Medicine, Faculty of
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UBCV
Arostegui, Martin
Doctor of Philosophy - PhD
University of British Columbia
Tissue development and regeneration rely on the cooperation of multiple mesenchymal progenitor (MP) subpopulations. Recently, the gene Hypermethylated in cancer 1 ( Hic1) has been identified as a marker of quiescent MPs in multiple adult tissues. Hic1⁺ MPs have since been implicated in the repair and regeneration of several organs such as skeletal muscle (SkM), skin, heart, and brain. While the roles that Hic1⁺ MPs play in tissue maintenance are currently being elucidated, knowledge surrounding their developmental origin and fate is lacking. Using the developing forelimb as a model system, we describe the embryonic origin of Hic1⁺ MPs and demonstrate that they first appear in the developing limb ~48 hours after limb bud initiation at embryonic day (E) 11.5 and represent a rare subset of the total mesenchymal cell population. Time-resolved single cell-omics analyses coupled with lineage tracing reveal that Hic1⁺ cells generate a unique MP hierarchy, that includes both recently identified adult universal fibroblast populations (Dpt⁺, Pi16⁺ and Dpt⁺, Col15a1⁺) and more specialized mesenchymal derivatives such as, peri- and endo-neural cells, pericytes, bone marrow stromal cells (BMSCs), myotenocytes, tenocytes, and fascia-resident fibroblasts, with limited contributions to chondrocytes and osteocytes within the skeletal elements. We further demonstrate that, in the adult organism, MPs endure within these compartments, continue to express Hic1 and represent a critical reservoir to support post-natal growth and regeneration. Within the appendicular skeleton, Hic1⁺ descendants do not contribute to the primary skeletal anlagen but represent the elusive MP population that generate progeny which connect bone to tendon (entheses) and tendon to muscle (myotendinous junction). Furthermore, deletion of Hic1 leads to skeletal defects reflective of deficient muscle-bone coupling. Collectively, these findings demonstrate that Hic1 identifies a unique MP population that contributes to multiple cell lineages during limb development. Furthermore, Hic1⁺ MPs are crucial for appendicular skeletogenesis as their progeny primarily constitute several forelimb tendon-bone and tendon-muscle attachment regions. As parallels between tissue regeneration and embryonic development are being increasingly made, understanding the origin and fate of heterogenous tissue-resident MPs will support efforts aimed at promoting regeneration in mature tissues.
Origin, Fate, and Function of Embryonic Hic1-positive Mesenchymal Progenitors in the Developing Limb by Martin Arostegui B.Sc., The University of British Columbia, 2015 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Cell and Developmental Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) July 2022 © Martin Arostegui, 2022 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Origin, Fate, and Function of Embryonic Hic1-positive Mesenchymal Progenitors in the Developing Limb submitted by Martin Arostegui in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Cell and Developmental Biology Examining Committee: Michael Underhill, Professor, Cell and Developmental Biology, UBC Supervisor Timothy O’Connor, Professor, Cell and Developmental Biology, UBC Supervisory Committee Member Louis Lefebvre, Associate Professor, Medical Genetics, UBC University Examiner Francis Lynn, Associate Professor, Surgery, UBC University Examiner Additional Supervisory Committee Members: Douglas Allan, Associate Professor, Cell and Developmental Biology, UBC Supervisory Committee Member Pamela Hoodless, Professor, Medical Genetics, UBC Supervisory Committee Member iii Abstract Tissue development and regeneration rely on the cooperation of multiple mesenchymal progenitor (MP) subpopulations. Recently, the gene Hypermethylated in cancer 1 ( Hic1) has been identified as a marker of quiescent MPs in multiple adult tissues. Hic1+ MPs have since been implicated in the repair and regeneration of several organs such as skeletal muscle (SkM), skin, heart, and brain. While the roles that Hic1+ MPs play in tissue maintenance are currently being elucidated, knowledge surrounding their developmental origin and fate is lacking. Using the developing forelimb as a model system, we describe the embryonic origin of Hic1+ MPs and demonstrate that they first appear in the developing limb ~48 hours after limb bud initiation at embryonic day (E) 11.5 and represent a rare subset of the total mesenchymal cell population. Time-resolved single cell-omics analyses coupled with lineage tracing reveal that Hic1+ cells generate a unique MP hierarchy, that includes both recently identified adult universal fibroblast populations (Dpt+, Pi16+ and Dpt+, Col15a1+) and more specialized mesenchymal derivatives such as, peri- and endo-neural cells, pericytes, bone marrow stromal cells (BMSCs), myotenocytes, tenocytes, and fascia-resident fibroblasts, with limited contributions to chondrocytes and osteocytes within the skeletal elements. We further demonstrate that, in the adult organism, MPs endure within these compartments, continue to express Hic1 and represent a critical reservoir to support post-natal growth and regeneration. Within the appendicular skeleton, Hic1+ descendants do not contribute to the primary skeletal anlagen but represent the elusive MP population that generate progeny which connect bone to tendon (entheses) and tendon to muscle (myotendinous junction). Furthermore, deletion of Hic1 leads to skeletal defects reflective of deficient muscle-bone coupling. Collectively, these findings demonstrate that Hic1 identifies a unique MP population that contributes to multiple cell lineages during limb development. Furthermore, Hic1+ MPs are crucial iv for appendicular skeletogenesis as their progeny primarily constitute several forelimb tendon-bone and tendon-muscle attachment regions. As parallels between tissue regeneration and embryonic development are being increasingly made, understanding the origin and fate of heterogenous tissue-resident MPs will support efforts aimed at promoting regeneration in mature tissues. v Lay Summary In humans, most organs are capable of a remarkable degree of regeneration or repair. When an organ is injured, groups of specialized cells called mesenchymal progenitors (MPs) that are found within the affected area become activated and initiate a response to fix the damage. Several of the processes that are involved in the activation of these cells are related to programs that are originally needed during embryonic development. Therefore, understanding the behavior of MPs during development such as where they come from and what kind of cells they give rise to is crucial for generating drugs and/or treatments that may accelerate tissue regeneration in an adult. The research described here has determined where a subset of embryonic MPs originate, and what cells and structures they are responsible for creating in the developing limb. This work represents the foundation for understanding the roles that MPs play during development, some aspects of which could become applicable in regenerative medicine. vi Preface A version of chapter 2 has been published. Arostegui, M., Underhill, T.M. (2021) Murine Limb Bud Organ Cultures for Studying Musculoskeletal Development. Methods In Molecular Biology 2021;2230:115-137. I optimized all methodologies described in this publication. Michael Underhill and I wrote the manuscript for the published chapter. A version of chapter 3 has been submitted and is currently in revision. Cellular taxonomy of Hic1+ mesenchymal progenitor derivatives in the limb; from embryo to adult. Martin Arostegui, R. Wilder Scott, Kerstin Bose, and T. Michael Underhill. All experiments described in this paper were performed my Wilder Scott and me. Kerstin Bose characterized the Hic1F/F mouse model. The submitted manuscript was written and edited by Wilder Scott, Michael Underhill, and me. Michael Underhill and I were responsible for the experimental design and data interpretation. Chapter 4 is a modified version of a manuscript in final stages of preparation. Hic1+ progenitors contribute to tendon-bone and tendon-muscle attachment sites during limb development. Martin Arostegui, R. Wilder Scott, and T. Michael Underhill. All experiments were performed by Wilder Scott and me. I wrote the initial version of the manuscript. Michael Underhill and Wilder Scott edited the manuscript text. Michael Underhill and I were responsible for the experimental design and data interpretation. All studies in this thesis were conceived and designed by Michael Underhill and me. I performed all experiments and bioinformatic data analysis, with the exception of the Cicero and Signac analyses presented in Figure 4.15 which were generated by Wilder Scott. vii Animal studies were reviewed and approved by the University of British Columbia Committee on Animal Care under protocols A17-0273, A19-0221 and A19-0279. viii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ....................................................................................................................... viii List of Tables ............................................................................................................................... xii List of Figures ............................................................................................................................. xiii List of Abbreviations ............................................................................................................... xviii Acknowledgements .................................................................................................................... xxi Dedication .................................................................................................................................. xxii Chapter 1: Introduction ................................................................................................................1 1.1 Development of the appendicular musculoskeletal system ............................................ 1 1.2 Development of tendon-bone attachment units .............................................................. 4 1.3 Origin and function of muscle connective tissue fibroblasts and FAPs ......................... 6 1.4 Origin and function of dermal fibroblast populations ..................................................... 9 1.5 Origin and function of pericytes ................................................................................... 11 1.6 Origin and function of bone marrow stromal cells ....................................................... 13 1.7 Development and function of nerve-associated fibroblast populations ........................ 15 1.8 Hypermethylated in Cancer-1 (Hic1) ............................................................................ 18 1.9 Mesenchymal progenitors or mesenchymal stem cells? ............................................... 19 1.10 MPs in skeletal muscle repair and regeneration ........................................................... 21 1.11 Key technological advances .......................................................................................... 23 ix 1.11.1 Embryonic and postnatal lineage tracing of Hic1CreERT2; R26tdTomato mice ........... 23 1.11.2 Combined use of micromass, limb culture explants, and wholemount immunofluorescence ............................................................................................. 25 1.12 Objectives of this work ................................................................................................. 28 Chapter 2: Methods .....................................................................................................................30 2.1 Mice .............................................................................................................................. 30 2.2 Muscle Injury ................................................................................................................ 34 2.3 Immunofluorescence staining ....................................................................................... 35 2.4 Generation and detection of anti-HIC1 antibody .......................................................... 37 2.5 Wholemount immunofluorescence staining and imaging ............................................. 38 2.6 Flow cytometry and fluorescence-activated cell sorting (FACS) methodology ........... 39 2.7 Cell cycle analysis ......................................................................................................... 40 2.8 Single cell RNA-Seq ..................................................................................................... 41 2.9 Single-cell fate trajectory reconstruction ...................................................................... 44 2.10 Single cell ATAC-seq ................................................................................................... 45 2.11 Ex-vivo Forelimb Cultures ............................................................................................ 47 2.12 Micromass Cultures ...................................................................................................... 47 2.13 Wholemount and cryosection X-gal staining ................................................................ 49 2.14 Whole-mount skeletal preparations .............................................................................. 50 2.15 Quantification and Statistical analysis .......................................................................... 51 Chapter 3: Cellular taxonomy of Hic1+ mesenchymal progenitor derivatives in the limb from embryo to adult ...................................................................................................................55 3.1 Introduction ................................................................................................................... 55 x 3.2 Results ........................................................................................................................... 56 3.2.1 Hic1 expression patterns in the mouse embryo .................................................... 56 3.2.2 Hic1 identifies a rare MP population within the limb bud .................................... 61 3.2.3 ScRNA-seq and histological analyses reveal a unique MP hierarchy .................. 80 3.2.4 Lineage potential of Hic1+ MPs is embedded in the chromatin landscape ........... 91 3.2.5 Embryonic Hic1+ MP post-natal lineage contributions ........................................ 97 3.2.6 Embryonic Hic1+ fibroblasts persist into adulthood and have roles in SkM regeneration ......................................................................................................... 104 Chapter 4: Hic1 identifies a population of embryonic chondro-tenoprogenitors responsible for secondary bone morphogenesis ..........................................................................................108 4.1 Introduction ................................................................................................................. 108 4.2 Results ......................................................................................................................... 111 4.2.1 Hic1 identifies a subpopulation of ScxGFP+ MPs in the developing limb bud .. 111 4.2.2 Hic1+ MP derivatives populate musculoskeletal attachment units ..................... 121 4.2.3 Chondrocytes and tenocytes emerge from an embryonic subpopulation of bipotent Hic1+ MPs ........................................................................................................... 129 4.2.4 Hic1 deletion negatively impacts skeletal morphogenesis ................................. 137 Chapter 5: Conclusion ...............................................................................................................146 5.1 Overview ..................................................................................................................... 146 5.2 On the Embryonic Origin of Hic1+ MPs ..................................................................... 150 5.3 Hic1+ MPs and “classic” mesenchymal cell types ...................................................... 154 5.4 Hic1+ MP derivatives contribute to fibroblast and peripheral nerve mesenchymal components during development ................................................................................ 156 xi 5.5 Embryonic Hic1+ MPs represent a source of mural cells ........................................... 158 5.6 Embryonic Hic1+ MP progeny in enthesis and MTJ development ............................. 160 5.7 Embryonic Hic1+ MP progeny in adult tissue repair and regeneration ...................... 162 5.8 Final remarks .............................................................................................................. 163 References ...................................................................................................................................165 xii List of Tables Table 2.1 Summary of antibodies used ......................................................................................... 52 Table 4.1 List of observed morphological deficiencies in long bones of Prrx1-Cre; Hic1f/f mice...................................................................................................................................................... 144 xiii List of Figures Figure 1.1 Somite partitions. ........................................................................................................... 2 Figure 1.2 Schematic of a mouse forelimb highlighting the embryonic origin of musculoskeletal tissues. ............................................................................................................................................. 3 Figure 1.3 Schematic representation of a tendon-bone attachment and myotendinous junction .... 5 Figure 2.1 Schematic representation of Hic1 expression patterns in Hic1nLacZ embryos. ............ 31 Figure 2.2 Hic1 alleles and derived mouse models. ..................................................................... 32 Figure 2.3 Schematic representation of tdTomato expression patterns in Hic1CreERT2/R26tdtomato embryos recombined at E10. ......................................................................................................... 33 Figure 3.1 Hic1 expression dynamics in early embryonic development visualized via the Hic1nLacZ mouse line. .................................................................................................................... 57 Figure 3.2 Determination of optimal embryonic timepoint for Cre-induced recombination in Hic1CreERT2; R26tdTomato embryos. .................................................................................................. 58 Figure 3.3 Analysis of tdTomato+ cell distribution in somitic mesoderm at E11.5. ..................... 61 Figure 3.4 Quantitative analysis of Hic1 expression in the proximal-distal axis of E10.5 and E11.5 forelimb buds. ..................................................................................................................... 62 Figure 3.5 Hic1 identifies a discrete MP population within the limb mesenchyme. .................... 64 Figure 3.6 Quantitative analysis of tdTomato+ cells in Hic1CreERT2; R26tdTomato forelimbs at sequential developmental time points. .......................................................................................... 65 Figure 3.7 TdTomato+ MPs are distinct from limb LPM and localize with nerves and myogenic progenitors. ................................................................................................................................... 66 xiv Figure 3.8 Live imaging of Hic1CreERT2; R26tdtomato forelimbs identifies a highly dynamic tdTomato+ population. .................................................................................................................. 69 Figure 3.9 TdTomato+ MPs are localized to the proximal forelimb during early limb morphogenesis. ............................................................................................................................. 70 Figure 3.10 Live imaging of Hic1CreERT2; R26tdTomato micromass cultures highlights the movement of tdTomato+ MPs throughout internodular regions. .................................................................... 71 Figure 3.11 TdTomato+ MPs do not overlap with myotome and neural crest-derived migratory populations at the forelimb level. .................................................................................................. 72 Figure 3.12 TdTomato+ MPs partially overlap with sclerotome and syndetome populations at the forelimb level. ............................................................................................................................... 