Medicine, Faculty of
Medicine, Department of
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UBCV
Coulombe, Patrick
Doctor of Philosophy - PhD
University of British Columbia
Embryonic development is a highly coordinated process during which cells undergo regulated division, differentiation, and migration to ultimately shape specialized tissues that perform particular functions. Throughout this course, cell fate is driven by intrinsic cell decisions in response to microenvironmental cues from their surroundings and understanding what governs these processes is at the core of developmental biology. Aberrant expression of essential developmental genes typically elicits apparent embryonic defects due to disruption of normal developmental processes. Thus, investigation of the resulting phenotype in animal models provides valuable insights into their role and overall biological pathways driving organogenesis. The work presented in this dissertation investigates the role of such developmental genes and uncovers significant signaling interactions involving endothelial cells to promote cell specification and maturation of non-vascular cells. First, this study explores the contribution of Sash1 during organogenesis. Examination of Sash1-knockout mouse embryos revealed cell non-autonomous functions in regulating both prenatal lung development and embryonic hematopoiesis. On one hand, endothelial Sash1 signaling in the lung promoted nitric oxide production which plays a critical role in the maturation of alveolar epithelial cells prior to birth. On the other hand, Sash1 expression in non-endothelial cells contributed to the emergence of hematopoietic stem and progenitor cells during the endothelial-to-hematopoietic transition (EHT) in the early embryo. Our findings suggest that Sash1 in the microenvironment impacts the progression of specified hemogenic endothelial cells into hematopoietic clusters and the work presented in chapter 4 identifies multiple candidate cell types as the origin of Sash1 signaling. In addition, early specification of a hemogenic endothelium distinct from its arterial counterpart is a critical event at the origin of EHT. Therefore, the last part of the study explored factors influencing endothelial cell fate and provided evidences to support a role for the developmental gene Meis1 in driving early hematopoietic commitment of the endothelium while showing that loss of its expression significantly impairs EHT. Altogether, the data presented in this dissertation highlights critical functions for Sash1 and Meis1 in embryogenesis and showcases examples of communication from, to, and within endothelial cells to influence cell fate in organogenesis.
ENDOTHELIAL REGULATION IN CELL FATE SPECIFICATION DURING EMBRYONIC DEVELOPMENT OF THE LUNG AND THE HEMATOPOIETIC SYSTEM by PATRICK COULOMBE B.Sc. McGill University, 2013 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Experimental Medicine) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) March 2021 © Patrick Coulombe, 2021 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Endothelial regulation in cell fate specification during embryonic development of the lung and the hematopoietic system submitted by Patrick Coulombe in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Experimental Medicine Examining Committee: Dr. Aly Karsan, Pathology Supervisor Dr. Connie Eaves, Medical Genetics Supervisory Committee Member Dr. Kelly McNagny, Medical Genetics Supervisory Committee Member Dr. Fabio Rossi, Medical Genetics University Examiner Dr. Peter Zandstra, Medical Genetics University Examiner Additional Supervisory Committee Members: Dr. Keith Humphries, Medical Genetics Supervisory Committee Member iii ABSTRACT Embryonic development is a highly coordinated process during which cells undergo regulated division, differentiation, and migration to ultimately shape specialized tissues that perform particular functions. Throughout this course, cell fate is driven by intrinsic cell decisions in response to microenvironmental cues from their surroundings and understanding what governs these processes is at the core of developmental biology. Aberrant expression of essential developmental genes typically elicits apparent embryonic defects due to disruption of normal developmental processes. Thus, investigation of the resulting phenotype in animal models provides valuable insights into their role and overall biological pathways driving organogenesis. The work presented in this dissertation investigates the role of such developmental genes and uncovers significant signaling interactions involving endothelial cells to promote cell specification and maturation of non-vascular cells. First, this study explores the contribution of Sash1 during organogenesis. Examination of Sash1-knockout mouse embryos revealed cell non-autonomous functions in regulating both prenatal lung development and embryonic hematopoiesis. On one hand, endothelial Sash1 signaling in the lung promoted nitric oxide production which plays a critical role in the maturation of alveolar epithelial cells prior to birth. On the other hand, Sash1 expression in non-endothelial cells contributed to the emergence of hematopoietic stem and progenitor cells during the endothelial-to-hematopoietic transition (EHT) in the early embryo. Our findings suggest that Sash1 in the microenvironment impacts the progression of specified hemogenic endothelial cells into hematopoietic clusters and the work presented in chapter 4 identifies multiple candidate cell types as the origin of Sash1 signaling. In addition, early specification of a hemogenic endothelium distinct from its arterial counterpart is a critical event at the origin of EHT. Therefore, the last part of the study explored factors influencing endothelial cell fate and provided evidences to support a role for the developmental gene Meis1 in driving early hematopoietic commitment of the endothelium while showing that loss of its expression significantly impairs EHT. Altogether, the data presented in this dissertation highlights critical functions for Sash1 and Meis1 in embryogenesis and showcases examples of communication from, to, and within endothelial cells to influence cell fate in organogenesis. iv LAY SUMMARY How mammals grow from a single fertilized egg into a complex organism is a remarkable feat of life. To build the variety of functional organs that compose our body, each cell needs to make a series of decisions that results in a specific fate. To understand what controls these decisions, we investigated the function of specific genes required during development. In this study, we show that lung maturation is supported by signals coming from endothelial cells (cells that form blood vessels) prior to birth. Some endothelial cells are also known to be capable of giving rise to blood cells in the early embryo. Here, we demonstrated that this decision to become blood cells is influenced by cues that originate both within the cells and from its surrounding. Altogether, the data presented in this dissertation reveals critical factors influencing a cell’s destiny towards the proper generation of several functional organs. v PREFACE All experiments and data analysis conducted for this study together with all written work and figures presented in this thesis are the original work of Patrick Coulombe (P.C.), with the following exceptions mentioned below. The work presented in Chapter 3 has been published in Cell Reports (full citation below) as part of a collaborative study. • Coulombe P, Paliouras GN, Clayton A, Hussainkhel A, Fuller M, Jovanovic V, Dauphinee S, Umlandt P, Xiang P, Kyle AH, Minchinton AI, Humphries RK, Hoodless PA, Parker JDK, Wright JL, Karsan A. Endothelial Sash1 Is Required for Lung Maturation through Nitric Oxide Signaling. Cell Rep. 2019 May 7;27(6):1769-1780.e4. doi: 10.1016/j.celrep.2019.04.039. PMID: 31067462. For this publication, P.C. wrote the manuscript and performed the experiments and analyses for the data presented in figures 2E, 3C-D, 5C, 5E, 6C-D, 7A-B, 7D-G, and supplemental figures S3B-E, S4, S6, and S7. In addition, P.C. analyzed and interpreted the data presented in figures 3A-B, 4B-I, and S3A. The graphical abstract was created by P.C. using Biorender.com. Some additional parts of this collaborative publication were included in Chapter 3 of this dissertation to assist with the flow of the overall thesis. Megan Fuller (M.F.) and P.C. both contributed to the immunostaining and quantification of CD31 in Fig. 3.1C-E and Jenny Li helped with plotting the analysis for Fig. 3.1E. Dr. Grigorios Paliouras (G.P.) collected the tissue samples for electron microscopy (Fig. 3.1F) that were submitted to Derrick Horne at the UBC Bioimaging facility. The micrographs were analyzed by P.C. thanks to the guidance of Dr. David Walker. G.P. performed the quantification of surfactant expression in Fig. 3.2A (replotted by P.C. for the manuscript). G.P. collected the cells used in the microarray experiment (Fig. 3.2F, 3.3) and submitted the samples to The Centre for Applied Genomics at The Hospital for Sick Children (Toronto, Canada) who performed the microarray. The microarray data was analyzed in Gene Set Enrichment Analysis (GSEA) by P.C. and Dr. Jeremy Parker. vi M.F. performed immunostaining and quantification of NO in WT and Sash1-/- lungs (Fig. 3.5A, B). M.F. also performed intraperitoneal injections for in vivo DAR4M-AM and SNAP treatment required in Fig. 3.5 and Fig. 3.7D, E. Oral gavage of Riociguat for the experiment in Fig. 3.7F, G was performed by Animal Resource Centre staff at the BC Cancer Research Centre. Ashley R. Clayton designed and performed the experiment in Fig 3.8A. Shauna Dauphinee (S.D.) created the constructs used in Fig. 3.8B. Sukhbir Manku cloned the mutant constructs used in Fig. 3.8D from vectors previously generated in the lab by S.D. The work in presented in Chapter 4 was performed by P.C., in some cases assisted by Katherine Bilan (K.B.) during her coop term in the Karsan lab. K.B. and P.C. both contributed to the immunostaining in Fig. 4.1A and the quantification of Sash1 excision in Fig. 4.9B. In addition, K.B. performed LacZ staining and took the image shown in Fig. 4.7A. K.B. also helped with genotyping of the embryos used for various experiments in chapter 4. The Biomedical Research Centre sequencing core at the University of British Columbia (Vancouver, British Columbia) performed the library preparation and sequencing of the scRNA-seq and CITE-seq samples. Derek Tam helped with the pre-processing of this data. All analyses and figures derived from the scRNA-seq/CITE-seq data are the work of P.C. In Chapter 5, the ChIP-seq dataset used for the analysis in Fig 5.3 was published by: • Wilson NK, Foster SD, Wang X, Knezevic K, Schütte J, Kaimakis P, Chilarska PM, Kinston S, Ouwehand WH, Dzierzak E, Pimanda JE, de Bruijn MF, Göttgens B. Combinatorial transcriptional control in blood stem/progenitor cells: genome-wide analysis of ten major transcriptional regulators. Cell Stem Cell. 2010 Oct 8;7(4):532-44. doi: 10.1016/j.stem.2010.07.016. PMID: 20887958. The schematics presented in this dissertation (Fig. 1.1, 1.2, 3.2C, 3.9, 4.3A, 4.13, and 5.8) were created using BioRender.com. Maggie Ling drew the embryo in Fig. 4.5E. All experiments involving mice were approved by UBC Animal Care Committee (protocols A16-0082, A18-0156, and A20-0116) vii TABLE OF CONTENTS ABSTRACT ................................................................................................................................... iii LAY SUMMARY .......................................................................................................................... iv PREFACE ....................................................................................................................................... v TABLE OF CONTENTS .............................................................................................................. vii LIST OF TABLES .......................................................................................................................... x LIST OF FIGURES ....................................................................................................................... xi LIST OF ABBREVIATIONS ...................................................................................................... xiii ACKNOWLEDGEMENTS .......................................................................................................... xv DEDICATION ............................................................................................................................. xvi CHAPTER 1: INTRODUCTION ................................................................................................... 1 1.1 Brief introduction to embryology ..................................................................................... 1 1.2 Vascular development ...................................................................................................... 2 1.3 Endothelial contribution during lung development .......................................................... 3 1.4 Endothelial cell heterogeneity and specification .............................................................. 5 1.5 Embryonic hematopoiesis and EHT ................................................................................. 6 1.5.1 Definitive hematopoiesis and hematopoietic stem cells ........................................... 7 1.5.2 Endothelial-to-hematopoietic transition.................................................................... 8 1.5.3 Defining the hemogenic endothelium ....................................................................... 9 1.5.4 Transcriptional regulation of EHT .......................................................................... 10 1.5.5 Inflammatory signaling and microenvironmental factors influencing EHT ........... 11 1.5.6 The endothelium as a continued support for hematopoiesis ................................... 13 1.6 Nitric Oxide .................................................................................................................... 14 1.7 Sash1 .............................................................................................................................. 15 1.8 Meis1 .............................................................................................................................. 16 1.9 AIM OF THE STUDY ................................................................................................... 17 viii CHAPTER 2: METHODOLOGY ................................................................................................ 19 2.1 Mouse strains and embryo collection ............................................................................. 19 2.2 Genotyping ..................................................................................................................... 20 2.3 Cell culture ..................................................................................................................... 21 2.4 Flow cytometry .............................................................................................................. 21 2.5 mRNA expression analysis (RT-qPCR and ddPCR) ..................................................... 22 2.6 Protein expression analysis ............................................................................................ 24 2.7 Immunostaining .............................................................................................................. 25 2.8 LacZ Staining ................................................................................................................. 27 2.9 Nitric oxide rescue experiments ..................................................................................... 27 2.10 Single-cell RNA-sequencing and CITE-seq .................................................................. 27 2.11 Data analysis .................................................................................................................. 28 2.11.1 Pathway Analysis .................................................................................................... 28 2.11.2 Single-cell data........................................................................................................ 28 2.11.3 Statistical analyses .................................................................................................. 29 CHAPTER 3: Endothelial Sash1 signaling contributes to lung maturation via nitric oxide secretion ........................................................................................................................................ 30 3.1 Introduction .................................................................................................................... 30 3.2 Results ............................................................................................................................ 31 3.2.1 Loss of Sash1 does not significantly affect the lung vasculature ........................... 31 3.2.2 Endothelial-deletion of Sash1 impairs alveolar maturation .................................... 33 3.2.3 Sash1 signaling contributes to nitric oxide production in the developing lung ...... 36 3.2.4 A nitric oxide donor can rescue the Sash1-knockout phenotype ............................ 42 3.2.5 Sash1 interacts with β-Arrestin1 to activate the Akt-eNOS pathway ..................... 44 3.3 Discussion ...................................................................................................................... 46 ix CHAPTER 4: Sash1-deletion affects the emergence of hematopoietic stem cells during the endothelial-to-hematopoietic transition ........................................................................................ 48 4.1 Introduction .................................................................................................................... 48 4.2 Results ............................................................................................................................ 49 4.2.1 Constitutive loss of Sash1 reduces intra-aortic hematopoietic cluster formation during EHT ............................................................................................................................ 49 4.2.2 Sash1 is required for cluster emergence, not early specification of the hemogenic endothelium ........................................................................................................................... 53 4.2.3 Sash1 supports EHT progression via a cell non-autonomous mechanism ............. 57 4.2.4 Identification of candidate pathways to explain the blockade of HE cells during EHT ................................................................................................................................. 67 4.3 Discussion ...................................................................................................................... 73 CHAPTER 5: Identification of Meis1 as an early driver of hemogenic endothelium commitment....................................................................................................................................................... 77 5.1 Introduction .................................................................................................................... 77 5.2 Results ............................................................................................................................ 78 5.2.1 Characterization of CITE-seq cell populations identifies Meis1 as differentially expressed in pre-hemogenic endothelial cells ....................................................................... 78 5.2.2 Meis1 expression enriches for the hemogenic subset of endothelial cells in vivo .. 84 5.2.3 Endothelial-deletion of Meis1 impairs EHT initiation............................................ 89 5.3 Discussion ...................................................................................................................... 93 CHAPTER 6: SUMMARY, PERSPECTIVES, AND SIGNIFICANCE ..................................... 96 6.1 Endothelial Sash1 in the developing lung ...................................................................... 96 6.2 Regulation of the endothelial-to-hematopoietic transition ............................................. 98 6.3 Sash1 contribution to embryonic development ............................................................ 103 REFERENCES ........................................................................................................................... 107 x LIST OF TABLES Table 2.1 List of Genotyping Primers ....................................................................................... 21 Table 2.2 List of FACS antibodies ............................................................................................. 22 Table 2.3 List of qPCR/ddPCR primers ................................................................................... 24 Table 2.4 List of immunostaining antibodies ........................................................................... 26 xi LIST OF FIGURES Figure 1.1 Timeline of hematopoietic development during embryogenesis in mice ............... 8 Figure 1.2 Schematic of IAHC formation in the AGM of E10.5 mouse embryos ................. 13 Figure 3.1 Sash1-deletion does not significantly affect pulmonary vascularization ............. 32 Figure 3.2 Loss of endothelial Sash1 impairs differentiation of alveolar epithelial cells ..... 35 Figure 3.3 Identification of Akt and nitric oxide pathways downregulation in Sash1-/- lung endothelial cells ........................................................................................................................... 37 Figure 3.4 eNOS-null embryos showed reduced surfactant transcript prior to birth.......... 38 Figure 3.5 Endothelial Sash1-deletion reduces nitric oxide secretion in lungs in vivo ......... 40 Figure 3.6 Endothelial Sash1-deletion decreases NO level in alveolar epithelial cells ......... 41 Figure 3.7 NO and cGMP activation can rescue the differentiation phenotype in Sash1-deficient embryos ........................................................................................................................ 43 Figure 3.8 Sash1 activates the Akt-eNOS pathway via interaction with beta-arrestin1 ...... 45 Figure 3.9 Schematic summary of Sash1 pathway in lung maturation ................................. 47 Figure 4.1 Fewer pre-HSPC emerge from Sash1-deficient embryos at E10.5 ...................... 51 Figure 4.2 Reduced expression of EHT-related genes in Sash1-/- AGM at E10.5 ................. 52 Figure 4.3 Ex vivo culture showed hemogenic potential in Sash1-/- E9.5 endothelial cells .. 55 Figure 4.4 Increase number of non-IAHC hemogenic endothelial cells in Sash1-/- embryos56 Figure 4.5 Endothelial-deletion of Sash1 does not impair pre-HSPC .................................... 59 Figure 4.6 Detection of NO in E10.5 AGM cells....................................................................... 60 Figure 4.7 LacZ staining for detection of Sash1-expressing cells ........................................... 63 Figure 4.8 Identification of Sash1-expressing cells by single-cell RNA-sequencing ............. 64 Figure 4.9 VE-Cadherin driven Cre excised Sash1 in blood cells present at E10.5 ............. 66 xii Figure 4.10 Single-cell RNA-sequencing and clustering of CD31+ cells ................................ 69 Figure 4.11 Comparison between WT and Sash1-/- (KO) cells in scRNA-seq ....................... 70 Figure 4.12 Differential gene expression analysis in WT vs Sash1-/- EHT cells .................... 72 Figure 4.13 Model for Sash1 role in EHT progression ............................................................ 76 Figure 5.1 CITE-seq analysis of CD31+ cells using EHT-related surface markers .............. 79 Figure 5.2 Transcriptional differences between pre-HE and the vascular endothelial subset....................................................................................................................................................... 82 Figure 5.3 Meis1 targets hematopoietic-related genes in the pre-HE and HE/pro-HSPC ... 83 Figure 5.4 Meis1 expression in the dorsal aorta of E10.5 embryos ........................................ 87 Figure 5.5 Meis1 expression enriches for the hemogenic endothelium in vivo ..................... 88 Figure 5.6 Endothelial-deletion of Meis1 reduces hemogenic potential of the dorsal aorta endothelium ................................................................................................................................. 91 Figure 5.7 Loss of Meis1 impairs pre-HSPC emergence ......................................................... 92 Figure 5.8 Meis1 drives the endothelium towards a hematopoietic fate ................................ 