-CreERT2/+; flox/+ or Wt1-CreERT2/+; +/∆˙embryos, indicating that this phenotype is not due to Cre toxicity and is only observed in a scenario where complete gene deletion can take place. In addition, this dam was injected at E10.5 and E12.5, which may reduce the exposure of embryos to elevated tamoxifen levels when compared to animals injected on consecutive days (i.e. E10.5 and E11.5). However, in order to draw any interpretations from these embryos, we must confirm that their potential epicardial knock out status translates to a loss of NMIIB protein in the epicardium. We have previously demonstrated that this can be achieved by performing immunohistochemistry for NMIIB on tissue sections. Only then will we be able to conclude whether or not epicardial-Myh10 ablation results in cardiac defects.

In the event that loss of epicardial NMIIB is shown to cause the aforementioned cardiac abnormalities, we would seek to repeat the coronary vessel characterisation and analysis of epicardial cell function we performed in the EHC mutants - as described in Chapters 3 and 4 respectively – and compare these data. This research would help to definitively establish the effect of Myh10 ablation on epicardial cell biology and the programme of coronary vessel formation.

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5.4 Conclusion To conclude, we have made significant inroads into generating robust and reliable tissue- specific Myh10 knock out animals. We have preliminary data to suggest that we have generated myocardial-specific Myh10 knock out adult mice. From this, we strongly suspect that loss of NMIIB in cardiomyocytes during development does not disrupt correct cardiogenesis, at least in respect to gross anatomical defects causing embryonic lethality, and therefore does not underpin the epicardial dysfunction observed in globally ablated Myh10 animals. The correct development of the coronary vasculature and epicardial function in myocardial-specific Myh10 null mice has not previously been reported in the literature. This reinforces our original hypothesis that NMIIB is specifically required in the epicardium to regulate coronary vessel formation. The direct testing of this hypothesis can be achieved by validating the loss of epicardial NMIIB by immunohistochemistry in the Wt1-CreERT2/+; flox/∆ embryo that showed cardiac defects, whilst simultaneously generating additional embryos on which to conduct an evaluation of coronary vessel architecture and epicardial function, as described in the previous two chapters. Similarly, the generation of epicardial-specific Myh10 mutant animals has not been reported previously. If we are able to definitively link loss of epicardial NMIIB to cardiac defects, we would seek to further examine the epicardial-specific Myh10 null phenotype to delineate the precise role(s) of NMIIB during normal cardiogenesis. This work is described in detail in the future objectives section of this thesis.

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Chapter 6: Final Discussion

This thesis research has made substantial inroads into enhancing our understanding of the processes that underpin epicardial cell function, specifically in relation to coronary vessel formation. We have established that an ENU-induced point mutation in Myh10 results in catastrophic cardiac defects and embryonic lethality. We have presented strong evidence to indicate that the pathology of these cardiac malformations is likely to be caused by compromised epicardial cell function, due to the loss of NMIIB from this tissue. To the author’s knowledge, the effect of NMIIB ablation on epicardial cell behaviour is yet to be specifically investigated, providing the opportunity to disseminate novel discoveries in this field and advance our understanding of epicardial cell biology.

6.1 Phenotypic Analysis of the EHC Mouse This work has expanded upon the undertakings of previous laboratory members who first isolated and sequenced the EHC point mutation in Myh10 (Mitchell, K., PhD Thesis, University of Manchester; Clowes, C., PhD Thesis, University of Manchester). In addition, these investigators provided the first phenotypic characterisation of the EHC mouse, primarily establishing that the EHC mutant displays abnormal cardiac development. Concurrent with data from previously reported studies on the Myh10 ablated mouse (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003), it was found that EHC mutants displayed profound and widely varying cardiac defects; mutant hearts develop DORV, and show evidence of VSDs. In addition, mutants display a hypoplastic myocardium, with a number of multinucleated cardiomyocytes, indicative of cytokinesis defects. Similarly, the EHC embryo develops embryonic hydrocephalus, as reported for the Myh10 deficient mouse (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003; Ma et al., 2007). Heterozygous EHC animals are viable, and do not display a detrimental phenotype. Of particular interest, it was noted that EHC mutants present an irregular epicardial cell morphology that has not been previously described for the Myh10 ablated mouse (Mitchell, K., PhD Thesis, University of Manchester). Epicardial defects have not previously been reported or investigated in the Myh10 knock out mouse.

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6.2 Confirmation that a Mutation in Myh10 Causes the EHC Phenotype Building upon early mutation mapping studies, the present thesis has crucially established that the point mutation in Myh10 is the causative mutation of the EHC phenotype by performing a complementation test with a known Myh10 null allele. A global Myh10 knock out line, denoted at Myh10∆, was generated by deleting Myh10 exon 2 with Cre-loxP technology. Homozygous Myh10∆ mutants show embryonic lethality, with brain and cardiac malformations. We found that EHC/Myh10∆ embryos similarly displayed embryonic lethality, with strikingly similar cardiac defects to the EHC homozygous mutants. We have not been able to determine whether or not the aberrant EHC Myh10 transcript is translated; however, the failure of the EHC allele to rescue the Myh10 null phenotype categorically demonstrates that the EHC mutant does not produce wild type functional NMHC IIB. We have therefore validated that the EHC mouse is an accurate model in which to study the Myh10 null phenotype, and addressed the first aim of this research project. I will discuss potential experimentation to investigate NMHC IIB protein production in the EHC mouse in the future directions section of this thesis.

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6.3 Defective Brain Development in Myh10 Knock Out Mice The EHC embryo develops embryonic hydrocephalus, consistent with previous observations (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003; Ma et al., 2007). In line with the expertise and interests of the Hentges laboratory, we did not extensively examine the defective processes that may underpin this aspect of the EHC phenotype. However, we have previously shown by in situ hybridization that the central nervous system (indicated by Shh expression), prosenchephic and mesencephalic ventricles (indicated by Otx-2 expression), and hindbrain (indicated by En-2 expression) (Mizuseki et al., 2003), are correctly specified in the EHC mutants (Mitchell, K., PhD Thesis, University of Manchester; Stephen, L., PhD Thesis, University of Manchester, data not shown). This suggests that hydrocephalus does not arise as an inherent property of abnormal brain segment structure, and furthermore, that NMIIB is not required for specification of the brain ventricles.