73 Figure 3.13 Immunofluorescence detection of HIC1 shows that HIC1+ MPs partially overlap with sclerotome and syndetome as early as E10.5. ....................................................................... 75 Figure 3.14 Immunofluorescence detection of HIC1 shows that HIC1+ MPs and myogenic cells represent separate progenitor populations. ................................................................................... 76 Figure 3.15 Immunofluorescence detection of HIC1 shows that HIC1+ MPs are in close proximity to extending nerve axons and are distinct from migratory myogenic progenitors. ...... 77 Figure 3.16 HIC1+ MPs are distinct from SOX10+ neural crest progenitors and PAX7+ myogenic precursors. ..................................................................................................................................... 78 Figure 3.17 Distribution of tdTomato+ cells in E12.5 and E13.5 forelimbs. ................................ 79 Figure 3.18 Proliferation dynamics of tdTomato+ cells in Hic1CreERT2; R26tdTomato embryonic forelimbs. ...................................................................................................................................... 80 Figure 3.19 Single-cell RNA-sequencing reveals multiple cell fates for tdTomato+ MPs throughout limb development. ...................................................................................................... 81 xv Figure 3.20 URD-generated t-SNE plot of E16.5 tdTomato+ cells used to identify terminal cell fates. .............................................................................................................................................. 82 Figure 3.21 ScRNA-seq analyses define a mesenchymal progenitor hierarchy. .......................... 84 Figure 3.22 Cells located at the root express progenitor markers. ................................................ 85 Figure 3.23 Characterization of embryonic Hic1+ MP-derived fibroblast populations. ............... 86 Figure 3.24 Lineage tracing analyses confirm computationally inferred lineage potential. ......... 88 Figure 3.25 Embryonic Hic1+ progenitors do not contribute to skeletal muscle nor endothelial lineages. ........................................................................................................................................ 89 Figure 3.26 Adipogenic potential of tdTomato+ MPs at the single cell level. .............................. 90 Figure 3.27 A broad mesenchymal lineage potential is embedded in the epigenome of early tdTomato+ MPs. ............................................................................................................................ 92 Figure 3.28 Hic1+ MPs become compartmentalized by gene expression at E16.5. ..................... 95 Figure 3.29 MP specification and differentiation are reflected in chromatin and transcriptome dynamics during their developmental trajectories. ....................................................................... 96 Figure 3.30 Embryonic Hic1+ MPs give rise to distinct populations of adult stromal cells. ........ 99 Figure 3.31 Characterization of mural cell fates. ........................................................................ 100 Figure 3.32 Histological confirmation of perineural fate for Hic1-lineage MPs. ....................... 102 Figure 3.33 Embryonic Hic1+ MPs contribute to diverse cell types that persist into adulthood. 103 Figure 3.34 Lineage tracing of embryonic Hic1+ MPs into adulthood reveals development-specific cell types. ....................................................................................................................... 106 Figure 3.35 Embryonic Hic1-lineage FAPs persist postnatally and support muscle regeneration...................................................................................................................................................... 107 xvi Figure 4.1 Hic1nLacZ embryos show strips of LacZ+ cells extending from the hypaxial somites into the forelimb bud. .................................................................................................................. 113 Figure 4.2 Characterization of Hic1 expression patterns in the embryonic limb bud. ............... 114 Figure 4.3 Hic1+ MPs localize to the internodular mesenchyme in limb micromass cultures. .. 115 Figure 4.4 Characterization and quantification of tdTomato/ScxGFP double-positive cells in Hic1CreERT2; R26tdTomato; ScxGFP forelimbs. ............................................................................... 117 Figure 4.5 Spatial characterization of tdTomato+/ScxGFP+ cells in E11.5 Hic1CreERT2; R26tdTomato; ScxGFP embryos. ....................................................................................................................... 118 Figure 4.6 TdTomato+ MPs and myocytes localize to internodular regions in PLM cultures. .. 119 Figure 4.7 Quantification of ScxGFP+ cells within the tdTomato+ MP population. .................. 122 Figure 4.8 Early forelimb mesenchyme contains Hic1+ chondro-tenogenic MPs. ..................... 123 Figure 4.9 Hic1+ MPs contribute specifically to the bone-tendon attachment units but not the tendon body nor primary cartilage anlagen. ............................................................................... 126 Figure 4.10 Wholemount visualization of forelimbs reveals localization of tdTomato+ cartilage and tendons. ................................................................................................................................ 127 Figure 4.11 A subset of tdTomato+ MPs are ScxGFP/SOX9 double-positive. .......................... 130 Figure 4.12 TdTomato+ MPs contribute to the myotenocyte compartment during development of the MTJ. ...................................................................................................................................... 131 Figure 4.13 Analysis of the Hic1CreERT2; R26tdTomato embryonic forelimb scRNA-seq dataset with Harmony identifies four general fates. ........................................................................................ 134 Figure 4.14 ScRNA-seq analyses reveal differentiation trajectories for chondrogenic and tenogenic progenitors. ................................................................................................................. 135 Figure 4.15 A subpopulation of Hic1+ MPs are bipotent chondro-tenoprogenitors. .................. 136 xvii Figure 4.16 Characterization of Hic1nLacZ/nLacZ skeletal deformities. .......................................... 139 Figure 4.17 Hic1 deletion severely impacts secondary skeletal morphogenesis. ....................... 141 Figure 4.18 Consequences of Hic1 deletion on forelimb long bone morphology. ..................... 143 Figure 5.1 Fate of Hic1+ MPs in limb development. .................................................................. 149 xviii List of Abbreviations 4-OHT - 4-hydroxytamoxifen BABB - Benzoic acid benzyl benzoate BMSC - Bone marrow stromal cell BSA - Bovine serum albumin CFUf - Colony forming unit fibroblast cKO - Conventional knockout CNS - Central nervous system CreERT2 - Tamoxifen inducible P1 bacteriophage derived Cre recombinase v. 2 DAPI - 4′,6-diamidino-2-phenylindole DF - Dermal fibroblast DMEM - Dulbecco's modified eagle medium DNA - Deoxyribonucleic acid DTR - Diphtheria toxin receptor E - Embryonic day ECM - Extracellular matrix EdU - 5-ethynyl-2’-deoxyuridine f – Floxed allele FACS - Fluorescence-activated cell sorting FAPs - Fibro-adipogenic progenitors FCM - Flow cytometry Fibro1 - Fibroblast cluster 1 xix Fibro2 - Fibroblast cluster 2 HiRE - Hic1 response element HSC - Hematopoietic stem cell IRES - Internal ribosome entry site KO - Knockout LPM - Lateral plate mesoderm MCs - Mitotic cells MCT - Muscle connective tissue MEF - Mouse embryonic fibroblast MP - Mesenchymal progenitor mRNA - Messenger ribonucleic acid MSC - Mesenchymal stem cell MTJ - Myotendinous tendinous junction NLS - Nuclear localization signal NTX - Notexin snake venom phospholipase extract OCR - Open chromatin region OCT - Optimal cutting temperature P - Postnatal day PBS - Phosphate buffered saline PDFP - Papillary dermal fibroblast progenitor PET - Polyester PFA - Paraformaldehyde xx PLM - Primary limb mesenchyme PNS - Peripheral nervous system R26 - ROSA26 RA - Retinoic acid RAR - Retinoic acid receptor RDFP - Reticular dermal fibroblast progenitor RNA - Ribonucleic acid scATAC-seq - Single cell assay for transposase-accessible chromatin SCPs - Schwann cell precursors scRNA-seq - Single cell RNA-sequencing SD - Standard deviation SkM - Skeletal muscle SMCPs - Smooth muscle cell progenitors t-SNE - t-distributed stochastic neighbor embedding TA - Tibialis anterior TAM - Tamoxifen UMAP - Uniform manifold approximation and projection UMI - Unique molecular identifier VSMCs - Vascular smooth muscle cells X-gal - 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside µCT - Micro-computed tomography xxi Acknowledgements Over the course of my PhD studies I have been lucky enough to have accumulated a wealth of friends and colleagues to whom I am indebted for making my time as a graduate student truly memorable. First, I would like to thank my supervisor Dr. Michael Underhill for the opportunity to conduct research in his lab. This work would not have been possible without Michael’s exceptional mentorship and guidance throughout the past several years. Similarly, I am grateful for my supervisory committee, Drs. Pamela Hoodless, Doug Allan, and Tim O’Connor for providing continuous support and helpful discussions during committee meetings. I would like to give a special thanks to Dr. Le Su, who showed me the ways of tissue culture and never doubted the quality of my western blots. Another special thanks has to go to Dr. Wilder Scott who has mentored me since day one, in laboratory techniques, early 2000’s comedy, and how to deal with conflict in the workplace. I am also thankful to the Underhill lab members, past and present, for sharing the highs and lows of graduate school with me and providing a sense of camaraderie in the lab. In no particular order, I am grateful to Petra Schreiner, Kim Weigand, George Davenport III, Tiffany Lam, Danial Taušan, Kathleen Martin, Germain Ho, Amanda Yang, Howell Liu, Lauren Martin, Adam Diett, Atefeh Samani, Cindy Crichlow, Lesley Hill, and William Bastedo. I would also like to acknowledge my program, Cell and Developmental Biology as well as the UBC Faculty of Medicine for providing funds to support myself during my studies. Getting through graduate school required more than academic and financial support, I would like to thank my family, Carlos E. Aróstegui Raviella, Luz María Vargas García, Carlos Arostegui, and Nayeli Arostegui, as well as, Ms. Geunhye Song for their continued support and patience throughout the entirety of my studies. With all sincerity, thank you. xxii Dedication For my mother and father who always believed in me1 Chapter 1: Introduction 1.1 Development of the appendicular musculoskeletal system The limb musculoskeletal system is composed of muscle, bone, and associated connective tissues such as tendon, fascia, and ligament. Several other physiological systems are required for limbs to become functional units. The integumentary, circulatory, and nervous systems all comprise essential components of a functional limb. These organs and associated tissues interact cooperatively to bestow the organism with locomotion and structural stability1. Although the mature limb is a prime example of cross-tissue communication and cooperation, most tissues that comprise an adult limb are derived from different embryological sources and, during development, undergo a series of migratory, proliferative, and apoptotic events that ultimately generate an integrated and functional appendage. Studies performed on vertebrate model organisms such as birds and mice have shown that two germ layers provide most of the cell types needed for limb development; the mesoderm and ectoderm. The mesoderm is generated during gastrulation and is further subdivided into lateral plate, somitic, and intermediate mesoderm. Lateral plate mesoderm (LPM) surrounds the embryonic cavities and is the source of ventral and limb dermis as well as the appendicular and axial cartilage, bones, and tendons. Somitic mesoderm compartmentalizes in an anterior-posterior fashion into discrete units called somites. Different somite regions are identified depending on the tissue that they give rise to (Figure 1.1). The dermomyotome is the dorsolateral portion of each somite and gives rise to the dorsal skin dermis and underlying components thereof, as well as, to the dorsal, ventral, and appendicular musculature. The syndetome is a source of tenogenic progenitors; however, their contributions seem to be restricted to the axial skeleton exclusively2. 2 The sclerotome is the ventral-most portion of the somite and gives rise to vertebral cartilage. The ectoderm is the outermost germ layer and is the source of skin epithelium and the central nervous system. Neural crest cells (sometimes referred to as a “fourth germ layer”) delaminate from the dorsal portion of the neural tube and migrate in a dorsal-ventral direction giving rise to several tissues including the peripheral and enteric nervous system. Figure 1.1 Somite partitions. (a-b) Schematic representations of somite subdivisions. In mice, forelimb morphogenesis begins at E9.5 with a thickening of the ectoderm overlying the prospective limb fields and expansion of LPM mesenchyme below it to form an outgrowth from the embryonic trunk3. As the limb bud extends along the proximal-distal axis, mesenchymal cells in its core begin to condense and differentiate to form a continuous cartilaginous anlagen which will afterwards be subdivided into discrete skeletal regions via the emergence of synovial joints4 and endochondral ossification5. At E10.5 somite-derived muscle progenitors begin to migrate into the forelimb buds as streams of cells in conjunction with nerve axon projections6. 3 Once in the limb bud, myogenic progenitors segregate into dorsal and ventral masses and begin to self-organize into an array of differentiated myotubes that follow the direction of the future mature muscle fibers7,8. These muscle bundles will sometimes split to allow for the formation of smaller individual muscle groups9. Simultaneously, tendon primordia begin to form between muscle masses. These extend and individuate into specific tendons which then sequentially attach each to their respective cartilaginous elements. As development proceeds, mature tendons connect to adjoining myotubes and form the myotendinous junction (MTJ)10. Little is known about the mechanisms and series of events involved in MTJ formation; however, recent publications have suggested an important role for BMP-signaling and muscle connective fibroblasts in the establishment of functional MTJs during development11,12. In summary, the limb musculoskeletal system has a dual origin; somite-derived muscles in compliment with LPM-derived bone, tendon, and associated connective tissues13,14. Figure 1.2 Schematic of a mouse forelimb highlighting the embryonic origin of musculoskeletal tissues. For simplicity, only select muscles and tendons are illustrated. 4 1.2 Development of tendon-bone attachment units For limbs to create motion, muscle must transmit its generated forces onto the skeletal framework via tendons (Figure 1.3). Tendons are specialized connective tissues formed primarily by type I collagen fibrils organized in parallel to the tendon axis15. The discovery of Scleraxis in the early 2000’s, a basic helix-loop-helix transcription factor, as a marker of connective tissues that mediate the attachment of muscle to bone16, has been instrumental in uncovering the cellular and molecular mechanisms underlying tenogenesis in the developing embryo. Limb tendon progenitors reside within the LPM of the forming limb bud and differentiate into mature tendons upon interactions with cartilage and muscle precursors17. Interestingly, the formation of proximal tendons diverges from their distal counterparts. Proximal elements develop in tandem with their associated musculature. In contrast, distal tendons are formed in coordination with neighboring skeletal structures and in separation from their respective muscles16. Tendon development overall is closely associated with that of musculature, the examination of muscleless limbs has shown that tendon progenitors initiate their differentiation but further formation of mature tendons requires the presence of muscle16,18. As limb growth proceeds, increasingly specialized skeletal structures are formed that require the precise coordination of various progenitor cell populations. An example is the formation of the tendon-to-bone attachment unit, termed the enthesis, which requires tendon, cartilage, and muscle progenitors to cooperate either directly or indirectly in its genesis19–21. Entheses are critical in dissipating and transmitting forces generated by associated muscles22,23. Our ability to move relies on muscles to generate contractile forces which are transmitted first to tendon and subsequently to bone via entheses. There are two types of entheses: Fibrous, in which 5 the tendon attaches directly to bone, and fibrocartilaginous where the tendon attaches to a bone through a fibrocartilage intermediary. Typically, a fibrocartilaginous tendon-bone attachment is composed of four zones: dense fibrous connective tissue, uncalcified fibrocartilage, calcified fibrocartilage, and bone. This four-zone arrangement allows the enthesis to become a mechanically efficient force-transmitting structure and to protect the attachment site from failure caused by a sudden release of energy originating from a mechanically flexible material (muscle and tendon) to a relatively stiff one (bone)24. Figure 1.3 Schematic representation of a tendon-bone attachment and myotendinous junction 6 It has recently been shown that tendon-bone attachment units of the mouse limbs form in a modular fashion by a unique pool of Scx+ Sox9+ progenitors following the development of the main cartilaginous template of the long bones25. In early limb skeletogenesis, LPM-derived mesenchymal cells first condense and differentiate into chondrocytes to form the cartilage anlagen of what will eventually become skeletal elements. At this stage, all chondrocytes express the basic helix-loop-helix transcription factor Sox926–28. Enthesis formation begins at this stage, when the primary cartilage template is morphologically modified by a distinct population of Scx+ Sox9+ progenitors. These cells give the premature cylindrical template a mature form via the addition of tendon-bone attachment units. The embryonic origin of this unique pool of mesenchymal progenitors is currently unknown. Mechanistically, TGF-b is required for the specification of enthesis progenitors, Tgfbr2 knockouts lack Sox9 lineage progenitors in and around sites of prospective entheses compared to wild type controls. Following specification, BMP4 is required for the differentiation of Scx+/Sox9+ enthesis progenitors into chondrocytes. Ablation of BMP4 in developing embryos result in the failure to express Col2a1 (a mostly cartilage-specific collagen) in mutant limb tendon attachment units29. 1.3 Origin and function of muscle connective tissue fibroblasts and FAPs All biological tissues contain an extra cellular matrix (ECM) that serves as a scaffolding and signal transduction medium for the surrounding parenchymal and non-parenchymal cells30. The ECM is mainly composed of protein fibers, proteoglycans, and a mix thereof 31. Fibroblasts are cells found in the stroma of tissues and are responsible for depositing basic units of the ECM such as collagens, fibronectin, elastin, among other components32–34. The development and maintenance of skeletal musculature requires the support of stromal cell populations such as 7 muscle connective tissue (MCT) fibroblasts and fibro-adipogenic progenitors (FAPs)35–37. During limb development, Tcf4 (Tcf7l2)-positive connective tissue fibroblasts are responsible for laying down a prepatterned ECM in which muscle progenitors will organize and differentiate8. These connective tissue cells have also been shown to be responsible for crucial steps in the myogenic differentiation program such as muscle fiber type development and maturation38. Furthermore, a modest ablation of Tcf4+ fibroblasts (~40% reduction) in adults is sufficient to produce deficient muscle regeneration resulting in lower-volume muscle fibers due to premature differentiation and reduced proliferation of Pax7+ satellite cells39. Satellite cells are myogenic progenitors that maintain SkM tissue homeostasis. FAPs are another stromal cell population that have been linked with supporting muscle repair and regeneration. FAPs were initially identified by expression of Pdgfra, Ly6a, and Cd34 as well as their differentiation potential into adipogenic and fibrogenic lineages40,41. Indeed, FAPs have been shown to be responsible for excessive fibro-fatty deposition giving rise to fibrosis in ageing or chronically damaged muscles40,41. Genetic ablation of FAPs results in skeletal muscle atrophy and impaired regenerative potential42. Similarly, depletion of a subset of proinflammatory FAPs after acute injury limits the production of profibrotic cells and reduces interstitial collagen accumulation43. During normal muscle regeneration, FAPs proliferate, release chemokines and cytokines to facilitate an early immune response44,45, localize to ailing myofibers and secrete factors that promote myogenic differentiation40,46,47. Once the injury has resolved, excess FAP derivatives are cleared via apoptosis and tissue integrity is restored48,49. However, in myopathies such as Duchenne muscular dystrophy, muscles are not able to regenerate efficiently due to the chronic nature of the disease. In these cases, muscle repair is employed to maintain tissue integrity at the expense of functionality, the result is fibro-fatty lesions that compromise muscular contraction50,51. 