95 xiii LIST OF ABBREVIATIONS AE1C Alveolar epithelial type 1 cells AE2C Alveolar epithelial type 2 cells AGM Aorta-gonad-mesonephros bFGF Basic fibroblast growth factor BPD Bronchopulmonary dysplasia cDNA Complimentary DNA CFU Colony-forming unit ChIP-seq Chromatin immunoprecipitation sequencing CITE-seq Cellular Indexing of Transcriptomes and Epitopes by sequencing cKO Conditional-knockout DAPI 4',6-diamidino-2-phenylindole DAR-4M Diaminorhodamine-4M ddPCR Droplet digital PCR DEG Differentially expressed genes DMSO Dimethyl sulfoxide DNA Deoxyribonucleic acid DNase Deoxyribonuclease E Embryonic day EC Endothelial cells EHT Endothelial-to-hematopoietic transition eNOS Endothelial nitric oxide synthase Epo Erythropoietin FACS Fluorescence-activated cell sorting FBS Fetal bovine serum FDG fluorescein Di-β-D-Galactopyranoside G-CSF Granulocyte-colony stimulating factor GFP Green fluorescence protein gMFI Geometric mean fluorescence intensity GSEA Gene set enrichment analysis xiv HE Hemogenic endothelial HMEC Human microvascular endothelial cells HSC Hematopoietic stem cells HSPC Hematopoietic stem and progenitor cells IAHC Intra-aortic hematopoietic cluster IL-3 Interleukin 3 KO Knockout LPS Lipopolysaccharide Meis1 Myeloid ecotropic viral integration site 1 mRNA Messenger RNA NO Nitric oxide OCT Optimal cutting temperature PBS Phosphate buffer saline PCR Polymerase chain reaction qPCR Quantitative PCR RNA Ribonucleic acid RNase Ribonuclease RT-qPCR Reverse transcriptase quantitative PCR Sash1 SAM and SH3 containing protein 1 SCF Stem cell factor scRNA-seq Single-cell RNA sequencing sGC soluble guanylate cyclases shRNA Short hairpin RNA SNAP S-Nitroso-N-Acetyl-D,L-Penicillamine Sp Surfactant-associated protein TEM Transmission electron microscopy TF Transcription factor UGR Urogenital ridge UMAP Uniform Manifold Approximation and Projection VEC Vascular endothelial cadherin WT Wildtype xv ACKNOWLEDGEMENTS First, I would like to express my sincere gratitude to my supervisor, Dr. Aly Karsan, for giving me the opportunity to pursue this journey in your laboratory. Thank you for being supportive and staying optimistic throughout the process. I would also like to thank the members of my supervisory committee, Drs. Connie Eaves, Keith Humphries, and Kelly McNagny; all of whom I hold in high esteem. You always knew the right questions to ask and your comments are always enlightening. I had the privilege to work with many wonderful people during my time in Karsan lab and you shall not be forgotten. A special thank goes to the former technicians Patricia Umlandt, Megan Fuller, Kristy Dockstader, and Grace Cole whose help with the maintenance of the animal colonies was truly invaluable. I am also thankful to Deborah Deng and former lab members Linda Chang, Jeffrey Lam, and Ping Xian for your helpfulness and generous guidance, you all make a positive impact around you. I would like to acknowledge the coop students that I had the pleasure to mentor, Katherine Bilan and Maggie Ling. Thank you for your hard work, curiosity, and willingness to learn. I wish you both success in your respective future steps. While having supportive and inspiring co-workers makes the journey somewhat easier at times, friends are what makes it enjoyable and unforgettable. To Angela Mo, Patrick Lac, Samantha Mar, Xuan Wang, and Aparna Gopal, I am grateful to have shared this journey with you all, both inside and outside of the lab. To Eva Yap, I have been blessed with your friendship since our first day in the Karsan lab and I could always count on you, thank you for everything. Finally, to all my family who, even from thousands of miles away, will always be there for me: thank you for being an important part of my life. Knowing that you are always there to support me is what allows me to pursue my dreams and that means the world to me. Financial support was provided by a Graduate Student Award from the Centre for Blood Research, a UBC Four Year Doctoral Fellowship, and a Doctoral Research Award from the Canadian Institutes of Health Research. xvi DEDICATION To Justin and Philippe … So young and already having a big impact, you deserve a better world 1 CHAPTER 1: INTRODUCTION 1.1 Brief introduction to embryology Biology is the study of life and stems from the fundamental questions of understanding who we are and where we come from. But in addition to more philosophical motivations, developmental biology, and embryology, is the study of the building blocks of every organ and organ system that make up our bodies. Understanding how each element is formed and functions under normal conditions is the first step to identify what is “abnormal” during disease states. Furthermore, understanding the mechanisms driving normal development can provide the key to correcting dysregulated pathways or cell behaviors that are often seen in congenital disorders, cancer, and other diseases. Studying human development comes with several challenges, including access to tissue and ethical considerations. As a consequence, we often rely on model organisms such as the worm (C. elegans), the zebrafish, the chick, and the mouse. In the mouse, the gestation period lasts 19-21 days and developmental timing has been conveniently categorized into half embryonic days (E), counting from fertilization (or post-coitum) [1]. Given that the exact timing of fertilization in mice is impossible to determine, detection of a vaginal plug the morning following fertilization is referred to as E0.5 [1]. From the moment the egg is fertilized, it undergoes a highly coordinated series of events driven by cellular division and differentiation. Post-implantation, gastrulation (around E6.5 - 7.5 in mice) is a major milestone characterized by the formation of the three germ layers (endoderm, mesoderm, and ectoderm) from the single-layered blastula [1]. This is the beginning of differentiation to establish cell lineages with restricted potential: the endoderm will give rise to internal organs such as the gastrointestinal tract and the lung, the mesoderm is the precursor to the circulatory system and muscle, and the ectoderm will form the nervous system and skin. This phase is followed by organogenesis from E8 until birth, during which specific organs are formed from each of the three germ layers. In this dissertation, my studies focus on aspects of organogenesis. 2 1.2 Vascular development Every living cell requires a minimum of metabolites to survive. As the number of cells continues to multiply rapidly with an increasingly complex structure following gastrulation, the need for a conduit to distribute oxygen and nutrients in exchange for metabolic waste becomes fundamental. Hence, the cardiovascular system is the first organ to become fully functional during organogenesis [2]. Blood vessels, composed of endothelial cells, are formed by two distinct mechanisms; vasculogenesis and angiogenesis. Vasculogenesis involves the formation of de novo endothelial cells from the differentiation of a mesodermal precursor and occurs mainly during early embryonic development [3]. On the other hand, angiogenesis is the process of blood vessel formation from existing endothelial cells, responsible for remodeling and expanding the vascular network, and is driven mostly by cell migration and proliferation [4]. Construction of a vascular system is initiated at the end of gastrulation in the mouse embryo. A primitive vascular plexus is first formed extra-embryonically in the yolk sac as mesodermal cells give rise to aggregates of cells referred to as “blood islands” by E7.5 [5, 6]. Endothelial cells define the periphery of the blood islands with hematopoietic cells (mainly erythroid precursors) located in the center of what will become the lumen [5]. Simultaneously, E7-E8 is an active period of vasculogenesis in the embryo proper as a vascular network is also being developed from the cranial region [5]. Local proliferation of endothelial precursors expressing Flk1 (also known as vascular endothelial growth factor receptor 2, VEGFR2) and Scl/Tal-1 leads to the emergence of endothelial cell clusters at E8.0, first as loose networks that then proceed to enclose a lumen forming paired dorsal aortae [5-7]. The anterior ends of the dorsal aortae connect to the cardiac outflow tract while the elongating posterior ends will fuse with the vitelline vein, connecting the yolk sac to the embryo proper [8]. Around E9.5, the primary dorsal aortae merge at the midline to form a single dorsal aorta descending along the trunk, the largest blood vessel in the embryo [8]. The need for vascularization means that endothelial cells are a specialized cell type present in every organ of the body. A unique feature of endothelial cells is that they remained associated with the growing organ [6]. Therefore, every organ in the body can be described as the sum of endothelial and nonvascular cells. The notion that endothelial cells play more than a passive role in the regulation of different organs is now well supported, including examples in the liver, 3 pancreas, and kidney [4, 6, 9, 10]. Not only do these cells serve as the lining of vessels to transport nutrients, hormones, and cells, but the endothelium can exert specific function via angiocrine signaling. For example, endothelial cells secrete IGF1 during craniofacial cartilage development to regulate jaw growth by driving proliferation of chondrocytes [11]. In the liver, activation of VEGFR1 on sinusoidal endothelial cells promotes the growth of hepatic cells via a paracrine mechanism and endothelial expression of Itgb1 also supports liver growth concomitant with an increase in hepatic growth factor expression [12, 13]. While the fact that endothelial-derived signals are required for proper organogenesis is now widely appreciated, understanding the scope of what those signals are, and how each signal regulates organ development, remains a challenge to be addressed. 1.3 Endothelial contribution during lung development Lung development begins with the formation of a primary lung bud from the anterior foregut endoderm around E9.5 in the mouse embryo and can be divided into five major morphologically distinct phases. The embryonic stage initiated at E9.5 gives rise to the trachea and separate lung buds which then undergo branching morphogenesis during the pseudoglandular stage (E12-E16.5) to form a tree-like network extending to terminal bronchioles [14]. Then, the canalicular stage (E16.5-17.5) leads to the formation of distal airway and the beginning of differentiation of the epithelium that allows distinction of conducting vs respiratory airways [14-16]. It is followed by the saccular stage which extends to postnatal days in the mouse [14-16]. The saccular stage is characterized by the formation of terminal saccules, increase vascularization, and further differentiation of alveolar epithelial cells into type 1 (AE1C) and type 2 (AE2C) cells, which is concomitant with a thinning of the septa by elongation of AE1C and initiation of surfactant secretion by AE2C [15, 16]. Even after birth, the lung keeps maturing during the alveolar stage of development leading to an increase in the number of fully mature alveoli and pulmonary capacity [16]. The main function of the lung is oxygenation which relies on two major processes: oxygen intake into the pulmonary alveoli and gas-exchange with the adjacent blood vessels which will circulate oxygen throughout the body. Therefore, it is not surprising that during lung organogenesis the processes of alveolarization and angiogenesis are interdependent. A close association of endothelial and epithelial cells is essential for respiration. From the very 4 beginning, the lung bud develops in the proximity of newly form blood vessels derived from the mesoderm [17, 18]. While the lack of vascular structures does not impede the formation of the early lung bud and progenitor cells during the embryonic phase [19], the endothelium plays a crucial role in lung morphogenesis from the pseudoglandular phase onwards. In a lung explant culture system, recombinant vascular endothelial growth factor (VEGF) was shown to induce branching events leading to an increase in the number of terminal branches, an effect mediated by the receptor Flk-1 (VEGFR2) expressed on endothelial cells [20]. Similarly, using a VEGF decoy receptor Lazarus et al. showed that VEGF inhibition reduces vascularization of the early lung (E11.5-E12.5) which consequently impacted branching morphogenesis [21]. Blocking vascularization does not impact epithelial cell proliferation but leads to larger airways with fewer branching events particularly impacting bifurcation when a rotation in the branching plane is involved [21]. Most studies focused on early stages of lung development, but the endothelium is continuously required to shape the lung. In newborn rats, Pecam1 (CD31) signaling affects the alveolar structure in vivo [22]. Unlike with VEGF inhibition, Pecam1 blockade using a blocking antibody did not impact endothelial cell proliferation or survival but disrupted their migration pattern [22]. Still, blocking endothelial cell migration by Pecam1 inhibition is sufficient to cause larger tubular structures instead of properly formed alveoli [22], reinforcing once again the role of the endothelium in defining the pulmonary architecture. Most known examples of the endothelium function on lung development involve patterning. However, in adult lung injury models, stem cell differentiation and expansion of alveolar epithelial cell populations are also influenced by signals coming from the endothelium [23, 24]. The colony-forming ability of bronchioalveolar stem cells can be maintained in vitro during co-culture with primary lung endothelial cells but knockout of Thrombospondin 1 (Tsp1) in endothelial cells impairs the formation of alveolar colonies to favor bronchiolar colonies in this system, a phenotype that can be rescued by addition of exogenous Tsp1 [23]. Concordantly,Tsp1 expression increases in vivo following selective injury to AE2C, via bleomycin administration, to promote alveolar recovery while decreasing in response to naphthalene injection which models bronchiolar injury [23]. Meanwhile, in a pneumonectomy model of lung regeneration, MMP14 activation downstream of VEGFR2 and FGFR1 was also implicated in alveoli restoration by 5 promoting AE2C expansion [24]. These studies suggest that endothelial-epithelial cross-talk might constantly be regulated, even after the main pulmonary architecture is established. Therefore, endothelial cells may have additional functions in lung development that were not identified in the models used in the aforementioned studies. 1.4 Endothelial cell heterogeneity and specification Endothelial cells are part of organs with vastly different functions in the body, and this is associated with microvascular endothelial heterogeneity across various anatomical sites. One of the best-known examples of heterogeneity includes the tight junctions formed by endothelial cells of the blood-brain barrier in contrast to fenestrated and more permeable endothelial cells in organs such as the kidney [4]. This reflects a change in endothelial cell morphology based on the organ-specific required function. At the transcriptional level, endothelial cells isolated from various tissue of origin also show distinct gene signatures, supporting the concept of endothelium-organ interdependency [25]. Distinct angiocrine factors are expressed based on the organ from which endothelial cells are isolated which likely impacts the surrounding tissue in specific ways [25]. Interestingly, mouse embryonic stem cell (mESC)-derived endothelial cells in vitro adopt different gene expression profiles depending on the organ in which they engraft in vivo [25]. For example, mESC-derived endothelial cells that engraft in the liver upregulate VEGFR3 while the same cells upregulate CD34 expression if they engraft in the kidney, essentially reproducing the organ-specific pattern already present [25]. This shows that endothelial cells have plasticity and that the microenvironment in which they reside influences their fate. Therefore, not only does the endothelium provide essential information to the surrounding tissues, but it also receives cues from the local microenvironment. There is also apparent morphological heterogeneity between macrovascular endothelial cells, with arterial cells being typically longer and narrower than venous cells [26]. Arterial, venous, and lymphatic endothelial cells become molecularly distinct as they differentiate from the emerging primordial vasculature, even before the onset of circulation. Arterial cells can be distinguished by expression of EphrinB2 while their venous counterparts express EphrinB4 and Nr2f2 (also known as COUP-TFII) [27, 28]. Lymphatic endothelial cells, which buds from veins, also express COUP-TFII but are further specified by upregulation of Prox1 [29]. The balance between arterial and venous fate is established in part by modulation of the Notch signaling 6 pathway. For instance, COUP-TFII can inhibit Notch1 and Jag1 signaling to promote venous identity [28]. On the other hand, Sox17 is a key regulator of arterial identity acting upstream of the Notch pathway to promote Dll4, Notch1, and Notch4 expression [30]. In both cases, knockout or enforced expression was shown to alter cell identity supporting that specification is tightly regulated by a fine balance between reciprocal signals. In addition to the three subtypes of endothelium mentioned above, some specialized endothelial cells are more transient during embryogenesis. This is the case of endothelial cells contributing to heart development by undergoing an endothelial-to-mesenchymal transition (EndMT) [31] and a population of hemogenic endothelial (HE) cells which have the potential to undergo an endothelial-to-hematopoietic transition (EHT) giving rise to blood progenitor cells [32]. HE cells resemble the arterial endothelium and share many common pathways including Notch signaling [33]. However, eventual repression of an arterial-like identity via downregulation of Sox17 and Notch1 is required for true hemogenic specification [34, 35]. 1.5 Embryonic hematopoiesis and EHT The co-emergence of endothelial and hematopoietic cells in blood islands was observed as early as 1920 [36] and experimental evidence in the chick (later supported in the mouse and zebrafish) have supported a common origin from a Bry+Flk-1+ “hemangioblast”. Cell labeling for lineage fate tracing corroborated the origin of blood cells from a precursor expressing endothelial markers [37, 38]. While the in vivo clonality of a hemangioblast giving rise to both an endothelial and a hematopoietic daughter cell might still be debatable [39], the mass of cells from which blood islands are derived clearly has both potentials and it is likely that the plasticity within individual cells is influenced by additional microenvironmental factors to determine fate. This first wave of blood cell production, taking place in the yolk sac, is referred to as primitive hematopoiesis (Fig. 1.1). It gives rise mainly to primitive erythrocytes; large erythroid cells that are still nucleated and express embryonic globins [40, 41], whose role is to quickly deliver oxygen. Some macrophage and megakaryocytes are also detected during this first wave. This is followed by the emergence of erythromyeloid progenitors (EMPs) which characterizes the second wave of blood formation. EMPs are detected starting around E8.25 in the yolk sac and are able to produce definitive erythrocytes and cells of myeloid lineage [40, 41]. 7 1.5.1 Definitive hematopoiesis and hematopoietic stem cells The second wave of hematopoiesis gives rise to adult-like blood cells which represent the start of definitive hematopoiesis [39]. However, within the embryo proper, hematopoiesis begins during the third wave which is defined by the emergence of hematopoietic stem cells (HSCs) primarily from the dorsal aorta in the aorta-gonad-mesonephros (AGM) region of the mouse embryo. The first HSC can be detected in the AGM as early as E10.5 with a slightly higher number by E11.5 [42, 43]. The process by which these first HSCs emerge is thought to be conserved in the human where HSCs also develop from the AGM around day 32-41 of gestation [44]. HSCs are defined by two major characteristics: the potential to differentiate giving rise to all mature blood cells in the organism, and a self-renewal ability that allows them to sustain hematopoiesis throughout the organism’s lifetime [45]. In contrast, the term “hematopoietic progenitor cells” is used to designate cells with a more limited self-renewal capacity with either multipotent or more restricted lineage potentials. Therefore, the most precise assays to detect HSCs need to show long-term functionality, with the most common and stringent one to date being the in vivo regeneration of progeny cells with regenerative activity in serial transplantation assays. In contrast, the immature pre-HSC found at early embryonic stages require further maturation steps, either in vivo or via ex vivo co-culture, prior to be able to engraft and repopulate a recipient [46]. Given that even the more primitive hematopoietic stem and progenitor cells display functional heterogeneity within defined compartments and represent a spectrum of cell trajectories, the terms “HSPC”, referring to both hematopoietic stem and progenitor cells, and “pre-HSPC” will be used throughout the rest of this dissertation. This is also consistent with the assays presented in this dissertation which do not allow for the distinction between emerging stem or progenitor cells. At E11.5, pre-HSPCs are also detected in extra-embryonic blood vessels (vitelline and umbilical arteries), the head, and the placenta [43]. Given that their detection follows the emergence of HSPC in the AGM, whether this observation is due to migration or de novo HSPC generation from these sites has proven hard to determine conclusively. Newly formed HSPCs migrate in the circulation to the fetal liver where they undergo massive expansion between E12.5-E16.5 before colonizing the bone marrow just prior to birth [47]. 8 Figure 1.1 Timeline of hematopoietic development during embryogenesis in mice Different anatomical sites act as the main stage for hematopoiesis in the embryo as the hematopoietic system is progressively established. Only the major sites are depicted in this diagram. 1.5.2 Endothelial-to-hematopoietic transition In both mouse and zebrafish, live-cell imaging has shown the emergence of cells with hematopoietic features from the endothelial wall of the dorsal aorta [32, 48, 49], providing strong support for the occurrence of EHT during embryonic hematopoiesis. In the zebrafish, transgenic reporter lines were used to demonstrate that kdr-expressing flat endothelial cells in the vascular layer of the dorsal aorta round up and simultaneously start expressing cmyb as they emerge ventrally into the caudal vein [48]. This is different from murine EHT during which the cells bud into the lumen of the dorsal aorta and form large intra-aortic hematopoietic clusters (IAHC) [32]. Murine IAHC contain cells co-expressing CD31 and c-kit (CD117), with the budding cells upregulating surface expression of CD41, an early maker of hematopoietic commitment [32]. Functionally, cells co-expressing endothelial markers (CD31 or VEC) with low levels of CD41 show HSPC potential based on transplantation assays in irradiated mice after co-culture ex vivo, and these cells were therefore defined as “pre-HSPC” [46]. Additional studies have contributed new markers to define cells at different maturation stages along the transition and it is widely accepted that cells undergoing EHT progressively express c-Kit, CD41, CD43, and CD45 towards a HSPC phenotype [46, 50, 51]. More recently, CD27 expression was shown to further enrich for progenitors with long-term repopulation activity thus defining the following 9 populations of CD31+VEC+cKit+CD41low IAHC cells: CD43-CD45- “pro-HSPC”, CD43+CD45-CD27- “pre-HSPC type I”, and CD43+CD45+CD27+ “pre-HSPC type II” [52]. Initial studies have reported that as few as 1-2 HSPCs and up to 60 pre-HSPCs emerge in the mouse AGM around E11 based on transplantation assay despite the presence of 600-700 IAHC cells [42, 43, 53]. However, lineage tracing using a Cre-inducible confetti reporter led to an estimate of up to 700 Flk1+ mesoderm precursor cells contributing to long term hematopoiesis in the mouse [54]. Similarly, more than 700 embryonic Tie2+ cells, labeled around E9.5, were shown to contribute to adult hematopoietic cells based on barcoding approaches in vivo [55]. This number is much higher than experimentally determined based on in vivo reconstitution ability at specific embryonic time points, even under the best conditions to detect immature pre-HSPCs [43]. While experimental manipulations may yield lower numbers of HSCs by affecting the cells directly or due to detection limit of various assays, this enormous discrepancy argues that we still have a lot more to learn about in vivo embryonic hematopoiesis. 1.5.3 Defining the hemogenic endothelium Not all endothelial cells have the potential to undergo EHT, with only roughly 1.3% of endothelial cells isolated from E10.5 dorsal aorta showing functional hematopoietic potential ex vivo [56, 57]. These cells have been termed “hemogenic endothelial” (HE) defined by their potential to become a hematopoietic cell. By definition, HE cells exhibit an endothelial phenotype. That includes the expression of some classical endothelial surface markers such as VE-Cadherin (Cdh5) and CD31 (Pecam1) but also morphological features [39]. Hence, HE refers to cells embedded in the vascular wall such as in the dorsal aorta and consequently can be difficult to distinguish from other non-hemogenic cells that comprise the blood vessel [39]. Given that HE cells represent a functionally distinct subset of endothelial cells, they can only be definitively identified retrospectively after producing hematopoietic cells. Nevertheless, a few key reporters were developed to prospectively enrich for endothelial cells with hemogenic potential in the mouse embryo based on the activity of regulatory elements that control expression of Sca1 (Ly6a-GFP), Runx1 (Runx1+23GFP), and Gfi1 (Gfi1+/tomato) [56, 58, 59]. As consequence, several definitions of the hemogenic endothelium exist and the degree to which these cell populations overlap is not always clearly defined. More recently, the term “pre-HE” emerged to define cells along the endothelial to hematopoietic trajectory that are closest to HE 10 cells but lack expression of Runx1 [60]. Better molecular and functional characterization of pre-HE cells is still required, along with investigating the events leading to both the emergence and progression of this pre-HE population. 1.5.4 Transcriptional regulation of EHT The transcription factor Runx1, also known as acute myeloid leukemia 1 (AML1) or core-binding factor alpha 2 (Cbfa2), is required for definitive hematopoiesis and embryos lacking Runx1 expression are devoid of IAHC and HSPC [61]. Runx1 is expressed in emerging IAHC cells in the AGM at E10.5 and its conditional-deletion in endothelial cells using a VEC-Cre system leads to a similar phenotype, showing a lack of IAHC and strong deficit in both c-kit+ cells within the AGM and hematopoietic progenitors based on colony-forming unit assays [62]. Given that Runx1 is critical for EHT, it is extensively studied and widely used as a marker of EHT potential. Runx1 expression is regulated by two promoters, distal and proximal, and a conserved non-coding enhancer element termed +23 (or sometimes referred as +24) enhancer, defined by its location from the distal promoter [63, 64]. While both promoters appear to be important in definitive hematopoiesis, the +23 enhancer is specifically active in HE cells and marks emerging IAHC [63, 64]. This enhancer was labeled with a GFP fusion protein to allow tracking of its activity in a mouse model and now represents a useful tool to enrich for a population of cells undergoing EHT [56, 64]. The exact mechanism by which Runx1 drives EHT is complex but appears to likely involve both the repression of the arterial phenotype and the upregulation of hematopoietic genes. Downregulation of arterial fate is mediated in part by negative feedback between Sox17 and Runx1. Whereas Sox17 expression is required for development of the arteries and still expressed in HE cells [30], a high Runx1 to Sox17 ratio was found in IAHC cells in the mouse dorsal aorta and Sox17 was identified as a downstream target of Runx1 by chromatin-immunoprecipitation [34, 35]. Runx1 directly exerts a repressive effect on Sox17 expression and temporal loss of Sox17 alone is sufficient to increase EHT-derived cells in the mouse embryo [34]. Interestingly, Sox17 also inhibits Runx1 expression [34], therefore the balance between those two genes may contribute to determining cell fate and additional factors may be required to initiate the transition to a Runx1high cell. 11 In addition, two other Runx1 targets promote EHT; Gfi1 and Gif1b. Like Runx1, Gfi1 expression marks HE cells in E10.5 AGM, whereas more differentiated pre-HSPC cells are both Gfi1+/Gfi1b+ [58, 65]. Homozygous deletion of both Gfi1 and Gfi1b blocks EHT whereas compensatory mechanisms can partially rescue the phenotype in single mutants [65]. Mechanistically, Gfi1/Gfi1b promotes EHT by repressing expression of endothelial genes and lack of Gfi1/Gfi1b expression in double mutant embryos prevents morphological changes associated with EHT, keeping HE cells embedded in the vasculature [65, 66]. This repressive effect of endothelial identity is in part mediated by recruitment of the histone demethylase Lsd1 (Kdm1a) [65], suggesting epigenetic remodeling during the transition. Amongst the many pathways investigated during embryonic hematopoiesis, Notch signaling pathway plays a critical role in both arterial specification of endothelial cells and EHT. Notch regulates Gata2 expression [67] and, similar to Runx1, complete loss of Gata2 has a strong inhibitory effect on the emergence of IAHC and HSPC from the AGM [68, 69]. Notch activation also regulates expression of the transcriptional repressor Hes1 which dampens Gata2 expression [67]. This antagonizing regulation might be responsible for the pulsative expression of Gata2 observed in emerging IAHC [70]. Interestingly, deletion of Hes1 leads to an overexpression of Gata2 and Runx1 concomitant with an increase in non-functional HSPCs [67]. Therefore, Gata2 expression needs to be carefully modulated. Similarly, the strength of the Notch signal needs to be tightly regulated with low levels of Notch activity being compatible with hematopoiesis [33]. The balance is achieved by Notch1 ligand Jag1 promoting hemogenic fate whereas Dll4 imposes an endothelial program through paracrine signaling [33]. Interestingly, Dll4 is expressed in IAHC to restrict the recruitment of adjacent HE cells and regulates cluster size in the dorsal aorta [71]. Opposing Dll4, Jag1 is expressed in the endothelium and stimulates the emergence of new clusters [71]. Hence, surrounding endothelial cells can drastically influence the fate of neighboring hemogenic cells in the dorsal aorta. 