Previous reports have suggested that the hydrocephalus pathology of Myh10 null mice arises following the ablation of NMIIB mediated adhesion in the neuroepithelium, in conjunction with disrupted migration of facial neurones, pontine neurones, and cerebellar granule cells (Ma et al., 2004; Ma et al., 2007). We have found that NMIIB is the only detectable NMII isoform in the neuroepithelium, indicating that NMIB plays a key role in the maintenance of neuroepithelial function (see Appendix 13). In addition, previous work in our laboratory showed that neuroepithelial cells obstruct the spinal canal of EHC mutant embryos (Mitchell, K., PhD Thesis, University of Manchester, data not shown), concurrent with the findings of Ma and colleagues (Ma et al., 2007). Interestingly, the NMII barrier can be restored by genetic replacement of NMIIB with NMIIA or a motor- impaired NMIIB hypomorph, indicating that the scaffolding function served by NMIIB in the neuroepithelium is shared with other NMII isoforms (Bao et al., 2007; Ma et al., 2007). In contrast, these experiments did not rescue disrupted neuronal migration, suggesting that impaired neurone migration is not the primary cause of the hydrocephalus phenotype. Interestingly, previous members of the Hentges group have shown that the CSF protein content is altered in the EHC embryo, although it is not clear whether or not this is a direct result of neural epithelial contamination of the spinal canal. (Mitchell, K., PhD Thesis, University of Manchester). CSF has been shown to be required for neuronal differentiation and cell proliferation (Gato et al., 2005; Miyan et al., 2006), opening the possibility that NMIIB loss of function in the developing brain may elicit the propagation of other

197 pathological mechanisms that manifest in the development of hydrocephalus. Further experimentation is required to resolve this knowledge gap.

The varying severity of brain defects presented by Myh10 ablated mice has been attributed to the influence of divergent genetic backgrounds. A spectrum of malformations varying from hydrocephalus to exencephaly has been previously reported in Apob knockout mice derived from different background strains (Homanics et al., 1995). This highlights the influence of background specific modifying genes on the manifestation of the Myh10 knock out phenotype.

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6.4 The EHC Mouse Shows Evidence of Epicardial Dysfunction Global ablation of Myh10 is known to play a role in the development of cardiac pathologies frequently observed in patients with congenital heart disease (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003). However, the existing paradigm to explain the pathologies of these defects is extremely underdeveloped. Previous work in our laboratory utilising histology and transmission electron microscopy established that the EHC heart displayed abnormal, rotund epicardial cells, suggesting that epicardial cell biology was disrupted. Numerous studies in a variety of model organisms have inextricably linked defective epicardial cell function with disrupted coronary vasculature development (Zamora et al., 2007; Mellgren et al., 2008; Hellstrom et al., 1999; Moore et al., 1999). Abnormal epicardial and coronary vessel development had not previously been reported for the Myh10 knock out mouse. By utilising immunohistochemistry, we characterised the vasculature of the EHC mutant heart at the molecular level, and conclude that whilst mutants form a primitive PECAM-1 positive endothelial plexus, they do not form mature vessels associated with positive SMαA staining of vascular smooth muscle cells. Drawing upon this, we suggest that the recruitment of vSMC is disrupted in the EHC heart. The known role of NMIIB in cell migration may be sufficient to explain this recruitment failure (Lo et al., 2004; Thomas et al., 2015). However, migration of mutant epicardial cells is unimpeded in culture conditions, suggesting that additional factors influence vSMC recruitment to the endothelial plexus. It is known that PDGF, FGF, and endothelin-1 promote vSMC recruitment (Rudijanto, 2007). Conversely, a number of recruitment inhibitors have been described, including heparin sulphate, nitric oxide, and TGF-β (Rudijanto, 2007). It is thought that the endothelial cells initiate vSMC recruitment in response to pressure changes that follow the initiation of blood flow; as NMIIB is implicated in the regulation of the molecular response to tension (Norstrom et al., 2010; Schiffhaurer et al., 2016), is it possible that NMIIB null endothelial cells cannot correctly respond to these pressure cues. Expanding upon this, it is known that NMIIB interacts with the actin cytoskeleton during the formation of VE-cadherin (vascular endothelial cadherin) adhesions; dysregulation of NMIIB contractility is associated with disruption of endothelial cell-cell adhesions, and pathological increase in microvessel permeability (Newell-Litwa et al., 2015). NMIIB may therefore function directly in the coronary endothelium.

Our group has previously shown that specification of epicardial precursor cells in the PEO and sinus venosus occurs correctly, as indicated by in situ hybridisation experiments showing correctly localised expression of Tbx18 and Shox2 respectively (Mitchell, K., PhD

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Thesis, University of Manchester, data not shown). This experiment demonstrates that Myh10 is not required for specification of epicardial precursors. However, here we provide compelling immunohistochemistry and western blot data that illustrates that NMIIB is preferentially expressed in the epicardium, and represents the predominant NMII isoform in this tissue during the peak of embryonic epicardial activity. Moreover, we show for the first time that NMIIB occupies a distinct sub-cellular localisation in cultured epicardial cells. Together, these data advocate that NMIIB plays an important and specialized role in epicardial cell function.

Expanding upon this, we have shown that the EHC epicardium displays evidence of disrupted epicardial EMT activation, and a reduced capacity to invade the underlying myocardium. These data suggest a role for NMIIB in the upstream signalling processes required for the initiation of epicardial EMT. These processes are essential for correct epicardial cell function and the formation of the coronary vasculature (reviewed in Olivey and Svensson, 2010; Peres-Pomares and de la Pompa, 2011; von Gise and Pu, 2012). We have therefore established specific aspects of epicardial function that are compromised in vivo, which are strongly suspected to underpin the vascular defects observed in the EHC mouse. A schematic model highlighting these key findings is shown in Fig 6.1. Accordingly, these findings have addressed the second aim of this research project.

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Figure 6.1: Revised Model Illustrating Defective Epicardial Function and Coronary Vessel Formation in the EHC Mouse Whilst the epicardium does form in globally Myh10 ablated mice, epicardial cells demonstrate an irregular rounded morphology. In addition, the composition of the sub- epicardial matrix is severely disrupted. The EHC heart shows a number of abnormalities in epicardial cell function, which detrimentally impact upon the formation of the coronary vasculature. (1) The EHC epicardium shows a reduction in epicardial EMT, and decreased migration of EPDCs into the myocardium. (2) Whilst EHC EPDCs can differentiate into vascular smooth muscle and cardiac fibroblasts, vascular smooth muscle cells are not recruited to developing vessel structures. (3) As a consequence, mature coronary vessels do not form in the EHC heart, and the primitive capillary plexus becomes vulnerable to rupture following the initiation of blood flow. (4) We have been unable to establish whether or not epicardial-myocardial or EPDC-myocardial reciprocal signalling is directly affected in the EHC heart, but defects in the development of the myocardium would appear to suggest so. CM – cardiomyocyte, ECM – extracellular matrix, EPDCs – epicardial-derived cells, PEO – proepicardial organ, VEC – vascular endothelial cell, VSMC – vascular smooth muscle cell.