8 In adult skeletal muscle (SkM), FAPs express Tcf445 and a subset of MCT fibroblasts express Pdgfra39, indicating a potential overlap between the two cell populations. Functionally, both MCT fibroblasts and FAPs respond to muscle injury in a similar fashion: an initial wave of proliferation, followed by the creation of a provisional ECM that promotes a regenerative microenvironment, and ending with a return of cell numbers back to homeostatic levels39,45. Furthermore, a recent study in which plastic-adherent MCT fibroblasts were isolated and characterized, determined that the phenotypic and biochemical properties of these cells were equivalent to those of FAPs52. Therefore, it seems that adult MCT fibroblasts and FAPs share common functions, genetic markers, and tissue localization; suggesting that these cell populations may be largely overlapping53. A common embryonic origin would help elucidate the lineage relationship between MCT fibroblasts and FAPs. While Tcf4+ cells in the limb have been studied in a developmental context and been suggested to be LPM-derived8; no such studies have been performed on FAPs. Although facial musculature-associated FAPs have been suggested to be neural crest-derived54, the embryonic origin of FAPs in other tissue compartments has not been addressed. Nonetheless, a population of embryonic FAP-like cells related to LPM-derived cells has been previously identified using the expression of Osr1 as a genetic marker55. Osr1+ mesenchymal cells in the developing mouse limb overlap partially (<60%) with Tcf4+ MCT fibroblasts and are located in slightly separate spatial domains (Osr1 more in the periphery of the limb and Tcf4 more centralized); interestingly, as development proceeds they are sequestered into separate muscle groups, for example: the biceps femoris muscle harbored almost exclusively Osr1+ cells whereas the flexor digitorum longus contained exclusively Tcf4+ MCT cells55. Lineage tracing of Osr1+ cells showed that this marker identifies only a limited subset of embryonic MCT cells that endure into adulthood as FAPs55. Even though the embryonic origin of FAPs has not 9 been well elucidated, the case can certainly be made that the biological functions, spatial localization, and cellular phenotype of Ly6a, Cd34 and Pdgfra triple-positive FAPs overlap significantly with that of Tcf4+ MCT fibroblasts and may therefore constitute a heterogenous population of interstitial cells with similar activities in muscle development, repair, and regeneration. 1.4 Origin and function of dermal fibroblast populations Similar to MCT cells, dermal fibroblasts (DFs) are known to be composed of heterogenous subpopulations56. Indeed, DFs from different regions of the body come from differing developmental sources. Those in the limb are derived from LPM; compared to those in the face and back, which are derived from the neural crest and somites respectively57,58. Furthermore, skin dermis is divided into upper (papillary), intermediate (reticular), and lower (hypodermis) layers each containing subsets of fibroblast populations with specialized functions56. The papillary dermis underlies the epithelium and harbors papillary fibroblasts (positive for Pdgfra, Prdm1, Dpp4, and Cd34 among other markers)59 which secrete ECM components that are prevalent in the upper dermis such as decorin, tenascin C, and collagen XVI60 and are required for hair follicle formation59. The reticular dermis is the middle skin layer and contains high concentrations of reticular fibroblasts. These fibroblasts express markers such as Ly6a, Dlk1, and Cldn1059,61 and are responsible for ECM production by secreting proteoglycans such as versican and fibrous proteins like collagen I, III, and V60. The reticular dermis also contains fibroblasts associated with hair follicles: follicular sheath cells and dermal papilla cells. Both of these populations express the transcription factor Sox262, and are involved in regenerating new hair follicles in a cyclic manner63. In this regard, hair follicles exhibit continuous cycles of regeneration which alternate between rest 10 (telogen) and growth (anagen) phases64. Follicular sheath cells have been suggested to contain a population of stem cells that repopulate not only the dermal sheath but also the dermal papilla at the onset of each hair cycle65. The dermal papilla is a cluster of mesenchymal cells found at the base of hair follicles and is required for hair growth. Indeed, laser ablation of the dermal papilla impairs initiation of hair regeneration66. The lower-most dermal layer, the hypodermis, contains a mixture of pre-adipocytes and mature adipocytes56. Although, this adipose-rich layer may be easy to overlook, it is functionally very significant for hair follicle regeneration. Pre-adipocyte cells within the hypodermis have been shown to be necessary and sufficient to drive follicular stem cell activation during hair follicle regeneration67. Superficial skin wounds elicits a regenerative response from tissue-resident DFs68. In dorsal dermis, a population of En1+ embryonic fibroblasts have been shown to be deposited during development and are the main cell type responsible for ECM secretion in wound healing, fibrosis, and tumor stroma formation69. Indeed, ablation of this cell lineage using a diphtheria toxin receptor (DTR)-expressing mouse line demonstrated that En1-lineage DFs are required for normal wound healing69. In most cases, human adult dermis heals via repair, a mechanism that results in scar tissue devoid of crucial epithelial appendages such as hair follicles, sweat glands, and sebaceous glands being deposited at the wound site70. However, neonatal mice have the capability of regenerating fully functional skin post-injury71, this regenerative potential has been attributed to a population of Lef1+ DF population that is extensive during embryonic development but recedes in adulthood72. Lef1+ fibroblasts have the ability to support de novo hair follicle generation during development and when Lef1 is expressed ectopically in adult DFs wounds exhibit less propensity to repair by the establishment of a pro-regenerative microenvironment72. 11 Lineage tracing experiments performed on embryonic and neonatal mouse back skin have revealed that DF populations emerge from common Pdgfra+, Dlk1+, and Lrig1+ multipotent precursors at E12.559. By E16.5, two populations of DFs have emerged from the common precursor: Pdgfra+, Prdm1+, Dlk1- papillary dermal fibroblast progenitors (PDFPs) and Pdgfra+, Prdm1-, Dlk1+ reticular/hypodermal dermal fibroblast progenitors (RDFPs). Expression of these genetic markers (such as Dlk1) is dynamic and, as development proceeds, can be observed in different cell populations at varying timepoints. Nevertheless, in postnatal skin (P2) the entire spectrum of DFs can be resolved and includes mature dermal papilla, sheath, reticular, and hypodermal fibroblasts59. Further studies are required to determine if this progenitor hierarchy applies to DFs from different body regions (and therefore different embryonic origins) such as those in the appendicular and facial dermis. 1.5 Origin and function of pericytes Blood vessels are composed of two main cell types: endothelial cells and mural cells73. Mural cells can be further subdivided into pericytes and smooth muscle cells. Pericytes are ubiquitous cells that are found wrapped around micro vessels (arterioles, capillaries and venules) and represent an often neglected area of vascular biology research74. Pericytes share a common basement membrane with endothelial cells and communicate with the underlying endothelium via gap junctions74. Similar to other mesenchymal cells such as DFs and FAPs, pericytes are also considered to represent a heterogenous cell population with varied ontogeny75. Although pericytes are found throughout most vascularized tissues, they are often present to varying degrees. For example, vasculature within neural tissues possess an extremely dense pericyte coverage with a 1:1 – 3:1 endothelial cell-to-pericyte ratio; on the opposite end, SkM has only a 100:1 ratio between 12 endothelial cells and pericytes75. Similarly, pericytes have a unique distribution pattern at the cellular level. They are more often found at capillary branchpoints76 and on the distal side of vessels adjacent to epithelial surfaces such as the choroid and the skin74. Several gene markers have been used to identify pericytes, examples include Acta277, Cspg4 (also known as NG2)78, Pdgfrb79, and Rgs580. However, these markers are dynamic and their expression may vary depending on developmental state, pathological conditions, and culturing protocols75. The variability in pericyte spatial localization, concentration (extent of blood vessel coverage), and gene markers suggests that pericytes may be composed of diverse subpopulations with specific functions. Indeed, pericytes have been implicated in several biological processes such as the development and maintenance of vascular permeability81,82, establishment and stabilization of endothelium basement membrane83, regulation of blood flow84, and as microenvironmental regulators during tissue repair85. Pericytes have been found to originate from multiple embryonic sources such as LPM, neural crest86–88, and paraxial mesoderm89. LPM-derived pericytes may share a common progenitor with endothelial cells, as it has been shown that Flk-1+ MPs within the LPM give rise to endothelial precursors90. Furthermore, Flk-1+ cells induced from embryonic stem cells can differentiate into endothelium, and mural cells such as vascular smooth muscle and pericytes in vivo91. Nevertheless, pericytes are recruited to nascent endothelial tubes during embryonic development and represent a hallmark of mature vasculature92. Pericytes from visceral organs such as the gut and liver have been suggested to derive from mesothelium93, a thin sheet-like monolayer of epithelial-like cells94. In this case, mesothelial cells undergo an epithelial to mesenchymal transition (EMT), delaminate, and migrate into adjacent organs75. Little is known about the 13 embryonic origin of pericytes populating appendicular tissues such as SkM, bone marrow, and dermis. Quail-chick transplantation experiments have demonstrated that vascular smooth muscle cells (VSMCs) and pericytes associated with limb blood vessels derive from the sclerotome and migrate into the limbs during embryonic development89. However, the ontogeny of mural cells in mammalian model organisms encompassing the spectrum of vascularized tissues in a functional limb remains unresolved. 1.6 Origin and function of bone marrow stromal cells Bone marrow is an essential and complex organ. Found protected between layers of cortical and/or trabecular bone, bone marrow is composed of multiple cell populations the most notorious of which, being hematopoietic stem cells (HSCs). HSCs generate all blood and immune cell types that are necessary to maintain hematological homeostasis and immunity95. HSCs are considered to be the adult stem cell par excellence, their fates and mechanisms have been very well characterized over the years96. However, an increasing focus has recently been aimed at stromal cells that support HSCs. These stromal cells create a microenvironment that allows and stimulates hematopoiesis: the hematopoietic niche. A niche is defined as a local tissue microenvironment that directly maintains and regulates a particular kind of stem cell or progenitor97. The bone marrow hematopoietic niche is localized in close proximity to sinusoidal endothelium98–100 and contains unique populations of bone marrow stromal cells (BMSCs) that are in part responsible for regulating HSC self-renewal, proliferation, and trafficking101. Several genetic models have been used to identify and classify BMSCs including, but not limited to: Cxcl12-GFP101, Nes-GFP102, Lepr-Cre103, Prrx1-Cre104, Sp7-Cre104, Mx1-Cre105, Grem1-CreERT106 and Ebf3-CreERT107. With the use of these marker genes, BMSCs have been characterized as heterogenous cell populations 14 with overlapping properties. For example, a subpopulation of BMSCs are leptin receptor-positive (Lepr+) and secrete factors (such as KITL, CXCL12, and PTN) that are required for HSC maintenance103,104,108,109; Nes+ stromal cells also secrete some of these factors albeit in a non-essential manner102–104. The advent of single cell RNA-sequencing (scRNA-seq) technology has brought Lepr+ stromal cells to the forefront of studies involving the bone marrow hematopoietic niche. Although relatively rare, comprising only < 0.3% of cells in the bone marrow110, several scRNA-seq experiments have corroborated that Lepr+ BMSCs express high levels of secreted factors essential for HSC maintenance such as KITL and CXCL12101; in comparison to Nes+ cells, endothelium, and Cspg4+ pericytes111–113. Interestingly, Lepr+ BMSCs are themselves heterogenous and comprise skeletal stem cells, osteogenic and adipogenic progenitors110,114,115. Recently, it has been shown that these subpopulations can be distinguished by spatial localization and expression of an osteogenic growth factor, osteolectin (Oln)116. Lepr+/Oln+ cells are short-lived osteoprogenitors that maintain a periarteriolar niche for lymphoid progenitors by secretion of factors such as KITL116. On the other hand, Lepr+/Oln- cells are adipogenic progenitors (with latent osteogenic potential) that create a perisinusoidal niche for HSCs116. Lepr+/Cxcl12+ BMSCs have also been shown to be responsible for the generation of cortical bone osteoblasts during the regenerative process and are critical in the maintenance of the hematopoietic niche115, further illustrating their importance within the adult bone marrow. In mice, a subset of BMSC progenitors invade the bone anlagen from a perichondral location in a “pericyte-like fashion”117 during early fetal stages of development alongside branching endothelium118,119. It can be reasoned that BMSCs may originate from one of the 15 mesoderm derivative populations such as the somatic LPM or paraxial mesoderm120. Experimental evidence for an LPM-origin of BMSCs exists in the shared expression of markers such as Acta2 and Twist1/2 between BMSCs and LPM121–124, as well as in the inherent trilineage differentiation potential (adipogenic, chondrogenic, and osteogenic) of LPM-derived MPs in the embryonic limb bud125. However, several studies have identified neural-crest derived cells within the bone marrow stroma126–128. Indeed, it’s been shown that a subset of BMSCs with in vitro multilineage differentiation potential (bone, cartilage, and fat) and self-renewing capability are derived from the neural crest during embryonic development129. However, the same study found cell populations in the adult bone marrow niche with identical BMSC properties that were not neural crest-derived. Furthermore, Nestin+ BMSCs have been suggested to be neural-crest derived and localize to the bone marrow by migrating along developing nerves130. Determining whether Nestin+ perivascular BMSCs share developmental origins and functional properties with perivascular Lepr-Cre+ cells has been challenging because there is currently no single marker that separates them75. To this day, the ontogeny of BMSCs remains an avenue in need of further investigation in the field of bone marrow biology. 1.7 Development and function of nerve-associated fibroblast populations The nervous system is a critical component of musculoskeletal physiology in both the developing and mature limb. Muscles rely on the peripheral nervous system (PNS) for chemical signals that are relayed from the central nervous system (CNS) via motor axons and neuromuscular junctions that ultimately result in muscular contraction and overall locomotion of the skeletal framework131. Within a limb, the nervous system extends axons that project through the appendage 16 and into the most distal regions to provide surrounding tissues with afferent (toward the CNS) and efferent (away from the CNS) signals. Neuron axons cluster together to form nerve bundles (fascicles) composed of not only sensory and motor axons, but also supporting neural, vascular132,133, immune133,134, and connective tissue cells135,136. The connective tissue components of nerves are arranged in three layers that cover and compartmentalize the nervous tissue137. The epineurium is the outermost layer and is composed of a dense, irregular, collagenous coat that completely surrounds the nerve. Each fascicle is then surrounded by the perineurium, a sleeve formed by layers of flattened epithelium-like cells. Due to the tight junctions that characterize these cells, the perineurium serves as a barrier to the passage of most macromolecules, as well as, protecting the nerve fibers from damage138. The innermost layer of connective tissue is termed the endoneurium and it envelops individual nerve axons in direct contact with the basal lamina of Schwann cells. This layer is an important factor in regulating the microenvironment of the nerve fiber as it is sandwiched between glial cells and the perineurium137. Within all three of these nerve-associated connective tissue components (epineurium, perineurium, and endoneurium), Pdgfra+ mesenchymal cells are present in varying quantities132,135,139,140. Although the embryonic origin and development of the nervous tissue component of the PNS has been extensively studied and well-characterized141–144, little is known about the ontogeny of its supporting stromal cell populations. A study carried out by Joseph and colleagues135 identified endoneurial fibroblasts as neural crest-derived. They further demonstrated that a multipotent neural crest progenitor gives rise to both endoneurial fibroblasts and glial progenitors in the developing mouse sciatic nerve and that perineurial cells, pericytes, and endothelium were not neural crest-derived and likely generated from mesodermal derivatives. The embryonic 17 origin(s) of epineurial and perineurial fibroblast populations has not been directly addressed due to the lack of a cell type specific marker, although several markers have been proposed, none are highly specific. THY1 (Cd90) has been traditionally used as a selective marker of nerve-associated fibroblasts145,146, but it is lost in all three connective tissue compartments post-natally139 and is also expressed in neuronal axons132,135. More recent histological characterization of endoneurial fibroblasts identified Cd34, Cspg4, and P4hb as genetic markers for this population in rat sciatic nerves147. Using transgenic mouse models and immunofluorescence microscopy, fibroblasts in epi-, peri-, and endo-neurium have been shown to express Pdgfra, and Cd34132. Single-cell sequencing technologies have allowed for a more thorough transcriptomic profiling of mesenchymal cells within nerve connective tissues132,136. However, further research is needed to create lineage-tracing tools that will take advantage of the gene markers unraveled via scRNA-seq to determine the ontogeny of epineurial and perineurial mesenchymal cells. Peripheral nerves have been proven to be indispensable for tissue regeneration in organisms such as amphibians148,149. Similarly, the importance of nerves in mammalian regenerative models such as skin wounds and digit tip amputations, is paramount as denervated wounds do not undergo complete regeneration150–152. The importance of nerves in digit tip regeneration has been attributed, in part, to the dedifferentiation and subsequent migration of Schwann cell precursors (SCPs) from within the bisected nerve towards the injury site152,153. Once at the blastema, SCPs secrete factors such as PDGFA and Oncostatin-M that promote expansion of MPs within this area152. Moreover, a recent study in digit tip regeneration has brought attention to the fibroblast populations present within the nerve connective tissue132. Long thought of as a simple scaffolding and insulator for axon bundles, nerve sheath mesenchymal cells have proven 18 to be a dynamic cell population post-injury, forming a cooperative network with SCPs to restore nerve structure and support an optimal microenvironment for axonal outgrowth154,155. More interestingly, nerve-associated mesenchymal cells also contribute directly to bone formation and dermal repair during digit tip regeneration132. This indicated that nerves may harbor subsets of quiescent MPs amongst their surrounding connective tissue. 