1.5.5 Inflammatory signaling and microenvironmental factors influencing EHT Although some evidence suggests the contribution of additional embryonic sites (head, vitelline, placenta) in HSPC emergence [43], the fact that EHT occurs mainly at a specific location during a narrow developmental window in embryogenesis raises an important question: what makes the 12 AGM the ideal location for EHT? Interestingly, even “HE” cells isolated from the yolk sac or the dorsal aorta based on the same selection of markers are molecularly distinct at the transcriptomic level [72]. This heterogeneity based on anatomical site suggests that not all hemogenic cells are created equally and perhaps contribute in different ways to hematopoiesis. Within the AGM, HSPC emerge from the ventral side of the dorsal aorta in most species which suggests polarizing signals that emerge from the ventral (supportive) and dorsal (antagonizing) mesoderm. In addition to a gradient of signaling molecules in the surrounding tissues, circulating immune cells also contribute to EHT. Macrophages are recruited to IAHC sites and represent a significant component of the niche [73]. Depletion of macrophages results in lower HSPC output from the AGM based on colony-forming unit and transplant assays [73]. A potential function of macrophages in HSPC emergence is to promote HSPC mobilization by expressing matrix metalloproteinase (Mmp9 and Mmp13) to degrade the extracellular matrix [73, 74]. But this does not necessarily explain the reduction in colony-forming unit observed and macrophages may also be a source of proinflammatory signaling, which was shown to stimulate EHT [75-77]. Other than macrophages, neutrophils are an important source of TNFα in the zebrafish [75]. TNFα activates Tnfr2 on endothelial cells to induce Jag1 expression, thereby modulating Notch signals in the HE. [75]. Similarly, pro-inflammatory signaling via Toll-like receptor 4 (TLR4) also activates the NF-κB-Notch axis in support of EHT [76]. The zebrafish constitutes a great model to more efficiently track cells over time. In doing so, Damm and Clements recently showed that neural crest cells also physically associate with emerging HSPCs during EHT in vivo [78]. While the migration of these neural crest cells is driven by Pdgf signaling, the authors did not identify the pro-hematopoietic signals that these cells transmit to EHT cells [78]. This illustrates the complexity of the in vivo microenvironment and how it may host a large network of interactions that we have yet to discover and fully understand. Therefore, it is critical to examine how the vascular and hemogenic endothelium are influenced by external stimuli during specification and differentiation. 13 Figure 1.2 Schematic of IAHC formation in the AGM of E10.5 mouse embryos EHT is regulated by a fine-tuned balance of opposing factors from both cell intrinsic and extrinsic origin. The contribution of neutrophils was shown in the zebrafish, however evidence in the mouse also supports a role for TNFα in promoting cluster formation. 1.5.6 The endothelium as a continued support for hematopoiesis The role of the endothelium on the hematopoietic system is not limited to a small window during development. After emerging via EHT in the dorsal aorta, HSPCs migrate to the fetal liver which constitutes the major HSPC niche prior to bone formation. In the fetal liver, HSPCs localize in perivascular niche close to pericytes adjacent to portal vessels [79]. Upon arrival, HSPCs trigger a remodeling of the vascular niche which was referred to as “endothelial cuddling”, where HSPCs are found anchored to a mesenchymal stromal cell and become surrounded by endothelial cells before undergoing cell division [80]. In addition, expansion of the portal vessel correlates with HSPC expansion in the fetal liver between E12 to E14.5 hinting at a potential regulation of the HSPC pool by the perivascular niche [79]. During adulthood, HSPCs also reside in the perivascular niche around sinusoidal vessels in the bone marrow [81, 82]. While 14 several components of the microenvironment (including mesenchymal stromal cells and other cells types) secrete factors to regulate various aspects of HSPC maintenance [83], the endothelium is critical in establishing that niche. Restoration of vascular networks is necessary for hematopoietic reconstitution of the bone marrow following irradiation [84]. Furthermore, Akt activation in endothelial cells of adult mice increases HSPC numbers [85]. Therefore, HSPCs are intimately connected to the endothelium throughout their life cycle. 1.6 Nitric Oxide Nitric oxide (NO) is a gas with significant biological implications by acting as a small signaling molecule in cellular processes. Biologically, it is synthesized by one of three isoforms of nitric oxide synthase (NOS); neuronal NOS (nNOS, NOS1), inducible NOS (iNOS, NOS2), or endothelial NOS (eNOS or NOS3) [86]. In the cardiovascular system, NO production by eNOS plays important functions in angiogenesis and vasodilation. When NO is released to adjacent smooth muscle cells surrounding the blood vessel, it binds soluble guanylyl cyclase (sGC) which stimulates cyclic guanosine monophosphate (cGMP) production [86]. Increase in cGMP levels activates cGMP-dependent protein kinase (PKG) to reduce intracellular calcium and lead to relaxation [86]. Shear stress, such as induced by laminar blood flow in an artery, is a mechanism to activate eNOS and thus leads to a decrease in blood pressure. In E8.5 embryos, onset of circulation creates shear stress which amounts to some level of NO secretion. In the dorsal aorta, this shear-stress induced NO affects the emergence of hematopoietic cells. Inhibition of NO production by injection of nitro-L-arginine methyl ester (L-NAME) decreases Runx1 expression in zebrafish AGM while also reducing the number of hematopoietic colony-forming units in the mouse [87, 88]. eNOS-null embryos display a similar phenotype to L-NAME treatment supporting that NO originates from eNOS activation [88]. Given that eNOS is expressed in both emerging IAHC and surrounding endothelial cells in the dorsal aorta, whether this represents a cell autonomous effect of NO remains undefined. However, in vitro assays suggest that shear stress is at the origin of this NO-mediated effect on hematopoiesis [87] hence further supporting that the microenvironment influences endothelial cell fate during EHT. 15 1.7 Sash1 SAM and SH3 domain containing protein 1 (SASH1) was initially described as a candidate tumor suppressor in breast cancer [89]. Both the human and mouse gene are comprised of 20 exons, with approximately 85% conserved sequence identity between the two species [89]. As its name suggests, this >1200 amino acid large protein contains several binding domains including 2 sterile alpha module (SAM) domains and a Src homology domain 3 (SH3) [89], which are characteristic of scaffolding proteins that can interact with multiple binding partners to assemble complexes and relay signal transduction. In addition, studies from our lab showed that Sash1 interacts with TRAF-6, via another domain (TRAF-6 binding domain) not originally described [90] and therefore Sash1 likely contains additional functional domains that are not as well characterized. Further studies conducted in our lab unveiled a role for Sash1 in immune signaling downstream of TLR4 in non-immune cells [90]. Specifically, knockdown of Sash1 impairs LPS-induced activation of NF-κB in human microvascular endothelial cells (HMEC) resulting in a decrease in IL-6 production, an effect that is mediated by the interaction of Sash1 with TAK1 and TRAF-6 [90]. Similarly, Sash1 interaction with TAK1 also leads to MAPK pathway activation [90], defining Sash1 as a molecule that can regulate multiple signaling cascades within the cell. In the last decade, several studies have investigated the role of Sash1 across many tumor types including cervical cancer [91, 92], colorectal cancer [93], hepatocarcinoma [94], oesophageal carcinoma [95], gastric cancer [96], osteosarcoma [97], lung cancer [98], and glioma [99]. The vast array of cancer types involved is supported by the fact that Sash1 was shown to be ubiquitously expressed at low levels in all human tissues, with higher expression in breast, lung, thymus, thyroid, placenta, and spleen [89]. Similarly, Sash1 is expressed in microvascular endothelium of multiple organs in the adult mouse including lung, thymus, spleen, liver, kidney, and the brain [90]. On the other hand, the expression pattern of Sash1 during embryogenesis has not been extensively described. In cancer studies, Sash1 has been associated with tumor suppressor activity where low Sash1 expression typically correlates with poor prognosis related to increased tumor size and metastasis. Across multiple cancer cell lines, Sash1 overexpression increased the fraction of cells 16 in G0/G1 phase of the cell cycle, increased apoptosis, and decreased cell invasion in a transwell assay. These phenotypic differences correlate with a decrease in cyclin D1 expression, an increase in pro-apoptotic caspase-3 concomitant with a decrease in the anti-apoptotic protein, Bcl-2, and decrease in MMP-9/-2 expression [91, 97, 98, 100]. While all in vitro studies converge on the notion that Sash1 expression reduces proliferation and migration, very few studies so far have supported this role in vivo. Sash1 overexpression was found to reduce tumor volume in mice in a model of esophageal squamous cell carcinoma [95] while Sash1-deficient HCT116 colon cancer cells lead to an increase in metastasis compared to WT cells [93]. More evidence of the role of Sash1 in vivo is required, both during homeostasis and under disease conditions. 1.8 Meis1 Myeloid Ecotropic Integration Site 1 (Meis1) was first identified in murine myeloid leukemic cells [101] and has been studied mostly for its role in leukemogenesis. Homeodomain protein, Meis1 can interact with other TALE-family members such as Hox proteins and Pbx1 to increase their DNA-binding affinity and regulate transcription [102]. In adult bone marrow, Meis1 expression is highest in the most primitive compartment compared to more differentiated hematopoietic cells and loss of Meis1 contributes to exhaustion of the HSPC pool in adult bone marrow [103]. Meis1 is required for HSPC maintenance in part by regulating quiescence by limiting reactive oxygen species (ROS) levels through activation of hypoxia-induced factors Hif1α and Hif2α [104, 105]. Furthermore, Meis1 is an essential developmental gene as evidenced by the embryonic lethality between E11.5-E14.5 of Meis1-null embryos, associated with defects in fetal hematopoiesis and hemorrhage [106, 107]. At E12.5, Meis1-null fetal liver hematopoietic cells show impaired colony-forming activity and are unable to repopulate irradiated mice, suggesting that Meis1-null mice are unable to produce or maintain HSPC [107]. Other observations include a smaller liver, lack of megakaryocytes, and defects in angiogenesis [106, 107]. Although multiple embryonic abnormalities were described in Meis1-null mice, the function of Meis1 in embryonic hematopoiesis still remains poorly characterized in particular during EHT. The spatial and temporal requirements for Meis1 expression are controversial and its regulatory network is poorly understood. 17 1.9 AIM OF THE STUDY Embryonic development involves a highly coordinated series of events that need to be faithfully replicated. Deep knowledge and understanding of the normal developmental processes essential for proper organogenesis is fundamental not only to advance “basic” biology and research but also to provide insights into disease mechanisms and advance clinical science. Developmental biology (along with stem cell biology) is at the root of regenerative medicine, a field in rapid emergence partly due to progress in technology and engineering. But the challenge to design technologies that can detect, prevent, and cure diseases must be pushed forward first by our understanding of normal biological systems. Recent years have also been characterized by an emergence of organoids and other 3D culture systems in an attempt to more faithfully replicate biological processes in vitro [108]. As a model system, organoids are still very much under development as a promising technology with vast potential [108]. A significant component and challenge with developing these systems is understanding how cells interact with each other and which cell types are important components influencing the overall biology of the organ. As discussed in previous sections, endothelial cells are a fundamental component of any organ and can have substantial effect through direct cell-cell contact or paracrine signaling. Similarly, the microenvironment impacts the fate of endothelial cells and equally needs to be considered when manipulating and specifying these cells in vitro. The aim of this study is to investigate the role of selected essential developmental genes in order to describe novel fundamental pathways in organogenesis, focusing on endothelial signaling to drive differentiation and cell fate. Deletion of essential developmental genes often leads to embryonic lethality in mouse model. Constitutive loss of Sash1 in mice leads to mortality at birth due to respiratory distress, pointing towards a lung defect [109]. However, Sash1 was reported to be preferentially expressed in endothelial cells of various organs in adult mice [90]. Chapter 3 will focus on the role of Sash1 signaling from the endothelium to promote lung maturation required for pulmonary compliance and proper ventilation at birth. Specifically, we showed that endothelial Sash1 signaling is an important source of nitric oxide (NO) which affects alveolar cell maturation. 18 The emergence of the vascular and blood systems is intimately connected. In chapter 4, we seek to characterize the role of Sash1 in the establishment of the hematopoietic system. Although not responsible for the lethality observed in Sash1-null mice, defects in hematopoiesis were also noted. Given that hematopoietic stem and progenitor cells (HSPCs) emerge from an endothelial-to-hematopoietic transition, we hypothesized that loss of endothelial expression of Sash1 might affect EHT and investigated the role of Sash1 in this developmental process. EHT itself represents an interesting participation of endothelial cells in the formation of another system, namely the hematopoietic system, where specification of specialized endothelium needs to be regulated by a number of factors. In addition, as hematopoietic clusters emerge, remodeling of the vascular wall is required and neighboring endothelial cells are believed to control IAHC size via Notch signaling [71]. What drives the initial cell fate trajectory towards hemogenic endothelium, rather than the vascular endothelium, is a central question in hematopoiesis. Given that EHT gives rise to de novo hematopoietic stem cells (HSCs) in vivo, there is a lot of hype around replicating this process in vitro to produce HSCs for clinical use. Chapter 5 attempts to address this question and identifies the gene Meis1 as an early marker of HE cells and potential regulator of HE specification. 19 CHAPTER 2: METHODOLOGY 2.1 Mouse strains and embryo collection All mouse strains used in timed mating were bred and maintained in-house on a C57BL/6 background, except for the eNOS-null mice that were directly purchased from The Jackson Laboratory. VEC-Cre males were also purchased from The Jackson Laboratory [62] to cross with our floxed animals and maintained as hybrids. For experiments in this dissertation, the Cre allele was carried by heterozygous males to allow for consistent tracking of Cre efficiency by repeated mating of each stud. Sash1 gene-trap mice [90] were maintained as heterozygous (Sash1+/-) by breeding with wildtype (WT) animals. Heterozygous mates were used in timed mating to generate litters of WT, heterozygous, and Sash1-null (Sash1-/-) embryos for experiments. For experiments with conditional deletion of Sash1, the Sash1-floxed (Sash1fl/fl) mouse model was generated in-house with help of the Transgenics Core (Terry Fox Laboratory, Vancouver, British Columbia). Briefly, a p-loxP-2FRT-PGKneo construct with floxed exon 3 and 4 of the Sash1 gene followed by stop codon at exon 5 was electroporated into embryonic stem cells. Sash1fl/fl females were crossed twice with VEC-Cre males to generate Sash1fl/fl;VEC-Cre embryos and WT littermates that did not receive the Cre allele (Sash1fl/fl or Sash1fl/+). The same strategy was employed to maintain the Meis1-floxed (Meis1fl/fl) and Meis1-VEC-Cre (Meis1fl/+;VEC-Cre) colonies. Meis1fl/fl mice with loxP sites flanking the exon 8 were generously provided by Dr. Keith Humphries and originally generated by Drs Nancy Jenkins and Neal Copeland [103, 105, 110]. The Meis1-GFP strain was generously provided by Dr. Keith Humphries [111] and maintained as heterozygous by crossing with WT C57BL/6 mice. The Meis1;Runx1 strain (Meis1fl/fl;Runx1+23GFP) was generated in-house by crossing our Meis1-floxed mice with Runx1+23mCNE-GFP mice generously provided by Dr. Motomi Osato [64]. To generate embryos with both the Runx1+23GFP reporter and conditional excision of Meis1, Meis1;Runx1 females were used in timed matings with Meis1-VEC-Cre males. Timed matings were set up by pairing 2 females with 1 stud male to generate embryos of known embryonic time points. The morning after timed mating, females were checked for the presence of a vaginal plug and separated into a new cage. Embryos were staged according to the number of days post-coitus, starting with E0.5 the morning of a vaginal plug detection. At the relevant time point as determined for each experiment, dams were first anaesthetized with isoflurane and 20 then euthanized by carbon dioxide inhalation followed by cervical dislocation. An incision was made on the abdomen of the dam to collect the embryos. For experiments at late time points (E17.5-E18.5), decapitation was performed as soon as the embryo was taken out to ensure death. The rib cage was then carefully opened using forceps and the lung was extracted making sure to remove the heart and other tissues. For experiments involving earlier embryonic stages (E9.5-E10.5), the uterus containing embryos was taken out and placed in phosphate buffer saline (PBS). Embryos were dissected one at a time under a dissecting scope in a different vessel containing PBS to avoid contamination. Forceps were used to initially separate each embryo from the placenta and collect the yolk sac. Subsequently the AGM was dissected using 30g ½ needles attached to a 1ml syringe. All animal experiments were carried in accordance to approved protocols by the Animal Care Committee of the University of British Columbia (Vancouver, British Columbia) 2.2 Genotyping Every embryo used in experiments was genotyped by PCR followed by gel electrophoresis. For E9.5 or E10.5 embryos, the yolk sac was collected in a 200 µL PCR strip tube during dissection while a piece of the tail was collected for older embryos at E17.5 or E18.5. The tissue was digested in 100 µL buffer with 0.4 mg/mL proteinase K (Sigma-Aldrich #P6556) for 16 hours at 55oC to release genomic DNA. The targeted genomic regions were amplified from 2 µL of digested tissue by PCR using Taq DNA Polymerase (New England Biolabs #M0267) or Platinum™ HiFi Taq Polymerase (Invitrogen #11304011) depending on the sequence. All primers (Table 2.1) were obtained from Integrated DNA Technologies (Coralville, Iowa). Amplified DNA fragments were separated on a 2% agarose gel for 30-45 minutes at 120 V. For Meis1-GFP, embryos were first observed under a fluorescent microscope (Leica MZ16FA) to detect GFP expression prior to dissecting the AGM. Genotyping by PCR was also carried after the experiment to confirm the genotype. Given that Runx1+23GFP expression was not as strong and widespread, this approach could not be used reliably to genotype embryos from this strain at the start of the experiment. 21 Table 2.1 List of Genotyping Primers Target Forward Primer Reverse Primer Cre GTTTCGAATTTACTGACCG CGCCGCATAACCAGTGAAAC GFP CACATGAAGCAGCACGACTT TGCTCAGGTAGTGGTTGTCG LacZ GCTGGATGCGGCGTGCGGT CTGGATCAAATCTGTCGATCCTT Meis1-flox CCAAAGTAGCCACCAATATCATGA AGCGTCACTTGGAAAAGCAATGAT Sash1-flox CCATGCTGACACGAGAACCT TGTTTTGGGATGGCGTACCT Sash1 (WT) TCGAGAACTTCCATGCCCATCCTT AGTCAGGTTGCCTATCTTGGCTGT 2.3 Cell culture Human microvascular endothelial cells (HMEC) were obtained from the Centre for Disease Control and Prevention (Atlanta, GA) and cultured in MCDB 131 medium. OP9 cells were purchased from ATCC and cultured in alpha Modified Eagle Medium (α-MEM). Both culture media were supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2 mM glutamine, and 100 U each of penicillin and streptomycin. For experiments involving stimulation with 100 ng/mL of lipopolysaccharide (LPS) (Sigma-Aldrich, St. Louis, MO), HMEC were first washed with serum-free medium and incubated overnight in MCDB 131 containing 2% FBS. For the OP9 assay, cells were plated 24h prior the experiment at a density of 5,000 cells per well in a 96-well plate or 50,000 cells per well in a 4-well culture chamber slide. Embryonic cells were isolated by FACS directly onto the OP9 stromal layer and incubated for 5 days in medium supplemented with SCF (100 ng/ml), IL-3 (200 U/ml), G-CSF (100 ng/ml), Epo (2 U/ml), bFGF (1 ng/ml), and β-mercaptoethanol (50 μM). 2.4 Flow cytometry Following dissection, lungs or AGMs were dissociated into single cell suspension in PBS supplemented with 2% FBS, 1mg/ml collagenase II (Sigma-Aldrich #C6885), and 1 mg/ml DNaseI (Sigma-Aldrich #D4513). Lungs were subsequently treated with ammonium chloride for 10 minutes on ice to lyse red blood cells. Dissociated cells were then spun at 400g for 5 minutes at 4oC and resuspended in FACS buffer (PBS + 2% FBS + 1 mg/ml DNaseI) supplemented with anti-CD16/32 (7 µL/mL) for blocking. After 5 minutes of incubation on ice, FACS antibodies 22 were added at the appropriate concentration (Table 2.2) to stain the cells for 30 minutes covered from light. Cells were transferred to FACS tubes, washed by adding PBS and resuspended into 200 µL of FACS buffer with DAPI for viability. Cells were sorted on a BD FACSAria III or BD FACSAria Fusion instrument and flow cytometry data was analyzed using FlowJo software (v7.6 or v10) after the experiment. For detection of the β-Galactosidase gene-trap in Sash1+/- or Sash1-/- embryos by flow cytometry, cells were incubated at 37oC for 5 minutes following dissociation. An equal volume of 2 mM fluorescein Di-β-D-Galactopyranoside (FDG, Invitrogen #F1179) (diluted in DMEM) was added and cells were incubated for 2 minutes at 37oC in the dark. 9 volumes of ice-cold PBS containing 2% FBS, 200 mM chloroquine, and 200 mM verapamil was quickly added to quench the reaction. Cells were incubated on ice for 30 minutes, then resuspended into FACS buffer with DAPI for sorting. Table 2.2 List of FACS antibodies Antibody Source Identifier # Dilution (uL/uL) CD31-PerCP710 eBioscience 46-0311-82 1/200 CD31-BV605 BioLegend 102427 1/400 CD45.2-APC780 eBioscience 47-0454-82 1/400 EpCAM-Alexa488 BioLegend 118210 1/200 Pdpn-APC BioLegend 127410 1/200 CD41-PE BD Biosciences 558040 1/400 CD206-Alexa488 BioLegend 141709 1/400 CD11b-APC BD Biosciences 553312 1/400 CD44-APC BioLegend 103011 1/400 2.5 mRNA expression analysis (RT-qPCR and ddPCR) Whole lung tissues used for gene expression analysis were snap frozen and ground with a mortar and pestle in liquid nitrogen on dry ice. mRNA was extracted using 1 mL of Trizol (Invitrogen # 15596-018) following the manufacturer’s protocol. For experiments with FACS-isolated cells from the AGM, total RNA and DNA was extracted using an AllPrep DNA/RNA Micro Kit 23 (Qiagen #80284) or RNA was extracted with a RNeasy Micro Kit (Qiagen #74004) as per manufacturer’s protocol. RNA was eluted with 14 µL of RNase-free water and DNA was eluted in 30 µL of the supplied buffer. Up to 1 µg of RNA was used to synthesize cDNA. Reverse transcription was carried out with SuperScript II enzyme (Invitrogen #18064-071) and random primers (Invitrogen #48190011) at 42oC for 1 hour or Maxima H enzyme (Invitrogen #EP0752) and oligo(dT)18 (ThermoFisher # SO132) at 50oC for 30 minutes. The resulting cDNA was stored short-term at -20oC until further analysis. Quantification by real-time PCR (qPCR) was performed on a StepOnePlus instrument (ThermoFisher) and each reaction was set up in triplicates using Power SybrGreen Master Mix (ThermoFisher #4368708), 1 µM of specific primers (Table 2.3), and 2.5 µL of an appropriate cDNA dilution. Threshold cycle (CT) values were normalized against Gapdh expression and fold change was calculated using the delta-delta-CT method. For samples with low RNA input, gene expression was quantified using Bio-Rad’s QX200 droplet digital PCR (ddPCR) system. Between 2-5 µL of sample cDNA was used as input and mixed with 100 nM of specific primers (Table 2.3) and EvaGreen Supermix (Bio-Rad #186-4034) to generate the droplets. Data was analyzed with Bio-Rad’s QuantaSoft software. 24 Table 2.3 List of qPCR/ddPCR primers Gene Forward Primer Reverse Primer Aqp5 TATCCATTGGCTTGTCGGTC GAAAGATCGGGCTGGGTTC Gapdh TGCAGTGGCAAAGTGGAGAT TTTGCCGTGAGTGGAGTCATA Pdpn CGTTTGGTTCTGGGACTCTG CATTAAGCCCTCCAGTAGCA Sftpa AATGGGAGTCCTCAGCTTGC CCGGCTCTGGTACACATCTC Sftpb TGCTTGATGTCTACCTGCCC AGCAGGAGAACTGTGTAGCG Sftpc TGATGGAGAGTCCACCGGAT CCACCACAACCACGATGAGA Sftpd CGAGCCTGACAAACAGAGGT GGAGAGAAAGGGCAGCATGT Sash1 exon3-4 GAGAAACCCGACGCCAGC TCCACTTCTGGTGTTGACAC Sash1 exon7-8 GAAGCGCTCGCCAGGCTGAAG GGTAGGAGCCATCGGGCCAGT Sash1 exon19-20 TCGGATGGCGATCCCGAGTGG GGACGCAACACAGCCGGGAG Meis1 ATATCATGAGGGCGTGGCTG CCTGTATCTTGTGCCAACTGC Runx1 GCCTAGAAGAACCAGGTAGC TGGTAGGTGGCAACTTGTGG Gfi1 CCTGGTCAAGAGCAAGAAGG CTCCATTTTCGACTCGCCTG Spi1 AACAGATGCACGTCCTCGATA CATCCGGGGCATGTAGGAA Gata2 GCTTCACCCCTAAGCAGAGA TGGCACCACAGTTGACACA Gpr56 GCAGAACACCAAAGTCACCA TGTCTCTGCTCACTGTCTCG Pecam1 AGCTAGCAAGAAGCAGGAAGGACA TAAGGTGGCGATGACCACTCCAAT Efnb2 GCCAAATCCAGGTTCTAGCA ATGCGATCCCTGCGAATAAG 2.6 Protein expression analysis To detect Sash1 protein expression in distinct cell types, E18.5 lung endothelial cells and alveolar epithelial cells were FACS-isolated into RIPA buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% NP-40, 0.25% sodium deoxycholate, plus protease inhibitors). Protein concentration in the total lysate was quantified using the colorimetric Bio-Rad DC™ Protein Assay Kit (Bio-Rad #500-0112) as per protocol. 