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Surprisingly, we have shown that Myh10 ablated epicardial cells do not show migration or proliferation abnormalities in culture conditions, suggesting that the repercussions of NMIIB deficiency on epicardial cell function are entirely dependent upon the context of the epicardial extracellular environment. In agreement with this, previous work from our laboratory has demonstrated that the sub-epicardial ECM is disrupted in the EHC mutants; immunohistochemical analysis of the matrix components fibronectin and laminin, in addition to pan-glycosaminoglycan staining with alcian blue (a marker of proteoglycans), is substantially reduced in this biological substrate (Ridge et al., Manuscript submitted, PLoS Genetics). Concurrently, NMIIB has been associated with playing a key role in the secretion of important matrix components (Kalson et al., 2013), and is linked to epicardial fibronectin deposition via its regulation of the EGFR-Bves signaling pathway (Lin et al., 2007; Benesh et al., 2013; Kim et al., 2012).

The sub-epicardial matrix is known to contribute substantially to correct epicardial cell function (Riley and Smart, 2011; Olivey and Svensson, 2010; von Gise and Pu, 2012; Perez-Pomares and de la Pompa, 2011). This contribution is known to involve the generation of substrate tension, which is implicated in the regulation of EMT through mechano-transduction of activating signals (O’Connor et al., 2015; Trembley et al., 2015). The data we present here strongly suggests that NMIIB may function in a dual capacity to facilitate both the secretion of sub-epicardial ECM, and simultaneously regulate the epicardial mechanical response to substrate-induced tension. Interestingly, bioengineered ECM implants have recently shown to enhance vasculogenesis and myocardial repair following ischemic heart damage (Mewhort et al., 2016). This highlights the importance of the epicardial-ECM-myocardial molecular relationship, and further characterisation of this in the EHC mouse would help to elucidate the precise role of NMIIB during this complex signalling interplay.

The generation and continued study of tissue-specific Myh10 knock out animals will help to address outstanding questions regarding the precise role served by NMIIB in cardiogenesis, and address the third and final aim of this project. We have accrued preliminary data that implies that we have successfully generated myocardial-specific Myh10 knock out mice. Consistent with previously reported studies, we have found that these mice are viable, and do not recapitulate any of the gross morphological defects associated with the EHC phenotype (Ma et al., 2009). This demonstrates that cardiomyocyte NMIIB expression is dispensable for normal cardiogenesis, specifically in relation to the formation of the coronary vasculature and the correct development of the outflow tract and interventricular

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septum. Thus, the congenital heart defects observed in globally ablated Myh10 mice is a direct result of NMIIB loss in other cardiac cell populations. Further characterisation of the myocardial-specific Myh10 knock out mouse is required by our laboratory to identify whether or not cardiomyocytes display cytokinesis defects, as previously reported (Ma et al., 2009). However, the absence of DORV in these animals suggests that any cardiomyocyte cytokinesis defect would not be sufficient in isolation to significantly impede outflow tract myocardialisation and impact upon the positioning of the great arteries.

We have proposed the hypothesis that NMIIB is required specifically in the epicardium during mammalian cardiogenesis. It is known that epicardial-myocardial reciprocal signalling is essential for appropriate myocardial proliferation and development (reviewed in von Gise and Pu, 2012). Thus, defective epicardial cell function may therefore contribute directly to the myocardial defects that underpin the DORV and VSD observed in Myh10 disrupted mice. The generation of epicardial-specific Myh10 knock out animals is fundamental to confirm or refute this hypothesis. We have potentially generated an epicardial-specific Myh10 knock out embryo that displays abnormalities in coronary vessel formation and positioning of the great arteries. However, we are yet to determine whether epicardial NMIIB ablation has been successful, and more detailed experiments are required to characterise the cardiac defects presented in this animal at the cellular and molecular level. This validation is essential to assess the contribution of epicardial NMIIB to the formation of the coronary vessels and outflow tract development. For example, epicardial specific deletion of Wt1 has been shown to result in myocardial malformations, suggesting that epicardial-derived signals are essential for correct development of the myocardium (Martinez-Estrada et al., 2010). From this, the observed outflow tract malformations in the potential epicardial-specific Myh10 knock out embryo may be a direct consequence of loss of epicardial-derived signals that propagate myocardial proliferation; this may impact myocardialisation of the proximal outflow tract, or the generation of signalling cues required to guide the recruitment of CNCCs. Alternatively, the DORV phenotype presented by global Myh10 null animals may arise directly due to compromised CNCC migration following ablation of NMIIB in these cells. Further studies to characterise the expression of NMIIB in CNCCs will similarly help to address this outstanding question.

Additional experimentation to delineate the precise function(s) of NMIIB in the processes discussed above is outlined in the future directions of this thesis. Knowledge gained from these investigations will provide valuable insight into the mechanisms underpinning the

203 cardiac developmental programme, and ultimately pathological disease processes in the heart.

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6.5 Final Conclusion Continued effort to increase our understanding of the molecular mechanisms that underpin embryonic epicardial cell biology may facilitate the development of therapeutic strategies to repair and regenerate the injured heart in the context of both congenital and acquired cardiac disease. Indeed, the therapeutic potential of epicardial dependent cardiac regeneration in response to injury has already been demonstrated (Smart et al., 2007; Smart et al., 2011). Here, we have provided substantial evidence that NMIIB plays a crucial and multifaceted function during mammalian heart development, and we have shown a novel link between NMIIB and epicardial functionality. Our results are consistent with previously described NMIIB function in cell adhesion, migration, and cytokinesis (reviewed in Vincente-Manzanares et al., 2009; Conti and Adelstein, 2008). Moreover, we present novel data implicating NMIIB in epicardial EMT regulation, concurrent with a pleiotropic role in extracellular matrix secretion (Kalson et al., 2013), tension generation (Norstrom et al., 2010), and signal transduction (Kim et al., 2012; Schiffhauer et al., 2016).

It is also prudent to consider that a wealth of our understanding concerning the biological function of NMIIB has been gleaned from cell lines with considerable different cytoskeletal properties compared to those within a 3D tissue environment (Conti and Adelstein, 2008). Studying the effects of the EHC mutation in the context of the mammalian embryo will improve the physiological relevance of our interpretations of NMIIB function, and ultimately its role in cardiogenesis. The findings presented in this thesis have therefore illustrated that ENU mutagenesis screens are a powerful tool to identify novel disease associated mutations, and aid the unbiased elucidation of gene function in animal models, which can be subsequently extrapolated to facilitate the medical battle against human illness.