1.8 Hypermethylated in Cancer-1 (Hic1) HIC1 is a transcriptional repressor that has been found to be epigenetically silenced via DNA hypermethylation in several types of cancers such as breast, prostate, and lung156–159. Hic1 was initially identified as a candidate tumor suppressor gene160 and was later discovered to be crucial for proper embryonic development to occur161. Homozygous disruption of Hic1 in mice results in impaired development and embryonic lethality usually within E13.5 to E16.5162. Mice heterozygous for Hic1 are viable and develop normally; however, they have an increased susceptibility to age-related tumors as a result of hypermethylation of the remaining wildtype Hic1 allele162. Indeed, ectopic expression of HIC1 has been demonstrated to induce growth arrest in cancer cell lines160. Other studies have linked HIC1 to the G1/S cell cycle transition via regulation of cell cycle genes encoding CCND1 and CDKN1A163–165. Hic1 expression has been characterized in the developing mouse embryo. During early embryogenesis, Hic1 is expressed in sclerotomal, lateral body wall, limb bud and cranio-facial mesenchymes166. During fetal development Hic1 was also observed in the perichondral and perineural mesenchyme of the limbs166. Murine and human HIC1 proteins share highly similar amino acid sequences and contain a Broad complex, Tramtrack and Bric à brac/POxviruses and Zinc finger (BTB/POZ) domain 19 which facilitates homodimerization and transcriptional repression, along with an internal C-terminal binding protein (CtBP) binding region that participates in gene repression via histone deacetylation166–168. At the C-terminal end, HIC1 possesses five Krüppel-like C2H2 zinc fingers which bind to a specific DNA sequence, termed “HiRE” (HIC1 responsive element)169. HIC1 gene targets include, but are not limited to: Sirt1 (which codes for a histone deacetylase)170, Fgfbp1 (Fibroblast Growth Factor-Binding Protein 1)171 a protein involved in angiogenesis pathways, and Atoh1 (Atonal Homologue 1) a transcription factor required for the proper development of various neural progenitor cells172. Due to the presence of a protein-binding motif, HIC1 could also bind to other transcription factors and indirectly repress transcription of genes that lack a HiRE sequence. Evidence of this exists with respect to the interaction of HIC1 with TCF4 resulting in down-regulation of the Wnt signaling pathway173. 1.9 Mesenchymal progenitors or mesenchymal stem cells? In the early 1970s, Friedenstein and colleagues identified a population of plastic-adhering, colony-forming fibroblasts present in spleen and bone marrow after removal of hematopoietic and lymphoid cells174–176. Due to their morphology, colony-forming and collagen-synthesizing abilities, they were originally termed colony-forming unit fibroblasts (CFUf) and were found to be a proliferative yet rare mesenchymal cell population found in lymphoid tissues177. In culture, bone marrow CFUf-derived colonies were also observed to spontaneously mineralize and generate adipocytes or chondrocyte nodules (if seeded at very high densities)178. This trilineage differentiation potential was later confirmed in vivo by heterotopic transplantation of stromal cells isolated from mouse bone marrow179,180. The capacity of CFUf colonies to differentiate into several 20 mesenchymal lineages such as cartilage, bone, and fat suggested that they may be derived from mesenchymal stem cells (MSCs)181,182 . However, in reference to adult stem cells, the term “MSC” was not widely adopted until several years later when stromal cells with similar properties were isolated from human bone marrow183. It is important to mention that in the 1980s through to the 1990s, the scientific consensus was that HSCs were the only stem cells present in the adult body184. In this way, the notion of an MSC was forever intertwined with non-hematopoietic adult stromal cells found in the bone marrow. In the years following the discovery of multipotent progenitors in human bone marrow, research on MSCs increased exponentially creating a renaissance period in stem cell biology. MSC-like cells were identified at perivascular locations first in bone marrow185 and later on in virtually all other adult tissues such as skeletal muscle, white adipose tissue, thymus, lung, liver, and spleen186–188. Around this time, controversies regarding whether MSCs were in fact ubiquitous or bona fide stem cells began to impact the field. The minimum criteria for a “stem cell” designation based in part on our understanding of HSCs, involves experimental proof of extensive self-renewal capability and uni- or multi-lineage differentiation potential. So far, only bone marrow derived MSCs have been rigorously tested and proven to meet these criteria102,118,185. However, to more properly reflect their location and functionality, some groups have proposed that subpopulations of bone marrow-derived MSCs be termed skeletal stem cells (SSCs), as this more accurately reflects their self-renewal capabilities and lineage potential189. Until now, the relationship between perivascular MSC-like cells from bone marrow and other tissues remains to be further defined, but early studies indicate that various organs contain distinct populations of perivascular cells that differ in lineage potential and transcriptomic features190. In addition, the 21 capacity for self-renewal of these extra-medullary MSC-like cells has not been fully determined. The summation of controversies reflected in contradictory findings regarding MSCs ontogeny, lineage restriction, physiological functions, and therapeutic potential led researchers to propose a substitution of the MSC term in favor of a more rigorous classification system for an evidently heterogeneous cell population184,191,192. Pending consensus on the nature and relationship between various tissue-resident mesenchymal cells, and for the sake of clarity, this body of work will refer to stromal cells with CFUf-forming activity and mesenchymal lineage potential as mesenchymal progenitors or MPs. 1.10 MPs in skeletal muscle repair and regeneration Tissue-resident MPs can be found to varying extents in all tissues and play vital roles in tissue homeostasis, renewal and regeneration193,194. In adult appendicular SkM, heterogeneity within the MP population has been described, with a minimum of three distinct populations that exhibit restricted lineage potential45,195. After injury, a subset of these MPs, termed FAPs and identified as Pdgfra+, Ly6a+ (also known as Sca1), Cd34+, generate a transient population that orchestrates multiple facets of the regenerative program. These include immune cell infiltration, vascularization, trophic factor secretion, and both extracellular matrix (ECM) and basement membrane production and remodeling. Once an injury has resolved, “excess” FAPs are cleared via apoptosis and tissue integrity is restored48,49. Although Pax7+ muscle satellite stem cells are responsible for de novo muscle synthesis, MPs and their derivatives are crucial components of the regenerative program by secreting promyogenic cytokines and ECM that will orient and organize the developing myotubes39–41. Indeed, it has been shown that genetic ablation of FAPs results in 22 SkM atrophy and impaired regenerative potential42. Based on transcriptomic profiles carried out under homeostatic conditions, FAPs can be subdivided into two major sub-groups, FAP1 and 2. The FAP1 cells express genes associated with the ECM, such as Col4a1 and a2, various Col6 transcripts, Col15a1 and other ECM genes such as Bgn, Mgp and Podn. In contrast, FAP2 cells are enriched for transcripts involved in signaling and trophic factor production, such as Dact2, Sfrp4, Wnt2, Sema3c, Pi16 and Tgfrbr2. The FAP1 and 2 subsets correspond to the recently described two universal fibroblast profiles, Dpt+ Col15a1+ and Dpt+ Pi16+, described in mouse and human45,196. Several markers have been utilized to identify MPs, most have been used specifically in a bone marrow context such as Nestin102,197, Lepr110,198, and Grem1106,199. Regarding extra-medullary MPs, markers include Gli1200, Pdgfra and Sca1129,201,202. However, the efficacy of these genes as MP markers varies depending on the tissue of interest. In the brain for example, Pdgfra is also expressed by oligodendrocytes and this is a similar case for Gli1. Recently, Hic1 has come to the forefront as a marker for quiescent MPs in several tissues such as SkM45, skin dermis203, and heart204. In adult SkM, Hic1+ MPs have been thoroughly characterized. Interestingly, in contrast to Hic1+ fibroblasts, other specialized Hic1+ MP subsets produce enduring progeny that regenerate fibrous connective tissues which link muscle to bone, such as tenocytes and myotenocytes in addition to epimysium45. Hic1 also identifies laminin-encased Rgs5+, Pdgfrb+, Kcnj8+ and Abcc9+ pericytes. Indicative of their progenitor status, all of the described populations exhibit open chromatin regions (OCRs) across numerous cell cycle loci and have been shown to enter the cell cycle following injury-induced activation45. 23 1.11 Key technological advances Methods in cell and developmental biology have evolved over time and are now capable of interrogating biological systems at the single cell level in a quantitative and qualitative manner. The use of emerging techniques in tandem with more traditional methods provides a holistic approach to the study of tissue and organ development. Some of the combined strategies used in this thesis are highlighted here. 1.11.1 Embryonic and postnatal lineage tracing of Hic1CreERT2; R26tdTomato mice Determining the fate of progenitor cell populations in an embryo is one of the focal points of studies in developmental biology. From where does a particular adult cell type emerge? Likewise, to what cell type(s) does a particular embryonic progenitor give rise? These questions have been historically addressed by embryologists utilizing an array of techniques to achieve the same goal: lineage tracing of progenitor cells through embryonic development. Initially, fate mapping studies were performed in model organisms such as nematodes and fish, where cells could be observed directly (with the aid of a light microscope)205,206. More complex organisms required the use of dyes207–209, viral transfection of genetic markers210,211, or inter-species transplantation212, which would allow for the identification of single cells and their progeny. Although some of the aforementioned lineage tracing techniques are applicable to mammalian model organisms, the efficacy, complexity, and reproducibility are far exceeded by modern methods of genetic recombination in mice. 24 Since the advent of mammalian genetic manipulation, transgenic mouse lines have enabled researchers to query gene function, cellular fate, and associated molecular mechanisms at an unprecedented scale. Conditional reporter mouse models quickly became crucial components of the developmental biologist’s toolkit to investigate the fate of embryonic progenitor cells. Indeed, lineage tracing studies have heavily relied on precise knock-in mouse models that allowed for spatial control of reporter gene expression via Cre/LoxP recombination system213–216. In these models, the expression of Cre recombinase is controlled by a cell-specific promoter. This protein will, in turn, induce the stable expression of a reporter gene(s) via genetic recombination of “floxed” sites217,218. In this way, progenitor populations can be strategically labelled to identify their differentiated progeny at a later time point by identifying reporter gene expression. In the late 1990’s an extra dimension was added to Cre/Lox-mediated lineage tracing by the fusion of a modified estrogen ligand-binding domain with enhanced selectively for the synthetic estrogen tamoxifen to Cre recombinase (CreER)219,220. This advancement allowed for the spatial and temporal control of reporter gene expression by the requirement of an estrogen analogue metabolite (4-hydroxytamoxifen, 4-OHT) for nuclear translocation of the CreER protein and DNA recombination to take place. 4-OHT is not a physiological metabolite; therefore, its precursor (tamoxifen, TAM) must be administered exogenously. The end result being that Cre-mediated recombination can be induced through TAM administration, specifically in target cells expressing the CreER construct. This inducible Cre system provides the major benefit in being able to carry out pulse-chase experiments in which animals are briefly exposed to TAM at any age (even in utero) and the fate of labeled cells can be assessed several days, months or years later221. 25 The Hic1CreERT2; R26tdTomato mouse line has allowed several research groups to perform lineage tracing of quiescent Hic1+ MPs before and, in some cases, after injury in various tissues such as SkM45, cardiac muscle204, skin203, bone222, brain, and testes223. The bulk of these experiments have focused on Hic1+ MP fate and function in adult tissue homeostasis and repair, and neonatal organ maturation. Within this thesis, I describe for the first time the lineage tracing of embryonic Hic1+ MPs using the Hic1CreERT2; R26tdTomato mouse line. These experiments have allowed the fate mapping of this progenitor population and its derivatives throughout embryonic and postnatal development. Through trial and error, optimal TAM-administration conditions were developed to enable labelling of Hic1+ cells as soon as Hic1 transcripts first became detectable. These conditions included TAM dose, embryonic timepoint at which to administer TAM, TAM delivery method, interval between TAM dosage and litter collection, among others. The end result being the ability to label the earliest Hic1-expressing cells in an embryo and identify their progeny; not only at later stages of embryonic and fetal development, but also postnatally and into adulthood. Due to TAM’s deleterious hormonal effects on pregnant mice, postnatal lineage tracing of embryonic Hic1+ MPs required cross-fostering of Hic1CreERT2; R26tdTomato litters with outbred CD-1 mothers. These experiments also required optimization of fostering techniques to achieve reliable outcomes. 1.11.2 Combined use of micromass, limb culture explants, and wholemount immunofluorescence The vertebrate appendicular skeleton forms through endochondral ossification, a process in which cartilage templates prefigure bones224. These cartilages begin as condensations of 26 mesenchymal cells which increase in density and ultimately chondrocyte differentiation ensues within the cores of these aggregates. Chondrocytes are responsible for secreting components of a typical cartilaginous ECM such as type II collagen and several proteoglycans, including aggrecan5. Once formed, the cartilage template becomes invaded by a mixture of vascular and perichondral cells that establish ossification centers. In these areas, mineralized bone will gradually replace cartilage until a mature bone and its associated components (bone marrow, cortical and trabecular bone, growth plates, etc.) is realized225,226. It is evident that endochondral ossification constitutes a crucial component of limb morphogenesis during fetal development. However, the study of cartilage biology is relevant for an array of postnatal applications such as bone homeostasis and repair, degenerative diseases such as osteoarthritis, and certain cancers including chondrosarcomas. The establishment of in vitro models of cartilage development allowed for the advancement of understanding the molecular and cellular mechanisms involved in chondrocyte specification, differentiation, and hypertrophy227–230. Primary limb mesenchyme (PLM) culture, also known as micromass cultures, are an example of such models. Micromass cultures consist of cells derived from dissociated rodent or avian limb buds cultured at extremely high cellular densities. After 2-3 days, undifferentiated limb mesenchyme within the spot culture will spontaneously condense and form cartilage231,232, the cartilage nodules will also be surrounded by fibroblastic cells and myoblasts233. When this method was developed in the late 1970’s, these cultures provided a technical advantage that allowed a 20- to 25-fold increase in the number of biological replicates attainable from the same amount of isolated limb bud cells234. Furthermore, micromass cultures recapitulate in vivo chondrogenesis with high fidelity and can be easily manipulated to interrogate physiological responses to small molecules 27 and other factors235. Indeed, since its establishment as an in vitro model of chondrogenesis, several groups have used PLM cultures to unravel the cellular and molecular intricacies of cartilage development236–240. Although a technical advancement in its own right, micromass cultures possess implicit limitations that commonly plague 2D culture systems241. The developing limb exhibits molecular and genetic gradients that provide cells with positional information242, these positional cues are difficult to convey in 2D cultures, as are the complexities associated with other cell types in the developing limb that shape morphogenesis. Ex vivo limb culture systems are a reliable method to maintain cellular positioning systems in place and limb morphogenesis in three dimensions. In contrast to culturing dissociated tissues, limb bud cultures are not only a good model for studying cell differentiation and dynamics but also provide valuable information on organ morphogenesis in three dimensions243. Ex vivo culture systems are robust, easy to manipulate, and more ethically acceptable than in vivo systems244. Although culturing limb buds ex vivo leads to some skeletal abnormalities, cartilage anlagen of the major bone elements appear in a clearly recognizable fashion. In the context of musculoskeletal malformations or diseases, limb bud cultures serve as a robust experimental model to explore the molecular basis of such deficiencies. In this manner, the system is highly amenable to genetic and chemical manipulation such that gene and/or function and fate can be efficiently interrogated. The use of appropriate experimental perturbation systems in conjunction with limb cultures routinely give rise to 90% of skeletal abnormalities seen in vivo245. To visualize limbs post-culture, samples may be fixed, stained with fluorescent antibodies for proteins of interest, cleared using any of the available protocols for tissue clearing246, imaged and rendered in 3D with a confocal or light sheet microscope and its associated 28 software. In this way, mammalian limb musculoskeletogenesis can be studied in three dimensions not only in ex vivo cultures but also during “normal” embryonic development247. The work described within this thesis effectively utilizes the three techniques described above (micromass cultures, ex vivo forelimb cultures, and wholemount tissue clearing and imaging) to provide a comprehensive picture of the cellular fate and function of Hic1+ or Hic1-lineage MPs in the developing mouse forelimb. Although several more cutting-edge techniques were used to complement those describe above (for example: scRNA-seq, single cell assay for transposase-accessible chromatin (scATAC-seq), and µCT imaging). I posit that for purposes of visualization and manipulation of MPs during forelimb musculoskeletogenesis, the aforementioned cell culture and histological methods that are uniquely combined in this work provide ample context to delineate spatial distribution, molecular mechanisms and genetic programs involved in MP fate and function during limb development. Although there is certainly more work to be done, especially regarding the molecular mechanisms that govern Hic1+ MP biology, the techniques described herein provide an experimental framework that has proven to be useful in the study of embryonic MPs and could therefore be translated into other organs and systems. 