20 µg of protein diluted in Laemmli sample 25 buffer was denatured at 95oC for 10 minutes and loaded on a 10% polyacrylamide gel to separate the proteins by size. Proteins were then transferred onto a nitrocellulose membrane, blocked with 5% skim milk in TBS-T buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 0.05% Tween-20) for 1 hour at room temperature, and incubated overnight at 4oC in the presence of 1:1000 anti-Sash1 (Novus #NBP1-26650) or 1:10000 anti-Gapdh (Cell Signaling Technology (CST) #2118S) antibodies. The following day, membranes were washed three times with TBS-T and incubated at room temperature for 1h in 5% skim milk TBS-T with 1:5000 anti-rabbit HRP-conjugated secondary antibody (Sigma-Aldrich #A9169). Membranes were washed three more times to removed the secondary antibody and incubated with enhanced chemiluminescence (ECL, PerkinElmer #NEL105) reagents for 1 minute to detect the signal using a film developer. The Simple Western system along with Compass software (ProteinSimple) was used for protein quantification by capillary immunoassay. Proteins were separated and detected using Wes Separation Capillary Cartridge 12-230kDa (ProteinSimple #SM-W004) and Wes Anti-Rabbit Detection Module (ProteinSimple #DM-001) following the manufacturer protocol. 0.2 mg/mL of protein lysate was loaded per lane and rabbit antibodies were used against phospho-AKT Ser473 (CST #4060S), phospho-eNOS Ser1177 (CST #9571S), and beta-arrestin1 (Abcam #32099) at a 1:10 dilution or against GAPDH (CST #2118S) and AKT (CST #9272S) at a 1:50 dilution. 2.7 Immunostaining In vivo DAR4M-AM (Santa Cruz Biotechnology #sc-221530) staining was performed by intraperitoneal injection 1h prior to euthanasia of the dam. DAR4M-AM stained lung samples were frozen in Tissue-Tek® Optimal Cutting Temperature (OCT) compound and cryosectioned. 10µm sections were fixed in acetone:methanol (1:1) for 10 minutes, washed, and blocked with 5% bovine serum albumin (BSA) in PBS + 0.1% Tween-20 for 1 hour. Samples were incubated with primary CD31 (2H8) antibody (Table 2.4) for 1h followed by 3 washes with PBS and 1 hour incubation with secondary antibody. Sections were washed overnight in PBS + 0.1% Tween-20 and stained with 5 µg/mL Hoescht 33342 for 1 minute prior to mounting in PBS. A custom robotic microscope [112] was used to image fluorescence at 546nm (DAR4M) and 647nm (CD31). 26 Embryos/AGMs collected for imaging were fixed in 4% paraformaldehyde for 4 hours at 4oC. Embryos were rinsed 3 times with PBS and dehydrated overnight in 30% sucrose solution. The following day, embryos were transferred to a cryomold, embedded in OCT, flashed frozen on dry ice, and kept at -80oC. Frozen tissues were sliced into 10 µm sections and collected on a microscopy slide. OCT-embedded sections were washed with PBS, incubated 30 minutes in blocking buffer (PBS + 0.2% Triton-X + 5% chicken/goat serum) and stained overnight with appropriate dilution of the primary antibodies (Table 2.4) in blocking buffer. Following 3 washes with PBS, samples were incubated at room temperature for 1 hour with 1/200 dilution of the fluorochrome-conjugated secondary antibodies, further washed to remove unbound antibodies, and mounted in Prolong Gold mounting reagent with DAPI (ThermoFisher #P36941). For c-kit staining, sections were blocked overnight and incubated with anti-c-Kit antibody for 1h to reduce background. For cells directly on slides, cells were fixed in 4% paraformaldehyde for 5 minutes and stained as described above. Stained slides were scanned on an Axioplan II Zeiss microscope and chosen sections were imaged on Zeiss AxioImager M2 microscope with ZEN 3.2 software. Table 2.4 List of immunostaining antibodies Primary Antibodies Antibody Host species Source Identifier Dilution GFP-Alexa488 Rabbit ThermoFisher A-21311 1:100 c-Kit (D13A2) Rabbit Cell Signaling Technology 3074S 1:200 CD31 (MEC 13.3) Rat BD Bioscience 550274 1:200 CD45 (30-F11) Rat BD Bioscience 550539 1:100 CD31 (2H8) Hamster ThermoFisher MA3105 1:200 SP-C Rabbit Santa Cruz Biotechnology Sc-13979 1:200 Secondary Antibodies Antibody Host species Source Identifier Dilution Anti-Rabbit Alexa-488 Chicken ThermoFisher A-21441 1:200 Anti-Rat Alexa-594 Goat ThermoFisher A-11007 1:200 Anti-Hamster Alexa-647 Goat ThermoFisher A-21451 1:500 27 2.8 LacZ Staining To detect Sash1 expression in vivo, Sash1+/- OCT-embedded embryos were cryosectioned and fixed in 0.2% glutaraldehyde for 10 minutes. Samples were washed with 0.1 M sodium phosphate buffer containing 2 mM magnesium chloride, 0.01% sodium deoxycolate, and 0.02% nonidet-P40. Samples were stained overnight at 37oC in wash buffer supplemented with 1 mg/mL of X-gal, 2.12 g/L of potassium ferrocyanide, and 1.64 g/L of potassium ferricyanide. Slides were washed in PBS and mounted in Prolong Gold mounting reagent with DAPI (ThermoFisher #P36941) for fluorescent nuclear detection which allowed for additional co-staining with antibodies prior to mounting. 2.9 Nitric oxide rescue experiments S-Nitroso-N-Acetyl-D,L-Penicillamine (SNAP, Santa Cruz Biotechnology #sc-200319B) was injected intraperitoneally at 5 mg/kg 24h prior to harvesting the embryos. Riociguat (BAY63-2521, Selleckchem #S8135) was dissolved in DMSO and diluted 30 times in 2% methylcellulose to be administered at a dose of 10 mg/kg via oral gavage 24h prior to harvesting the embryos. Control animals received either PBS intraperitoneally or 2% methycellulose orally, respectively. Ex vivo experiments were performed on FACS-isolated AE2 cells plated in Dulbecco’s Modified Eagle Medium supplemented with 10% FBS and penicillin-streptomycin/glutamine in 48-well adherent plate. Cells were incubated at 37oC overnight in the presence of 10 mg/ml BAY41-2272 (Tocris Bioscience #4430) or DMSO (SigmaAldrich, St. Louis, MO), or 1 mM SNAP or PBS. 2.10 Single-cell RNA-sequencing and CITE-seq Embryos were dissected and cells of interest were FACS-isolated in a 1.5 mL Eppendorf tube. Cells were then pooled based on their genotype following FACS and resuspended in 40 µL of PBS + 10% FBS. Cells from 2-5 embryos were pooled for any given samples. Single-cell RNA-sequencing (scRNA-seq) and CITE-seq was performed by the Biomedical Research Centre sequencing core at the University of British Columbia (Vancouver, British Columbia). Briefly, libraries were prepared using the 10X Chromium Single Cell 3’ Reagent v3 chemistry kit as per the standard protocol from 10X Genomics (Pleasanton, California) and sequenced on an Illumina Nextseq instrument. 28 For CITE-seq, cells were conjugated with the oligo-tagged antibodies of interest before FACS during the antibody staining step of the flow cytometry protocol. Each of the following antibodies was used at a final concentration of 0.2 ug/100 µL: TotalSeq-A0012 anti-mouse CD117 (BioLegend #105843), TotalSeq-A0443 anti-mouse CD41 (BioLegend #133937), TotalSeq-A0110 anti-mouse CD43 (BioLegend #143211), TotalSeq-A0157 anti-mouse CD45.2 (BioLegend #109853), and TotalSeq-A0191 anti-mouse/rat/human CD27 (BioLegend #124235). 2.11 Data analysis 2.11.1 Pathway Analysis Microarray data from WT and Sash1-/- CD31+ lung cells (GEO: GSE123889) [109] was analyzed using desktop GSEA (version 3.0) to compare gene expression profiles against pathways from MSigDB hallmark, canonical, and curated datasets. Custom genesets derived from top differentially expressed genes at various developmental stages [113, 114] were also included in the analysis. GSEA comparing WT and Sash1-/- cells in cluster of the scRNA-seq dataset was performed in R using the fgsea package [115] and the hallmark pathways. Genes up/down-regulated during normal EHT progression based on our scRNA-seq dataset were used to create custom gene signatures that were included in the analysis. Pathways with a Benjamini-Hochberg adjusted p-value < 0.1 were reported. To identify biological processes related to gene profiles, gene ontology (GO) term analysis was performed in R using gprofiler2 [116]. Selected terms were reported as significantly enriched based on hypergeometric p-value < 0.05 after correction for multiple testing (using the default g:SCS algorithm). 2.11.2 Single-cell data scRNA-seq and CITE-seq data was analyzed in R using the Seurat package [117]. Cells were filtered based on the number of features (genes) detected (> 1500) and mitochondrial content (< 10%) for quality control purposes. Uniform Manifold Approximation and Projection (UMAP) was used to reduce the dataset using the top 10 dimensions based on principal component analysis. Each cluster was then manually assigned to a most likely cell type based on the top 29 genes specifically enriched in that cluster as identified by the FindMarker function in Seurat, which uses the non-parametric Wilcoxon rank sum test. Genes were considered differentially expressed at adjusted p-value < 0.1 based on Bonferroni correction. Monocle3 [118] was used in R to reconstruct the EHT trajectory along a pseudotime, according to default settings. DESeq2 package [119] was used for analysis of differentially expressed genes (DEG) between the two genotypes within specific cell clusters. The gene expression matrix was first filtered for genes expressed in at least 10 cells and applied a zero-inflated binomial distribution model (ZINB-WaVe, [120]) to account for many zero values in single-cell dataset. DESeq2 was run using recommended settings. Genes were retained as differentially expressed based on a Wald test p-value < 0.1 cutoff after Benjamini-Hochberg multiple testing correction. 2.11.3 Statistical analyses Unless stated otherwise in the text or figure legend, p-values were calculated using a two-tailed Student’s t-test in GraphPad Prism 7.0 and considered to be statistically significant below a threshold of 0.05. The frequency of HE cells was approximated using Extreme Limiting Dilution Assay (ELDA) calculation from the online tool [121]. 30 CHAPTER 3: Endothelial Sash1 signaling contributes to lung maturation via nitric oxide secretion 3.1 Introduction A classical approach to studying gene function is to knockout the gene of interest and look for changes in phenotype. In cases of essential genes, this creates a lethal phenotype. Homozygous deletion of the Sash1 gene in mice (Sash1-/-) leads to perinatal lethality in less than an hour after birth due to respiratory distress [109]. Pups from Sash1+/- crosses are born 19 days after the detection of the vaginal plug (around E19-19.25). At that time, newborn Sash1-/- pups are cyanotic and appear to be gasping for air until they finally succumb [109]. Work from our group revealed that the lungs of Sash1-/- pups failed to inflate based on a buoyancy test which explains the respiratory distress phenotype [109]. Therefore, Sash1 appears to have an essential role in pulmonary development. However, Sash1 contribution during embryogenesis has not been reported before and its vital function is unknown. Lung organogenesis can be categorized into distinct morphological stages. In the mouse, the timing of birth occurs during the saccular stage which is initiated around E17.5 and lasts until post-natal day (P)5 [15, 122]. This stage is characterized by the differentiation of type 1 and type 2 alveolar epithelial cells, as the number of saccules and vascularization increases [122]. Alveolar epithelial type 1 cells (AE1C) are thin and elongated cells that cover most of the alveolar surface, forming the air-blood barrier along with the capillary endothelium [123]. On the other hand, alveolar epithelial type 2 cells (AE2C) have a more cubical shape and are responsible for secreting surfactants that reduce surface tension and facilitate alveoli inflation [123]. Thus, the saccular phase is setting the stage for proper oxygen transfer during respiration at birth by thinning of the alveolar wall and initiating surfactant production. Unlike in the mouse, the saccular stage is mostly pre-natal in human, at week 24-38 of gestation, and corresponds to the limit of viability for premature babies reflecting the importance of this maturation step [124]. Indeed, one of the major complications of preterm births in human is respiratory distress due to incomplete lung maturation which often leads to complication such as bronchopulmonary dysplasia [125]. While management of premature delivery has improved over the last decades, treatments such as surfactant therapy do not address the underlying pulmonary immaturity [126]. 31 Therefore, identifying major pathways involved in late-stage lung development may be of clinical significance. In this study, we investigated how endothelial expression of Sash1 affects alveolar development. First, we determined that the defect was not structural and then demonstrated that nitric oxide secretion downstream of Sash1 signaling in endothelial cells is an important modulator of lung maturation. 3.2 Results 3.2.1 Loss of Sash1 does not significantly affect the lung vasculature Sash1 was reported to be mainly expressed in endothelial cells of the adult lung in mice [90] and a recent scRNA-seq dataset suggested a similar pattern in the embryonic lung, although some club cells and fibroblasts might express it at low levels [127]. To confirm Sash1 restriction to lung endothelial cells, endothelial cells (CD31+CD45-EpCAM-) and alveolar epithelial cells (EpCAM+CD31-CD45-, which enriches for both type 1 and type 2 cells) were FACS-isolated from E18.5 WT lungs and western blotting was performed using an antibody against Sash1. As hypothesized, Sash1 protein expression was detected in the lysate of the endothelial cell fraction but not in alveolar cells (Fig. 3.1A). Therefore, we hypothesized that pulmonary dysfunction at birth in Sash1-/- pups originates from a defect within the endothelium. However, loss of Sash1 expression had no obvious significant effect on the lung vasculature itself. A similar number of endothelial cells were found in the lungs of WT and Sash1-/- embryos at E18.5, the latest time point we can study in utero before birth, as quantified by flow cytometry analysis (Fig. 3.1B). Similarly, cross-sections of Sash1-/- lungs stained for CD31 showed endothelial cells density, determined by the fraction of positive CD31 pixels per surface area, and distribution pattern to be comparable to WT lungs at both E17.5 and E18.5 (Fig. 3.1C, D, E). Transmission electron microscopy (TEM) revealed perfusion of the blood vessels, as defined by the presence of circulating red blood cells, around Sash1-/- alveoli and showed continuous segments of elongated endothelial cells juxtaposed next to alveolar cells separated by the basal lamina in both WT and Sash1-/- samples (Fig. 3.1F). Altogether, this data supports that the expression of Sash1 is mainly restricted to endothelial cells, but that there is no evidence of gross anatomical or architectural defect of the lung. This suggests that the defect might be in the function of those cells and reside in the signals that endothelial cells can provide to the surrounding environment. 32 Figure 3.1 Sash1-deletion does not significantly affect pulmonary vascularization A) Western blot for Sash1 expression in E18.5 lung alveolar epithelial cells (AEC) and endothelial cells (EC). B) Quantification of CD31+CD45-EpCAM- endothelial cells in E18.5 lungs by flow cytometry (n = 5 per genotype). C) Immunostaining of the endothelial marker Pecam1 (CD31) on E18.5 lung tissue section and quantification of D) endothelial density per surface area and E) distribution of endothelial cells relative to each other. F) Electron micrographs of E18.5 lung alveoli. Asterix (*) labels red blood cells, basal membrane between EC and AE2C is identified by black arrows, and white arrows show glycogen accumulation. (ns = non-significant p-value based on Student t-test) 33 3.2.2 Endothelial-deletion of Sash1 impairs alveolar maturation Pulmonary surfactant, that is composed of lipids, surfactant-associated proteins, and cholesterol, is responsible for reducing surface tension and facilitates lung inflation [128-130]. Four types of surfactant-associated proteins (Sp) are secreted by alveolar epithelial type 2 cells (AE2C): Sp-A, Sp-B, Sp-C, Sp-D [128, 131, 132]. Primarily, Sp-B and Sp-C are responsible for reducing surface tension and their expression is mainly limited to mature AE2C, with some Clara cells also expressing Sp-B [128, 131]. Therefore, Sp-C is often used as a marker for differentiation into mature AE2C. Given that Sash1-/- lungs failed to inflate at birth, we looked at expression of surfactant mRNA transcripts (Sftpa, Sftpb, Sftpc, and Sftpd) in whole lungs lysate. Interestingly, a significant reduction in the levels of all surfactant transcripts was noted in Sash1-/- samples compared to WT littermates at E18.5 (Fig. 3.2A). Therefore, the lack of surfactant secretion by AE2C could partially explain the respiratory distress observed at birth. In addition to the surfactant defect, apparent increase in glycogen accumulation concomitant with decrease of properly formed lamellar bodies was observed in AE2C (Fig. 3.1F and [109]), which is a clear indication of immaturity. During development, AE2C are derived from a bipotent progenitor that gives rise to both AE2C and alveolar epithelial type 1 cells (AE1C) [133]. Therefore, a lack of differentiation into both mature AE1C and AE2C could reflect a broader defect in lung maturation as opposed to a specific role of Sash1 on activation of surfactant transcription. Expression of the AE1C-specific gene Aqp5 was also significantly reduced in Sash1-/- lungs whereas the AE1C marker Pdpn showed a trend that was not statistically significant for all the samples (Fig. 3.2B). To validate that this phenotype was caused by a lack of Sash1 expression in endothelial cells, we created a conditional-knockout mouse model using a VE-Cadherin (VEC) driven Cre recombinase to excise the Sash1 gene in VEC-expressing endothelial cells. Specifically, VEC-cre mice purchased from The Jackson Laboratory were crossed twice with a Sash1-floxed mice (described in Chapter 2.1) to generate Sash1fl/fl;VEC-Cre embryos. This Cre-loxP system is a powerful tool to achieve tissue-specific deletion of genes and particularly useful to investigate a context-specific function, especially when a constitutive knockout might trigger multiple defects. Our mice were kept at heterozygous levels for Cre to avoid potential toxicity or unwanted effect associated with high Cre expression [134]. While the versatile Cre-loxP system is widely used, 34 the potential for inconsistent deletion, leading to mosaicism, and specificity of the promoter remain the main limitations [134]. Uncharacterized expression of the promoter, sometimes at earlier stages of development, can lead to deletion in untargeted tissue, which prompt us to use a well-studied Cre-reporter line. Additionally, incomplete bi-allelic recombination can sometimes result in remaining expression of the targeted gene. Here, we were able to validate complete excision of the floxed allele through regular testing by qPCR in FACS-isolated endothelial cells from embryonic lungs in this model (Fig. 3.2.C). Curiously, Sash1fl/fl;VEC-Cre pups generated from these crosses were viable despite excellent excision in endothelial cells. Nonetheless, lungs from E18.5 Sash1fl/fl;VEC-Cre embryos also had reduced expression of surfactants, including the AE2C specific marker Sfptc, and showed a significant decrease in both AE1C-related genes Aqp5 and Pdpn when compared to WT littermates (Sash1fl/fl) (Fig. 3.2D, E), as expected from the endothelial expression of Sash1 (shown in Fig. 3.1A).While lethality at birth was not observed, this immature epithelial phenotype was consistent with the observation made in the constitutive-knockout mice, with the exception that the reduction in Pdpn transcript levels also reached significance in the conditional-knockout. This suggests that other factors important for lung inflation (such as compliance of the chest wall or mechanical forces of the diaphragm) might contribute to the pulmonary distress at birth in Sash1-/- neonates, in addition to the lack of surfactant production by immature AE2C. The fact that the two AE1C markers do not always correlate in these experiments could be due to slightly different expression patterns where Pdpn is already expressed in some bipotent progenitors [133, 135]. Nevertheless, with a loss of Aqp5 and Sftpc as specific markers for differentiated AE1C and AE2C respectively, this experiment demonstrates that endothelial-restricted expression of Sash1 impacts alveolar epithelial development. However, how this endothelial-epithelial interaction is regulated is unknown. 35 Figure 3.2 Loss of endothelial Sash1 impairs differentiation of alveolar epithelial cells (A, B) Expression of (A) AE2C and (B) AE1C specific genes in total RNA extract from E18.5 lungs by qPCR, normalized to Gapdh (n= 7 WT, 10 Sash1+/-, 6 Sash1-/-). (C) Validation of Sash1 deletion at the mRNA levels in FACS-isolated endothelial cells in Sash1fl/fl;VEC-Cre E18.5 lungs (n = 4 Sash1fl/fl and 5 Sash1fl/fl;VEC-Cre) (D, E) Expression of (D) AE2C and (E) AE1C specific genes in total RNA extract from E18.5 lungs by qPCR, normalized to Gapdh (n = 5 Sash1fl/fl and 6 Sash1fl/fl;VEC-Cre) (F) Enrichment plot for gene signatures enriched at E18 vs E17 [114] and enriched in the saccular stage compared to canalicular [113]. (Student t-test, p-value: * < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001). Dr. Grigorios Paliouras performed the experiment in panel (A). 36 3.2.3 Sash1 signaling contributes to nitric oxide production in the developing lung Ding et al. showed that pulmonary endothelial cells play a major role in adult lung regeneration via activation of VEGFR2 and FGFR1 to secrete MMP14 which promotes expansion of AE2C and AE1C [24]. Whether a similar mechanism is used by endothelial cells to drive alveolar maturation during developmental stage is not known. In order to better understand the role of Sash1 in the pulmonary endothelium, microarray analysis was performed on FACS-isolated CD31+ endothelial cells from E18.5 WT and Sash1-/- lungs. Gene set enrichment analysis (GSEA) was performed to identify pathways that were differentially regulated. Interestingly, when compared to transcriptomic datasets of lung developmental stages [113, 114], the gene expression pattern of E18.5 Sash1-/- lung endothelial cells was more similar to an earlier time point (E17 and associated with the canalicular stage) whereas WT cells were enriched for gene signatures representative of E18 and the saccular stage of development as expected (Fig. 3.2F). This observation further supported a developmental delay in the absence of Sash1 expression. Amongst the most significant pathways enriched in WT cells, we found signatures related to NF-κB and innate immune signaling and (Fig. 3.3A, B) which was consistent with findings that our group previously described [90]. On the other hand, VEGFR and FGR signaling pathways were not significantly impacted by Sash1-deletion (Fig. 3.3C, D) in our model. Similarly, MMP14 was not significantly differentially expressed in our microarray dataset. Alternatively, genes related to Akt and nitric oxide (NO) signaling were significantly enriched in WT cells (Fig. 3.3E, F). This suggested an interesting avenue to explore since mice deficient for eNOS also display pulmonary defects at birth with roughly 40±5% of the pups dying of respiratory distress [136]. eNOS-null pups that die within a few hours after birth are cyanotic and have poorly developed saccules with a lack of CD31+ cells by immunostaining [136]. Although this phenotype does not completely recapitulate the defect observed in Sash1-/- pups, given that we could not detect ultrastructural abnormalities, we were able to show that eNOS-null pups also have reduced surfactant expression at the mRNA level prior to birth (Fig. 3.4). eNOS is an important factor in angiogenesis and it is possible that complete abolition of eNOS triggers additional and more pronounced defects than what could possibly be a dose-dependent and/or timing-dependent effect in Sash1 signaling that we observed. 37 In addition to NO, Akt has been reported to play a role in lung development. Akt is activated upon phosphorylation of its Thr308 and Ser473 residues by phosphatidylinositide-3-kinase (PI3K) [137]. Pharmacological inhibition of Akt activation using PI3K inhibitors LY294002 and wortmannin showed reduced SP-C expression in an early fetal lung explant culture system [138]. Furthermore, Akt is known to activate eNOS through phosphorylation of Ser1177 which leads to NO production [139]. Therefore, the implication of the Akt-eNOS pathway in Sash1-driven lung maturation was further explored. Figure 3.3 Identification of Akt and nitric oxide pathways downregulation in Sash1-/- lung endothelial cells (A-F) Enrichment plots of selected gene sets in gene set enrichment analysis (GSEA) on microarray data from E18.5 WT and Sash1-/-endothelial cells 38 Figure 3.4 eNOS-null embryos showed reduced surfactant transcript prior to birth Expression of surfactant transcripts in total RNA extract from E17.5 lungs (one day prior birth in that mouse strain) by qPCR, normalized to Gapdh (n= 8 for each genotype) (Student t-test, p-value: * < 0.05, ** < 0.01, *** < 0.001) Nitric oxide (NO) is a molecule with a very short physiological half-life (typically 0.1-10 seconds) in the body [140]. It reacts readily with oxygen, free radical species, and oxyhemoglobin which maintains it at low steady-stead concentration [140] making it challenging to accurately detect and quantify, particularly with high sensitivity when present at very low levels such as in an embryo. Fortunately, several tools have been developed including the use of cell-permeable fluorescent dyes. Diaminorhodamine-4M (DAR-4M)-AM reacts with NO and oxygen intracellularly to form the triazolo-rhodamine analog (DAR-4M T) that exhibits 840-fold greater fluorescence when excited by a 550nm light source [141] and can therefore be detected by microscopy. Pregnant dams were injected intraperitoneally with DAR-4M AM 1h hour before harvesting the embryos at E17.5, one day before we see a significant change in surfactant levels. Embryos were then frozen in OCT and later cryosectioned for analysis by microscopy. Both Sash1-/- and Sash1fl/fl;VEC-Cre embryos showed significant reduction in DAR-4M staining compared to their respective WT littermates (Fig. 3.5) supporting that less nitric oxide is being secreted in the lungs when Sash1 expression is lost. 39 Colocalization of NO with AE2C specifically was challenging to achieve by microscopy given that the best marker of AE2C described in the literature is SP-C, which we knew was lost in Sash1-/- cells. Nevertheless, we developed a strategy to enrich for AE2C by flow cytometry by depletion of endothelial cells (CD31) and hematopoietic cells (CD45) and further enriching for epithelial cells (EpCAM+) that expressed low levels of the AE1C marker Pdpn (also T1α) as shown in Fig 3.6A. To validate that this gating strategy enriches for AE2C, cells from WT E18.5 embryos were FACS-isolated directly onto a microscopy slide, fixed, and stained with an antibody against SP-C. About 80% of the cells isolated within the EpCAM+T1αlo gate expressed some level of SP-C protein by immunostaining (Fig. 3.6A) whereas SP-C was not expressed in any of the cells contained within the AE1C-enriched gate (EpCAM+T1αhi). Given that DAR-4M emits a strong fluorescent signal, we were able to detect it by flow cytometry using a 561nm laser with a 586/15 filter (Fig. 3.6B), akin to the common fluorochrome phycoerythrin (PE). Therefore, following intraperitoneal injection of DAR-4M AM, cells from Sash1fl/fl;VEC-Cre lungs and Sash1fl/fl (WT) littermates were stained with the antibody cocktail and events acquired by flow cytometry to quantify DAR-4M intensity specifically in AE2C. Interestingly, a small but significant reduction (p = 0.03) in DAR-4M geometric mean fluorescence intensity (gMFI) was observed in the AE2C-enriched population of Sash1fl/fl;VEC-Cre cells (Fig. 3.6C). The difference in DAR-4M gMFI within the AE1C-enriched gate was not statistically significant (p = 0.08, Student-t-test) between the two genotypes although a downward trend was also observed (Fig. 3.6D). If taken together, there was overall a significant decrease in DAR-4M gMFI in total EpCAM+ alveolar epithelial cells (Fig. 3.6E) which might mean that AE2C are slightly more affected or that increasing the sample size could make the trend statistically significant in AE1C given that there appeared to be a larger standard deviation in this epithelial subpopulation. Nevertheless, showing a direct transfer of NO from one cell to another is very challenging given the short half-life and high reactivity of NO as mentioned before. This experiment, using an endothelial-specific deletion of Sash1, supports that the presence of NO in AE2C in vivo is in part impacted by signaling coming from endothelial cells. 