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6.6 Limitations There are a number of limitations associated with the work presented in this thesis. Primarily, experimentation with animal models of embryogenesis presents overt challenges with regard to interspecies variation and the diversification of developmental programmes. Countless mutant mouse lines that display cardiac abnormalities strongly resembling congenital heart defects in the human have been generated. Indeed, the overall anatomy and programme of development of the murine and human heart is remarkably similar (Wessels and Sedmera, 2003; Krishnan et al., 2014). However, it is not yet possible to determine whether or not the molecular processes that drive both normal and pathological heart formation are conserved between mouse and man due to the obvious ethical considerations associated with the study of human embryogenesis. This problem underpins the difficulties in translating advances in basic science into medical advances for human patients. This issue is seemingly compounded specifically in relation to the study of globally ablated Myh10 mouse models generated on different genetic backgrounds. The development of exencephaly, rather than hydrocephalus, in mice derived from the Myh10∆ line, perfectly illustrates this. A spectrum of neural tube closure defects ranging from hydrocephalus to exencephaly previously described in the Apob mutated mouse have been attributed to the influence of mixed genetic background (Homanics et al., 1995). This suggests that strain specific modifying elements may impact Myh10 expression and ultimately function, forewarning potential discrepancies between the mouse and human.

There is scarce evidence in the literature linking Myh10 to human disease, let alone congenital heart conditions. A single individual presenting with defects associated with intrauterine growth restriction, including hydrocephalus, has been officially reported (Tuzovic et al., 2013). Whole exome sequencing revealed an E908X nonsense mutation in the NMHC IIB coiled-coil rod domain, and is predicted to cause the deletion of the majority of the myosin tail region. This glutamate residue at position 908 is conserved between human, mouse, chicken and zebrafish, and the phenotypic similarity to the Myh10 ablated mouse model with regard to disrupted CNS development indicates conserved NMIIB function. However, the lack of cardiac phenotype in this patient is intriguing, and may indicate NMIIB functional redundancy in the human heart, or a divergence from the cardiac specific function of NMIIB displayed in the mouse. Alternatively, it is known that a fragment of NMII - termed heavy meromyosin and consisting of the globular head domain, a pair of light chains and a portion of the rod domain – retains enough of the myosin coiled-coil rod domain to enable dimerisation (Vincente-Manzanares et al., 2009). It

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may be that the NMHC IIB resulting from the E908X mutation retains enough of its rod domain to support dimer polymerisation; this may be sufficient to elicit adequate NMIIB function in the human heart. Indeed, recent data suggests that NMIIB monomers may represent functional forms of NMII in motile cells, circumventing the requirement to polymerise in order to evoke biological function (Shutova et al., 2014). Further biochemical studies are required to characterise the consequence of the E908X mutation on NMHC IIB protein function. Moreover, an explanation as to the absence of clinical data linking MYH10 mutations with human congenital heart defects may be implied by the severity of the NMIIB null phenotype in animal models. The embryonic lethality associated with NMIIB loss of function may dictate that de novo loss of function mutations in MYH10 are incompatible with life, resulting in spontaneous abortion of the human embryo.

We have been unable to delineate the precise molecular mechanisms that underpin the EHC phenotype. Concurrently, in spite of condensing our research focus to the epicardium, we have not yet established the precise function(s) served by NMIIB in cardiogenesis. Due to time and financial constraints, we have been unable to perform analysis that would enable the identification of differentially expressed genes in the EHC mutant heart. We have preliminary data suggesting that the expression of NMIIA is reduced in EHC mutants. This finding opens up the possibility that other transcriptional alterations are incurred by the ablation of NMIIB. Transcriptome modifications are plausible due to the implied role of NMIIB in tension sensing and the transduction of molecular signalling pathways (Lee and Nelson 2012; Tajik et al., 2016). The classical means of identifying gene expression changes in a mutant organism is to construct a DNA microarray; constructing an array containing cDNA fragments corresponding to genes known to play a role in epicardial development and function would help to determine downstream effectors of NMIIB ablation. Alternatively, performing RNA-Seq would similarly enable the profiling of RNA expression changes, but with greater transcript read accuracies, and help to elucidate the function(s) played by NMIIB in this process (Wang et al., 2009). Taking this a step further, performing this analysis on isolated epicardial cells, ideally epicardial cells isolated in situ (i.e. by FACS), rather than cultured cells, would delineate transcriptome modifications specific to the epicardium, and facilitate a greater understanding of molecular changes induced by the loss of NMIIB.

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6.7 Future Directions

6.7.1 Further Characterisation of the Myh10 Null Phenotype We have made pertinent observations that have enhanced our understanding of the processes dependent upon correct NMIIB function during mammalian embryogenesis. These findings would be complimented by further experimentation.

We have not yet established the dysfunctional processes that underpin the DORV defect observed in Myh10 ablated embryos. Initial examination of the myocardial-specific Myh10 knock out line suggest that cardiomyocyte NMIIB expression is not required for correct outflow tract development. As DORV manifestation is frequently associated with defects in outflow tract elongation, measurements of the outflow tract in EHC and Myh10∆ mutant embryos would help to determine whether or not this process is disrupted following ablation of Myh10. Expanding upon this, we have not yet assessed CNCC migration and recruitment to the outflow tract. Performing in situ hybridization experiments for neural crest markers, such as Sox9 and Twist1 (Ishii et al., 2012; Achilleos and Trainor, 2012), would identify whether neural crest specification occurs correctly in mutant embryos. Although specific markers of CNCCs are lacking (van den Hoff and Moorman, 2000), Conway and colleagues (2000) have previously used mouse-chick chimeras to track CNCC migration in vivo (Conway et al., 2000). In addition, it has been reported that Pax3 expression can serve as a marker of CNCCs in the mouse embryo (Conway et al., 1997). By utilising immunohistochemistry for Pax3, we could profile the localisation of CNCCs and ascertain whether their migration to the outflow tract is disrupted. Similarly, the differentiation of CNCCs into the smooth muscle cell components of the outflow tract could be tested by immunohistochemistry for SMαA. This would provide a foundation to ask further questions, such as; is CNCC migration directly affected in a cell autonomous manner, or in a non-cell autonomous fashion by the differential expression of CNCC recruiting factors?

As correct outflow tract development is dependent upon the population of the proximal region by proliferating cardiomyocytes, performing immunohistochemical analysis of the proliferation marker, PHH3, or markers of cell death, such as activated caspase-3 (Krysko et al., 2008), would help to establish whether or not sufficient cell populations reach this region, and further develop a clearer picture of the defective processes that underpin DORV formation in the Myh10 ablated heart.