1.12 Objectives of this work MPs figure prominently in adult physiology and pathology, and Hic1+ MPs regulate multiple facets of the regenerative program across multiple cell types45,203,204. However, our understanding of their embryonic origin and subsequent fates is unclear. The origin and fate of 29 other tissue-resident stem and progenitor populations such as satellite cells within striated muscle has been well defined due to the specificity of the marker Pax7/PAX7. In contrast, due to the lack of effective markers of putative mesenchymal subsets in the early limb, it has been more challenging to resolve the origin, fate and cellular hierarchies within this cellular compartment. Here, I introduce Hic1 as a marker to study MP ontogeny and fate during embryonic development and cellular function postnatally. This concept can be divided into four main aims: 1. To investigate the embryonic origin of Hic1+ MPs in the developing forelimb. Specifically, I wanted to determine if Hic1+ MPs originate from limb bud LPM or another embryonic compartment. 2. To elucidate the cellular fates of Hic1+ MPs in the developing forelimb. I aim to corroborate, within an embryological context, the cell lineages that Hic1+ MPs contribute to during adult tissue repair and regeneration (fibroblasts, myotenocytes, tenocytes, and chondrocytes). 3. To characterize the cellular fates of embryonic Hic1+ MPs in postnatal skeletal muscle. More critically, this aim will determine whether embryonic Hic1+ MPs provide a postnatal reservoir of MPs to support adult tissue homeostasis and restoration. 4. To determine the functional consequences of Hic1 perturbation in appendicular skeletogenesis. As previously mentioned, Hic1+ mesenchyme surrounds the appendicular cartilage anlagen during embryonic development. This aim will address the impact of Hic1 deletion on skeletal development. 30 Chapter 2: Methods 2.1 Mice The generation of Hic1floxed (Hic1f) and Hic1CreERT2 mice have been previously described45. Both modified Hic1 alleles were conceptualized by the Underhill lab at the University of British Columbia and generated by GenOway Inc. under contract. The Hic1f mice were produced as follows: gene targeting methodology was used to insert a vector containing LoxP sites flanking the major coding exon 2 of Hic1 and an IRES-NLS-LacZ cassette distal to the 3’ LoxP site and downstream of exon 2. Upon Cre-mediated recombination, the major coding exon is excised and cells in which the Hic1 promoter is active are identifiable by LacZ expression. In this way, the Hic1f mouse can function as an MP reporter and a Hic1 knockout model by staining with chromogenic substrates such as 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-gal) and breeding with Cre-expressing mice, respectively (Figure 2.1). Most loss-of-function experiments described in this thesis involved the utilization of mice containing a Hic1 null allele (Hic1nLacZ). To generate this allele, Hic1f/f were bred with UBC-CreERT2 mice to generate UBC-CreERT2; Hic1f/+ mice. TAM was administered at 8-weeks of age and recombined animals were interbred into a C57BL/6 background to generate Hic1nLacZ/+ mice. 31 Figure 2.1 Schematic representation of Hic1 expression patterns in Hic1nLacZ embryos. Blue highlights LacZ+ areas. The Hic1CreERT2 mice required similar gene targeting methodology which involved the knock-in of an IRES-CreERT2 cassette into the 3’ untranslated region of the Hic1 gene. Other experimental mouse lines used in this work include: B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J (Jax stock number 007914; herein referred to as RosaTom), C57BL/6J (Jax stock number 000664), ScxGFP248, B6.Cg-Tg(Prrx1-Cre)1Cjt/J (Jax stock number 005584; herein referred to as Prrx1-Cre), Crl:CD1(ICR) (Charles River strain code 022), B6.129S4-Pdgfratm11(EGFP)Sor/J (Jax stock number 007669), and Ptch1tm1Mps/J (Jax stock number 003081). For lineage tracing experiments, Hic1CreERT2 mice were interbred with Rosa26tdTomato mice to generate Hic1CreERT2/CreERT2; Rosa26tdTomato/tdTomato progeny (Figure 2.2). This genotype was found to be ideal for fate determination experiments and, for simplicity, will be referred to as “Hic1CreERT2; R26tdTomato” throughout this body of work. To induce CreERT2 nuclear translocation, timed-pregnant females were administered 7.5 mg of TAM in sunflower oil by oral gavage. Unless otherwise stated, Cre-32 mediated recombination was induced the night of embryonic (E) day 10.0 for all experiments (Figure 2.3). Figure 2.2 Hic1 alleles and derived mouse models. Schematic representations of alleles used to generate the relevant mouse models for this work. 33 Figure 2.3 Schematic representation of tdTomato expression patterns in Hic1CreERT2/R26tdtomato embryos recombined at E10. Red areas highlight tdTomato+ regions. For lineage tracing Hic1CreERT2; R26tdTomato cell progeny into adulthood, timed-pregnant females were administered a single dose of 7.5 mg TAM along with 3.75 mg progesterone (Sigma P3972-5G) in sunflower oil via oral gavage. CD1 timed pregnancies were set up in parallel, CD1 females reliably give birth the night of E20. Therefore, the morning of E20.5, the Hic1CreERT2; R26tdTomato females were sacrificed and pups were collected via caesarean section and cross-fostered with a wildtype CD-1 litter. For experiments involving deletion of Hic1 in embryonic limb mesenchymal cells, Hic1f/f mice were interbred with heterozygous Prrx1-Cre mice to generate Prrx1-Cre; Hic1f/f mice. The Prrx1-Cre allele was always maintained in heterozygous male breeders which were mated with homozygous Hic1f/f females to generate Prrx1-Cre; Hic1f/+ progeny. Afterwards, the generation of Prrx1-Cre; Hic1f/f experimental animals was achieved by interbreeding Prrx1-Cre; Hic1f/+ males with Hic1f/f females. 34 For visualization of tendon and tendon progenitors during fate mapping experiments, Hic1CreERT2; R26tdTomato mice were interbred with ScxGFP homozygous mice to generate Hic1CreERT2; R26tdTomato; ScxGFP mice. To improve timed mating success rates and embryo quantities, these mice were outbred to a CD-1 background for 5 generations. Mice were housed under standard conditions (12 hr light/dark cycle) and provided food and water ad libitum. Animals were maintained and experimental protocols were conducted in accordance with approved and ethical treatment standards of the Animal Care Committee at the University of British Columbia. 2.2 Muscle Injury Muscle injury of the tibialis anterior (TA) muscles was induced by intramuscular injection of 0.2 µg of Notexin (NTX) snake venom extract (Latoxan) in 20 µL phosphate buffered saline (PBS) (10 µg/mL)40. Muscles were harvested four or fourteen days post-injury and processed for immunofluorescence staining as described below. Contralateral limbs were used as uninjured controls. Quantification of linage traced tdTomato+ cell expansion after NTX-induced muscle injury was carried out as previously described45. Six representative images were taken from three injury timepoints (day 0, day 3, and day 14) and two TAs were harvested per timepoint. tdTomato+ cells were enumerated using the ImageJ count function ensuring localization with DAPI to identify bona fide cells. Results were tabulated and plotted as the mean ± standard deviation. 35 2.3 Immunofluorescence staining Embryos were collected at indicated time points and fixed in 2% paraformaldehyde (PFA) for 72 hours. Following fixation, samples were cryoprotected by processing through increasing concentrations of sucrose up to 50%. Samples were subsequently embedded in Optimal Cutting Temperature (OCT – Tissue Tek 4583) embedding medium by use of an isopentane cold bath within liquid nitrogen. Histological cryosections were prepared at a thickness of 10-20µm on Superfrost Plus slides (VWR 48311-703) and stored at -20oC until further use. To begin the process of immunofluorescence staining, slides were retrieved from -20oC freezer and thawed on the surface of a hotplate (37oC) for 20 minutes or 1 minute per micron thickness. Once thawed, OCT medium was removed by washing in room temperature PBS for 30 minutes, changing into fresh solution every 10 minutes. To reduce endogenous tissue autofluorescence, slides were submerged in a freshly made solution containing sodium borohydride (NaBH4, 10 mg/ml in PBS) for 45 minutes. To remove excess NaBH4, slides were washed twice in PBS for 5 minutes. If the antigen being detected was localized to the nucleus, a cell permeabilization step consisting of treatment with a saponin-based reagent (Invitrogen C10636) at a 1:20 ratio in PBS for 20 minutes was necessary. Similarly, if the primary antibody was raised in mouse, a mouse blocking step was done in which slides were treated with mouse blocking reagent (Vector laboratories MKB-2213) for one hour. Following PBS washes, slides were incubated with blocking solution [2.5% goat serum (Gemini 100-190); 2.5% BSA (Sigma A7030) in PBS] for one hour. Primary antibodies were diluted in blocking solution at appropriate concentrations and incubated on slides overnight within a humidity chamber at 4oC. Secondary antibodies were diluted (1:500) similarly and incubated for 45 minutes at room temperature. All slides were counterstained with DAPI (600 nM) and 36 coverslipped with Aqua Polymount (Polysciences 18606) mounting media. This aqueous mounting medium solidifies overnight and therefore samples were analyzed within 24 hours or stored in a cool, dry location. Certain embryonic timepoints (mainly E10.5-E12.5) contained high quantities of erythrocyte-derived autofluorescence on Hic1CreERT2; R26tdTomato tissue sections. To minimize this autofluorescence, Vector TrueVIEW autofluorescence quenching kit (Vector laboratories SP-8400) was used as per the manufacturer’s instructions. Unless otherwise stated, all immunofluorescence experiments were replicated on a minimum of 3 embryos from different litters. Staining of embryonic cryo-embedded samples with a-PAX3 was technically challenging and required a compromise between maintaining tissue integrity and maximizing fluorescence signal. Through multiple rounds of trial and error, it was determined that antigen retrieval in 10 mM citrate buffer (pH 6.0) at 60 0C overnight (>12 hours) allowed for epitope exposure with minimal impact on the quality of the tissue section. After antigen retrieval, slides were rinse 3x in PBS and the immunofluorescence staining procedure was then followed as described above with one exception, NaBH4 quenching was not performed on slides that were exposed to a-PAX3 as this compromises signal detection. When combining a-PAX3 staining with the a-HIC1 tyramide amplification protocol a four-day protocol is required to achieve signal detection. On day 1, antigen retrieval for the PAX3 epitope was carried out overnight, fortunately this does not affect binding of the a-HIC1 antibody. On Day 2, the immunofluorescence protocol (ensure NaBH4 step is omitted) was initiated and incubation with a-PAX3 was performed overnight at 4 0C. Incubation in primary antibody overnight is paramount to achieve a strong signal. Day 3 involves completion 37 of the a-PAX3 staining with an appropriate secondary antibody followed by 3x PBS washes and then incubating in a-HIC1 antibody overnight. Again, overnight incubation was required for to achieve a detectable signal. On Day 4, the final step involves completing the tyramide signal amplification protocol as described in section 2.1.4. 2.4 Generation and detection of anti-HIC1 antibody To achieve direct detection of HIC1 protein in mouse tissues, the Underhill lab generated a rabbit polyclonal antibody to HIC1 based on 25 amino acids located at the carboxyl-terminus of HIC1 using ProSci Inc. (Poway, CA). This region exhibits 96% amino acid identity between mouse and human. A peptide (C25; AEVLSQGAHLAAGPDSRTIDRF) was conjugated to keyhole limpet hemocyanin and used to immunize 2 rabbits in complete Freund’s adjuvant. Collected serum was subsequently affinity purified on a C25 peptide-containing affinity column. For detection of HIC1 with anti-HIC1 a tyramide signal amplification kit was used (Thermo Fisher T20926) with the following modifications: To quench endogenous peroxidase activity, slides were treated with stabilized 3% hydrogen peroxide solution (Thermo Fisher B40926) for 30 minutes followed by two PBS rinses. Samples were then permeabilized using 0.5% Triton X-100 (Millipore 9410-OP) for 20 minutes and blocked for one hour in blocking solution. Anti-HIC1 was diluted 1:10,000 in blocking solution and incubated on slides overnight at 4o C. The following day, primary antibody was removed and slides were washed 3 X 10 minutes in room temperature PBS. Secondary anti-rabbit antibody (provided in the kit) was added for 1 hour at room temperature. Following 3 X 10 minute PBS washes, slides were treated with tyramide 38 working solution for 10 minutes followed by stop-solution and 3 quick PBS rinses. After this step, slides were counterstained with DAPI, and cover slipped as described in the previous section. For simultaneous immunodetection of HIC1 and other antigens of interest (i.e., PAX7), a three day protocol was necessary in which autofluorescence quenching, permeabilization, and antigen blocking steps were done on day 1, followed by incubation of antibodies against the first target protein overnight. Day 2 consisted of secondary antibody incubation, followed by 3 PBS washes and application of a-HIC1 overnight. Lastly, on day 3 the tyramide signal amplification and DAPI counterstaining steps were carried out as described above. 2.5 Wholemount immunofluorescence staining and imaging Depending on the end goal of each experiment, two wholemount clearing techniques were employed. For young embryos (< E14.5), a modified version of the CUBIC aqueous tissue clearing protocol249 was used. Post-dissection, embryos were fixed in 2% PFA for 72 hours, followed by 3 X 10 minute washed with PBS. At this point, samples were immersed in CUBIC solution overnight at room temperature or long as necessary to reach the desired transparency. For imaging or storing, samples were transferred into ScaleA2 solution250 and maintained at 4 oC. The main benefit of utilizing aqueous clearing reagents such as CUBIC and ScaleA2 is the maintenance of fluorescence activity of native fluorescent proteins (i.e., GFP or tdTomato) that are then amenable to imaging. Wholemount samples were imaged using a Nikon A1R inverted confocal microscope in glass-bottom polystyrene dishes (MatTek Corporation, P35G-1.5-14-C). 39 For more mature fetal samples (E14.5+), only forelimbs were collected and processed in the same way as described above for younger embryos. However, to achieve optimal transparency, limbs were subjected to an organic Benzoic Acid Benzyl Benzoate (BABB)-based protocol. The Kardon laboratory has developed an optimized version of this protocol247 (http://www.kardonlab.org/protocols; retrieved January 3rd, 2022). All organic-based clearing and wholemount immunofluorescence staining experiments visualized were carried out in accordance with the Kardon lab protocol, with the exception of the post-fix step which was found to be unnecessary for our samples. Imaging of these BABB-cleared limbs was performed on a Nikon A1R inverted confocal microscope. However, due to the nature of the clearing reagent, samples were imaged within custom-built metal containers with a #1.5 coverslip. After imaging, cleared samples can be stored in a glass vial containing BABB at 4oC for weeks or months. 2.6 Flow cytometry and fluorescence-activated cell sorting (FACS) methodology To isolate cells for flow cytometric analyses or FACS, a timed-pregnant dam was sacrificed at the developmental timepoint of interest and embryos were quickly harvested into ice-cold PBS. Freshly dissected E11.5 and E12.5 Hic1CreERT2; R26tdTomato or Hic1CreERT2; R26tdTomato; ScxGFP embryo forelimbs were digested in a Dispase enzyme-based solution [Dispase II 10 units/ml (Roche 4942078001); 10% fetal bovine serum (FBS) in Puck’s saline A]. Two-three forelimb pairs were pooled into 3 mL reactions. For E13.5, E14.5 and E16.5 forelimbs, the dissociation cocktail consisted of Collagenase A (Roche, 10103586001; 1.5mg/ml), Pronase (Roche, 10165921001; 0.5mg/ml), DNase (Roche, 11284932001; 100µg) in Dulbecco’s Modified Eagle Medium (DMEM; Gibco, 11960-044). Single forelimb pairs were dissociated in 2 ml reactions. In both 40 cases, enzymatic digestion was carried out at 37o C with gentle rotation for 1.5 hours with vortexing every 30 minutes. Digested material was then triturated by pipetting and larger tissue debris removed by filtration through a 40 µm cell strainer (Falcon, 352350). The resultant cell suspension was washed with FACS buffer (2mM EDTA, 2% FBS in PBS) and centrifuged at 500 x g for 5 minutes. To further enrich for MPs, an antibody panel in combination with tdTomato fluorescence, was designed to label and exclude lineage-positive cells [anti-CD45-647 (UBC Ablab 67-0047-01, 1:400), anti-CD117-APC (eBiosciences 17-1172-82, 1:500), anti-CD31-APC (BD Biosciences, 551262, 1:400), anti-CD11b-647 (UBC Ablab 67-0055-01, 1:500), anti-F4/80-647 (UBC Ablab, 67-0035-05, 1:500), and anti-Ter119-647 (UBC-Ablab, 67-0031-01, 1:200)]. Cells were incubated in staining cocktail for 30 minutes on ice in the dark. Subsequently, cells were washed in 15 mls FACS buffer and centrifuged at 500 x g for 5 minutes. The cell pellet was resuspended in FACS buffer containing Hoechst 33342 (Sigma B2261; final concentration 4 mM) and propidium iodide (Thermo Fisher, P1304MP; 1 mg/ml). For FCM, the single cell suspension was subsequently analyzed on a Beckman Coulter CytoFLEX flow cytometer. For FACS, cells were enriched using a BD Influx cell sorter and a combination of Hoechst, forward/side scatter parameters and propidium iodide were used to identify viable cells. Sorted cells were collected into sort media [20% FBS, L-glutamine (2 mM), penicillin (100 units/ml), and streptomycin (100 µg/ml in DMEM)] in cooled siliconized microcentrifuge tubes (Fisher Scientific; 02-681-320). 2.7 Cell cycle analysis To detect proliferating cells in embryonic forelimbs, standard methodology involving analysis of the incorporation of the nucleoside triphosphate analogue 5-ethynyl-2’-deoxyuridine 41 (EdU) was used. For this purpose, pregnant dams were injected intraperitoneally with a single dose of 1.5 mg of EdU (Thermo Fisher E10415) in PBS. Embryos were collected 24 hours post-injection and forelimb buds were processed for histology or flow cytometry using the Click-iT Plus EdU Alexa Fluor 647 Imaging kit (Thermo Fisher C10640) and the Click-iT Plus EdU Pacific Blue Flow Cytometry Assay kit (Thermo Fisher C10636), respectively, as per manufacturer instructions. 