40 Figure 3.5 Endothelial Sash1-deletion reduces nitric oxide secretion in lungs in vivo Nitric oxide detection by DAR-4M AM injection and quantification in (A, B) WT lungs (n =4) compared to Sash1+/- (n = 5) and Sash1-/- (n = 9) and in (C, D) conditional-knockout embryos (n= 3 Sash1fl/f, 4 Sash1fl/fl;VEC-Cre). (Student t-test, p-value: * < 0.05). Megan Fuller contributed to the immunostaining in panel (A). 41 Figure 3.6 Endothelial Sash1-deletion decreases NO level in alveolar epithelial cells (A) Flow cytometry strategy to gate for AE2C and validation by immunostaining for SP-C (green) against DAPI (blue). (B) Detection of DAR-4M signal in AE2C-enriched cell population by flow cytometry as proxy for nitric oxide (C-E) Quantification of DAR-4M geometric mean fluorescence intensity (gMFI) in alveolar epithelial cells based on gating strategy (n= 3 for each genotype) (Student t-test, p-value: * < 0.05) 42 3.2.4 A nitric oxide donor can rescue the Sash1-knockout phenotype NO can react to produce secondary reactive species but also acts as an important signaling molecule via activation of soluble guanylate cyclases (sGC), which in turns lead to cyclic guanosine monophosphate (cGMP) production [86]. To assess whether NO directly affects maturation of AE2C, we FACS-isolated a population of AE2C-enriched cells (using the gating shown in Fig. 3.6A) and treated the cells in ex vivo culture with the NO donor S-Nitroso-N-actetylpenicillamine (SNAP) or PBS as the vehicle control. This experiment was repeated using the compound BAY41-2272, which activates sGC independently of NO [142], or DMSO vehicle control. Primary AE2C do not keep well in culture and tend to differentiate rapidly. However, after being treated overnight ex vivo, cells that received either SNAP or BAY-41-2272 showed higher expression of Sftpb and Sftpc as compared to vehicle-treated cells (Fig. 3.7A, B). This suggests that NO signaling can support AE2C maturation or differentiation independently of the endothelium. Therefore, we tested whether treatment with the NO-donor SNAP or an sGC activator could rescue the Sash1-/- phenotype in vivo. For this in vivo experiment, the analog BAY63-2521 (Riociguat) was used instead of BAY41-2272 as it was readily bioavailable via oral gavage with better characterized pharmacokinetics [143]. In addition, Riociguat could be dissolved in 2% methycellulose as opposed to DMSO which could reduce any additional toxicity in vivo. Both treatment with SNAP and Riociguat rescued expression of surfactants (as markers for AE2C) and Aqp5 or Pdpn (as markers for AE1C) to levels comparable to WT non-treated embryos (Fig. 3.7D-G). Curiously, treatment of WT cells seemed to have the opposite effect, showing reduced surfactant protein levels especially with the NO-donor SNAP, suggesting that the level of NO might need to be tightly regulated. 43 Figure 3.7 NO and cGMP activation can rescue the differentiation phenotype in Sash1-deficient embryos (A-B) mRNA expression of Sftpb and Sftpc in AE2C cultured ex vivo following treatment with (A) BAY41-2272 or (B) SNAP measured by ddPCR and normalized to Gapdh. (D-G) mRNA expression of (D, F) surfactant and (E, G) AE1C related-genes in whole lung at E18.5 after in vivo treatment with (D-E) SNAP or (F-G) Riociguat measured by RT-qPCR and normalized to Gapdh. (Student t-test, p-value: * < 0.05, ** < 0.01, *** < 0.001, ns = non-significant) 44 3.2.5 Sash1 interacts with β-Arrestin1 to activate the Akt-eNOS pathway A goal of this project was to characterize novel pathways to better understand the various steps that can be dysregulated or modulated. Using a yeast-two-hybrid approach with the full-length human protein SASH1, our group previously identified beta-arrestin1 as a Sash1-binding partner, which was further validated by co-immunoprecipitation experiments [109, 144]. Interestingly, mice deficient for both beta-arrestin1 and beta-arrestin2 exhibit pulmonary defects leading to neonatal mortality with a phenotype similar to Sash1-/- pups including lack of differentiated AE1C and AE2C marked by lower Aqp5 and T1α (Pdpn) expression, reduced SP-C, and glycogen accumulation at the expense of properly organized lamellar bodies [145]. In addition, no change in branching morphogenesis was noted in that model [145], akin to our model. However, a pathway was not defined in this aforementioned study. Therefore, we decided to test whether the interaction between Sash1 and beta-arrestin1 could affect Akt-eNOS signaling to better understand the molecular pathway involved. Both Sash1 and beta-arrestin1 were knocked down in human microvascular endothelial cells (HMEC) using specific shRNA. Lipopolysaccharide (LPS) was used as a known Tlr4 ligand to activate Sash1 signaling [90], and western blot or capillary immunoassay (WES by ProteinSimple) was performed to assess the activation of Akt-eNOS based on phosphorylation. Both knockdown of Sash1 or beta-arrestin1 was shown to block phosphorylation of Akt and eNOS in response to LPS stimulation (Fig. 3.8A, B). To test the requirement of Sash1-beta-arrestin1 interaction for the activation of Akt-eNOS in that cell line, truncated proteins for Sash1 and beta-arrestin1, expressing only their cognate interacting domains [144], were transduced into HMEC to act as dominant-negative molecules. Overexpression of either the SAM1 domain of Sash1 (which binds beta-arrestin1) or the C-terminal domain of beta-arrestin1 (amino acid 319-418, which binds Sash1) blocked the phosphorylation of Akt and eNOS induced by LPS in HMEC (Fig. 3.8C, D). 45 Figure 3.8 Sash1 activates the Akt-eNOS pathway via interaction with beta-arrestin1 (A, B, D) HMEC were transduced with (A) shRNA targeting SASH1 or (B) beta-arrestin1 or (D) constructs overexpressing the SAM1 domain of SASH1 (aa 633-697) or the C-terminus of beta-arrestin1 (aa 319-418). Cells were induced with 100ng/mL LPS and phosphorylation of AKT (Ser473) and eNOS (Ser1177) was analyzed by (A) western blot or (B, D) capillary immunoassay. Quantification was performed by (A) densitometric scan or (B, D) using the area under the peak. (C) Schematic of SASH1 and ARRB1 interaction and the mutant construct overexpressed in (D). (3 < n < 5 for the various conditions) (Student t-test, p-value: * < 0.05, ** < 0.01, *** < 0.001). Experiment in panel (A) was performed by Ashley R. Clayton. 46 3.3 Discussion In trying to understand the respiratory distress phenotype in Sash1-/- pups, this study provided evidence for a cell non-autonomous signaling pathway, with an endothelial origin, to support alveolar maturation during late-stage lung development. The findings presented in this chapter show that NO promotes alveolar epithelial cell differentiation and that activation of the Akt-eNOS signaling axis downstream of the interaction between Sash1 and beta-arrestin1 in endothelial cells is a major source of pulmonary NO in utero (Fig. 3.9). Interestingly, deletion of Sash1 did not significantly impact vascularization of the lung despite its endothelial expression. Thus, the work described in this chapter outlines a novel paracrine function of the endothelium in lung organogenesis. Although it would be difficult to rule out a potential indirect effect of this endothelial signaling on epithelial cells (for example, via the mesenchyme), the ex vivo data strongly support a direct effect of NO on AE2C in the absence of additional cell types (Fig. 3.7B) while the in vivo data points to a reduction in NO transfer in the absence of endothelial Sash1 signaling (Fig. 3.6C). However, too much NO could potentially be detrimental based on the decrease in surfactant levels observed in WT embryos treated with SNAP (Fig. 3.7D). Although this study does not provide experimental evidence for the mechanism behind this effect, it is plausible that the dosage of NO is highly important. Excessive NO can cause cell death by increasing nitrosative stress [146] which could alter the gene expression profile. In addition, NO is well known to regulate vascular tone and inhaled NO is used to treat pulmonary hypertension [147]. Therefore, systemic administration of SNAP and Riociguat might have additional beneficial or undesirable side effects which should be further investigated. In addition to surfactant deficiency, Sash1-/- lungs exhibit morphologically immature AE2C, characterized by a lack of lamellar bodies and cytoplasmic glycogen accumulation, and have lost expression of AE1C markers. Hence, the pathway identified by studying this model revealed an underlying mechanism behind a broader alveolar immaturity. Whether NO is acting on a common progenitor, directly on both nascent AE1C and AE2C independently, or more broadly delaying development is not clear from this study. There is still a need to better understand what drives terminal differentiation of the alveoli, but the endothelium cannot be ignored in this process. 47 Figure 3.9 Schematic summary of Sash1 pathway in lung maturation Our results suggest that the interaction between Sash1 and beta-arrestin1 can lead to eNOS activation via the intermediate Akt in endothelial cells. NO secretion in the lung contributes to maturation of alveolar epithelial cells which is required for surfactant secretion and proper respiration. 48 CHAPTER 4: Sash1-deletion affects the emergence of hematopoietic stem cells during the endothelial-to-hematopoietic transition 4.1 Introduction Sash1 expression is essential during late embryogenesis for mice to survive and Chapter 3 explored how Sash1 plays a critical role in lung development. However, whether Sash1 also significantly impacts on the development of other organs/systems has not been previously studied. As discussed in previous chapters, the endothelium is part of every developing organs and Sash1 expression was found in many adult tissues. Therefore, could there be additional defects in Sash1-/- pups that are not lethal but could have an embryonic origin? Such developmental defects could potentially manifest later in life but in this case the effect would be masked by the neonatal lethality of the lung phenotype. Preliminary work performed in our lab (Paliouras, G., unpublished) showed that Sash1-/- embryos have a lower frequency of hematopoietic stem cells (HSCs) in the fetal liver at E14.5 when compared to WT littermates. This was best evidenced by a limiting dilution transplant assay where different numbers of fetal liver cells were transplanted into a recipient mouse that had been irradiated to ablate its own hematopoietic cells. Using a range of cell doses for transplantation allowed for inference of HSC frequency based on the number of recipients that showed long-term (24 weeks) reconstitution of the hematopoietic system. In this experiment, fewer recipients of Sash1-/- cells showed reconstitution at low doses of cells transplanted, reflecting a lower frequency of HSC (estimated frequency of 1 HSC in 59,280 Sash1-/- cells compared to 1 in 12,880 WT cells (p-value = 0.012); Paliouras, G., unpublished). However, at high doses even Sash1-/- liver cells were able to provide long-term multilineage reconstitution in an adult mouse suggesting that these few HSCs would most certainly be able to sustain the needs of the embryo until birth. Given that HSCs sustain life-long hematopoiesis in the organism, it is of interest to understand what might limit their formation in the embryo. In the embryo proper, definitive hematopoiesis is characterized by the emergence of hematopoietic stem and progenitor cells (HSPCs) from an hemogenic endothelium precursor via the endothelial-to-hematopoietic transition (EHT) [32, 42]. While the expression and function of Sash1 in hematopoiesis is unknown, Sash1 is expressed in endothelial cells. Therefore, could 49 endothelial Sash1 expression during EHT affect the establishment of a definitive hematopoietic system? In this chapter, we investigated the role of Sash1 in HSPC formation in the mouse AGM during early embryogenesis. Specifically, we identified a strong phenotype on EHT induced by the loss of Sash1 and show evidence to support that Sash1 acts from the microenvironment in a cell non-autonomous manner to promote EHT progression. 4.2 Results 4.2.1 Constitutive loss of Sash1 reduces intra-aortic hematopoietic cluster formation during EHT EHT is characterized by the emergence of intra-aortic hematopoietic clusters (IAHC), which contain pre-HSPCs, from the dorsal aorta in the aorta-gonad-mesonephros (AGM) region of the mouse embryo. To address whether studying Sash1 in the context of EHT is relevant, we first performed immunofluorescent staining of serial cross-sections of the full dorsal aorta from E10.5 Sash1-/- embryos and WT littermates. Using an antibody against CD31 allowed to readily identify endothelial cells forming the dorsal aorta whereas cells within IAHC can be labeled using CD117 (c-Kit) as previously reported [32]. By analyzing serial 10µm sections to cover the entire width of the dorsal aorta, we were able to quantify the number of IAHC present in each embryo. Interestingly, the average number of large IAHC (> 4 CD31+c-Kit+ cells in a cluster) per dorsal aorta was drastically reduced in Sash1-/- embryos to about half the number seen in WT controls (Fig. 4.1A, B). Additionally, most clusters noted in Sash1-/- aortas appeared visually smaller in size (Fig. 4.1A), although this was not thoroughly quantified. This simple experiment strongly suggests that emergence of IAHC is impaired to some extent in embryos with constitutive deletion of Sash1. To further validate this point, cell composition of the AGM was analyzed by flow cytometry. Consistent with the immunostaining data, fewer pre-HSPCs (defined by surface co-expression of CD31 and CD41low but lacking the pan-hematopoietic marker CD45 [46, 148]) were observed in Sash1-/- samples (Fig. 4.1C, D) despite no difference in the number of CD31+ endothelial cells isolated from those dorsal aorta (Fig. 4.1E). Runx1 is the best characterized regulator of EHT and embryos deficient for the Runx1 gene (Runx1-/-) are completely devoid of IAHC [61]. Under normal conditions, Runx1 is expressed starting from the hemogenic endothelium and throughout all the transition stages to HSCs, therefore its expression in E10.5 AGM is particularly enriched in IAHC [56, 64]. Gene 50 expression analysis by RT-qPCR on bulk cells from dissected E10.5 AGM showed reduced Runx1 expression at the transcript level in Sash1-/- embryos when compared to WT littermates (Fig. 4.2A). Similarly, expression of additional hematopoietic related-genes such as Meis1 [103, 106] (whose role in EHT will be further discussed in the next Chapter 5) and Gata2 [68, 69] also showed this trend (Fig. 4.2B). As a control, expression of the endothelial marker Pecam1 (CD31) and arterial marker Efnb2 were not differentially expressed in WT and Sash1-/- samples (Fig. 4.2B). This experiment was performed on unsorted dissected AGMs which represents a heterogeneous mixture of cell types. Therefore, it does not distinguish between lower expression per cell or lack of a certain cell population in this context. Nevertheless, combined with a lack of IAHC formation (Fig. 4.1A, B), the fact that Runx1 expression is mainly restricted to IAHC, and the definitive requirement for Runx1 during EHT, this significant decrease observed (Fig. 4.2A) supports a defect in hematopoietic potential of the dorsal aorta in Sash1-/- embryos. 51 Figure 4.1 Fewer pre-HSPC emerge from Sash1-deficient embryos at E10.5 (A) Immunostaining and (B) quantification of IAHC in the dorsal aorta of E10.5 embryos based on CD31 (red) and c-Kit (green) expression (DAPI in blue). (n = 4 WT and 4 Sash1-/-) (C) Representative flow cytometry plot of CD41 expressing “pre-HSPC” and quantification of (D) pre-HSPC and (E) total endothelial cells in E10.5 AGM based on flow cytometry analysis. (n = 7 WT, 10 Sash1+/-, 9 Sash1-/-) (Student t-test, p-value: ** < 0.01, *** < 0.001, ns =non-significant) 52 Figure 4.2 Reduced expression of EHT-related genes in Sash1-/- AGM at E10.5 (A-C) Expression of selected EHT-related genes in bulk AGM dissected from (A, B) E10.5 and (C) E9.5 embryos, normalized to Gapdh and compared to littermates (n = 3WT, 11 Sash1+/-, 6 Sash1-/- at E10.5 and n = 4 WT, 8 Sash1-/- at E9.5) (Student t-test, p-value: * < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001, ns = non-significant) 53 4.2.2 Sash1 is required for cluster emergence, not early specification of the hemogenic endothelium The data presented in figure 4.1 clearly indicates that the progression of EHT is altered or hindered in Sash1-/- embryos. In order to better understand the function of Sash1 in this developmental process and investigate potential pathways, it is necessary to first better define where the defect lies. Given that IAHC formation is compromised, the defect seems to originate in cells that precede HSPCs themselves. Specification of a specialized endothelium with hemogenic potential, the hemogenic endothelium that gives rise to IAHC, is the first step towards hematopoietic commitment. In an early specification model, HE is defined before E9.5 whereas IAHC are not detected prior to E10 in the mouse aorta [32, 56]. Whereas most studies of EHT have focused on “pre-HSPCs” at E10.5, properly defining and characterizing early hemogenic endothelium is more challenging. However, the fact that HE cells upregulate Runx1 expression is broadly accepted. In contrast to E10.5, at E9.5 Runx1 was not differentially expressed in Sash1-/- bulk AGM cells when compared to WT littermates and the same observation was made for Meis1 and Gata2 (Fig. 4.2C). In order to better assess the formation of a functional hemogenic endothelium in Sash1-/- embryos, endothelial cells (CD31+CD45-) were FACS-isolated from E9.5 AGM at a limiting dilution of 100 cells per well in a 96-well plate pre-seeded with OP9 stromal cells (Fig. 4.3A). OP9 co-culture of HE/pre-HSPCs in media supplemented with cytokines (IL-3, SCF, G-CSF, bFGF, and Epo) promotes HSPC maturation and hematopoietic differentiation leading to the formation of visible hematopoietic colonies. By plating only 100 cells per well, we found that each well will produce either no colonies or only 1 colony (Fig. 4.3B) due to the low frequency of HE relative to endothelial cells. Therefore, this experimental design allowed for easier quantification of HE frequency. Interestingly, no difference was observed in the frequency of cells able to form hematopoietic colonies ex vivo (Fig. 4.3C). This strongly suggests that an equal number of HE cells are present per dorsal aorta at E9.5 in both WT and Sash1-/- embryos and that these cells have similar potential. These observations led to the hypothesis that individual HE cells are specified early on but are unable to emerge into large cluster at a later stage in vivo. To determine whether this hypothesis could be observed, we looked at the CD31/c-Kit immunostaining of E10.5 dorsal aorta (Fig. 4.1A). Whereas c-kit staining was ubiquitous in large 54 IAHC, some cells that also expressed c-Kit appeared “isolated” and embedded in the vascular wall (ie: a single c-Kit+ cell surrounded by CD31+c-Kit- endothelial cells) (Fig. 4.4A). The number of “single c-Kit+ cells” was higher in Sash1-/- dorsal aortas (Fig. 4.4B), balancing the decrease in large IAHC quantified previously and consistent with the hypothesis of a specified HE that cannot form large clusters. To further investigate this hypothesis, endothelial cells were FACS-isolated from E10.5 AGM and analyzed. The sorted population comprised both vascular endothelial cells and the hemogenic endothelium but was depleted of pre-HSPC by gating out CD41+ cells. Gene expression analysis by ddPCR showed a slight increase in Runx1 expression in Sash1-/- cells compared to WT littermates (Fig. 4.4C). This was the opposite of what we observed in bulk AGM samples but consistent with the hypothesis that specified HE is blocked from progressing to IAHC. Similarly, expression of Gfi1, a hematopoietic gene expressed early during EHT [58, 65], was also increased in Sash1-/- endothelial cells (Fig. 4.4C). Therefore, in Sash1-/- embryos, reduced expression of hematopoietic genes in the bulk AGM seemed to correlate with a lack of IAHC cells while a greater number of HE cells remained in the vascular layer and still upregulated some EHT genes (ie: Runx1 and Gfi1). Overall, the data suggest a model where Sash1 expression is required post HE specification to promote emergence of clusters in the dorsal aorta. 55 Figure 4.3 Ex vivo culture showed hemogenic potential in Sash1-/- E9.5 endothelial cells (A) Schematic of the OP9 co-culture assay and (B) representative image of positive and negative outcome at the end of the co-culture assay. (C) Quantification of the frequency of cells plated that produced hematopoietic colonies (positive outcome). (n = 4 WT, 10 Sash1+/-, and 7 Sash1-/-) (p-value based on Student t-test). 56 Figure 4.4 Increase number of non-IAHC hemogenic endothelial cells in Sash1-/- embryos (A) Microscopy image of “single c-Kit+ cells” in the vascular layer of Sash1-/- dorsal aorta at E10.5 and (B) quantification of the total number of “single c-Kit+ cells” in each embryo (n = 3 WT and 3 Sash1-/-). (C) Expression of hematopoietic-related genes (associated with hemogenic endothelium) in FACS-isolated endothelial cells from E10.5 AGM. Quantification performed by ddPCR and normalized to Gapdh (n = 4 WT and 5 Sash1-/-) (Student t-test, p-value: * < 0.05, **** < 0.0001, ns = non-significant) 57 4.2.3 Sash1 supports EHT progression via a cell non-autonomous mechanism An important result of the ex vivo OP9 co-culture assay described in Fig 4.3 was the fact that E9.5 Sash1-/- HE cells were able to produce colonies just as well as WT cells in those culture conditions. Starting with the same frequency of HE cells should have shown a different number of output colonies had there been a cell intrinsic defect that blocked the progression of EHT. The fact that fewer clusters are formed in vivo in the dorsal aorta but the same number of colonies are derived ex vivo suggests that microenvironmental factors may play a role in this process. Sash1 is primarily expressed in endothelial cells, but whether it is required in the hemogenic endothelium or surrounding vascular cells is unknown. To confirm that Sash1-dependent signaling was coming from the endothelium, we used the endothelial-specific Sash1 deletion mouse model (Sash1fl/fl;VEC-Cre) generated in our lab from the lung study (as shown in Chapter 3). Surprisingly, we found that conditional-deletion of Sash1 restricted to endothelial cells did not replicate the phenotype seen in Sash1-/- embryos. Specifically, no difference in pre-HSPC count was observed by flow cytometry (Fig. 4.5A) and the overall level of Runx1 expression was not altered in bulk AGM cells (Fig. 4.5B). Importantly, this lack of phenotypic difference was noted despite excellent Cre excision to knockout Sash1 alleles (Fig. 4.5C). This surprising result clearly demonstrates that another source of Sash1 signaling from the microenvironment likely influences IAHC formation. Supporting additional origins of Sash1 expression, only about 50% of its total expression in the bulk AGM was lost upon conditional-knockout in endothelial cells (as opposed to > 80% reduction in FACS-isolated endothelium) (Fig. 4.5D). A cell non-autonomous function of Sash1 would suggest that its expression is not required in emerging pre-HSPC/HSPC. To further investigate whether this is the case, the ability of CD31+CD41+ cells isolated from E10.5 AGM to differentiate and proliferate into hematopoietic colonies, as previously shown [46], was tested in an ex vivo colony-forming unit (CFU) assay (Fig. 4.5E). This experiment was performed using the Sash1fl/fl;VEC-Cre mouse model to assess a potential function for Sash1 in a cell-specific manner. To validate that any cells giving rise to colonies in the CFU assay did not escape Cre excision (which would result in expansion of a clonal population with a WT Sash1 allele) and therefore truly lacked Sash1 expression, cells were harvested at the end point of the assay after counting colonies and the genomic DNA was analyzed by ddPCR. Once again, homozygous loss of Sash1 did not significantly affect 58 hematopoietic potential ex vivo (Fig. 4.5F) and all progeny showed complete deletion of the floxed allele (Fig. 4.5G). Altogether, these experiments demonstrated that Sash1 is dispensable in HE cells or pre-HSPCs for hematopoiesis and supported a cell non-autonomous mechanism. The effect of the AGM niche during HSPC emergence is of important significance and not a novel concept although the mechanisms are still poorly understood. A potential indirect function of the microenvironment is the presence of blood flow. Two reports suggested that NO, linked to blood flow and shear-stress, positively affects IAHC emergence in the dorsal aorta [87, 88]. In light of the findings outlined in chapter 3, where Sash1 was shown to promote NO production which can affect surrounding cell types in a cell non-autonomous manner, this represented an interesting avenue to explore. Specifically, a previous study showed that intrauterine inhibition of eNOS by N-nitro-L-arginine methyl ester (L-NAME) reduces the number of c-Kit+ cells by FACS and the number of CFU present in the AGM at E11.5 [88], suggesting that fewer hematopoietic progenitors emerge from AGM IAHC in the absence of NO signaling. However, the authors also mentioned expression of eNOS mainly in emerging HSPC and surrounding endothelium [88]. Since Sash1fl/fl;VEC-Cre embryos did not display a similar phenotype as the constitutive knockout model, Sash1 may be acting via a different effector in this context. Nevertheless, preliminary experiments were performed to look at NO levels using DAR-4M AM staining. Technical difficulties were encountered with intrauterine staining where no significant signal could be detected by fluorescent microscopy despite successful injection. This could be due to poorer diffusion of the dye at earlier time points, when the embryo proper is much less accessible, or much lower levels of NO as compared to what we previously observed with the lungs at E17.5. Instead, AGM were harvested at E10.5 and immediately stained in vitro prior to performing flow cytometry. No difference was observed between Sash1+/- and Sash1-/- endothelial cells in the AGM at E10.5 based on a flow cytometry analysis (Fig. 4.6). Unfortunately, no WT embryos were present in the litter collected for this experiment to compare with Sash1-/- samples. However, previous characterization of the phenotype had not shown a significant difference between WT and Sash1+/- embryos (Fig. 4.1). Although this data is limited to a single pilot experiment, and therefore a possible involvement of NO signaling in this model cannot yet be definitely ruled out, we opted to focus on investigating a possible non-endothelial origin of the defect given the rationale mentioned earlier. 