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We are yet to identify the underlying cause of the deviation in severity of brain malformations observed between EHC and previously reported Myh10 knock out mice (Tullio et al., 1997; Uren et al., 2000; Takeda et al., 2003; Ma et al., 2007) versus mice derived from the Myh10∆ line. Experiments to determine the expression domains of Shh (floorplate and notochord marker), Otx-2 (prosencephalon and mesencephalon marker) and En-2 (hind brain marker) would help to determine if brain segment specification occurs correctly in Myh10∆ derived mice, as observed in the EHC line. This would help to identify the mechanisms underpinning the development of exencephaly that may occur correctly in the EHC mouse, and indicate the elements of NMIIB function that may be susceptible to genetic background influence.

Considering the profound effects on the formation of the coronary vasculature, it seems appropriate to investigate the formation of the cardiac lymphatic system. The lymphatics are essential to maintain interstitial fluid homeostasis, and are thought to arise from vascular endothelial cells (Yang and Oliver, 2014). However, recent data suggests that the cardiac lymphatics have a heterogeneous extracardiac cellular origin, and closely align with the coronary veins (Klotz et al., 2015). Initial immunohistochemical experiments examining the localisation of the transcription factor Prox1 - a marker of lymphatic precursor cells - would facilitate the identification of possible early defects in the formation of the lymphatics (Flaht-Zabost et al., 2014). Indeed, Prox1 disrupted mice bear a degree of phenotypic similarity to Myh10 knock out animals; the global Prox1 knock out develops severe oedema (Wigle and Oliver, 1999), and cardiac-specific Prox1 disruption results in abnormalities including reduced cardiac growth, thinning of the myocardial wall and VSDs (Risebro et al., 2009; Petchey et al., 2014).

6.7.2 Exploring Epicardial Dysfunction We have made novel discoveries in establishing that the loss of NMIIB function translates into defects in epicardial cell biology. However, the precise mechanisms that underpin epicardial dysfunction remain elusive. The experiments suggested below may offer unique insight into these dysfunctional processes.

NMIIB has been implicated in the establishment of planar cell polarity during cardiogenesis, and failure to activate NMIIB is associated with cardiac malformations (Phillips et al., 2005). Planar cell polarity signalling is critical in the epicardium, and has been shown to regulate the properties of cell adhesion (Oteiza et al., 2010). Moreover,

209 mouse knock out studies of Par3, which is involved in the internalisation of β1 integrin, have demonstrated its requirement for the correct establishment of epicardial progenitor cell polarity (Hirose et al., 2006). This suggests that correct β1 integrin localisation is critical for normal epicardial development. Moreover, the fibronectin binding α5β1 integrin heterodimer has been demonstrated to activate myosin II contractility (Schiller et al., 2013). Conversely, the cycling of integrins is closely related to Rho GTPase mediated myosin contractility (Caswell et al., 2009). Myosin contractility can promote the polymerization of fibronectin in the extracellular matrix via binding to the α5β1 integrin ((Zhong et al., 1998) cited in Caswell et al., 2009). Interestingly, Heller and colleagues (2014) have recently shown that NMIIA is required to facilitate α5β1 integrin/fibronectin mediated migration of eye lid epithelial cells during development, setting a precedent for the involvement of NMII with these adhesion molecules (Heller et al., 2014). This finding is of particular interest; α 4β 1 integrin and fibronectin are expressed in the epicardium and sub-epicardial space (Yang et al., 1995). To investigate a possible mechanism of Myh10 function in epicardial cells, it would be useful to perform antibody staining on embryonic heart tissue sections to identify whether EHC epicardial cells have altered β1 integrin localisation profiles in vivo. Further experimentation to examine the localisation of important epicardial adhesion molecules, such as E-cadherin (Mahtab et al., 2008) and VCAM-1 (Kwee et al., 1995) would help to catalogue whether or not adhesion is adversely affected in Myh10 ablated hearts.

Our finding that the EHC heart shows dramatically disrupted sub-epicardial ECM composition is especially interesting. The specific matrix components fibronectin and laminin are largely lost from this matrix layer. In addition, it appears that glycosaminoglycans (GAGs), and therefore various proteoglycan components of the ECM, are similarly less abundant in the EHC heart. It is known that disruption of the genetic pathways that underpin ECM secretion result in cardiac malformations frequently observed in human patients with congenital heart disease (Lockhart et al., 2011). Initial experiments to repeat our histological and immunohistochemical characterisation of matrix components (specifically fibronectin, laminin, and GAGs) in the Myh10∆ homozygous mutants would help to establish whether these aberrations are recapitulated in an Myh10 null model derived from a different genetic background. In addition, hyaluronan is a major GAG component of the ECM, and mice deficient in hyaluronan synthase 2 (Has2), which produces hyaluronic acid (HA), demonstrate phenotypic similarity to the EHC mutants (Lockhart et al., 2011). Moreover, expression of HA has been directly associated with

210 enabling epicardial EMT during heart regeneration in the zebrafish; suppression of HA production blocks cardiac regeneration (Missinato et al., 2015). In addition, as collagen I, III IV and VI are known to be a major constituent of chick, mouse and human sub- epicardial matrix (Kalman et al., 1995; Tidball, 1992), it would be informative to profile collagen expression. This would help to determine whether or not extracellular matrix defects are central to epicardial cell function, and guide future experiments to delineate NMIIB function in the epicardium. Expanding upon this, we have also generated preliminary data that suggests that fibronectin deposition is impeded in the epicardial cell culture system (data not shown). It would be extremely interesting to establish whether or not this preliminary observation holds true, and extends to the secretion of other key ECM components. As many of the defective epicardial cell functions reported in the EHC embryo are seemingly corrected in culture, this experiment would identify whether the secretion of matrix is dependent upon substrate tension.

The conflicting data we have from our in vivo observations and our findings from the in vitro epicardial cell culture system, strongly suggest that the function and behaviour of NMIIB ablated epicardial cells is highly reliant on the cellular microenvironment on which these cells reside. This finding should encourage other investigators to refocus their attention on the sub-epicardial matrix, and the role of tension generating proteins in embryonic epicardial cell function, which is yet to be fully explored in the literature. It is possible to physiologically assess the effects of altered matrix composition on tension within the epicardium by utilising atomic force microscopy. This technique has previously been used to investigate tissue specific tension properties, as well as generating high-resolution images of the ECM structure (Graham et al., 2010). This technique would allow us to determine if loss of NMIIB alters both the ultrastructure of the sub-epicardial ECM, and the physical stresses imposed upon the epicardium, which have important consequences in the activation of downstream molecular signalling pathways.