2.8 Single cell RNA-Seq Forelimb pairs were collected, pooled and processed from Hic1CreERT2; R26tdTomato litters at five different developmental timepoints with the following number of embryos at each time point: 14 (E11.5); 7 (E12.5,); 6 (E13.5,); and 6 (E14.5). At E16.5, 6 forelimb segments containing only the distal stylopod, and proximal zeugopod and their articulation (elbow joint) were collected. For E11.5 whole 1 and 2 samples, whole Hic1CreERT2; R26tdTomato limbs were dissociated and total lineage negative cells captured. Single cell suspensions were generated and target cells were enriched by FACS (see above) into 0.22μm vacuum filtered SC collection media (DMEM containing 5% FBS). Cells were counted and quality control was evaluated by hemocytometer. If the single cell suspension was confirmed to contain >98% tdTomato+ cells, the solution was input into a Chromium Controller (10X Genomics). Cell capture and library generation was carried out with the Chromium single cell 3’ reagent kit v2 (10X Genomics) for E11.5, E12.5, and E13.5 samples and v3 for E14.5 and E16.5. cDNA libraries were sequenced on a Nextseq500 (Illumina) to a depth > 40,000 reads per cell. The generated reads were aligned to a modified mm10 reference genome45 which included transcripts for tdTomato, GFP, etc. Demultiplexing and sequence 42 alignment was performed using the Cell Ranger count pipeline (10X Genomics). Quality metrics for the scRNA-seq were obtained from the cellranger-count output. Aggregated library sets were generated using the Cell Ranger aggr pipeline. Principal component analysis, non-linear dimensionality reduction (t-distributed stochastic neighbor embedding, t-SNE and uniform manifold approximation and projection, UMAP), and clustering were done using the Monocle3 R-package251,252. To ensure only high-quality cells were integrated into further bioinformatic analyses, cells with low total unique molecular identifier (UMI) count (< 3,000) and high mitochondrial RNA content were filtered. Also, the E16.5 forelimb dataset contained a small cluster of contaminating epithelial cells (Krt1+ or Krt14+), these cells were filtered from the aggregated dataset. A parallel set of principal component and downstream analyses were performed using the Seurat R package253. Monocle3 clustering results were subset into the Seurat object to generate violin plots and heatmaps. For scRNA-seq of MPs from adult muscle, single cell suspensions were generated as previously described45. TA muscles pooled from two P56 Hic1CreERT2; R26tdTomato hindlimbs and were sequentially digested in a volume of 250 µL per TA containing 500 U/mL Collagenase II (Sigma C-6685-5G) for 30 min followed by a cocktail of 1.5 U/mL Collagenase D (Roche 11088882001) and 2.4 U/mL Dispase II (Roche 04942078001) for 60 min. Digested material was subsequently stained and sorted as described above with the following modification: after the initial sort, viable target cells were further purified by sorting a second time. If the single cell suspension was confirmed to contain >98% tdTomato+ cells, the solution was input into a Chromium Controller (10X Genomics). Cell capture and library generation was carried out with the Chromium single cell 3’ reagent kit v3. cDNA libraries were sequenced on a Nextseq500 43 (Illumina) to a depth > 50,000 reads per cell. The generated reads were aligned to the aforementioned modified mm10 reference genome. Demultiplexing and sequence alignment was performed using the Cell Ranger count pipeline (10X Genomics). Quality metrics for the scRNA-seq were obtained from the cellranger-count output. Cells containing low total UMI counts (< 1,500), high mitochondrial RNA content and small clusters containing cycling cells (Birc5+, 90 cells), and contaminating cell types (Mpz+, 146 cells; Pecam1+, 19 cells) were removed from further downstream analyses. Principal component analysis, non-linear dimensionality reduction (t-SNE and UMAP), and clustering were done using the Seurat R package. To generate force-directed layouts for purposes of visualizing and characterizing cell fate diversity, the python package ‘Harmony’ was used254. The calculated affinity matrices were then used to calculate single cell differentiation trajectories, gene expression trends and associated pseudotime via the python package ‘Palantir’255. ‘Start cells’ were determined by high expression of manually curated genes characteristic of embryonic MPs (Snai1, Id1). Likewise, ‘end cells’ were determined with gene markers for each cell fate: fibroblasts (Dpt, Pi16), DFs (Crabp1, Twist2), chondrocytes (Col2a1, Sox9) and tenocytes (Tnmd, Thbs4). A focus on tenogenic and chondrogenic lineages was made possible by filtering out cells of interest according to the emergence of tenogenic and chondrogenic gene programs, respectively. Gene expression heatmaps were overlayed on the force-directed layout after imputation with the MAGIC algorithm which denoises data and fills dropouts256. Representative genes for chondrogenic (Sox9, Acan) or tenogenic (Scx, Tnmd) cell fate as well as for the embryonic MP state (Id1, Snai1) were selected and used to illustrate the change in gene expression level as a function of pseudotime using the gene expression trend function in Palantir. 44 2.9 Single-cell fate trajectory reconstruction The URD R package which involves a simulated diffusion-based approach was used to computationally generate differentiation trajectories in pseudotime at the single cell level257. Cells undergoing mitosis confounded analysis and were removed from subsequent analyses. Transition probabilities and pseudotime were calculated as per the URD tutorial (https://github.com/farrellja/URD). Cells originating from E11.5 forelimbs were assigned as the root. Tip populations were determined manually through analysis of differentially expressed marker genes in the E16.5 dataset. To generate the tree structure, visitation frequencies were derived based on biased random walks performed from random tip cells towards the root. URD functions were mostly performed with default parameters with the following exceptions: for the diffusion map calculation, the number of nearest neighbors (knn) was optimized to 200 and the kernel width to use (sigma.use) was set to 8. Regarding pseudotime calculations, a total of 150 simulations (floods) were performed and averaged. To determine tips for the tree structure, the optimal clustering of the E16.5 scRNA-seq data was found to be Louvain-Jaccard clustering with number of nearest neighbors (num.nn) set to 25. Clusters containing cells expressing progenitor markers were omitted from tip selection. To determine cellular trajectories in pseudotime, 10,000 biased random walks (n) were simulated. To build the tree, divergence method was set to “ks”; the remaining parameters were changed as follows: min.cells.per.segment = 10, p.thresh = 0.025, and minimum.visits = 1. To generate a force directed layout, the method was set to “fr”, and the following arguments were modified: num.nn = 120, and min.final.neighbours = 4. Force-directed layout rendition of the URD tree then was optimized for visualization in 2D using the 45 “treeForceStretchCoords”, “treeForceRotateCoords”, and “treeForceTranslateCoords” functions from the URD package. For generating gene expression cascades, representative genes for each trajectory were selected from a list of differentially expressed genes generated by the URD R package. Beginning with each tip population, URD compared cell transcriptomes in each segment pairwise with cells from each of that segment’s siblings and children. Genes were then considered differentially expressed if they were expressed in at least 10% of the cells in the segment under consideration, their mean expression was upregulated 1.5x compared to the sibling branch and the gene was 1.25x better than a random classifier for the population. For integration into the population’s cascade, a gene must be considered differential in at least 60% of their siblings and not be differentially upregulated in a different trajectory downstream of the branchpoint at any given branchpoint. To generate a cascade heatmap for every trajectory, the expression pattern of each marker gene was fit into a linear, sigmoid, or double sigmoid relationship to describe at what time expression begins and ends. Then, cells in each trajectory were grouped using a moving window through pseudotime and the mean gene expression was plotted for each window, scaled to the maximum gene expression per window. 2.10 Single cell ATAC-seq For single cell ATAC-seq capture, ~ 100,000 FACS-purified tdTomato+ cells from Hic1CreERT2; R26tdTomato were collected from E12.5, E14.5, and E16.5 forelimbs as per scRNA-seq sorting conditions described above. Nuclei were isolated by lysing the cells according to the 46 described protocol (10X Genomics, https://support.10xgenomics.com/single-cell-atac) for 5 minutes. Nuclei were quantified using a Countess II FL automated cell counter (Thermo Fisher) and ~ 10,000 nuclei were targeted for transposition and captured using a 10x Chromium controller. ScATAC-seq libraries were prepared according to the Chromium Single Cell ATAC Reagent Kits User Guide (10x Genomics; CG000168 Rev B). Single cell libraries were sequenced on a Nextseq500 (Illumina). The cellranger-atac single cell ATAC pipeline version 1.1.0 was used to generate fastq files from the sequencer output bcl files and further perform read filtering and alignment, detection of accessible chromatin peaks, dimensionality reduction, cell clustering and differential accessibility analyses. Quality metrics for the scATAC-seq such as insert size distribution, enrichment around the transcriptional start site and a t-SNE heatmap of fragments per cell were obtained from the cellranger-atac output. Track plots were generated using Loupe Cell Browser (10x genomics, v. 4.1.0). The 10x genomics output was further processed using the standard Signac workflow to compute quality control metrics, normalization, latent semantic indexing and subsequent UMAP calculation. Cells were then clustered using the Leiden algorithm and integrated with scRNA-seq data using cross-modality integration and label transfer. For pseudotime and differential accessibility analysis, the 10x filtered peak barcode matrix output was used to generate a cell data set-class object using monocle3. To generate pseudotime trajectories of chondrogenic, tenogenic, and DF cell commitment, separate subset objects were created by using MP combined with chondrocyte, tenocyte, or DF clusters, respectively. The cicero package was used to construct trajectories with accessibility data, perform differential accessibility analysis, and plot accessibility changes in pseudotime. The cicero function was then used to construct cis-regulatory, and compute cis-co-accessibility, networks258. To plot genomic ranges 47 including cicero connections, links were incorporated into the Seurat object and visualized using the coverage plot function259. 2.11 Ex-vivo Forelimb Cultures Ex-vivo culture of embryonic forelimbs was carried out as previously described243. Limb culture incubation chambers were prepared by placing a stainless steel mesh into the inner well of a 60 mm organ culture dish. The stainless steel mesh was previously punctured to allow for the placement of a polyester (PET) membrane on its surface. The outer trough of the dish was filled with distilled water, and forelimb culture media (10% FBS in BGJb medium containing 100 mM penicillin/streptomycin and 100 mM L-glutamine) was added to the inner well. Embryonic limb buds were collected at E11.5 by dissection at the limb-trunk interface and placed on the PET membrane surface within a limb culture incubation chamber. Limbs were cultured in a well-humidified incubator (37 0C, 5% CO2) for 1-3 days. When live imaging was necessary, limbs were similarly cultured in a glass-bottom 35mm dish atop a PET membrane and stainless steel mesh. 2.12 Micromass Cultures Preparation of PLM cultures was done as previously described235. Embryos from timed-pregnant females were harvested at E11.5 and limb buds were dissected followed by enzymatic dissociation using a cocktail containing 10% FBS and Dispase I (10 units/ml) in Puck’s saline A. Limb buds were removed by pinching at the body wall with a pair of Dumont #5 forceps (Fine Sciences Tools, 11251-20) or by taking the distal ¼ for distal tip cultures. Following dissection, 48 limbs were minced with sharp forceps. Minced limbs were then transferred into a 15mL conical tube ensuring complete transfer by rinsing the mincing tube several times with PBS. Limb buds were incubated in enzyme cocktail for 80 minutes at 370 C with slight shaking (~70 rpm) and gently vortexed every 20 minutes. Following digestion, cells were washed in micromass media (10% FBS in 40% DMEM/60% F12 nutrient mixture with 100 mM penicillin/streptomycin and 100 mM L-glutamine), pelleted at 1,000 x g for 5 minutes, filtered through a 40 µm cell strainer (this step removes the ectoderm and cell clumps, generating a reasonably good single cell suspension), and quantified using a hemocytometer. Micromass cultures were generated by spotting 10 µl of cell solution at a density of 1-2 x 107 cells/ml into the center of a well within a 24 well plate (Nunc or Grenier plates are optimal to minimize droplet spread and reduce evaporation). Cultures were maintained in a humidified incubator (5% CO2) for an hour before adding 1 ml of micromass media to each well. Media was changed every other day thereafter. Cartilage condensation usually become visible between days 1 and 2, cartilage nodules should start to appear by late day 2 or early day 3. For formation of hypertrophic cartilage, media changes were supplemented with 10 mM b-glycerol phosphate and 50 µg/mL ascorbate and incubated for 18-24 days. For staining of micromass cultures with Alcian blue, media was removed from day 2 or day 3 -old cultures and washed twice with warm PBS. Cultures were then fixed with 1X formalin for 15 minutes followed by a PBS and 0.2M HCl wash. Alcian blue stain was then added to each well (4:1 ratio of 0.2 M HCl and 0.5% Alcian blue in 95% ethanol) and incubated overnight at room temperature in a plastic container with damp paper towels. To remove excess stain the following day, cultures were washed with 70% ethanol and allowed to dry (can be stored dry). 49 Stained micromass cultures were imaged in 70% ethanol on a Zeiss or Nikon dissection light microscope. 2.13 Wholemount and cryosection X-gal staining b-galactosidase enzymatic activity at different stages of embryonic limb development was detected using established whole-mount X-gal staining protocols260. Embryos were collected at various developmental time points ranging from E11.0 to E14.5. Dissected limbs were fixed in LacZ fixative (100 mM MgCl2, 0.2% glutaraldehyde, 5 mM ethylene glycol tetra-acetic acid in 2% PFA). Duration of immersion was dependent on the time point collected: E11.0 and E11.5 limbs were fixed for one hour, E12.5 for 1.5 hours, E13.5 for 2 hours, and E14.5 for 2.5 hours. All fixative incubation periods were carried out in the dark at 4o C. Fixed samples were washed 3X in cold PBS for 30 minutes each followed by overnight incubation in LacZ staining solution (2 mM MgCl2, 0.01 % Deoxycholate, 0.02% NP40, 5 mM potassium ferricyanide, 5 mM potassium ferrocyanide and 1mg/mL 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside) at 37o C. Samples were post-fixed in 4% PFA for an hour at 4o C prior to imaging. To generate X-gal-stained histological embryonic limb sections, pregnant mice were sacrificed and embryos were isolated in cold PBS. Limbs were subsequently dissected from the trunk and fixed in LacZ fixative as described above. Fixed limbs were washed 3 times and processed for cryosectioning as described above. Samples were subsequently processed for X-gal staining by cryosectioning the frozen OCT blocks with a Leica CM 1950 cryostat at 6-10 µm. For staining, slides were thawed at room temperature and then washed 3 X 10 minutes in PBS to 50 remove residual OCT compound followed by incubation overnight in LacZ staining solution (see previous section) at 37o C. Once stained, slides were washed in PBS 3 X 5 minutes, counterstained with Nuclear Fast Red (NFR) for 2 minutes, rinsed in water and dehydrated through a series of solutions as follows: 2 X 2 minutes in 70% ethanol, 2 X 2 minutes in 95% ethanol, 2 X 5 minutes in 100% ethanol, and 2 X 10 minutes in xylene. Slides were then cover-slipped with Cytoseal 60 mounting media (Thermo Scientific Cat. # 831016). 2.14 Whole-mount skeletal preparations Alcian blue and Alizarin red staining of Hic1nLacZ skeletons was performed as described in Rigueur and Lyons261. E16.5 and E17.5 embryos derived from Hic1nLacZ/+ x Hic1nLacZ/+ timed pregnancies were collected and scalded in 65o C water to facilitate removal of the epidermis. Embryos were subsequently eviscerated. Fixation was achieved by sequential overnight incubations in 95% ethanol and acetone. Cartilage staining was done by overnight incubation in 0.03% Alcian blue solution in a 4:1 mixture of 80% ethanol and glacial acetic acid, respectively. Excess stain was removed by washing in 70% ethanol twice and 95% ethanol overnight. Embryos were then pre-cleared in 1% KOH for one hour. Mineralized tissues were stained by incubation in 0.005% Alizarin red in 1% KOH for 4 hours. The solution was then replaced by a 50:50 mixture of glycerol: 1% KOH. For long term storage, samples were kept in 100% glycerol. Stained skeletal samples were imaged on a Leica MZ FL III dissection microscope. For micro-computed tomography (µCT) analysis of skeletons, Prrx1Cre/+; Hic1f/f mice were sacrificed at P56, skinned, eviscerated and stored in 70% ethanol prior to imaging. µCT scanning 51 was carried out in a vivaCT 40 at a 35µm voxel resolution for whole skeletons or µCT 40 (Scanco Medical AG, Bassersdorf, Switzerland) at a 20µm resolution for forelimb-focused images. Images were visualized using Scanco analysis software (µCT Ray V4.0-4). To ensure proper controls, seven samples were imaged in total: Prrx1Cre/+; Hic1f/f (n = 3), Prrx1Cre/+; Hic1f/+ (n = 2), Prrx+/+; Hic1f/+ (n = 1), and Prrx1+/+; Hic1+/+ (n = 1). 2.15 Quantification and Statistical analysis All data are represented as mean ± SD, the sample number and statistical analyses are indicated in the figure legends. Sample size determination was based on anticipated variability and effect size that was observed for similar experiments. For pairwise comparisons, unpaired two-tailed t tests were used to calculate P values, p-value < 0.05. For comparison of >2 means, one-way ANOVA with Bonferroni’s correction and post hoc analyses were employed, p-value < 0.05. Both statistical methods were carried out using Prism (GraphPad Software v. 8). 52 Table 2.1 Summary of antibodies used Flow Cytometry Vendor Catalogue Number Dilution anti-CD31-APC BD Biosciences 551262 1:400 anti-Mo CD117 (c-Kit) -APC eBioscience 17-1172-82 1:500 anti-mCD11b-647 UBC Ablab 67-0055-01 1:500 anti-mF4/80-647 UBC Ablab 67-0035-05 1:500 anti-mCD45 Pan -647 UBC Ablab 67-0047-01 1:400 anti-mTer119-647 UBC Ablab 67-0031-01 1:200 Primary Antibodies for Histology Supplier Catalogue Number Dilution anti-Sox9 Abcam ab185966 1:100 anti-CD31 BD Pharmingen 550274 1:50 anti-myosin (skeletal, fast) Sigma M4276 1:1000 anti-Pax7 UBC Ablab AB0000457 1:500 anti-Pax3 DSHB Pax3 1:100 anti-myosin heavy chain, sarcomere DSHB MF 20 1:50 anti-Neurofilament Heavy Chain Abcam ab4680 1:5000 anti-RFP Abcam ab62341 1:250 anti-GFP Abcam ab13970 1:500 53 anti-Laminin Abcam ab11575 1:100 anti-Tagln Abcam ab14106 1:100 anti-Sox2 Abcam ab97959 1:100 anti-Perilipin 1 Cell Signaling Technology 9349s 1:250 anti-VE Cadherin Abcam ab205336 1:100 anti-Myod1 Abcam ab133627 1:100 anti-Sox10 Abcam ab155279 1:100 anti-Pax1 Abcam ab252847 1:100 anti-Calponin Abcam ab46794 1:100 anti-Hic1 Underhill Lab N/A 1:10,000 Secondary Antibodies for Histology Supplier Catalogue Number Dilution Alexa Fluor 488 goat anti-mouse IgG (H+L) Thermo Fisher Scientific A11029 1:500 Alexa Fluor 647 goat anti-mouse IgG (H+L) Thermo Fisher Scientific A32728 1:500 Alexa Fluor 647 goat anti-rat IgG (H+L) Thermo Fisher Scientific A21247 1:500 Alexa Fluor 594 goat anti-rabbit IgG (H+L) Thermo Fisher Scientific A11012 1:500 54 Alexa Fluor 488 goat anti-rabbit IgG (H+L) Thermo Fisher Scientific A11034 1:500 Alexa Fluor 647 goat anti-rabbbit IgG (H+L) Thermo Fisher Scientific A27040 1:500 Alexa Fluor 488 goat anti-rat IgG (H+L) Thermo Fisher Scientific A11006 1:500 Alexa Fluor 488 goat anti-Chicken IgG (H+L) Thermo Fisher Scientific A11039 1:500 55 Chapter 3: Cellular taxonomy of Hic1+ mesenchymal progenitor derivatives in the limb from embryo to adult 3.1 Introduction Tissue-resident mesenchymal progenitors (MPs) can be found to varying extents in all tissues and play vital roles in tissue homeostasis, renewal and regeneration193,194. The gene Hic1 has been shown to identify MPs in adult skeletal muscle (SkM)45. Lineage tracing and single cell transcriptomics have revealed that Hic1+ cells in SkM are comprised of three subpopulations: fibro-adipogenic progenitors, pericytes, and tenogenic progenitors45. FAPs generate a transient population that orchestrates multiple facets of the regenerative program. Once an injury has resolved, ‘excess’ FAPs are cleared via apoptosis and tissue integrity is restored48,49. Tenogenic progenitors produce enduring progeny that regenerate fibrous connective tissues which link muscle to bone, such as tenocytes and myotenocytes45. Hic1+ pericytes are laminin-encased and express bona fide pericyte markers such as Rgs5+, Pdgfrb+, Kcnj8+ and Abcc9+. Indicative of their progenitor status, all of the described populations exhibit open chromatin regions (OCRs) across numerous cell cycle loci and enter the cell cycle following injury-induced activation45. Collectively, Hic1+ MPs regulate multiple facets of the regenerative program across multiple cell types45,203,204. However, our understanding of their embryonic origin and subsequent fates is unclear. The origin and fate of other tissue-resident stem and progenitor populations such as satellite cells within striated muscle has been well defined due to the specificity of the marker Pax7/PAX7. Fate mapping experiments performed in birds and mice have demonstrated that 56 satellite cells originate in the somitic dermomyotome and migrate to form the appendicular and axial skeletal musculature262–264, where they persist to form a reservoir of post-natal stem cells265. In contrast, due to the lack of effective markers of putative mesenchymal subsets in the early limb, it has been challenging to resolve the origin, fate and cellular hierarchies within the MP cellular compartment. In this chapter, I show that the E11.5 limb mesenchyme exhibits significant heterogeneity and that Hic1 defines a distinct mesenchymal subset which contributes to limb fibroblasts and other specialized mesenchymal derivatives, some of which persist to support adult tissue regeneration. 