59 Figure 4.5 Endothelial-deletion of Sash1 does not impair pre-HSPC (A) Quantification of pre-HSPC in E10.5 AGM by flow cytometry (n = 13 for each genotype). (B) Runx1 expression in dissected bulk AGM at E10.5 by RT-qPCR, normalized to Gapdh (n = 4 for each genotype). (C) Excision of Sash1 floxed allele (exon 3-4) at the genomic level (gDNA) and changes in mRNA expression (cDNA) in FACS-isolated endothelial cells from E10.5 AGM. Quantification was performed by ddPCR, normalized to an exogenous locus (gDNA) or Gapdh (cDNA) (n = 7 and 9). (D) Sash1 expression at the mRNA level in bulk AGM dissected at E10.5, analyzed by RT-qPCR and normalized to Gapdh (n = 4 and 5). (E) Schematic of the CFU experiment followed by validation of Cre excision. (F) Number of colonies formed from FACS-isolated E10.5 pre-HSPC after 10 days of culture in MethocultTM and (G) validation of Sash1 excision at the genomic level in progeny cells from the CFU assay, quantified by ddPCR (n = 4 for each genotype). (Student t-test, p-value: **** < 0.0001, ns = non-significant) 60 Figure 4.6 Detection of NO in E10.5 AGM cells Flow cytometry quantification of DAR-4M mean fluorescence intensity (MFI) as a proxy for nitric oxide level in (A) endothelial cells and (B) pre-HSPC from E10.5 dissected AGM. (n = 5 Sash1+/- and 2 Sash1-/-, non-significant p-value based on Student t-test) 61 To investigate the cell non-autonomous effect of Sash1, we first needed to determine which cell types, in addition to endothelial cells, express Sash1 in the AGM. The Sash1-knockout model used in this study contains a LacZ gene-trap that leads to the expression of a truncated Sash1 transcript along with β-galactosidase. Thus, we took advantage of this β-galactosidase expression in Sash1+/- embryos as a proxy to detect Sash1-expressing cells. First, we developed a protocol to perform X-Gal staining in combination with immunostaining to detect Sash1 expression in the AGM. While this showed widespread expression of Sash1, no specific pattern of expression was obvious (Fig. 4.7A). To isolate and better define the Sash1-expressing cells, broadly dissected AGMs were dissociated into single cell suspension and incubated in the presence of fluorescein Di-β-D-Galactopyranoside (FDG), a substrate that is hydrolyzed by β-galactosidase to produce a highly fluorescent fluorescein molecule. Cells that internalized and hydrolyzed FDG thus produce a signal that is detectable by flow cytometry (Fig. 4.7B) and were FACS-isolated for single-cell RNA-sequencing (scRNA-seq). Interestingly, FDG did not co-stain with CD31+CD41lowCD45- pre-HSPC in the samples that we analyzed (Fig. 4.7C) reinforcing the hypothesis of a cell non-autonomous regulation of EHT by Sash1. In total, transcriptomes of 4772 FDG+ cells from 3 different pools, each pool composed of 3-5 Sash1+/- embryos, was obtained by scRNA-seq. ScRNA-seq provides an unbiased approach to analyze gene expression patterns and find markers that can classify each single cell into a grouping (or cluster of cells). Using principal component analysis (PCA) and uniform manifold approximation and projection (UMAP) as dimensionality reduction and visualization tool, we were able to identify 14 clusters of Sash1-expressing cells in the broadly dissected AGM region analyzed (Fig. 4.8A, B). Using the distinctive gene signatures associated with each cluster, clusters were labeled as specific cell type (Fig. 4.8C). For instance, a large population of cells with high enrichment of endothelial-specific genes (Fig.4.8B, C) was observed as expected from previous knowledge. In addition, Sash1 was found in immune cells (macrophages and neutrophils), in the developing nervous system (neural crest, neural stem cells (NSC), neurons), some epithelial cells, and mesenchymal cell populations that likely include smooth muscle cells (Fig.4.8B, C). A small population of megakaryocytes and erythrocytes were detected, however none of the cells corresponding to those clusters showed detectable levels of Sash1 by scRNA-seq (Fig. 4.8D) and we believe that those cells might have contaminated the 62 FACS fraction. Different mesenchymal populations were found with distinct signatures. The top 50 genes enriched in each of those cluster was used to performed gene ontology (GO term) analysis. One subset seemed to be derived from lateral plate mesoderm (Mesenchyme_LP) and expressed genes related to the development of both the cardiac outflow tract and limbs (ie: Hand1, Pitx1, Postn, Gata6) [149-151] (Fig. 4.8C, E). Meox1 was shown to be widely expressed in the somite and AGM mesenchyme and at E10.5 [152] which was consistent with the output of the GO term analysis while the third mesenchymal population enriched for GO terms related to the kidney (therefore labeled as “Mesenchyme_Metanephric”) (Fig. 4.8C, E). The urogenital ridges develop closely together with the mesonephros and flanks the dorsal aorta in the AGM region of the E10.5 embryo, while the neural tube develops dorsally below the aorta. Therefore, Sash1 appears to be expressed in many cells that compose the microenvironment surrounding the dorsal aorta, as shown by beta-galactosidase staining of tissue sections (Fig. 4.7A). 63 Figure 4.7 LacZ staining for detection of Sash1-expressing cells (A) Microscopy image of X-Gal staining for LacZ detection (black) combined with CD31 immunofluorescence staining (red). DA; Dorsal Aorta, V; Ventral side. (B) Representative flow cytometry plots of cells stain with FDG for LacZ detection. (C) Representative flow cytometry plot of LacZ and CD41 detection in CD31+CD45- endothelial enriched cells. Panel (A) was generated by Katherine Bilan. 64 Figure 4.8 Identification of Sash1-expressing cells by single-cell RNA-sequencing (A) Schematic of the E10.5 AGM region used to isolate Sash1-expressing cells. DA: dorsal aorta, M: mesonephros, NT: neural tube, S: somite. (B) UMAP clustering of FACS-isolated FDG+ cells based on single-cell RNA expression. (C) Expression of selected marker genes associated with different clusters representing different cell populations. (D) Sash1 expression detected in every cluster except for erythrocytes and megakaryocytes. (E) Gene ontology (GO) term analysis on the top 50 genes differentiating the three mesenchymal population. 65 Interestingly, immune signaling from monocytes/neutrophils has been reported to play a role in HSC emergence in the zebrafish [75]. Additionally, CD206+ macrophage recruitment to IAHC sites was recently associated with HSC emergence in the mouse [73]. Therefore, these cells represented an attractive option to explain the cell non-autonomous effect of Sash1. Macrophages and neutrophils found around E10.5 are blood cells derived from earlier wave of hematopoiesis likely circulating from the yolk sac (Fig. 1.1). Since they are derived from a precursor expressing endothelial markers such as VEC, the first question to address was whether Sash1 was also excised in those cells in the Sash1fl/fl;VEC-Cre model. FACS-isolated CD45+CD206+ cells (to enrich for macrophages) and CD45+CD11b+ (to enrich for neutrophil and some monocytes) both showed significant loss of Sash1 expression in Sash1fl/fl;VEC-Cre AGM (Fig. 4.9A, B), a model that does not replicate the constitutive knockout phenotype. In addition, preliminary experiments suggested that the total number of CD206+ macrophages circulating in the AGM at E10.5 was not significantly different between WT and Sash1-/- embryos (Fig. 4.9C, D) hinting that lack of early macrophage production is not responsible for the phenotype observed. Whether or not these macrophages are being recruited to IAHC sites is a relevant question but hard to assess experimentally given the transient nature of this phenomenon. A “snap-shot” in time with fixed sections as done previously to quantify clusters may not provide accurate representation and Mariani et al. relied on live-imaging technology to track these very few cells over time [73]. While in vivo live cell imaging was not performed for this study, the lack of Sash1 expression in Sash1fl/fl;VEC-Cre macrophages was strong evidence to support investigating a different cell of origin. 66 Figure 4.9 VE-Cadherin driven Cre excised Sash1 in blood cells present at E10.5 (A) Expression of surface markers selected to isolated Sash1-expressing subpopulation of macrophages/neutrophils based on single-cell RNA-sequencing data. (B) Sash1 expression in FACS-isolated blood population from E10.5 AGM by ddPCR, normalized to Gapdh (n = 4 and 2 for CD206+ cells, n = 5 and 4 for CD11b+ cells). (C, D) Quantification of CD206+ macrophage in E10.5 AGM by flow cytometry in (C) absolute count and (D) as a fraction of live cells (n = 2 WT, 5 Sash1+/-, 2 Sash1-/-). 67 4.2.4 Identification of candidate pathways to explain the blockade of HE cells during EHT In order to better characterize the regulatory mechanisms of EHT in the AGM, it will be important to determine the cells from which Sash1 signaling originates. Nonetheless, the cells ultimately affected by this potential “paracrine” effect remain HE cells that fail to produce IAHC. In order to accurately modulate the signals sent to differentiate these HE cells, for instance in an ex vivo experimental assay or in reprogramming studies, there is a need to better understand how they respond to changes in their microenvironment. In an attempt to address this question, scRNA-seq was performed on FACS-isolated CD31+ cells from E10.5 Sash1-/- and WT AGM. CD31 marks all endothelial cells, HE, IAHC cells, and is still expressed on emerging HSCs, therefore encompassing the full spectrum of cells of interest. 7,788 CD31+ WT cells from 4 pooled samples and 4801 CD31+ Sash1-/- cells from 2 pooled samples (2-5 embryos per pool for each genotype) were sequenced. Clustering analysis identified the expected populations, with a large cluster of cells expressing endothelial markers alone (ie: Cdh5, Emcn, Tie2), some intermediate pre-hemogenic endothelial (pre-HE) cells (expressing Gata2, Cd44, Cd47, and Procr (CD201) which have been shown to enrich for a population with hematopoietic potential [60, 153, 154]), a spectrum of cells undergoing EHT (ie: with expression of Runx1, Gfi1, Spi1 in addition to endothelial genes), and blood cells expressing lineage markers but lacking endothelial gene signatures (Fig. 4.10A, B). Little to no expression of Sash1 was detected at the transcript level in the EHT subset (Fig. 4.10B), consistent with the lack of LacZ detection in pre-HSPC by flow cytometry (Fig. 4.7C). In addition, a population of cells with mesenchymal markers lacking Pecam1 (CD31) expression was found as part of the dataset. Although mesenchymal cells are of great interest in this context based on the work presented above, cells with mesenchymal markers in this experiment were isolated inadvertently and were mostly detected in 1 of the 6 samples sequenced (>53% were from sample WT_3) suggesting that this could be due to a bias in the staining/gating for that sample during FACS. Hence this mesenchymal population was excluded from further analyses. Of note, a small population of cells was found to express a distinct signature associated with pluripotency (Sox2, Nanog, Pou5f1 (also known as Oct4), and Dppa3) in addition to Pecam1, the 68 mesenchymal gene Crabp1, and the endodermal gene Epcam. These cells were also omitted from subsequent analyses focusing on the stages of EHT. Further clustering analysis performed on the “EHT” sub-population alone revealed additional heterogeneity amongst these cells. Cells could be divided into “HE/pro-HSPC” based on relatively higher expression of endothelial genes, “pre-HSPC” based on upregulation of Ptprc (CD45) and two progenitor populations with myeloid and erythroid bias (expressing Cd53/Itgam and Kfl1/Gata1, respectively) (Fig. 4.10C, D). Given the divergent profile of all these cell types, separating these subpopulations was an important step to carry out before performing any analysis to compare different clusters in our dataset. Overall, Sash1-/- cells clustered together with WT cells from the same identity, showing no striking differences between the two genotypes (Fig. 4.11A). While the cells sequenced have a similar gene profile, previous experiments described in this chapter suggested a reduction in the number of pre-HSPC likely due to a blockade during EHT progression in the absence of Sash1. Therefore, the cell composition of the Sash1-/- samples was compared to WT for the relevant EHT clusters. In this dataset, a higher proportion of Sash1-/- (KO) cells was found within the “pre-HE” and “HE/pro-HSPC” compartment compared to cells from the WT samples, at the expense of more committed hematopoietic subpopulations (Fig 4.11B). Specifically, the transition from “HE/pro-HSPC” to “pre-HSPC” was significantly affected (Fig. 4.11B). Given that the “HE/pro-HSPC” population is the first population to express Runx1 and Gfi1 in this scRNA-seq dataset (Fig 4.11C), this finding is consistent with a blockade in pre-HSPC progression post Runx1 and Gfi1 upregulation in CD41- cells (Fig. 4.4C). In addition, no significant difference was observed in the level of Runx1 and Gfi1 expression between WT and Sash1-/- cells in the scRNA-seq dataset (Fig.4.11C), suggesting that the difference observed in previous experiments might be reflective of changes in cell type representation. While cells clustered well together on UMAP, a closer look at the bottleneck population of “HE/pro-HSPC” revealed a potential enrichment of KO cells in a subcluster that appeared to run adjacent to the WT trajectory from “pre-HE” to “pre-HSPC” (Fig. 4.11A). Therefore, gene expression analysis was performed to explore the differences between the two genotypes. 69 Figure 4.10 Single-cell RNA-sequencing and clustering of CD31+ cells (A) UMAP clustering of FACS-isolated CD31+ cells based on single-cell RNA expression. (B) Expression of selected marker genes associated with different clusters representing different cell population. (C) UMAP representation of the EHT subset annotated based on reclustering. (D) Expression of selected markers genes in the reclustered EHT cells to identify subpopulation. 70 Figure 4.11 Comparison between WT and Sash1-/- (KO) cells in scRNA-seq (A) UMAP representation of FACS-isolated CD31+ cells based on single-cell RNA expression, colored by genotype. (B) Distribution of cells at each step of EHT in the scRNA-seq data for each genotype. Arrows indicate transition step between two adjacent subpopulations and the difference was analyzed using a two-proportions Z-test (* p < 0.05). (C) Expression of the EHT genes Runx1 and Gfi1 across the EHT subsets and annotated by genotype. 71 Based on clustering analysis using the top 10 principal components, Sash1-/- cells overlapped well with their WT counterpart based on cell type (Fig. 4.11A), suggesting that the differences between genotypes in specific cell types are fewer than in the signature that defines a given subset. To dissect transcriptional differences between the two genotypes within each individual population of cells undergoing EHT, differential gene expression analysis was performed using DESeq2 along with a zero-inflated negative binomial distribution (Zinbwave) to account for high rates of “zero counts” in single-cell data [120]. This analysis returned a relatively low number of differentially expressed genes (DEGs) at an adjusted p-value cutoff < 0.1. Specifically, 125 DEGs were identified in the “pre-HE” population and 170 and 205 DEGs in “HE/pro-HSPC” and “pre-HSPC” respectively (Fig. 4.12A). In an attempt to find pathways dysregulated during EHT in the absence of Sash1 expression, GSEA was performed against the hallmark pathways. Very few pathways were significant (adjusted p-value < 0.1). In the “HE/pro-HSPC” population of interest, the main distinction was highlighted by the enrichment of cell cycle pathways in Sash1-/- embryos relative to their WT counterpart (Fig. 4.12B). Pre-HSPC forming IAHC are typically thought to be highly proliferative [154, 155] but whether a physiological delay in cycling activity exists in Sash1-/- IAHC cells would have to be explored further. In addition, significant pathway enrichment was observed in the “pre-HE” but not “pre-HSPC” population. Specifically, pre-HE cells in Sash1-/- embryos appeared to be enriched in an oxidative phosphorylation signature whereas glycolysis was enriched in WT embryos (Fig. 4.12B). An mTOR signature which has previously been recognized in pre-HSPC [154] was also enriched in WT pre-HE cells. The transition from endothelial to hematopoietic cells is often represented as a step-wise progression involving intermediate cells. In our scRNA-seq dataset, 1026 genes were found differentially expressed between the WT cells labeled as “EC” and “pre-HE” based on the “FindMarkers” function in Seurat which uses the non-parametric Wilcoxon rank sum test. Similarly, 887 DEGs were identified between “pre-HE and “HE/pro_HSPC” and 266 DEGs between “HE/pro-HSPC” and “pre-HSPC”. To evaluate whether the DEGs identified between the two genotypes could reflect a delay in the progression of EHT, GSEA was performed against the homemade signatures from the genes that varied between each step. Interestingly, Sash1-/- cells in the “pre-HE” and “HE/pro-HSPC” cluster had a specific expression pattern that was 72 more similar to genes that would normally be downregulated during the transition from “EC” to “pre-HE” or “pre-HE” to “HE/pro-HSPC” respectively. In contrast, WT cells showed enrichment for genes that are upregulated at those specific transition steps (Fig. 4.12C), suggesting that Sash1-/- cells might have a slightly more immature gene expression profile than expected for the subpopulation where they are found. Figure 4.12 Differential gene expression analysis in WT vs Sash1-/- EHT cells (A) Volcano plot representing genes differentially expressed in WT and Sash1-/- cells from EHT-related subpopulations in the scRNA-seq dataset. (B) Enrichment of GSEA hallmark pathways in WT (blue) or Sash1-/- (KO; red) cells. (C) Enrichment of genes differentially expressed during EHT progression at specific stages in WT (blue) or Sash1-/- (KO; red) cells. For (B, C) GSEA was performed on significant differentially expressed genes based on adjusted p-value < 0.1. 73 4.3 Discussion Altogether, the data presented in this chapter reveal a defect in EHT progression in the AGM of E10.5 mouse embryos in the absence of Sash1 expression, thus describing a novel contribution of Sash1 in embryogenesis. While the deficiency in IAHC emergence is apparent in Sash1-/- embryos, the lack of phenotypic differences in an endothelial-restricted model of Sash1 deletion hinted at a cell non-autonomous mechanism involving the surrounding microenvironment in which those cells evolve. This was further supported by functional ex vivo assays where Sash1-/- cells appeared to perform just as well as their WT counterpart when cultured under the same conditions. Additionally, pre-HSPC did not express Sash1 based on LacZ staining by flow cytometry furthermore supporting a cell non-autonomous effect. Increasing evidences point towards a contribution of the AGM niche during EHT but much remains to be learned about its effect on HSPC emergence. The ventral polarization of HSPC emergence observed in the dorsal aorta of most species suggests that a gradient of signaling molecules from the microenvironment may regulate this process. While some IAHC are detected on both the ventral and dorsal side of the dorsal aorta in the mouse, the majority of HSPC still emerge from the ventral side [156] and this asymmetry was attributed in part to differences in BMP signaling [156, 157]. Additionally, transplant studies have suggested that the AGM provides cues important for HSPC maturation [156]. Specifically, contribution from at least three compartments of the AGM are required for efficient HSPC generation; both ventral and dorsal section of the dorsal aorta along with the urogenital ridges (UGR) [156]. Using a WT-GFP chimerism co-aggregate system, Souilhol and colleagues showed that the frequency of functional HSPC emerging from E10.5 dorsal aorta is enhanced by the presence of co-culture with the UGR prior to transplantation into irradiated mice [156]. While expression of the key regulator of HSPC SCF (Kitl) was found in the UGR [156], more molecular analyses are required to better understand the role of the UGR in HSPC emergence. Our scRNA-seq identified mesenchymal and epithelial cells from the UGR that express Sash1 and it would be of interest to further investigate the role of Sash1 in that region. In addition to being expressed in a subset of endothelial cells and cells from the UGR, scRNA-seq identified many cell types in the AGM microenvironment in which Sash1 is also expressed. The list included immune cells which have been shown to contribute to EHT regulation [73, 75]. 74 While immune cells could be mediators of an indirect effect, it seems unlikely that Sash1 is acting from within that cell population given that Sash1 excision in immune cells in our conditional knockout model did not recapitulate the pre-HSPC defect. Moreover, Sash1 was also found to be expressed in the mesenchyme and somite. In the zebrafish, Wnt16 expression in the somite has been reported to regulate Notch signaling in promoting HSPC emergence from the aorta [158]. This Wnt-Notch signal transduction was in part mediated by FGF signaling via Fgfr4 [159]. Likewise, studies in the mouse support a role for the subaortic mesenchyme, where high Noggin expression downregulates BMP4 signaling to promote HSPC development [156]. Those studies present evidences of mesenchymal signaling during EHT, thus providing a rationale to further investigate a potential role of Sash1 in these cells. Finally, Sash1 was expressed in cells from the nervous system which has also been linked to EHT progression to some extent. Secretion of catecholamines, downstream of Gata3, from sympathoadrenal cells in the subaortic mesenchyme promotes EHT in the mouse [160] and neural crest cells were shown to interact with HE cells in the zebrafish to promote HSPC specification [78], demonstrating that multiple cell types can be influencing IAHC formation. Interestingly, while disrupting neural crest cells development and migration impacts HSPC emergence in the zebrafish, the signaling between the two cells types was not identified [78]. Altogether, this indicates multiple potential avenues to explore regarding the role of Sash1 in the developing AGM. One approach to identify the source of Sash1 expression relevant in the context of EHT would be using various conditional knockout mouse model to excise its expression in distinct cell populations identified in our scRNA-seq. For instance, our Sash1-flox mice could be crossed with a Sox2-Cre mouse to delete Sash1 in cells from the nervous system, a Pdgfrb-Cre mouse targeting mesenchymal populations, or a Cldn6-Cre mouse to target epithelial cells. Altogether, these three Cre-reporter would cover all non-endothelial/hematopoietic cells present in our scRNA-seq of FDG+ cells. If any of these models recapitulate the Sash1-/- phenotype, which the VEC-Cre failed to do, this would provide clear evidence of the origin of the cell non-autonomous mechanism. Moreover, scRNA-seq could be performed on FDG cells FACS-isolated from Sash1-/- embryos to compare the gene expression profile to Sash1+/- cells which would allow the role of Sash1 to be addressed in a cell-specific manner in the embryo and help identify dysregulated pathways. While not many pathways were identified at a significant p-75 value in hemogenic endothelium or pre-HSPC cells from our dataset, analyzing genes and pathways differentially expressed in cells from the microenvironment might reveal some commonalities to explore in greater details. Our attempt at identifying pathways that are being affected in halted EHT cells yielded very few significant results. While some genes were significantly different, the expression fold change was modest. Perhaps sequencing more cells and/or sequencing at greater depth would help resolving those signals better. Based on the current findings, there could be metabolic differences between WT and Sash1+/- cells that would need to be further explored and validated experimentally. Formation of IAHC originates from the emergence of individual CD41+ cells which first undergo cell division [71]. Subsequently, large IAHC become polyclonal [54, 71] suggesting the eventual recruitment of adjacent cells, but initial proliferation seems to drive EHT [71, 155]. Giving that GSEA revealed enrichment of cell cycle pathways in HE/pro-HSPC from Sash1-/- embryos, whether these cells are blocked or delayed during cell cycle progression is an area that deserves further investigation. It is also possible that the majority of the cells sequenced were simply not very different from their WT counterpart at each individual step. Indeed, some IAHC are formed and some functional HSPC emerge from this process even in Sash1-/- embryos. If, for instance, the defect resides in cell proliferation rather than differentiation, the fewer cells present in the dorsal aorta might still exhibit a similar gene profile. The clustering algorithms used also means that cells in the dataset will be “moved” to aggregate with other cells that have similar gene expression profile. Hence, the difference between all cells labeled as one cluster in silico would be expected to be minimal. Again, sequencing more cells at a greater depth could possibly help identifying more subclusters induced by subtle differences, especially in that seemingly mixed population of HE and pro-HSPC. Nevertheless, the data presented a preferential accumulation of “HE/pro-HSPC” at the expense of more differentiated cells along the EHT trajectory in Sash1-deficient embryos. This finding corroborates the other observations presented in this chapter, such as the potential reduction in IAHC size, the accumulation of c-Kit+ cells embedded in the vascular wall, and higher levels of Runx1 and Gfi1 expression in CD41-depleted endothelial cells in contrast to lower expression in 76 the bulk AGM due to fewer IAHC overall. Together with similar frequencies of functional HE cells at E9.5 and lower numbers of phenotypic pre-HSPC at E10.5, all the evidences point to a hindrance in EHT progression at a post HE commitment stage in the absence of Sash1 signaling from a cell non-autonomous source. Figure 4.13 Model for Sash1 role in EHT progression Loss of Sash1 does not affect early specification of the HE but impairs emergence of large IAHC, an effect induced by Sash1 expression in cells from the microenvironment 77 CHAPTER 5: Identification of Meis1 as an early driver of hemogenic endothelium commitment 5.1 Introduction Chapter 4 explored the EHT from multiple angles, including cells that compose the microenvironment niche. At the origin of EHT is a cell decision that arises in early endothelial cells about an arterial or hemogenic fate. But as briefly alluded to in Chapter 4.2.2, defining and prospectively isolating endothelial cells with hemogenic potential at an early stage is still challenging. Although some groups have used transgenic reporter mice such as Runx1+23GFP or Ly6a-GFP to identify cells co-expressing endothelial and hematopoietic genes [56, 161], there is no ideal single marker to enrich for those cells [162] and whether this identifies the earliest cells primed towards hematopoietic lineage is debatable. Prospectively isolating these cells has proven difficult given that they represent a small fraction of the endothelial population, are transient in nature, and are defined based on their future nature [162]. Therefore, what specifies an early endothelial-like cells towards an hemogenic potential or fate is still not clearly understood and better characterization of the early transcriptional changes driving hemogenic endothelium transcriptional specification is still required. Single-cell technologies have enabled us to dissect heterogeneous populations and study rare cells to new extents. This is particularly useful in a context such as EHT where a relatively small number of heterogenous IAHC cells, composed of HE cells, pro-HSPC, pre-HSPC, committed progenitors, and other mature blood cells such as macrophages, can be detected in a single embryo around E10.5 [58, 73]. Therefore, the scRNA-seq experimental data on CD31+ cells from WT AGM, outlined in Chapter 4.2.4, constitute a great resource to investigate the normal progression of EHT. In addition to the transcriptomic data, Cellular Indexing of Transcriptomes and Epitopes by sequencing (CITE-seq) was performed on 3 of the 4 WT samples, providing phenotypic data for 4,292 cells after QC filtering. Therefore, we took advantage of this dataset to investigate transcriptional differences between the bulk endothelial cell population and the earliest population transitioning towards the hematopoietic lineage in our dataset. We identified Meis1 as a candidate regulator of early hematopoietic specification and showed that loss of its function impacts early EHT stages. 