Expanding upon this, we have identified MRTF-A as a potential tension-sensitive candidate molecule whose activity may be compromised in the EHC heart. Additional experiments utilising an antibody against MRTF-A that does not show the non-specific staining we have reported here, would reveal whether or not the correct nuclear translocation of MRTF-A in response to EMT inducing signals is disrupted in Myh10 ablated epicardial cells. In addition, as FAK phosphorylation has been previously shown to link changes in cell contractility with the initiation of EMT gene expression in the epicardium (Lee and Nelson, 2012; Artamonov et al., 2015), it would be extremely

211 informative to examine whether levels of phosphorylated FAK are altered in the EHC epicardium.

We have not yet assessed the ability of Myh10 mutant epicardial cells to invade a 3D culture substrate. Implementation of an invasion assay based on a Boyden chamber with 3D gel substrata would permit the examination of both the ability of NMIIB null cells to both invade and migrate within a 3D substrate (Benesh et al., 2013; Veevers-Lowe et al., 2011; Rhee et al., 2009). This study could be expanded to monitor the response of mutant epicardial cells to the application of exogenous stimuli, including molecular and biophysical cues. The in vitro epicardial cell culture system provides a model of epicardial function that is amenable to physical manipulation that is not possible in the context of the embryo. It would be possible to culture control and Myh10 mutant epicardial cells on substrates with different tension properties, such as varying concentrations of polyacrylamide gels (O’Connor et al., 2015). By using this method, it may be possible to better recapitulate the in vivo sub-epicardial matrix tension properties. One could then treat the epicardial cultures with EMT inducing factors, such as TGFβ1 (Sridurongrit et al., 2008), and monitor the response of mutant epicardial cells to identify if and how this response diverges from control cells. For example, fluorescent ECM substrates have been used to monitor proteolytic degradation of ECM by matrix metalloproteinases (Artym et al., 2009), a key process during cell invasion. Furthermore, conditioned media from these experiments could be used to assess alterations in epicardial-derived signalling molecules that act to propagate myocardial development (Wei et al., 2015). These experiments may allow us to elucidate how NMIIB co-ordinates correct epicardial cell invasion and migration, as well as responsive secretions. These experiments may shed light in the processes affected by loss of Myh10 during cardiogenesis.

6.7.3 Further Defining NMIIB Function in Cardiogenesis The continued study of the tissue-specific Myh10 knock out lines described in this thesis is essential to dissect the function of NMIIB during cardiogenesis. Confirming NMIIB deletion from these specific cells and the molecular characterisation of their coronary vessel architecture will allow us to conclude precisely in which cell types NMIIB is required to correctly generate the coronary vessels and contribute to normal cardiogenesis. In addition, due to the heterogeneous nature of the epicardium, it may be necessary to generate compound epicardial-specific Myh10 lines, using supplemental epicardial specific Cre drivers (i.e. Tbx18). This would better ensure the ablation of Myh10 in the organ wide epicardium.

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We are yet to determine specific transcriptional changes that arise from loss of NMIIB in the heart. The most direct means of identifying gene expression changes that impact upon the EHC phenotype would be to characterise the transcriptome of mutant epicardial cells and compare these to stage matched controls. The technical limitations associated with the isolation of epicardial cells for analysis of this nature has prohibited our efforts in this area to date. In addition, it is possible that the physical process required to isolate epicardial cells may inadvertently alter their gene expression. One method that could possibly overcome this challenge is laser capture micro-dissection, in which individual single cells can be physically removed from a section of heterogeneous tissue and isolated for downstream analysis, including both protein and RNA transcript profiling (Golubeva et al., 2013). By using this method, one could isolate individual epicardial cells and quantify gene expression changes induced by NMIIB deletion. This would facilitate the identification of molecular pathways dependent upon NMIIB function. In addition, this technique is amenable to optimisation with immunohistochemistry, making it theoretically possible to isolate distinct sub-populations of the epicardium (Katz et al., 2012).

Finally, it would be extremely beneficial to generate a panel of Myh10 mutants to disrupt individual functions of NMIIB and identify the specific NMIIB domains required in the epicardium. We hypothesise that NMIIB plays a multifaceted role in cardiac development. By generating hypomorphs, in which mutations can modify or cause loss of function in specific protein domains without totally ablating protein function, we may be able to further define the functions of NMIIB required in cardiogenesis, and for epicardial function in particular. For instance, engineering the only disease associated human Myh10 mutation (E908X) into the mouse would allow us to perform biochemical analysis on the mutant protein, which would help to ascertain why no cardiac phenotype was observed in this patient. Similarly, emerging uses of the CRISPR-Cas9 genomic editing technology, such as CRISPRi (CRISPR interference), could be utilised to generate an allelic series of mutations in Myh10 (Qi et al., 2013). Analysis of the resultant hypomorph phenotypes would help to classify functions of NMIIB in distinct populations of cardiac cells.

The provisional experiments outlined above will help to define the function served by NMIIB during mammalian heart formation. It may be possible to therapeutically reactivate embryonic NMIIB function in the diseased heart to enhance cardiac repair and regeneration. This would represent a small but vital progressive step towards realising the aims of cardiac regenerative medicine.

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Appendices

Appendix 1: Table of Genotyping Primers

Primer Name Forward 5’-3’ Reverse 5’-3’ TGC-TAG-ATC-AGT- ATC-CAG-ATG-TAG-TGG- EHC mutation AGG-CTG-TGC TGC-ATG GGT-CAT-CCA-GAA- TGG-TCT-TCT-ACT-CTT- Myh10∆ CTG-TGT-TGT CTT-TGC CTG-ACA-TGC-TTG- CAG-AGG-CAT-GCA-GAT- Myh10 flox AAC-GAA-GC CTC-TGT GAA-CCT-GAT-GGA- AGT-GCG-TTC-GAA-CGC- Cre Recombinase* CAT-GTT-CAG-G TAG-AGC-CTG-T Cre Internal Control TTA-CGT-CCA-TCG- TGG-GCT-GGG-TGT-TAG- (Myogenin)* TGG-ACA-GC CCT-TA All primers were obtained at a stock concentration of 10nmol from Eurogentec. Primers were diluted 1:10 with ddH2O to obtain working primer mix solution. *Primer design for Cre recombinase and Myogenin primer pairs obtained from Qiagen.

Appendix 2: List of Genotyping PCR Thermal Cycler Programmes

Initial Number Final Programme Denaturing Annealing Extension Denaturing of cycles extension EHC 94°C 94°C 58°C 72°C 72°C x40 mutation (5 min) (1 min) (1 min) (1 min) (5 min) ABI 96°C 96°C 50°C 60°C x25 N/A sequencing (1 min) (30 sec) (15 sec) (4 min) 94°C 94°C 56°C 72°C 72°C Myh10∆ x40 (5 min) (45 sec) (45 sec) (3 min) (5 m 30 s) 96°C 96°C 60°C 72°C 72°C Myh10 flox x35 (5 min) (1 min) (1 min) (2 m 30 s) (5 min) 95°C 95°C 62°C 72°C 72°C Cre* x35 (5 min) (30 sec) (30 sec) (30 sec) (5 min)

*Genotyping protocol for Cre and Myogenin primer pairs courtesy of Qiagen.