3.2 Results 3.2.1 Hic1 expression patterns in the mouse embryo The expression dynamics of Hic1 in the developing mouse embryo have been previously characterized through in situ hybridization in the late 1990s166. Grimm and colleagues found that detectable Hic1 expression begins at E9.5 in somites of the occipital cervical region as a striped pattern in the rostral-caudal direction. By E10.5, Hic1 expression is localized to the mesenchymal portion of the somites, underlying the epithelial dermomyotomes. This region, considered to be sclerotomal, contained highly spatially-restricted Hic1+ cells that were maintained until E11.5 and E12.5. In the case of the developing limbs, Hic1 was not expressed within the limb mesenchyme until E10.75 in the proximal dorsal and ventral surfaces166. With this knowledge in mind, I used the Hic1nLacZ reporter mouse line to define Hic1 expression patterns in early embryos (E9.5-12.5). Wholemount LacZ staining confirmed that Hic1 expression is not detectable prior to E10.5 (Figure 57 3.1a). Beginning at E10.75, a LacZ signal is evident within regions of the epaxial and hypaxial somitic mesenchyme (Figure 3.1b). At E11.0, LacZ+ cells can be observed faintly within cranio-facial regions (Figure 3.1c) as well as commencing to occupy areas of the limb mesenchyme (Figure 3.1d). Overall, these results confirm those reported by Grimm166, supporting our use of the Hic1nLacZ mouse line to study Hic1+ progenitors during embryogenesis. Figure 3.1 Hic1 expression dynamics in early embryonic development visualized via the Hic1nLacZ mouse line. (a-d) Hic1nLacZ/+ embryos were collected and X-gal stained at the specified embryonic timepoints. Scale bars, 1mm. Dotted lines outline forelimb bud. Arrowheads: Black, hypaxial somite; white, epaxial somite; red, facial mesenchyme. The Hic1CreERT2;R26tdTomato mouse line was introduced into these embryological studies to further expand on the study of Hic1+ progenitors during embryonic development and to bypass some of the limitations of the Hic1nLacZ mouse model, such as loss of LacZ/Hic1 expression upon cell differentiation45 and the inability to perform lineage tracing studies. These reporter mice allowed for lineage tracing of Hic1+ cell progeny regardless of Hic1 expression post-recombination and without the need for time-consuming histochemical techniques. Building on previous knowledge regarding the onset of Hic1 expression during embryogenesis, several TAM-induction windows were attempted with the goal of labelling the earliest Hic1+ cells possible but also maximizing the amount of recombination that occurs (Figure 3.2a). TAM was administered 58 to pregnant mice at E8.5, E9.5, and E10.5. At E13.5, embryos were collected and induction at E10.5 was found to yield the greatest concentration of tdTomato+ cells out of the three timepoints tested (Figure 3.2b-d, n = 3 embryos from independent litters). As a result, TAM-induced recombination was selected to be performed at E10.0 to label the earliest Hic1-expressing cells without compromising the quantity of cells labelled during the recombination window266. Figure 3.2 Determination of optimal embryonic timepoint for Cre-induced recombination in Hic1CreERT2; R26tdTomato embryos. (a) Schematic representation of TAM temporal pulse optimization. (b-d) Fluorescence microscopy images of E13.5 Hic1CreERT2; R26tdTomato embryos treated at the indicated timepoints with a single TAM dose. Scale bars, 500µm. h, heart; nt, neural tube; Rfl, right forelimb. 59 Wholemount confocal microscopy of Hic1CreERT2; R26tdTomato embryos TAM-treated at E10 (corresponding to the night of E9.5) and collected at E11.5 reveal similar patterns of tdTomato protein compared to that of b-galactosidase in Hic1nLacZ embryos of the same age with some notable exceptions. Although tdTomato+ cells were present within somitic mesenchyme and formed similar striped patterns in an anterior-posterior fashion, no cells were yet evident within the cranio-facial regions nor in the limb bud proper (Figure 3.3a). However, two elongated clusters of tdTomato+ cells were observed extending from the hypaxial region of the somites into the proximal region of the forelimb bud (Figure 3.3a, arrowheads). Based on histological examinations of Hic1nLacZ embryos and previous work done on embryonic Hic1 expression166, it is evident that Hic1+ MPs and their progeny were abundant within the somitic mesoderm; however, it was not yet clear which specific region of the somite is populated by these progenitors. To address this, I employed an immunofluorescence approach consisting of identification of somite boundaries by expression of specific markers, in combination with the Hic1CreERT2; R26tdTomato mouse line. By E11.5, most somites have undergone epithelial-mesenchymal transition and given rise to four distinct somitic regions: the dermatome, myotome, syndetome, and sclerotome (Figure 3.3b). To identify the relevant sub-compartments of the somite, the following antibody markers were used: MYOD1 and PAX7 for the myotome267,268; ScxGFP for the syndetome2,248, and PAX1 for the sclerotome269,270. TdTomato+ cells were located medial to the myotome (Figure 3.3c-d). No overlap between tdTomato+ and MYOD1+/PAX7+ cells was observed (Figure 3.3c-d). In contrast, a subset of the most medially-located tdTomato+ progenitors were also positive for PAX1, indicating a partial overlap with cells derived from the sclerotome (Figure 3.3e). Furthermore, there was an appreciable population of ScxGFP/tdTomato double-positive cells within the somitic 60 mesenchyme (Figure 3.3f). These results suggest that tdTomato+ progenitors form a subset of the somitic sclerotome and syndetome, but not the myotome. (Figure legend on next page) 61 Figure 3.3 Analysis of tdTomato+ cell distribution in somitic mesoderm at E11.5. (a) Lateral view of an E11.5 Hic1CreERT2;R26tdTomato embryo imaged in wholemount. White dotted-line highlights the forelimb; white box represents inset region on the right; white arrowheads indicate tdTomato+ cells within the proximal forelimb. Scale bars, 1 mm; inset 500µm. (b) Schematic representation of transverse section through an E11.5 embryo used to generate samples in panels c-f. (c) Representative immunofluorescence image of Hic1CreERT2;R26tdTomato embryo transverse section stained with a-PAX7 to identify myotomes. Scale bars, 100µm. (d) Representative immunofluorescence image of Hic1CreERT2;R26tdTomato embryo transverse section stained with a-MYOD1 to identify myotomes. Scale bars, 100µm. (e) Representative immunofluorescence image of Hic1CreERT2;R26tdTomato embryo transverse section stained with a-PAX1 to identify sclerotomes. Scale bars, 100µm. (f) Representative immunofluorescence image of Hic1CreERT2;R26tdTomato;ScxGFP used to identify sclerotomes. Scale bars, 100µm. (c’-f’) Schematics illustrating the respective regions of interest highlighted within a red box corresponding to adjacent panels. D, dorsal; L, lateral; M, medial; V, ventral. 3.2.2 Hic1 identifies a rare MP population within the limb bud In the developing murine embryo, forelimb bud outgrowth initiates at E9.5 and is composed of an ectodermal jacket encasing LPM-derived mesenchyme. Approximately 36 hours post-initiation, Hic1 transcripts become evident within the proximal limb mesenchyme166. Quantitative PCR analyses of E10.5 limb segments revealed low Hic1 expression. In contrast, Hic1 transcript was detected at E11.5 in a decreasing proximal to distal gradient (Figure 3.4a-b). To explore the nature and origin of Hic1+ cells in the limb, a lineage tracing strategy was employed involving a Hic1CreERT2 allele in conjunction with a Cre-dependent reporter, R26tdTomato (Figure 3.5a). Embryos that were pulse-labelled with TAM at E10 showed no trace of tdTomato+ cells in the forelimb buds until ~E11.5 (Figure 3.5b-c). Consistent with this, tdTomato+ cells were observed mostly restricted to the cultured proximal, but not distal, forelimb fragments and corresponding micromass cultures therefrom (Figure 3.5d). 62 Figure 3.4 Quantitative analysis of Hic1 expression in the proximal-distal axis of E10.5 and E11.5 forelimb buds. (a-b) Quantitative PCR analysis of Hic1 mRNA transcripts. Corresponding limb bud regions are shown in the left panel diagram. Replicates 1-3 are shown. To determine at which time the tdTomato+ cells populated the limb, E11 forelimb buds were analyzed and revealed the presence of two clusters at the body wall-limb bud interface (Figure 3.5c, E11.0). By E11.75, whole-mount imaging identified a band of tdTomato+ cells within the hypaxial somite that extended into the proximal forelimb bud (Figure 3.5b, E11.75). By E12.5, this band of cells extended to surround the chondrogenic core of the developing limb (Figure 3.5c, E12.5). Flow cytometry (FCM) analysis at E11.5 showed that tdTomato+ cells had populated the central limb mesenchyme and constituted 0.45 ± 0.16% of total forelimb mesenchymal cells (Figure 3.6a-b). The frequency of this population increased markedly to comprise 5.79 ± 1.1%, 12.1 ± 1.6% and 26.6 ± 0.5% of the limb mesenchymal cell compartment by E12.5, E13.5 and E14.5, respectively (Figure 3.6c). To determine if tdTomato+ cells represented a unique subset of mesenchymal cells, scRNA-seq was performed on E11.5 forelimb buds and on forelimb-derived FACS-purified 63 tdTomato+ cells. TdTomato+ cells clustered separately from other whole-limb mesenchymal cells (Figure 3.7a). Genes such as Hic1 and Aldh1a2 distinguished the tdTomato+ cluster from tdTomato- limb bud mesenchymal cells (Lhx9+, Msx2+), myogenic precursors (Myod1+, Pax7+) and epithelium (Krt5+, Krt14+) (Figure 3.7b). Co-expression of the retinoic acid (RA) synthesis gene, Aldh1a2 with Hic1 is consistent with direct regulation of Hic1 transcription by the nuclear receptors for RA, RARs271. The differential expression signature of tdTomato+ cells further reinforced the distinct nature of this population. Congruent with this, at E11.5 tdTomato+ cells were observed in close association with myogenic progenitors (PAX7+) and projecting nerve axons (Figure 3.7c-f) in the E11.5 and E12.5 proximal limb bud. 64 Figure 3.5 Hic1 identifies a discrete MP population within the limb mesenchyme. (a) Schematic of the experimental strategy used for Hic1CreERT2; R26tdTomato genetic recombination and experimentation. (b) Maximum intensity projections of wholemount E11.0, E11.5, and E11.75 Hic1CreERT2; R26tdTomato embryos. White dotted line indicates the forelimb bud. E11.5 is identical image used in Figure 3.3a. Scale bars, 500µm. (c) Representative images of Hic1CreERT2; R26tdTomato forelimb sections taken at sequential stages of development and counterstained with a-CD31 and DAPI. Scale bars, 100µm (left half), 20µm (right half). White boxed represent inset regions highlighted on adjacent panels, white arrowheads define tdTomato+ cells. (d) Representative maximum intensity projections of Hic1CreERT2; R26tdTomato micromass cultures. Limbs were harvested at E11.5 and partitioned into proximal or distal portions. Scale bar, 100µm. 65 Figure 3.6 Quantitative analysis of tdTomato+ cells in Hic1CreERT2; R26tdTomato forelimbs at sequential developmental time points. (a) Representative maximum intensity projections of Hic1CreERT2; R26tdTomato forelimbs acquired at the specified developmental stages. Scale bars, 1mm. (b) Representative FCM scatter plots from dissociated Hic1CreERT2; R26tdTomato forelimbs corresponding to timepoints in (a). (c) FCM analysis of enzymatically dissociated Hic1CreERT2; R26tdTomato embryo forelimbs. Data represent mean +/- SD. 66 Figure 3.7 TdTomato+ MPs are distinct from limb LPM and localize with nerves and myogenic progenitors. (a) UMAP of aggregated scRNA-seq data representing whole forelimb dissociations (whole 1 and whole 2) and FACS-purified tdTomato+ MPs from E11.5 forelimb buds. (b) UMAPs of representative genes for each identified cell population: tdTomato+ MPs (Hic1 and Aldh1a2), tdTomato- MPs (Lhx9 and Msx2), myogenic progenitors (Myod1 and Pax7), and epithelium (Krt5 and Krt14). (c) E11.5 Hic1CreERT2; R26tdTomato embryo imaged and stained in wholemount with antibodies against neurofilament heavy chain (NEFH), PAX7, and MF20. Scale bars, 500 µm and 100 mm. White box represents inset region highlighted in panel on right, arrowhead indicates tdTomato+ cells within the limb. (d) E12.5 Hic1CreERT2; R26tdTomato embryo imaged and stained in wholemount with indicated antibodies. Scale bars, 500 µm and 100 µm. White box represents inset region highlighted in panel on right. Arrow heads indicate clusters of tdTomato+ cells. (e) Region of interest captured in panels (f). (f) E11.5 Hic1CreERT2; R26tdTomato forelimb sections stained with antibodies against NEFH and PAX7. Scale bars, 100 µm and 20 mm inset. White box represents inset region highlighted in bottom-right panel. A, anterior; D, distal; Po, posterior; Pr, proximal. 67 The spatiotemporal localization of tdTomato+ MPs led to the hypothesis that this cell population could represent a subset of migratory progenitors which invade the developing limb bud akin to myogenic precursors and nerve axons. To test this, live imaging fluorescence microscopy was performed on dissected E11.5 Hic1CreERT2; R26tdTomato forelimb buds to observe tdTomato+ MP behavior in real time. TdTomato+ cells were evident within the proximal end of the limb bud at the moment of dissection (Figure 3.8). As time progressed, the cells appeared to ambulate throughout the forelimb in an overall proximal-distal direction (Figure 3.8). Furthermore, ex vivo organ culture of Hic1CreERT2; R26tdTomato limb buds revealed that movement of tdTomato+ cells into the most distal regions of the limb could be interrupted by physical separation of the limb bud into proximal and distal segments (Figure 3.9a-b). Similarly, live imaging of Hic1CreERT2; R26tdTomato embryo-derived PLM (micromass) cultures showed that tdTomato+ progenitors were highly mobile and concentrated within internodular regions with respect to the cartilaginous condensations (Figure 3.10). These results suggest that embryonic tdTomato+ MPs are a highly dynamic and migratory cell population. It is well established that several embryonic progenitor populations migrate from their tissue of origin into the developing limb buds272,273, examples include somite-derived myoblasts, blood vessels274,275, and neural crest derivatives144. To compare tdTomato+ MPs to other limb-bound migratory populations at the histological level, an immunofluorescence approach was employed in which cells derived from the myotome (PAX3+), sclerotome (PAX1+) and neural crest (SOX10+) could be identified by the specified protein markers and overlayed on a Hic1CreERT2; R26tdTomato embryo. As mentioned previously, tdTomato+ MPs do not co-localize with PAX7+ cells; however, myoblasts that are actively migrating during early forelimb morphogenesis do not 68 express Pax7, but rather express Pax3276. A subset of limb pericytes and VSMCs have been found to derive from the somitic sclerotome89 and therefore a-PAX1 was used to identify the sclerotome at the time of tdTomato-cell migration. Lastly, a-SOX10 was chosen to label neural-crest derived migratory cells277. These immunofluorescence experiments demonstrated that tdTomato+ cells do not overlap with migratory PAX3+ dermomyotome-derived progenitors (Figure 3.11a-b) nor with SOX10+ neural crest-derived cells (Figure 3.11c-d). Interestingly, PAX1/tdTomato double-positive cells were observed (Figure 3.12a,b) suggesting that a subset of Hic1+ progeny may be sclerotome-derived. Furthermore, the sclerotome contains a population of Scx+ tenogenic progenitors which also overlap with tdTomato+ cells at the forelimb level (Figure 3.12c,d). In summary, these results indicate that tdTomato+ MPs are migratory cells that do not derive from the dermomyotome nor neural crest, but likely have an origin within the sclerotome and syndetome compartments of the somite. The recombination time window that exists between TAM induction and de facto expression of tdTomato in Hic1CreERT2; R26tdTomato embryos presents a challenge for the determination of a specific origin of Hic1+ MPs in the limb. Specifically, in all lineage tracing experiments described in this text, 1.5 days are allotted (from E10 to E11.5) for Cre-recombinase nuclear translocation, DNA recombination, and tdTomato expression to occur. This time period represents a blind spot in which Hic1+ MPs may be located at different regions of the embryo or display variances in gene expression prior to their identification as tdTomato+ cells. To address this issue, an immunofluorescence histological approach was used wherein an a-HIC1 antibody would identify HIC1+ MPs and sub-compartments of somitic progenitors would be identified by immunodetection of select markers. 69 Figure 3.8 Live imaging of Hic1CreERT2; R26tdtomato forelimbs identifies a highly dynamic tdTomato+ population. Representative images with accompanying vector graphs showing movement direction of tdTomato+ cells in an E11.5 forelimb imaged overnight. Timescale (t) indicates number of hours after dissection. Scale bar, 500um. P, proximal; D; distal. 70 Figure 3.9 TdTomato+ MPs are localized to the proximal forelimb during early limb morphogenesis. (a) Schematic representation of experimental workflow. (b) Representative wholemount images of Hic1CreERT2; R26tdTomato forelimbs cultured whole, proximally or distally-truncated and imaged at 0, 24, and 48 hours post collection. Scale bars, 1 mm. 71 Figure 3.10 Live imaging of Hic1CreERT2; R26tdTomato micromass cultures highlights the movement of tdTomato+ MPs throughout internodular regions. Representative images and accompanying vector graphs showing general movement of tdTomato+ cells in a PLM culture imaged for 24 hours. Scale bar, 500um. Blue circles indicate mesenchymal condensations that precede the formation of cartilage nodules. 72 Figure 3.11 TdTomato+ MPs do not overlap with myotome and neural crest-derived migratory populations at the forelimb level. (a) Schematic of an E11.5 embryo cross-section. Red box encloses the area captured in panel (b). (b) Immunofluorescence staining with a-PAX3 on E11.5 Hic1CreERT2; R26tdTomato embryos at the forelimb level. (c) Schematic of an E11.5 embryo cross-section. Red box encloses the area captured in panel (d). (d) Immunofluorescence staining with a-SOX10 on E11.5 Hic1CreERT2; R26tdTomato embryos at the forelimb level. Scale bars, 100um. D, dorsal; L, lateral; M, medial; nt, neural tube; V, ventral. Dotted boxes highlight inset regions. 73 Figure 3.12 TdTomato+ MPs partially overlap with sclerotome and syndetome populations at the forelimb level. (a) Schematic of an E11.5 embryo cross-section. Red box encloses the area captured in panel (b). (b) Immunofluorescence staining with a-PAX1 on E11.5 Hic1CreERT2; R26tdTomato embryos at the forelimb level. (c) Schematic of an E11.5 embryo cross-section. Red box encloses the area captured in panel (d). (d) Representative image of E11.5 Hic1CreERT2; R26tdTomato; ScxGFP forelimb section. Scale bars, 100um. D, dorsal; L, lateral; M, medial; nt, neural tube; V, ventral. Dotted boxes highlight inset regions. 74 The results showed that at E10.5 HIC1+ progenitors could be found within the medial somitic mesenchyme, partially overlapping with the PAX1+ sclerotome (Figure 3.13a) and ScxGFP+ syndetome (Figure 3.13b). Immediately lateral to these HIC1+ cells was the PAX3/7-positive myotome. No HIC1+ cells were observed to overlap with PAX3 or PAX7-positive myotome cells (Figure 3.14a-b). HIC1+ cells that were located at proximal end of the limb bud were often found in close proximity with extending nerve axons (Figure 3.15a) and did not colocalize with PAX3+ migratory muscle progenitors (Figure 3.15b) nor with PAX7+ myoblasts (Figure 3.16a). It is important to note that by E10.5, the migration of PAX3+ myogenic progenitors toward the forelimbs is in full effect278 and by this stage many PAX3+ cells can be observed within the limb bud proper268 (Figure 3.15b). On the other hand, HIC1+ MPs are mostly concentrated in the embryonic trunk at E10.5 with a notable absence of any expression of HIC1 within the limb buds. This suggests that the migration of myogenic progenitors from the dermomyotome precedes and is distinct from that of HIC1+ MPs into the forelimb buds. Similar to Hic1 lineage-traced cells (tdTomato+), HIC1+ cells did not overlap with PAX7+ myogenic progenitors (Figure 3.16a) nor with neural crest-derived precursors (Figure 3.16b-c). Taken together, these results indicate that HIC1+ MPs originate within the sclerotomal/syndetomal compartments of the somite and represent a cell population distinct from PAX3+ myogenic progenitors. 75 Figure 3.13 Immunofluorescence detection of HIC1 shows that HIC1+ MPs partially overlap with sclerotome and syndetome as early as E10.5. (a) Immunofluorescence staining with a-PAX1 on E10.5 embryos at the forelimb level. (b) Representative image of E10.5 ScxGFP embryo at the forelimb section. Scale bars, 100µm, 20µm inset. White dotted boxes in (a) and (b) highlight inset regions in a’ and b’, respectively. 