78 5.2 Results 5.2.1 Characterization of CITE-seq cell populations identifies Meis1 as differentially expressed in pre-hemogenic endothelial cells Given that the transcriptome data used for these analyses represent a subset of the samples analyzed in the previous chapter (Fig. 4.10), the same populations emerged upon clustering (Fig. 5.1A). In order to identify markers from our dataset, the accurate labeling of the different cell populations represented a key first step for further analyses. For this purpose, in addition to the transcriptome data, cells were stained with specific antibodies conjugated to an oligo-tag prior to sequencing which allowed for surface profiling of epitopes. To better define the population of cells undergoing EHT, antibodies traditionally used in the literature were selected for CITE-seq. CD117 (or c-Kit) marks all IAHC cells in the embryo at E10.5 [32, 58]. CD41 is an early marker of hematopoietic commitment that is expressed early on pro-HSPC while CD43 is upregulated later as the cells transition towards an HSPC phenotype on the way to upregulating the pan-hematopoietic marker CD45 [46, 50, 51]. CD31+CD117+CD41+CD43+CD45+ cells can be further enriched for pre-HSPC with long-term potential by expression of CD27 [52]. For the purpose of this project, we were interested in identifying early genes that differentiate HE cells from their non-hemogenic vascular endothelial counterparts and are associated with EHT progression. Using antibody derived-tags (ADT), we were able to determine that the cells previously labeled as “pre-HE” (Fig. 5.1A) did not express any of the surface markers that were targeted in this experiment (Fig. 5.1B-D). In this regard, these pre-HE cells are essentially undistinguishable from the remaining endothelial cells, by traditional techniques such as flow cytometry, despite showing clear differences at the transcriptional level. In comparison, most cells labeled as “HE/pro-HSPC” already showed expression of CD117 and CD41, suggesting that this cluster is enriched in pro-HSPC. Meanwhile, our “pre-HSPC” population upregulates CD43, CD45, and could be divided into CD27+/- cells. Altogether, the ADT information supports the labeling of the different clusters according to previous knowledge (Fig. 5.1B-D). Pseudotime analysis to order the cells into a differentiation trajectory also showed a transition from “pre-HE” to “HE/pro-HSPC”, then “pre-HSPC” and branching at the two progenitor clusters with lineage biases (Fig. 5.1E). Altogether, this gave us confidence to use this dataset to understand normal progression of EHT. 79 Figure 5.1 CITE-seq analysis of CD31+ cells using EHT-related surface markers (A) UMAP clustering of FACS-isolated CD31+ cells based on single-cell RNA expression (B-C) Ridgeplot and (D) scatterplot showing expression of ADTs on cells within each cluster (E) Pseudotime analysis using monocle3 showing a progression from pre-HE cells towards lineage-biased progenitors 80 To explore what initially primes the cell towards a hemogenic fate, differential gene expression analysis was performed using the Seurat platform to identify genes that separate “pre-HE” cells from the endothelial cluster in our CITE-seq dataset. Compared to endothelial cells, 518 genes were upregulated and 508 genes were downregulated in “pre-HE” (adjusted p-value <0.1). The signature of upregulated genes was enriched for several pathways previously reported to affect EHT, such as TGF-β, NF-κB, and Wnt signaling (Fig. 5.2A). Of the 518 upregulated genes, 52 were transcription factors (TFs) (Fig. 5.2B). TFs bind DNA to modulate expression of other genes, thus they can act as upstream regulators and play a significant role in cell fate during development. Amongst the most significantly upregulated TFs, four groups emerged including genes related to the Notch pathway, TGF-beta pathway, and Gata family members, all of which have been implicated in EHT [163]. In addition, Meis1 was also significantly enriched in the “pre-HE” subset, along with its homolog Meis2 and its binding partners Pbx1 and Hoxa10 (Fig. 5.2B, C). Together, they represent 4 of the top 23 TFs (Fig 5.2B). Meis1 is well-known for its role in leukemogenesis and regulation of adult bone marrow HSPC [103, 164]. Moreover, it is an essential developmental gene. Constitutive deletion of Meis1 leads to embryonic lethality around E11.5-14.5 with defects in fetal liver hematopoiesis [106, 107]. Previously reported expression of Meis1 during EHT was mostly associated with an upregulation during maturation of pre-HSPCs in the mouse [58, 154, 161, 165]. Given its role in normal maintenance of adult HSPC, it is not surprising that it would also be required in embryonic HSPC. However, in a recent report Meis1 was suggested to act early during in vitro differentiation of human pluripotent stem cells [166]. Therefore, whether Meis1 drives specification of HE or is required only at a later stage is still unclear. Several chromatin immunoprecipitation (ChIP)-seq datasets are publicly available for Meis1 binding sites in the genome to identify target genes, including an experiment performed in a hematopoietic progenitor cell line (HPC7) [167]. Therefore, to further explore the relevance of investigating the role of Meis1 in pre-HE cells, we overlapped this dataset with the list of genes identified as upregulated in the Meis1-expressing “pre-HE” population from our CITE-seq dataset. This analysis revealed that 41.5% of “pre-HE” upregulated genes (215/518) contain a potential Meis1 binding site (Fig. 5.3A). Similarly, 29 out of the 52 (over 55%) TFs identified earlier have a Meis1 binding site (Fig. 5.3A). That list includes TFs that were previously reported 81 to be important in regulating hematopoiesis such as Gata2, Bcl11a, Hmga2, Mecom, and Lyl1 [69, 154, 167]. Gene ontology analysis showed an association of both the 215 genes and 29 TFs with biological processes relating to the development of the hematopoietic system (Fig. 5.3A). On the other hand, the remaining 303 genes upregulated in HE, that were not candidate targets of Meis1 based on the ChIP-seq dataset, were associated with GO terms relating to angiogenesis and blood vessel development. Genes that are upregulated in pre-hemogenic compared to vascular endothelium, and are also Meis1 targets are involved in hematopoiesis, suggesting that Meis1 might prime endothelial cells for hemogenic commitment by being an upstream regulator of hematopoietic genes expression. The “pre-HE” population identified in this dataset is an intermediate between endothelial cells and typically defined HE cells in that it lacked Runx1 and Gfi1 expression but expressed other markers shown to enrich for hemogenic cells (Fig. 5.2C). However, these populations are transient, and genes that truly confer hemogenic potential to early endothelial-like cells must be able to trigger or facilitate additional transcriptional changes leading to the next step in the progression. To evaluate whether Meis1 could fit that definition, we also tested for genes that are upregulated in “HE/pro-HSPC” compared to “pre-HE”. 497 genes were identified with 26.3% of them (131/497) containing a potential Meis1 binding site (Fig. 5.3B). Those 131 genes enriched for terms associated with hematopoiesis in gene ontology (GO) term analysis and include genes known to be associated with EHT progression such as the TFs Runx1, Gfi1, Kit, and Myb while the remaining 366 genes showed association with ubiquitous cellular processes such as transcription and translation (Fig. 5.3B). Therefore, it appears that Meis1 can regulate gene expression in pre-HE cells and facilitate a transcriptional shift associated with HE identity. Although the findings based on the ChIP-seq dataset are correlative and Meis1 might not be actively driving expression of every one of those genes in this specific context or at the same time, this identified Meis1 as a potentially significant upstream regulator of HE specification and early progression of EHT. 82 Figure 5.2 Transcriptional differences between pre-HE and the vascular endothelial subset (A) Selected enrichment plots from GSEA performed on DEGs between EC and pre-HE. (B) List of TFs enriched in pre-HE vs EC (adjusted p-value < 0.1 based on Wilcoxon rank sum test), arrows point to Meis genes and binding partners (C) Expression of HE-associated markers, Meis1-related TFs, and EHT regulators across all subpopulation in the CITE-seq dataset. 83 Figure 5.3 Meis1 targets hematopoietic-related genes in the pre-HE and HE/pro-HSPC (A-B) Overlapping of genes enriched in (A) pre-HE or (B) HE/pro-HSPC with potential Meis1 targets based on published ChIP-seq dataset along with GO term analysis for biological processes enriched in Meis1-bound genes vs unbound 84 5.2.2 Meis1 expression enriches for the hemogenic subset of endothelial cells in vivo The previous data was generated from in silico analysis of subsets of cells isolated from E10.5 AGM. To investigate Meis1 expression pattern in vivo, a green fluorescent protein (GFP)-reporter mouse model was used [111]. This mouse model, generated and generously shared by Dr. Keith Humphries, features insertion of the GFP sequence after the ATG start site of the Meis1 gene followed by a P2A cleavage site and the normal Meis1 sequence [111]. Therefore, GFP is co-transcribed with Meis1, but expressed as two different proteins upon translation which maintains the WT functions of Meis1 protein. This allowed for efficient tracking of Meis1 expression during embryonic development. GFP-expressing embryos can be imaged under fluorescent microscopy and showed high expression in specific regions such as the AGM and the eye (Fig. 5.4A), consistent with previous reports [107]. Fluorescent imaging was also performed on cryopreserved and sectioned E10.5 embryos to look more closely at the dorsal aorta. GFP was highly detected in all IAHC (Fig. 5.4B), consistent with Meis1 expression during EHT. Furthermore, GFP could also be detected in the endothelium around the IAHC sites but was mainly absent in endothelial cells in regions devoid of IAHC, where Meis1 expression was rather enriched in the subaortic mesenchyme (Fig. 5.4B). Similarly, flow cytometry analysis on dissected E10.5 AGM showed high and ubiquitous expression of GFP in pro-/pre-HSPC populations (marked by CD31+CD41lowCD45+/-) while two populations of endothelial cells (EC: CD31+CD41-CD45-) were observed; Meis1-GFP+ and Meis1-GFP- (Fig. 5.4C-E). Therefore, immunostaining and flow cytometry analyses validated the expression pattern observed in the CITE-seq dataset. Interestingly, the proportion of EC that expressed Meis1-GFP decreased over development time going from E9.5 to E11.5 (Fig. 5.4F, G) with E9.5 associated with early specification of HE and E11.5 marking the decline of EHT potential in the AGM [43, 56]. Remarkably, although the proportion decreased from E9.5 to E10.5, the absolute number of cells within the Meis1-GFP+ EC compartment did not change significantly based on flow cytometry quantification between those two time points (Fig. 5.4H). However, the fluorescence intensity of GFP was increased at E10.5 compared to E9.5 (Fig 5.4E). This shows that cells choose to express Meis1 as early as E9.5, consistent with early an specification of the hemogenic endothelium, and that, while their number does not change significantly by E10.5, they might progress towards a more “Meis1 committed” state associated 85 with a perhaps stronger Meis1-driven transcriptional program over that developmental window. This increase in GFP intensity was not observed in phenotypic pro-/pre-HSPC which appeared to required ubiquitous Meis1 expression (Fig. 5.4D, E). Based on our previous analysis of the CITE-seq data and these observations by flow cytometry, we hypothesized that only the Meis1-GFP+ endothelium has hemogenic potential and that Meis1 might be important in early specification of the HE. To further test that hypothesis, EC from E10.5 AGM were FACS-isolated based on GFP expression. First, expression of selected EHT-related genes was assessed by ddPCR for sensitive absolute quantification. Of the genes selected, expression of Runx1, Gfi1, and Spi1 was detected exclusively in Meis1-expressing EC, with Gpr56 also being significantly enriched in that population as compared to FACS-isolated GFP-negative EC (Fig. 5.5A). While Gata2 is also required for EHT [68, 69] and enriched in pre-HE (Fig. 5.2B-C), its expression was not restricted to Meis1-GFP+ EC (Fig. 5.5A). This is consistent with a slightly more widespread expression pattern observed at the transcriptomic level in the endothelial cluster of the CITE-seq data (Fig. 5.2C). Some low levels of Gata2 may have additional function in vascular cells [168, 169]. In contrast, expression of the endothelial marker Pecam1 (CD31) appeared to be slightly downregulated at the transcriptional level in Meis1-GFP+ cells compared to the remaining endothelium (Fig. 5.5A). Although CD31 is expressed on the surface of pre-HSPC and emerging HSPC, this transcriptional downregulation together with increase expression of hematopoietic genes indicates a shift of Meis1-GFP+ EC towards a hematopoietic fate at the expense of an endothelial trajectory. To confirm that only Meis1-GFP+ EC can undergo EHT, a functional differentiation assay was performed ex vivo. Cells were sorted in different wells of a 4-well chamber slide pre-plated with OP9 cells and allowed to differentiate over 5 days in the presence of cytokines (as described in Chapter 4.2.2). For this assay, four populations were isolated; Meis1-GFP+ EC, Meis1-GFP- EC, CD31+CD41+CD45- cells, and CD45+ cells. The last two populations were used as controls to confirm that this culture system can A) promote differentiation/ maturation of CD31+CD41+CD45- cells into CD45+ cells and B) maintain CD45+ hematopoietic cells in culture. Regardless of the initial number of cells plated, only Meis1-GFP+ cells were able to form small colonies of round hematopoietic cells and the acquisition of the surface marker CD45 was validated by antibody staining and fluorescent imaging (Fig. 5.5B). This experiment clearly 86 demonstrates that only Meis1-expressing cells are able to undergo EHT. In this system, GFP- cells could still upregulate Meis1 expression. However, GFP- cells were unable to differentiate in response to cytokines in the co-culture with OP9 cells, suggesting that Meis1 is required early to prime EC towards a hematopoietic fate. 87 Figure 5.4 Meis1 expression in the dorsal aorta of E10.5 embryos (A) Fluorescence imaging of a Meis1-GFP embryo at E10.5. (B) Immunostaining of IAHC and the dorsal aorta, showing CD31 (red) and GFP (green) expression. Ao: aorta, V: ventral. (C-H) Flow cytometry analysis of GFP distribution in cells from Meis1-GFP AGM. (C) Representative flow plot at E10.5. (D) Fraction of GFP+ cells in gated populations (EC: CD31+CD41-CD45-, pro-HSPC: CD31+CD41lowCD45-, pre-HSPC: CD31+CD41lowCD45+, HC: CD31-CD45+) and (E) associated mean fluorescence intensity (MFI). (F-H) Quantification of GFP+ cells in the EC compartment at various time points during development. (p-value: ** < 0.01, *** < 0.001, ns = non-significant; based on Student t-test) 88 Figure 5.5 Meis1 expression enriches for the hemogenic endothelium in vivo (A) Expression of selected EHT-related genes in Meis1-GFP+ and Meis1-GFP- EC quantified by ddPCR (n = 5). (p-value: * < 0.05, ** < 0.01, **** < 0.0001, ns = non-significant; based on Student t-test) (B) Ex vivo differentiation assay on OP9 to assess hemogenic potential. Representative brightfield images of hematopoietic colonies formed and confirmation of CD45 expression by immunostaining shown below the schematic. The table on the right shows the number of positive wells at the end point for the various initial cell inputs tested. 89 5.2.3 Endothelial-deletion of Meis1 impairs EHT initiation We identified a potential marker and candidate regulator of early stages of EHT in Meis1. However, whether its expression is a requirement for EHT initiation had not been explored yet. Given the multiple defects associated with constitutive homozygous deletion of the Meis1 gene in mice [106, 107], we opted to use a Cre-mediated conditional knockout (cKO) mouse model to excise Meis1 specifically in endothelial cells. VEC-Cre males were mated with females carrying Meis1-floxed alleles (Meis1fl/fl, floxed exon 8) twice to generate litter with a mix of WT (Meis1fl/fl or Meis1fl/+, hereafter referred to as Mf/f), heterozygous (Meis1fl/+;VEC-cre (Mf/+;Cre)), and cKO (Meis1fl/fl;VEC-cre (Mf/f;Cre)) embryos. Since only Meis1-expressing cells were able to differentiate into hematopoietic progeny in the OP9 co-culture system (Fig. 5.5B), the requirement for Meis1 in HE generation was further assessed functionally using this assay. E9.5 EC were plated at the limiting dilution of 100 cells per well in a 96-well plate to quantify the frequency of functional HE cells at an early stage (as performed previously in Fig. 4.3). Interestingly, the frequency of HE in E9.5 AGM significantly declined in Mf/f;Cre embryos compared to WT littermates (Fig. 5.6A) supporting the hypothesis that Meis1 regulates hemogenic specification of the endothelium. Further reinforcing this point, expression of Runx1, Gfi1, and Spi1 was lower in FACS-isolated AGM endothelium from E10.5 of Mf/f;Cre embryos relatively to WT littermates (Fig. 5.6B). All of these genes are associated with EHT progression and their expression was restricted to Meis1-GFP+ EC (Fig. 5.5A). Thus, this finding is consistent with a reduction in the proportion of hemogenic cells amongst the bulk endothelial population as evidenced by the ex vivo assay (Fig. 5.6A). This could be caused by a lower level of transcriptional activation of those specific genes in the absence of Meis1 in individual HE cells, which would also be supported by the ChIP-seq data. To strengthen the observation of an early defect, flow cytometry analysis was performed on dissected AGMs harvested at E9.5, before the emergence of visible large IAHC. The surface marker CD44 was included in the panel to enrich for pre-HE/HE cells prior to the acquisition of pro-/pre-HSPC markers [153]. Fewer CD44+ cells were detected per AGM in Mf/f;Cre embryos as a proportion of CD31+CD45- cells (Fig. 5.6C). Altogether, these findings demonstrated that functional Meis1 contributes to hemogenic potential within the AGM and that Meis1 promotes the specification of early pre-HE to Runx1/Gfi1 expressing HE. 90 Consequently, the reduced hemogenic potential of Mf/f;Cre dorsal aorta impaired the emergence of pre-HSPC. Flow cytometry analysis showed a lower number of phenotypic CD31+CD41lowCD45- pre-HSPC in the AGM of Mf/f;Cre embryos both as a proportion of endothelial cells and absolute cell count at E9.5 (Fig. 5.7A). While the number of WT pre-HSPC strongly increased by E10.5, the number of phenotypic pre-HSPC lagged behind in Mf/f;Cre embryos, maintaining a roughly 2-fold reduction compared to WT littermates (Fig. 5.7B, C). This suggests that the defect originates early and is propagated over the developmental process. Finally, we crossed the Runx1+23GFP reporter mouse that was previously characterized as labeling HE and IAHC cells in the dorsal aorta [56, 64] with our Meis1-cKO mice, generating transgenic embryos that expressed the +23GFP reporter and lacked endothelial expression of Meis1 (Meis1fl/fl;VEC-cre;+23GFP, abbreviated Mf/f;Cre;+23GFP). Based on Mendelian genetic and the restriction around viably breeding these mice, we could only generate Mf/f;Cre;+23GFP embryos at a ratio of 1 in 8 therefore not every litter produced these triple-transgenic embryos. Nevertheless, we were able to obtain some E10.5 embryos (n = 3 Mf/f;Cre;+23GFP) to look at IAHC by immunostaining cross-sections of the dorsal aorta using the +23GFP reporter as a marker. While +23GFP cells were detected in both WT (Mf/f;+23GFP or Mf/+;+23GFP) and Mf/f;Cre;+23GFP dorsal aorta, the number of large IAHC (>4 cells) was significantly reduced in Mf/f;Cre;+23GFP dorsal aortas (Fig. 5.7D, E). Visually, most +23GFP cells appeared to be still flat and embedded in the vascular wall of the dorsal aorta as opposed to bulging out into clusters as observed in WT counterparts (Fig. 5.7D). Therefore, this experiment also demonstrated that the early defects induced by Meis1-deletion prevented EHT progression. 91 Figure 5.6 Endothelial-deletion of Meis1 reduces hemogenic potential of the dorsal aorta endothelium (A) Frequency of hemogenic endothelial cells in E9.5 embryos quantified by ex vivo differentiation assay on OP9 (n = 8 Mf/f, 3 Mf/+;Cre, 4 Mf/f;Cre). (B) Expression of EHT-specific genes in FACS-isolated endothelial cells quantified by ddPCR (n = 3). (C) Quantification of CD44+ cells in E9.5 embryos along with representative flow cytometry plots (n = 11 Mf/f, 4 Mf/+;Cre, 7 Mf/f;Cre). (p-value: * < 0.05, ** < 0.01, *** < 0.001, ns = non-significant; based on Student t-test) 92 Figure 5.7 Loss of Meis1 impairs pre-HSPC emergence (A) Quantification of phenotypic pre-HSPC in (A) E9.5 (n = 19 Mf/f, 10 Mf/+;Cre, and 11 Mf/f;Cre) and (B) E10.5 embryos (n = 5 Mf/f and 6 Mf/f;Cre) along with (C) representative flow cytometry plots of CD41 expressing cells at E10.5 (D) Immunostaining of E10.5 IAHC in +23GFP embryos with WT or endothelial-deletion of Meis1 (Mf/f;Cre), showing GFP (green) and CD31 (purple) expression, and (E) quantification of large IAHC per dorsal aorta. (n = 4 Mf/f;+23GFP and 3 Mf/f;Cre;+23GFP). (p-value: * < 0.05, ** < 0.01, *** < 0.001; based on Student t-test) 93 5.3 Discussion In this chapter, we identified the transcription factor Meis1 as a potential early regulator of HE specification to initiate EHT. While Meis1 has been shown to regulate hematopoiesis [103, 105], its role in EHT is still ambiguous. Our data suggests that it is expressed in an earlier precursor cell (pre-HE) compared to other known hematopoietic factors that promotes the transition. One caveat when analyzing this type of dataset is that all the cells sequenced were taken from a “snap-shot” in time at E10.5 thus we must attempt to based our observations on a “pseudotime” progression of EHT rather than real time. Nevertheless, experiments performed with Meis1-GFP mice showed that only Meis1-expressing cells express EHT-related genes and have the potential to undergo EHT, supporting the idea that a Meis1-expressing precursor exists in vivo. While Meis1 expression keeps increasing from pre-HE to pre-HSPC, this places Meis1 closer to the initial hemogenic specification event than what was previously reported in some mouse studies [58, 154, 161]. Some evidence coming from hPSC differentiation studies argued that Meis1 affects commitment of mesoderm progenitors into functional HE progenitors in vitro [166]. While the author defined HE progenitors as CD31+CD34+ cells, which represents a heterogeneous population of endothelial cells [170], our in vivo study corroborates the idea that Meis1 controls specification of an early precursor cell during EHT. Indeed, the Meis1-expressing pre-HE cells that we identified in this dataset are devoid of Runx1 expression but expressed other markers reported to enrich for cells with hemogenic potential such as CD44 and Gata2. While our CITE-seq data showed that Meis1 is enriched in CD44+ cells, endothelial-deletion of Meis1 itself reduced the proportion of CD44+ endothelial cells in the embryo. Interestingly, Cd44 is a potential Meis1 target based on the ChIP-sequencing data [167] used in this study. This indicates that CD44 surface expression might be directly, although not necessarily uniquely, controlled by Meis1 during hemogenic commitment. As a transcription factor, Meis1 has the potential to regulate transcription of many other genes. The correlation observed between genomic loci bound by Meis1 and genes upregulated during EHT is consistent with a role in early activation of the hemogenic program in endothelial-like cells, however further experimental validation should be gathered using our conditional-knockout model. For instance, performing scRNA-seq experiments on WT and Meis1-cKO embryos could reveal on a large scale whether the expression of the 215 potential target genes 94 identified is altered. Based on our sequencing studies outlined in chapter 4, this would require a larger number of cells to be sequenced at a potentially greater depth than what has been done so far. Alternatively, a short-listed selection of genes could be validated by ddPCR, as in Fig. 5.6B, which would likely require a similar number of embryos. Given that Meis1 deletion impacts the emergence of IAHC and phenotypic pre-HSPC as shown in this chapter, further investigating the transcriptional network regulated by Meis1 in cells at each step of this transition could uncover critical mechanism driving hemogenic specification. While our focus is on the early specification event of the hemogenic endothelium, ubiquitous and elevated Meis1 expression in pre-HSPC also suggests that it might be indispensable at various steps of the transition. Along these lines, experimentally delineating the different functions of Meis1 at specific steps has proven challenging. This is due, in part, to the fact each of these subpopulations of HE cells and pre-HSPC are defined based on their future functional outcome that needs to be assayed. For instance, the ex vivo differentiation assay on OP9 revealed a decrease in HE frequency. However, because Meis1 is believed to be required intrinsically within the cells that are undergoing EHT, a caveat of interpreting these findings is that by itself, the assay does not distinguish between an inherent decrease in the number/frequency of HE plated at the start of the assay or an inability of those cells to progress through EHT to generate detectable colonies. To assess a potential functional role of Meis1 in pre-HSPC during the transition, CD31+CD41lowCD45- cells could be plated at the start of the assay instead. Therefore, unless each subpopulation during EHT is finely isolated and assayed individually, we