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Appendix 3: Commonly Used Reagents

10x Phosphate Buffered Saline

11g Na2HPO4, 2g KH2PO4, 80g NaCl2, 2g KCl, pH to 7.4 and make up to 1L with ddH2O.

50x TAE 242g Tris, 57.1mL acetic acid, 100mL 0.5M EDTA (p.H 8.0), pH to 8.5 and make up to 1L with ddH2O.

Appendix 4: X-gal Staining Solution

100µL potassium ferrocyanide (500mM stock solution), 100µL potassium ferricyanide

(500mM stock solution), 20µL 1M MgCl2, 100µL X-gal (Bioline) in dimethylformamide (100mg/mL stock solution), 20µL 10% (v/v) Igepal, 20µL 5% (w/v) sodium deoxycholate, 200µL 1M Tris pH7.8, make up to 10mL with PBS.

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Appendix 5: Tamoxifen Dosing Regime for Wt1-CreERT2 Derived Mice

Female Wt1 Male Tamoxifen Litter Notes Number Number Administration Size Flox∆122 154 E10.5 & E11.5 7 1 x (Wt1-CreERT2;flox/∆) Flox∆129 154 E10.5 & E11.5 3 All dead and resorbing Flox∆128 154 E10.5 & E11.5 4 All dead and resorbing 3 resorbed, 1x (Wt1- Wt1 242 227 E10.5 & E12.5 6 CreERT2;flox/flox), 1x (Wt1-CreERT2;flox/∆), 1x (+/+∆/∆) 2 resorbed, Flox∆ 139 154 E10.5 & E12.5 5 1x (Wt1-CreERT2;flox/∆) Not Wt1 218 150 E10.5 & E12.5 N/A Pregnant Not Flox∆ 130 150 E10.5 & E12.5 N/A Pregnant 1x (Wt1- CreERT2;flox/flox), 1x Flox∆ 125 155 E10.5 & E11.5 5 (Wt1-CreERT2;flox/∆) *bleeding Not Flox∆ 126 155 E10.5 & E11.5 N/A Pregnant Flox∆ 142 154 N/A Swollen uterus Flox∆ 170 258 E10.5 & E12.5 N/A All resorbed, fibrous uterus Flox∆ 171 258 E10.5 & E12.5 2 Both dead, 1 resorbed Not Flox∆ 172 258 E10.5 & E12.5 N/A Pregnant Flox∆ 175 154 E10.5 & E12.5 N/A All resorbed, fibrous uterus Flox∆ 178 154 E10.5 & E12.5 1 Recently dead Not Wt1 268 150 E10.5 & E12.5 N/A Pregnant Not Wt1 269 150 E10.5 & E12.5 N/A Pregnant Wt1 301 150 Non-injected 3(?) Approx. E6.5 Resorbed, uterus swollen Flox∆157 306 E11.5 (sesame oil) 2 with fluid Wt1 298 306 Non-injected N/A Uterus filled with fluid

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Appendix 6: List of Primary Antibodies

Product Application Target Type Supplier Code (dilution) Mouse β-actin-HRP Sigma A3854 WB (1:200,000) monoclonal Rabbit Santa Cruz MRTF-A SC-32909 Myh10∆ ICC (1:50) polyclonal Biotechnology Rabbit NMHC IIA Biolegend PRB-440P polyclonal Myh10∆ IHC (1:500) Rabbit NMHC IIB Biolegend PRB445P Myh10∆ ICC (1:500) polyclonal Myh10∆ WB (1:1000) Rabbit NMHC IIC Biolegend PRB-444P polyclonal EHC and Myh10∆ PECAM-1 Rat Ab Serotec MCA2388GA whole heart IHC (CD31) monoclonal (1:1000) Phospho- Rabbit EHC cryosections Merk Millipore 06-570 histoneH3 polyclonal IHC (1:300) Smooth EHC and Myh10∆ Mouse Muscle α- Sigma A5228 paraffin sections IHC monoclonal Actin (1:400) Goat EHC cryosections Snail Abcam Ab53519 polyclonal IHC (1:100) Rabbit EHC cryosections Vimentin Proteintech 10366-1-AP polyclonal IHC (1:50) EHC and Myh10∆ Rabbit Santa Cruz Wt1 SC-192 epicardial culture ICC polyclonal Biotechnology (1:300) Rabbit EHC cryosections Wt1 Calbiochem CA1026 polyclonal IHC (1:300) Rabbit Myh10∆ epicardial ZO-1 Invitrogen 40-2300 polyclonal culture ICC (1:100)

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Appendix 7: List of Secondary Antibodies and Amplification Reagents

Name Supplier Product Code ABC reagent Vector PK-6100 Biotinylated goat anti- rat Vector BA-9400 Biotinylated goat anti- Vector BA-1000 rabbit Biotinylated goat anti- Vector BA-9200 mouse Biotinylated horse anti- Vector BA-9500 goat Donkey anti- rabbit-HRP Santa Cruz Biotechnology SC-2313 Goat anti- rabbit-FITC Sigma F9887 Streptavidin- Invitrogen S11223 AlexaFluor488 Streptavidin-Cy3 GE Healthcare PA43001 Streptavidin-Cy5 GE Healthcare PA45001

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Appendix 8: Recipes for Western Blotting Solutions

10x Running Buffer

30.3g Tris, 144g glycine, 10g SDS, make up to 1L with ddH2O.

10x Transfer Buffer

30.3g Tris, 144g glycine, 20g SDS, pH to 8.3 and make up to 1L with ddH2O.

10x Tris Buffered Saline

88g NaCl2, 30.3g TRIS, pH to 7.4 and make up to 1L with ddH2O.

TBS-T TBS, 0.05% (v/v) Tween-20.

12% Acrylamide Resolving Gel

6mL ddH2O, 4.95mL 30% acrylamide, 3.75mL 1.5M Tris (pH 8.8), 150µL 10%(w/v) SDS, 150µL 10%(w/v) ammonium persulphate (APS), 50µL TEMED.

Acrylamide Stacking Gel

2.8mL ddH2O , 850µL 30% acrylamide, 1.25mL 0.5M Tris (pH 6.8), 50µL 10%(w/v) SDS, 50µL 10%(w/v) APS, 50µL TEMED.

Epicardial Extraction Buffer 10mL PBS, 1 protease inhibitor tablet (Roche, 04 693 159 001).

4x Sample Loading Buffer 40% (v/v) glycerol, 4(w/v) SDS, 200mM Tris pH6.8, 0.04% (w/v) bromophenol blue, 10% (v/v) β-mercaptoethanol.

RIPA Buffer 100µL triton X-100, 1µL Igepal, 1 protease inhibitor tablet (Roche, 04 693 159 001), make up to 10mL with ddH2O.

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Appendix 9: Table of Protein Extraction Volumes Used

RIPA Buffer Volume for SDS-PAGE Sample Tissue Type Protein Extraction Volume (per lane) E14.5 Ventricles 100µL/heart 15µL (+5µL 4x SLB) E10.5-12.5 Ventricles 50µL/heart 15µL (+5µL 4x SLB) Explants 50µL/genotype 10µL (+3.4µL 4x SLB) Enriched Epicardial Cells Resuspend in 50µL 4x SLB 15µL Adult Lung 200µL/2mm3 sample 15µL (+5µL 4x SLB)

Appendix 10: Peak Intensity Normalisation Calculation

푃푒푎푘 퐼푛푡푒푛푠푖푡푦 (푃푒푎푘 표푓 퐼푛푡푒푟푒푠푡) 푅푒푙푎푡푖푣푒 퐴푏푢푛푑푎푛푐푒 (%) = 푋 100 푃푒푎푘 퐼푛푡푒푛푠푖푡푦 (훽−푎푐푡푖푛)

Appendix 11: Table of Western Blot Densitometry Analysis Relative Abundance of NMII Isoforms in Different Heart Tissues

NMII Peak Volume β-actin Peak Volume Relative Abundance Tissue Type (AU) (AU) (%) A 2,782,150 3,078,565 90.37 Whole Heart B 3,239,196 1,374,916 235.59 (+/+ or ∆/+) C 97,776 5,708,520 1.71 A 956,830 2,935,345 32.59 Whole Heart B 0 1,502,064 0 (∆/∆) C 0 3,510,710 0 A 2,905,610 7,986,598 36.38 Explant B 4,052,396 2,984,216 135.79 (+/+ or ∆/+) C 3,451,175 8,175,927 42.21 A 639,064 7,190,422 8.89 Explant B 0 2,583,525 0 (∆/∆) C 0 6,643,128 0 Epicardial A 0 7,952,337 0 Cells B 260,208 7,653,420 3.39 (+/+ or ∆/+) C 0 5,908,912 0 Epicardial A 0 1,918,361 0 Cells B 0 2,799,312 0 (∆/∆) C 0 1,633,824 0

Control samples highlighted in blue. AU – Arbitrary Units

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Appendix 12: PCR Genotyping Analysis of Myh10 Exon 2 Deletion Genotyping results for control, Myh10∆ heterozygous and Myh10∆ homozygous animals. (A) Wild type animals (+/+) generate a single PCR product of approximately 1.1Kb. Homozygous Myh10∆ animals (∆/∆) generate a single band of approximately 600bp. The reduction in amplicon size is due to the deletion of the Myh10 exon 2 chromosomal sequence. As expected, heterozygous Myh10∆ animals (∆/+) generate both wild type and deleted PCR products. An additional band of approximately 1.3Kb representing the non- deleted floxed Myh10 chromosome is also generated (arrow). (B) Schematic representation showing the Myh10 exon 2 locus, mapped primer binding sites and approximate amplicon sizes in wild type, (C) floxed non-deleted, and (D) floxed deleted (Myh10∆) alleles.

~1.1Kb F primer R primer

B Myh10 exon 2

~1.3Kb F primer R primer

C Myh10 exon 2 loxP loxP

~600bp F primer R primer

D loxP

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Appendix 13: Expression of NMII Isoforms in the Brain and Descending Aorta of Myh10∆ Heterozygous Animals (A-C) NMII immunohistochemistry on paraformaldehyde-fixed transverse sections of E14.5 brain and (D-F) descending aorta from Myh10∆ heterozygous (∆/+, n=3) animals. NMIIB is the only detectable isofrom in the neuroepithelium (B, arrowheads), and the aorta (E, arrowheads). The punctate areas of intense staining (D, F, arrows) are autofluorescent erythrocytes. Scale bars = 100µm. DAo – descending aorta, E – esophagus, nt – neural tube.

Appendix 14: Attached DVD Movie files from the scratch wound imaging analysis (described in Chapter 4) are contained on the attached DVD. Individual files are labelled with their respective genotype. The coloured lines represent the tracking data for 10 individual cells in each field of view, which was subsequently interrogated to generate average cell migration speeds and directional persistence. The image files correspond to the still images depicted in Fig 4.5. Additional video files are available upon request.

Appendix 15: Myh10 Exon 2 Deletion in ‘First Generation’ α-MHC- Cre/+; flox/flox Adult Ear Punches (A) PCR analysis of Myh10 exon 2 deletion in DNA samples from the heart ventricles and (B) ear punches of α-MHC-Cre/+; flox/flox adult animals. The presence of the smaller PCR product in the ear punch DNA samples confirms non-specific Cre activity in this tissue. The Myh10 exon 2 deletion was also shown to occur in brain, liver and tail samples of α-MHC-Cre/+; flox/flox E14.5 embryos (data not shown, n>3). A tail sample from a heterozygous Myh10∆ (∆/+) animal was used as a positive control to compare band sizes.

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Appendix 16: Myh10 Exon 2 Deletion in ‘Second Generation’ α-MHC- Cre/+; flox/flox Embryonic Heart Ventricles PCR analysis of Myh10 exon 2 deletion in DNA samples prepared from the heart ventricles of α-MHC-Cre/+; flox/flox E18.5 embryos (n=3). The presence of the smaller PCR product confirms the specific Cre-mediated Myh10 exon 2 deletion in this tissue. The smaller PCR amplicon was not seen in Cre negative controls (+/+; flox/flox) (data not shown). The failure to detect the non-deleted band from non-myocytes cells may be due to the limited number of these cells in the heart at this developmental stage. A tail sample from a heterozygous Myh10∆ (∆/+) animal was used as a positive control to compare band sizes.

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Appendix 17: Preliminary Molecular Characterisation of the Coronary vasculature of ‘Second Generation’ Cardiomyocyte- Specific Myh10 Knock Out Animals (A) Whole mount PECAM-1 immunohistochemistry of control (+/+; flox/flox, n=2) and (B-C) myocardial-specific Myh10 knock out (α-MHC-Cre/+; flox/flox, n=2) E16.5 hearts. Vascular endothelial cells are organised into long vessel structures that traverse the ventricular surface in both control and mutant hearts (A, B, arrowheads). Detectable PECAM-1 staining delineates large, mature coronary vessels on the dorsal surface on the mutant heart (C, dashed line). Scale bars = 50µm.

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