must rely on other characteristics to infer the origin of the differences observed in those functional assays. Taken together with the changes in phenotypic populations observed by flow cytometry and changes in gene expression profile determined by ddPCR, the data presented in this chapter demonstrates the importance of Meis1 in HE specification. Altogether, it shows that Meis1 plays a significant role in hematopoietic stem cell emergence well before the first HSPC is actually formed and that the Meis1 GFP-reporter mouse model can be used to prospectively isolate endothelial cells with hemogenic potential from the AGM. 95 Figure 5.8 Meis1 drives the endothelium towards a hematopoietic fate In WT embryos, Meis1-expressing endothelium is primed to upregulate Runx1 and the EHT program giving rise to pre-HSPC. When Meis1 is deleted from the endothelium, the hemogenic potential is deeply affected resulting in overall fewer pre-HSPC. 96 CHAPTER 6: SUMMARY, PERSPECTIVES, AND SIGNIFICANCE 6.1 Endothelial Sash1 in the developing lung Chapters 3 and 4 established that Sash1 is an important developmental gene; essential in the context of lung development but also playing a significant role in the establishment of the hematopoietic system. The work presented in these chapters highlights previously unrecognized developmental roles for Sash1, a gene that has mainly been studied in the context of cancer. In this study, we first sought to explain the phenotypic defect observed in Sash1-deficient mice. By investigating the neonatal lethality resulting from Sash1 deletion, our data highlights the importance of endothelial signaling during late-stage lung maturation just prior to birth as shown in Chapter 3. The lung architecture is a highly organized network within which the vascular endothelium needs to grow in conjunction with the alveoli to form a functional unit [122, 124]. Importantly, no structural defect in the lung vascular architecture or in the juxtaposition of endothelial cells next to alveolar epithelial cells was observed in Sash1-/- embryos supporting the notion of cell-cell communication rather than an organizational issue with the overall 3D structure In the investigated model, endothelial-deletion of Sash1 leads to lower levels of surfactant being produced in E18.5 lungs as a result of immature alveolar epithelial cells. Morphologically, alveolar epithelial type 2 cells (AE2C) show features of incomplete differentiation in Sash1-deficient lungs, including the lack of lamellar bodies and a correlated increase in glycogen accumulation, despite not expressing Sash1 themselves. Similarly, expression of AE1C-specific markers was also reduced, reflecting a broader differentiation delay as opposed to a specific regulation of surfactant levels by Sash1. These findings indicate that paracrine signaling from the endothelium regulates terminal differentiation of alveolar epithelial cells. AE1C and AE2C start differentiating from a common bipotent precursor during the saccular stage of lung development, and their maturation process extends postnatally until the lung is fully formed [133]. This process is critical for respiration as it prepares the terminal saccules to receive air. Surfactant augments pulmonary compliance and thinning of the septa is crucial for gas-exchange. In humans, the saccular stage occurs at week 24-38 of gestation, meaning that the lungs of preterm babies (before 37 weeks) are still within that phase at the time of birth [124, 97 171]. As a consequence, incomplete lung maturation remains a major concern associated with premature birth and understanding how the alveoli fully develop is of clinical importance. Respiratory distress syndrome (RDS) at birth can progress to bronchopulmonary dysplasia (BPD), a chronic lung disease induced by lung injury. Mechanical ventilation is a factor that can contribute to the onset of BPD and up to 40% of premature babies develop BPD [125]. Antenatal corticosteroid and surfactant therapies have improved survival of preterm newborns but the incidence of BPD remains mostly unchanged [125, 171]. This reflects the clinical need for more effective and less invasive treatment strategies for RDS. The interdependency between pulmonary vascular and alveolar development has been recognized but identifying the various cell-cell interactions and mechanisms driving AE2C to maturity might help tailor therapies to improve outcome. In this study, we identified a pathway where the interaction of Sash1 with β-Arrestin1 induces NO secretion downstream of Akt activation. By modulating NO level using SNAP, surfactant levels were restored in prenatal lungs of Sash1-/- mouse embryos. Inhaled NO is already used to treat pulmonary hypertension however it is highly controversial as a treatment in premature infants. While some patients might have benefited from it under specific conditions in conjunction with antenatal corticosteroid [147], the lack of beneficial evidence in large-scale studies originally pushed the NIH to issue a statement against the use of inhaled NO in preterm newborns [172]. Our own data suggests a mixed effect of NO based on the state of the lung and/or overall NO levels. Normal lungs treated with a NO-donor at late embryonic stages showed a reduction in surfactant levels, in contrast to the rescue observed in NO-deficient Sash1-/- lungs but other potential systemic side effects of NO were not examined in this study. While NO-donors improve surfactant production by AE2C ex vivo, NO is a highly reactive compound in biological systems and both its concentration and localization may affect its effect in vivo. Better understanding of the signaling cascade involving AE2C maturation during the late saccular stage might unveil new components of the pathway that could be modulated to circumvent side effects. In the context of this study, Sash1 itself may not represent an actionable target, which is why we need to have a comprehensive picture of the developmental process. β-98 Arrestin mutants have embryonic lung defects similar to Sash1-/- embryos but a mechanism was not identified in that study [145]. Therefore, establishing connections between different models will contribute to drawing a clearer portrait of developmental processes involved in lung maturation. There are significant differences in the timing of alveolar maturation surrounding birth between mouse and human. The saccular stage is almost entirely prenatal in human whereas it extends up to 5 days after birth in the mouse [122]. The emergence of in vitro models of lung maturation using human cells should therefore be beneficial to the field. While promising, some of the recently developed in vitro models also have limitations and terminal differentiation is not always achieved [173, 174]. Most attempts at developing ex vivo organs focused on epithelial cells but proper vascularization is often limited or not achieved [4]. Does the endothelium hold the key? 6.2 Regulation of the endothelial-to-hematopoietic transition During embryogenesis of Sash1-/- mice, we also identified defects in hematopoiesis which prompted the investigation of a potential role of Sash1 in EHT, given its known expression in the endothelium. While the deficiency in hematopoiesis in the Sash1-null mouse model does not trigger gross abnormalities, careful investigation of the phenotype revealed interesting findings. Chapters 4 and 5 explored the regulation of EHT in the dorsal aorta of the mouse embryo as it leads to HSPC emergence during embryogenesis. Given the ability of HSPC to self-renew and sustain life-long hematopoiesis in the organism, the study of HSPC formation has been an area of great interest for many years. Nevertheless, the difficulty to generate HPSC de novo indicates the importance of further understanding the molecular requirement for EHT in vivo [175]. In Chapter 4 and 5, we used different mouse models to investigate the role of two genes in this process; Sash1 and Meis1. While the most robust assay to quantify HSPC remains multilineage engraftment into a recipient upon transplantation which was not performed in this study, the deficiency in functional HSPC has already been shown in transplant of liver cells from both the Sash1-/- (Paliouras G, unpublished) and Meis1-null [107] mouse models. Therefore, this study focused on the regulation of EHT and the early steps driving the process giving birth to HSPC. 99 At first glance, both Sash1 and Meis1 deficient embryos can appear to trigger a similar defect given that in both cases we observed a roughly 50% reduction in the number of IAHC present in the dorsal aorta in vivo. However, detailed investigation of the phenotypes revealed marked differences between the two models. On one hand, Meis1 is acting via a cell intrinsic transcriptional regulation to drive hemogenic endothelium specification at early stages of EHT. Indeed, ex vivo differentiation assay to quantify functional HE cells in the embryo showed strikingly lower HE frequency upon endothelial-deletion of Meis1. On the other hand, the frequency of HE was unaltered in Sash1-/- AGM and pre-HSPC did not require Sash1 expression to produce hematopoietic colonies ex vivo. Together with the lack of pre-HSPC defect in VEC-Cre driven excision of Sash1, all evidences corroborate a cell non-autonomous influence of Sash1 on EHT in sharp contrast with Meis1. Hence, this reflects different regulatory mechanisms affecting HSPC formation, with Meis1 expression being required in cells initiating EHT while Sash1 is dispensable in those cells but rather acts via the microenvironment. Those findings highlight the importance of taking into consideration both the cell intrinsic and extrinsic factors given that they can ultimately yield a similar phenotype when it comes to a reduction in HSPC outcome during EHT. A more challenging aspect to dissect experimentally is the step at which each factor is playing a critical role during the transition. Here, we showed that Meis1 and Sash1 are affecting different stages of EHT. While a similar defect in pre-HSPC number was observed at E10.5, Meis1-knockout embryos already showed phenotypic differences at E9.5. In fact, the reduction in IAHC in Meis1-deficient embryos appears to be a consequence of fewer HE cells illustrated by a reduction in the percentage of CD44+ endothelial cells and loss of Runx1 expression in the endothelium as early as E9.5. In contrast, Sash1 does not appear to influence HE specification. If anything, there seemed to be a slight increase in HE cells at the expense of IAHC, caused by a blockade in cluster formation. In an attempt to further explore the impact of Sash1 on different subset of cells during EHT progression, we performed single-cell RNA-sequencing (scRNA-seq). The first scRNA-seq study of EHT, published a couple years prior, analyzed very few cells and was performed at E11 therefore focusing on more committed pre-HSPC populations [154]. Since then, a few more single-cell studies have been published on the topic in both zebrafish [176] and mouse embryos 100 [58, 60, 153] , each focusing on slightly different aspects. By comparing cells from WT and Sash1-deficient embryos, our dataset presented in Chapter 4 was aimed at very specific questions that could not be addressed using data from published sources. Meanwhile, our CITE-seq approach for the WT samples complements other studies in the literature by including all cells of interest in the dorsal aorta. Rather than isolating a small subset of cells and potentially losing some precious cells, we opted for a more comprehensive approach using CITE-seq technology which allowed us to sequence an unbiased bulk population and to annotate phenotypic surface marker information in silico retrospectively. Given the increasing amount of data generated collectively by multiple sequencing studies, it becomes important to cross-validate new findings and critical information derived from a particular dataset. Ideally the wealth of new information generated by single-cell experiments should be integrated in a way that will provide a more accurate and complete picture of the underlying biological phenomenon. Interestingly, a new report [60] was published after we conducted the Meis1 study in which their scRNA-seq data seem to corroborate our CITE-seq results based on the interactive data explorer tool released online for that data, which allowed for quick comparison of the two datasets. For instance, the markers that we used to label our “pre-HE” population (ie: Cd44, Procr, Cd47, Gata2) also seem enriched in the “pre-HE” cells identified in their study making our two populations analogous in many ways and supporting the choice of terminology used in this dissertation. In addition, while not mentioned in their manuscript, the expression pattern appears to hold true for the TFs that we identified in this study such as Meis1, Meis2, Nkx2.3, and Hey2 thus confirming our findings. One strength of single-cell technologies is that it allows to finely dissect heterogeneous population. Therefore, as we identify more subpopulations, it will become especially important to define cells in analogous ways to facilitate the integration of multiple studies in advancing the field. While Zhu and colleagues outlined a clear population of “HE” cells [60], we found that most cells in our cluster that immediately followed “pre-HE” already expressed c-Kit and CD41 surface markers thus labeling the cluster “HE/pro-HSPC”. The distinction between the two studies emerged from the way we isolated and defined the cells. Here, we sequenced all CD31+ cells and added surface markers expression in silico based on antibody-derived tag (ADT) detection, whereas Zhu et al. FACS-isolated specific populations a priori. Therefore, we 101 retrospectively defined “HE” cells as c-Kit and CD41 negative cells and c-Kitlow and/or CD41low cells as “pro-HSPC”. However, their prospectively isolated “HE” population encompassed a mixture of c-kitlow/- and/or CD41low/mid/- cells [60]. It is thus likely that our two populations represent similar cell populations. While the new term “pre-HE” may represent a convenient way of labeling those early cells, what functionally limits them in terms of being truly “hemogenic” will be harder to define experimentally. In Zhu et al., in silico analysis hinted a bottleneck at the transition to Runx1-expressing HE cells [60] and the authors mentioned that the molecular regulation behind that developmental barrier is unknown. While we did not thoroughly explore this directly in our CITE-seq dataset, our data may reflect a similar trend. Indeed, if our first population to have significant upregulation of Runx1 (“HE/pro-HSPC”) already expressed the pre-HSPC surface markers c-Kit and CD41 based on antibody-derived tags, the presence of a Runx1-expressing HE (Runx1+c-Kit-CD41-) may be a small and rapidly transient population and we may need to sequence more cells to accurately detect it. Because there are only 48 “HE/pro-HSCP” cells in this CITE-seq dataset, this limits our ability to further investigate that phenomenon. Still, we found a greater proportion of cells in our “pre-HE” and “pre-HSPC” populations as compared to “HE/pro-HSPC” which aligns with Zhu and colleagues’ data. Therefore, what regulates this transition to Runx1-expressing cells becomes the important question. The data presented in Chapter 5 hints that, likely amongst other factors, Meis1 may play an important role in controlling that transition. From our CITE-seq analysis, we further focused on validating the role of Meis1. However, it is likely that Meis1 is not the only critical regulator of HE specification. Our analysis revealed 52 TFs upregulated in pre-hemogenic compared to vascular endothelial cells with Nkx2.3 as the most significantly differentially expressed. Interestingly, our dataset is not the only one to reflect this Nkx2.3 expression pattern, although not directly mentioned elsewhere [60, 161]. Nkx2.3 is a homeobox gene, akin to Hox genes and Meis1, that are typically described as master regulators of developmental processes. Nonetheless, very little is known about Nkx2.3 in hematopoiesis, in particular during embryonic development. Nkx2.3 is expressed in adult HSPC-enriched CD34+ primary cells [177] but is mostly known for its implication in spleen and gut development [178]. Endothelial expression of Nkx2.3 in lymphoid organs regulates leukocyte homing via Madcam1 102 expression [179] and knockdown of Nkx2.3 in cell lines was shown to affect cytokine and chemokine levels [180]. Whether it has a similar or different function in HE cells of the dorsal aorta would have to be determined experimentally. As exemplified in this dissertation, cell-cell interactions and the role of cell non-autonomous signaling are crucial factors during embryogenesis. However, there are still several unanswered questions that needs to be addressed regarding Sash1 signaling in the context of EHT. Mainly, where amongst all the possibilities listed in Chapter 4 is the signal coming from and what is the signaling axis involved? Advances in technologies may facilitate the exploration of such complex interactions as our ability to capture information increases. In the context of EHT, a report published on zebrafish hematopoiesis used laser capture microdissection to investigate six regions of the zebrafish caudal tissue and reconstruct their ligand-receptor interactions with HSPC [176]. A different group also recently performed tomo-seq on zebrafish, chicken, mouse, and human samples to examine the ventral-dorsal and anterior-posterior polarization of microenvironmental signals [181]. These kinds of studies provide a general idea of the secretory molecules that may be present in specific regions of the embryos but not at a single-cell resolution. As the sensitivity, accuracy, and feasibility of techniques such as mass-spectrometry imaging and spatial scRNA-seq become more refined, this could facilitate the identification of direct cell-cell interactions and cross-talk signaling (for instance ligand-receptor interaction) during embryogenesis. In addition, these techniques combining high-throughput expression data (protein or mRNA) with spatial information have the power to detect changes in cell type composition and localization in complex heterogeneous tissues. This could be useful in studying EHT but also in various tissues such as investigating patterning and/or differentiation in the developing lung. While there would still be limitations to these techniques in understanding some microenvironmental factors, such as mechanical forces that can also affect cell fate, it could get us closer to a more comprehensive view of what is happening in vivo. Ultimately, functional assays would nonetheless be required to validate any findings from high-throughput and in silico studies. 103 Regardless of the approach, solving complex interactions remains challenging but pursuing this avenue further might unveil necessary lessons because ultimately, the important question is whether we can leverage the knowledge gained from developmental studies. In the context of EHT, understanding microenvironmental factors means optimizing in vitro protocols that are supposed to mimic EHT, whether it is to develop better assays to characterize cells or more efficient induction of HSPC fate in reprogramming studies to generate HSC that can be used for stem cell transplants. One aspect in common between the most recent efforts to reprogram HSPC de novo has been the need for a proper niche to support HSPC maturation [182-184]. But these reprogramming efforts have relied heavily on knowledge derived from developmental studies from a cell intrinsic perspective. One of the most promising approaches to date used Fosb, Gfi1, Runx1, and Spi1 overexpression in adult mouse endothelial cells [182]. In this thesis, we have shown that loss of Meis1 affects the expression of at least 3 of these 4 genes (Gfi1, Runx1, and Spi1) in vivo during mouse EHT, all genes whose expression is contained entirely within Meis1-expressing endothelial cells. With 41% and 26% of upregulated genes during the EC to pre-HE and pre-HE to HE/pro-HSPC transitions respectively containing a Meis1 binding site, Meis1 could be regulating more than Runx1 to confer hemogenic potential to endothelial cells. 6.3 Sash1 contribution to embryonic development In the adult mouse, Sash1 expression is not restricted to the lung but rather generalized to many organs including the kidney and the brain [90]. As part of this study, we also found a widespread expression of Sash1 in tissue sections of early embryos based on LacZ staining, using the β-galactosidase gene-trap allele in Sash1+/- mice as a proxy for Sash1 transcription and translation. To identify these LacZ positive cells, scRNA-seq provided transcriptome-wide information that was used to infer cell types based on specific gene expression pattern. This approach revealed at least 12 clusters of cells with distinct gene signatures that express Sash1 in our samples at E10.5. These clusters correspond to cells from the mesenchyme, the nervous system, the mesonephros, and the hematopoietic system. The array of cell types detected by scRNA-seq reflects the ubiquitous expression of Sash1 in the dissected region of the embryo that was analyzed. For the purpose of this study, we focused on EHT and therefore, only cells from the AGM and surrounding tissue were collected in this process. Specifically, this includes the dorsal aorta (endothelial and blood cells) and the urogenital ridge (gonad and mesonephros) of the AGM, 104 together with the surrounding mesenchyme and the underlying neural tube. However, it is likely that Sash1 could be expressed in additional regions of the embryo such as the head, limbs, and perhaps fetal liver. Co-staining of LacZ with CD31 antibody confirmed the presence of Sash1 in a subset of endothelial cells in the dorsal aorta. It will be important in the future to perform this dual staining with markers from each of the other identified populations to validate the scRNA-seq findings and more importantly to localize these cells in relationship to the dorsal aorta and emerging IAHC. Cells interact with each other in different ways based on proximity. For instance, adjacent cells can participate in juxtacrine communications, such as the Notch pathway, whereas cells further away would need to migrate or send signals via paracrine or endocrine signaling which involves molecules such as cytokines, chemokines, and hormones. In the context of EHT, only endothelial cells, circulating immune cells, and adjacent pericytes/subaortic mesenchymal cells can participate in a juxtacrine regulation, and most of these cells (except for the mesenchymal subset) were shown to have excised Sash1 in Sash1fl/fl;VEC-Cre embryos. Therefore, understanding the 3D organizational structure nearing the IAHC in the AGM would be beneficial in understanding the type of interactions and candidate signaling pathways involved. Given the extensive expression of Sash1, it would be of interest to uncover whether Sash1 affects additional developmental processes. For example, Sash1 is clearly expressed in the nervous system and all around the neural tube at E10.5 (data not shown). Whether it is required for some aspect of neural development is unknown. As a large protein containing multiple binding domains, the signaling pathways regulated by Sash1 should be investigated in a cell-specific and context-dependent manner. Our lab described the activation of SASH1 downstream of TLR4 in microvascular endothelial cells [90]. However, amongst the Sash1-expressing cell populations identified in this study, only neutrophils, macrophages, and some endothelial cells had detectable Tlr4 expression in our dataset. Thus, Sash1 might also be acting downstream of alternative receptors/pathways. Additionally, data from our group has shown that Sash1 can drive distinct pathways based on its interacting partners; binding β-Arrestin1 to activate Akt or activating NF-κB in complex with TRAF-6 [90, 109]. Consequently, the availability of binding partners and perhaps its subcellular localization may affect Sash1 targets in the cell. While experiments 105 conducted in various cell lines converge towards a similar phenotype, the in vivo function of Sash1 remains to be studied across multiple cells types to determine its effect. A common observation between the roles of Sash1 in lung organogenesis and embryonic hematopoiesis is the implication of cell non-autonomous pathways. Yet, a surprising contrast between the two developmental processes is the origin and directionality of that interaction. While the surfactant and NO defect observed in the lung of Sash1-/- embryos was replicated by endothelial-restricted deletion of Sash1 in Sash1fl/fl;VEC-Cre embryos, the defect observed in pre-HSPC numbers at E10.5 was not. Together with the distinct expression pattern observed in each organ respectively, the data argues an endothelial origin in the lung as opposed to an extrinsic signaling in the AGM, which targets a subset of specialized endothelial cells, the HE. In both cases, this affects the differentiation of the receiving cell. Thus, the endothelium influences surrounding cells types (for example AE2C) but also receives cues from its surroundings reflecting the interdependency of biological systems. Regarding the effect of Sash1 on hematopoiesis, analysis of the types of cells that emerge from each IAHC was not performed which is a limitation of this study. Altering the ex vivo environment influences the type of cell outputs produced by HE in culture [57]. Therefore, it would be interesting to further investigate the effect of Sash1-deletion on the functional progenitor cells produced in the AGM. Our current data suggests that progenitors can form colonies in culture regardless of Sash1 expression but whether there is a bias in vivo could be explored more rigorously. Finally, the number of IAHC found in Sash1-/- E10.5 embryos is reduced by 2-fold compared to WT embryos which matches the reduction in the number of immunophenotypic pre-HSPC. However, the fold change initially observed in the fetal liver based on the repopulation assay was 4.6 times fewer HSC. Many factors can influence the number of HSPC present at a given point in time, including cell proliferation, cell viability, apoptosis, and cellular differentiation. The fetal liver is a major embryonic niche for HSPC and promotes their expansion. Thus, the differences observed between the magnitude of the defect at E10.5 and E14.5 could be attributed to fewer HSPC successfully colonizing the fetal liver or perhaps additional issues with expansion 106 in the fetal liver. Therefore, it would be interesting to see whether Sash1-deletion has an impact on the fetal liver niche, which also organizes around the endothelium. In conclusion, this study was initiated with the goal of defining the role of an essential developmental gene in embryogenesis and better characterizing critical developmental pathways during organogenesis. The findings presented in the chapters above illustrate the cell non-autonomous roles of Sash1 in the development of the lung and definitive hematopoietic system. In Chapter 3, we described a pathway originating within endothelial cells to promote alveolar cell maturation which exemplifies the importance of the endothelium in the regulation of organ formation. 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Thesis/Dissertation
eng
Experimental Medicine
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Endothelial regulation in cell fate specification during embryonic development of the lung and the hematopoietic system
Text