76 Figure 3.14 Immunofluorescence detection of HIC1 shows that HIC1+ MPs and myogenic cells represent separate progenitor populations. (a) Immunofluorescence staining with a-PAX3 on E10.5 embryos at the forelimb level. (b) Immunofluorescence staining with a-PAX7 on E10.5 embryos at the forelimb level. Scale bars, 100µm, 20µm inset. White dotted boxes in (a) and (b) highlight inset regions in a’ and b’, respectively. 77 Figure 3.15 Immunofluorescence detection of HIC1 shows that HIC1+ MPs are in close proximity to extending nerve axons and are distinct from migratory myogenic progenitors. (a) Immunofluorescence staining with a-PAX7 on E10.5 embryos at the forelimb level shows the relative position of HIC1+ MPs with nerves and muscle precursors. Yellow dotted-lines outline the location of nerve axons. (b) Immunofluorescence staining with a-PAX3 on E10.5 embryos at the forelimb level shows that migrating PAX3+ cells are distinct from HIC1+ MPs. Scale bars, 100µm, 20µm inset. White dotted boxes in (a) and (b) highlight inset regions in a’ and b’, respectively. 78 Figure 3.16 HIC1+ MPs are distinct from SOX10+ neural crest progenitors and PAX7+ myogenic precursors. (a) Immunofluorescence staining with a-PAX7 and a-HIC1 on E10.5 embryos does not show any double-positive cells HIC1+. Dotted box outlines inset region a’. (b-c) Immunofluorescence staining with a-HIC1 and a-SOX10 on serial tissue sections of E10.5 embryos indicates that HIC1+ MPs and SOX10+ neural crest cells are distinct populations. Scale bars, 100um, 20um inset. D, dorsal; V, ventral. Dotted boxes highlight inset regions. 79 Lineage tracing preformed on Hic1CreERT2; R26tdTomato embryos revealed that although very few tdTomato+ cells are present in the forelimb at E11.5 (Figure 3.6), expansion rapidly increases and by E12.5, tdTomato+ cells were interspersed amongst the developing dorso-ventral muscle masses and their accompanying nerves (Figure 3.17a). Indeed, immunofluorescence (IF) analysis at E13.5 revealed abundant tdTomato+ cells within the dorsal and ventral muscle masses underlying the epithelium and circumscribing the developing chondrogenic anlagen (Figure 3.17b). However, the tdTomato+ cells did not contribute to the primary cartilaginous elements which is derived from the LPM (Figure 3.17b). Furthermore, expansion of the tdTomato+ compartment was associated with a proliferative phase in which up to 45.2 ± 2.1% (E12.5) of the tdTomato+ cells had incorporated EdU (Figure 3.18a-b). Taken together, these results suggest that tdTomato+ cells represent a unique MP subset within the developing limb. Figure 3.17 Distribution of tdTomato+ cells in E12.5 and E13.5 forelimbs. (a) Maximum intensity projection of E12.5 Hic1CreERT2; R26tdTomato embryo imaged and stained in wholemount with antibodies against NEFH, PAX7, and MF20. Identical to figure 3.7d. Scale bars, 500 µm and 100 µm. White box represents inset region highlighted in panel on right. Arrow heads indicate clusters of tdTomato+ cells. (b) Representative images of Hic1CreERT2; R26tdTomato forelimbs at E13.5, counterstained with CD31 and DAPI. Dotted box highlight inset region (b’). Scale bars, 100µm (top), 20 mm (bottom). White box represents inset region highlighted in bottom panels. D, distal; Dmm, dorsal muscle mass; Hc, pre-cartilage primordium of humerus; Pr, proximal; URc, radio-ulnar pre-cartilage primordium; Vmm, ventral muscle mass. 80 Figure 3.18 Proliferation dynamics of tdTomato+ cells in Hic1CreERT2; R26tdTomato embryonic forelimbs. (a) Representative FCM plots of tdTomato+ cells following a 24-hour EdU pulse-trace in Hic1CreERT2; R26tdTomato forelimbs at indicated time points. (b) FCM analysis of EdU incorporation assay on Hic1CreERT2; R26tdTomato forelimbs. n = 4-6 forelimb pairs per timepoint, data represent mean +/- SD, one-way ANOVA followed by Tukey’s multiple comparison test, ***P<0.001, ****P<0.0001. 3.2.3 ScRNA-seq and histological analyses reveal a unique MP hierarchy To characterize the molecular nature and developmental fates of tdTomato+ cells, scRNA-seq was performed on FACS-purified limb populations at 5 developmental timepoints ranging from E11.5 to E16.5 (Figure 3.19a). In total, 32,225 cells were profiled. Louvain-Jaccard clustering identified 9 distinct cell populations within the aggregated dataset and select cell-type markers were used to annotate the clusters (Figure 3.19b-c). Chronologically, the earliest populations (MPI, MPII) were composed mainly of cells from E11.5 (32%) and E12.5 (62%) (Figure 3.19d-e) and expressed progenitor cell markers such as Id genes279 (Figure 3.19c). Mitotic cells (MCs) formed a distinct 81 cluster enriched for proliferation-associated genes such as Birc5, Mki67, and Ccnb1 (Figure 3.19c). The majority of cells from E14.5 and E16.5 expressed cell lineage defining markers indicative of a more differentiated phenotype. They formed 6 different groups based on gene expression: fibroblasts (Fap, Fbn1), chondrocytes (Lect1, Col2a1), DFs (Col7a1, Rspo1), tenocytes (Tnmd, Htra3), and pericytes (Rgs5, Kcnj8) (Figure 3.19b-c). An intermediate cluster between MPs and fibroblasts was termed “fibrogenic progenitors” and expressed markers such as Osr1, Rspo3, and Hmcn1 (Figure 3.19b-c). Figure 3.19 Single-cell RNA-sequencing reveals multiple cell fates for tdTomato+ MPs throughout limb development. (a) Schematic of Hic1CreERT2; R26tdTomato sample collection for scRNA-seq. (b) UMAP of aggregated scRNA-seq data from sorted forelimb tdTomato+ cells coloured/numbered according to cluster identity determined by Louvain clustering. (c) Heatmap of scRNA-seq data showing enriched genes within each cluster. (d) UMAP of aggregated scRNA-seq data from sorted forelimb tdTomato+ cells colored according to embryonic timepoint. (e) Stacked bar graph illustrating the composition of each cluster with respect to embryonic timepoints. 82 To further refine the tdTomato+ cell lineage tree, the URD analytical tool257 was used to generate differentiation trajectories for the aggregated (E11.5 – E16.5) dataset. The tips, or terminal populations, were manually curated based on marker analysis (Figure 3.20a-b). Figure 3.20 URD-generated t-SNE plot of E16.5 tdTomato+ cells used to identify terminal cell fates. (a) Cluster map of E16.5 Hic1CreERT2; R26tdTomato sorted tdTomato+ cells projected in URD-generated t-SNE space. (b) Representative genes used to assign URD cluster identities and further determine the tips of the developmental trajectories. The reconstructed trajectory tree recapitulated the identified clusters and effectively ordered each of the populations from the intermediate timepoints (E12.5, E13.5, E14.5) in chronological order without a priori annotation (Figure 3.21a-b). The tree segments closest to the root represent earlier stages on the pseudotime trajectory and were enriched for MP cell markers such as Crabp2, Hmga2, Id1, Osr1, and Sox11 (Figure 3.22). The developmental trajectories that emanated from the E11.5 tdTomato+ population were resolved into 8 major distinct fates: chondrocytes (Sox9, Col2a1; Figure 3.21c)280, tenocytes (Scx, Tnmd ; Figure 3.21d)281, fascia (C1qtnf3, Mfap5, Clec3b; Figure 3.21e and 3.23a-b)72, periskeletal cells (perichondrium, Sox5, 83 Sox6, Dkk3; Figure 3.21e and 3.23c)282, perineurial cells (Cldn1, Wnt6; Figure 3.21f)136, fibroblasts (Cd34, Dpt; Figure 3.21g)282,283, pericytes (Rgs5, Acta2; Figure 3.21h)75, DFs (Axin2, En1, Lef1, Nkd2; Figure 3.2i)56,61. Interestingly, fibroblasts were also ramified into two subpopulations one of which (Fibro1) exclusively expressed Cxcl14, Pappa2, and Ptgfr (Figure 2.23d); and the other branch (Fibro2) was specified by expression of Ly6c1, Pi16, and Tek (Figure 3.23e). Similarly, the DF branch contained a bifurcation further specifying this cluster into PDFPs which express genes such as Prdm1, Sox18, and Foxd1 (Figure 3.23f), and RDFPs identified by expression of Cldn10, Sp5, and Lum (Figure 3.23g). 84 Figure 3.21 ScRNA-seq analyses define a mesenchymal progenitor hierarchy. (a) Force-directed layout of Hic1-lineage labelled MP differentiation trajectory, coloured according to developmental timepoint and optimized for 2D visualization. (b) URD-inferred developmental trajectories of Hic1 lineage-labelled forelimb MPs, coloured by developmental timepoint. Chon, chondrocytes; fibro, fibroblasts; PDFPs, papillary dermal fibroblast progenitors; perichon, perichondrium; PnC, perineural cells; RDFPs, reticular dermal fibroblast progenitors; teno, tenocytes. (c-i) Representative gene expression heatmaps projected on layout from (a), showing transcript abundance of specified mesenchymal cell fate markers. Blue lines in each plot indicate trajectory and the red branch highlights the tip/cell fate. 85 Figure 3.22 Cells located at the root express progenitor markers. Dendrograms illustrating gene expression intensity of progenitor-associated genes projected on the mesenchymal lineage hierarchy. 86 Figure 3.23 Characterization of embryonic Hic1+ MP-derived fibroblast populations. (a) URD-inferred developmental trajectories of Hic1 lineage-labelled forelimb MPs, fibroblast populations are highlighted by colored boxes. (b-g) Gene expression heatmaps projected on dendrogram branchpoints for each fibroblast subpopulation. Fibro1, fibroblast 1; Fibro2, fibroblast 2, RDFPs, reticular dermal fibroblast progenitors; PDFPs, papillary dermal fibroblast progenitors. 87 To experimentally interrogate the computationally inferred lineage trajectories, the fate of tdTomato+ cells was examined in fetal forelimbs. By E18.5, tdTomato+ clusters of chondrocytes and perichondral cells were observed concentrated in specific areas of the developing bones, such as the distal humerus (Figure 3.24a). TdTomato+ ScxGFP+ tenocytes were found mostly in short limb tendons, for example at the connection point between the biceps brachii and the radius (Figure 3.24b). The MTJs of such short tendons also contained high concentrations of tdTomato+ cells (Figure 3.24b, arrowheads). Numerous stromal compartments containing varying contributions of tdTomato+ cells in alignment with the in silico-derived lineages were also observed. TdTomato+ TAGLN+ cells, indicative of pericytes associated with capillaries75, were identified throughout the E18.5 forelimb (Figure 3.24c). TAGLN- perivascular cells were also ubiquitously observed. Similarly, tdTomato+ cells were identified circumscribing and within nerve axons of the appendicular peripheral nervous system (Figure 3.24d), these cells represent endoneurial and perineurial fibroblasts. Numerous tdTomato+ cells populated connective tissues of the limb skin including SOX2+ dermal papilla fibroblasts and SOX2- reticular fibroblasts62,284 (Figure 3.24e). In SkM, tdTomato+ Pdgfra+ FAPs or fibroblasts were abundant within the fascia, SkM interstitium and associated connective tissues41 (Figure 3.24f). In agreement with histological data concerning the origin of Hic1+ MPs, tdTomato+ cells did not contribute to the myogenic (Figure 3.25a-b) nor endothelial lineages (Figure 3.25c). Although there was an intimate association between tdTomato+/PdgfraGFP+ fibroblasts and SkM, no tdTomato+ myotubes were observed (Figure 3.25d). Similarly, although Hic1-lineage MPs have been shown to harness adipogenic potential45, no tdTomato/PLIN double-positive adipocytes were 88 Figure 3.24 Lineage tracing analyses confirm computationally inferred lineage potential. (a-f) Representative immunofluorescence images of E18.5 Hic1CreERT2; R26tdTomato forelimbs of tdTomato+ MP-derived cells demonstrating contribution to the following lineages: (a) chondrocytes and perichondrium. White arrowheads indicate perichondrium, red arrowheads highlight tdTomato+ chondrocytes; (b) tenocytes, arrowheads indicate cells at the MTJ; (c) pericytes; (d) perineurium and endoneurium cells. White arrowheads indicate endoneurial cells; (e) PDFPs and RDFPs highlighted by white and red arrowheads respectively; and (f) fibroblasts. White boxes in (a, e) highlight regions of interest that are magnified in adjoining panels. Scale bars, 20µm. 89 Figure 3.25 Embryonic Hic1+ progenitors do not contribute to skeletal muscle nor endothelial lineages. (a) Representative immunofluorescence image of an E14.5 Hic1CreERT2; R26tdTomato embryo (TAM at E10) stained with a-MyHC. Scale bar, 100µm. (b) UMAP plots of aggregated scRNA-seq data from Hic1CreERT2; R26tdTomato sorted tdTomato+ cells showing no detectable expression of myogenic regulatory factors (Myod1, Myf5, and Pax7) and embryonic myosin (Myh3). (c) UMAP plots of aggregated scRNA-seq data from Hic1CreERT2; R26tdTomato sorted tdTomato+ cells showing no detectable expression of endothelial cell markers (Podxl and Sox17). (d) Immunostaining of E18.5 Hic1CreERT2; R26tdTomato; PdgfraH2B-eGFP forelimbs show the spatial proximity of eGFP+ fibroblasts and skeletal muscle. Scale bars, 100µm. Dotted boxes highlight inset regions. 90 observed histologically. However, genes indicative of adipogenic lineage potential were widely expressed and included in areas of open chromatin within tdTomato+ MPs (Figure 3.26). Collectively, these findings reinforce that early Hic1+ MPs represent a hitherto undescribed mesenchymal cell population with multi-lineage potential. Figure 3.26 Adipogenic potential of tdTomato+ MPs at the single cell level. Chromatin accessibility and mRNA expression for adipogenic genes in tdTomato+ cells assessed via scATAC-seq and scRNA-seq, respectively. Genome tracks of relevant promoter regions are shown with corresponding gene expression heatmaps overlayed on the URD-generated lineage tree. Chondro, chondrocytes; fibro 1, fibroblasts 1; fibro 2, fibroblasts 2; PDFPs, papillary dermal fibroblast progenitors; peri; pericytes; perichon, perichondrium; RDFPs, reticular dermal fibroblast progenitors; teno, tenocytes. 91 3.2.4 Lineage potential of Hic1+ MPs is embedded in the chromatin landscape To further interrogate the lineage potential underlying the proposed cell hierarchy, epigenetic profiling using scATAC-seq was performed on a total of 41,066 single nuclei from sorted tdTomato+ forelimb cells captured at E12.5, E14.5, and E16.5 forelimbs (Figure 3.27a-b). K-means clustering (k=7) projected onto a t-stochastic neighbor-embedding (t-SNE) dimensionality reduction plot identified 6 main cell populations (Figure 3.27c). This analysis yielded two MP clusters: MPs I and MPs II. The majority of cells comprising MPs I were derived from the E12.5 limb sample (97%); similarly, cells from MPs II were mostly derived from E14.5 limbs (85%) (Figure 3.27d). MPs were found to have OCRs at gene loci associated with the cell cycle and progenitor status, such as Hic145, Id1279 and Snai1285 (Figure 3.27e-f). OCRs within lineage-defining genes corresponding to the aforementioned 8 major cell fates were all evident within this population (Figure 3.27g). For example, gene ranges encompassing the Sox9 (chondrogenic), Scx (tenogenic), Osr1 (fibro/adipogenic)55, and Lrig1 (dermal fibrogenic) loci contained OCRs (Figure 3.27g) indicative of broad mesenchymal lineage potential. With time, loci associated with alternative cell fate trajectories became differentially accessible as lineage-restricted populations emerged (Figure 3.27h-k). 92 Figure 3.27 A broad mesenchymal lineage potential is embedded in the epigenome of early tdTomato+ MPs. (a) Schematic of Hic1CreERT2; R26tdTomato sample collection for scATAC-seq. (b) t-SNE plot of aggregated scATAC-seq data from sorted tdTomato+ MPs colored according to embryonic timepoint. (c) t-SNE plot of aggregated scATAC-seq data from sorted tdTomato+ MPs colored according to cluster identity. (d) Stacked bar graph illustrating the composition of each cluster with respect to embryo timepoints. (e-k) Genome browser tracks from scATAC-seq analyses displaying the promoter sum signal near the indicated gene loci from clusters defined in panel (c). Grey boxes highlight clusters exhibiting promoter sum signal relevant to the indicated cell function or fate. 93 The largest population was identified as a collection of fibrogenic cell types with OCRs identified in loci including Ly6a, Postn, and Cd34 (Figure 3.27h). A dermal fibrogenic cluster consisted of cells containing OCRs within the Col7a1286, Rspo161, and Irx1287 loci (Figure 3.27i). The chondrogenic and tenogenic progenitor clusters were classified based on OCRs within Acan, Col2a1, and Chad, or Tnmd, Thbs4, and Kera loci, respectively (Figure 3.27j-k). Distinct pericyte and perineural clusters were not detected, although respective underlying OCRs (Kcnj8, Rgs5, Cldn1, Wnt6) were identified and these populations were found to be interspersed throughout the clusters. This likely reflects the fact that these rare populations are less efficiently resolved at these time points. As would be expected from a multipotent cell population upstream of a lineage trajectory, limited transcription from OCRs associated with the committed mesenchymal lineages was observed in MPs at E12.5 (Figure 3.28a-b). This potential is only realized in more committed progenitors and differentiated cells at E16.5 with the detection of the corresponding transcripts (Figure 3.28c), indicating that Hic1+ MP lineage potential is embedded within the epigenome. Comparisons between independently clustered E12.5 and E16.5 scRNA and scATAC-seq datasets (Figure 3.29a-d) revealed that genes associated with a progenitor status were widely expressed and the respective gene loci were accessible throughout all clusters of Hic1 MPs at E12.5 (Figure 3.29e). On the other hand, expression and accessibility of these genes became restricted to specific progenitor clusters at E16.5 (Figure 3.29f). Chondroprogenitors became apparent both transcriptionally and by chromatin state as early as E12.5 (Figure 3.29g); this cluster becomes established at E16.5 with highly specific OCRs and largescale expression of chondrocyte markers such as Acan and Chad (Figure 3.29h). DFs (Figure 3.29i-j), fibroblasts (Figure 3.29k-l), and tenocytes (Fig. 3.29m-n) displayed a similar trend: RNA expression of cell type-specific genes 94 were absent at E12.5 and chromatin accessibility peaks were ubiquitous throughout scATAC-seq clusters (Figure 3.29i, k, m). At 16.5, cells have differentiated and this becomes apparent in both gene expression and chromatin landscape (Figure 3.29j, l, n). Collectively, these studies reveal that the early Hic1+ MPs represent a mesenchymal cell progenitor population capable of giving rise to various subsets of skeletal and stromal cells. 95 Figure 3.28 Hic1+ MPs become compartmentalized by gene expression at E16.5. (a) genome browser tracks displaying the promoter sum signals for diverse lineage-associated genes from scATAC-seq analyses of E12.5 and E16.5 samples. (b-c) UMAP plots of scRNA-seq expression data of genes in panel a. b, E12.5; c, E16.5. 96 Figure 3.29 MP specification and differentiation are reflected in chromatin and transcriptome dynamics during their developmental trajectories. (a) UMAP of scRNA-seq data from E12.5 sorted limb tdTomato+ cells colored according to cluster identity. (b) t-SNE plot of scATAC-seq data from E12.5 sorted tdTomato+ cells colored according to cluster identity. (c) UMAP of scRNA-seq data from E16.5 sorted limb tdTomato+ cells colored according to cluster identity. (d) t-SNE plot of scATAC-seq data from E16.5 sorted tdTomato+ cells colored according to cluster identity. (e-f) Violin plots and genome tracks from representative genes for identification of MPs. (g-h) Violin plots and genome tracks for identification of chondrocytes. (i-j) Violin plots and genome tracks for identification of DFs. (k-l) Violin plots and genome tracks for identification of fibroblasts. (m-n) Violin plots and genome tracks for identification of tenocytes. 97 3.2.5 Embryonic Hic1+ MP post-natal lineage contributions Hic1+ MP lineages emerge at various times during embryogenesis, where they contribute to a variety of mesenchymal cell types. To further interrogate Hic1+ MP lineage contributions, scRNA-seq was performed on tdTomato-sorted SkM samples collected at postnatal day (P) 56 following E10 TAM administration along with histological analysis of SkM and surrounding tissues (Figure 3.30a). This approach would allow lineage tracing of embryonic Hic1 MPs postnatally and into adulthood. Bioinformatic analyses revealed six major tdTomato+ populations present in SkM (Figure 3.30b-c), the largest of which comprised fibroblasts (Cd34+, Pdgfra+, Ly6a+, Dpt+) which contained Dpt+, Col15a1+ (fibroblast 1) and Dpt+, Pi16+ (fibroblast 2) subpopulations (Figure 3.20c). Fibroblasts were abundant in the SkM interstitium (Figure 3.30d). Tenocytes (Scx+, Tnmd+) and myotenocytes (Col22a1+) were also identified (Figure 3.30e). Three clusters constituted vasculature-associated cell types and were classified based on gene expression as: VSMCs (Acta2+, Cnn1+, Smtn+) which lined arteries (Figure 3.30f) pericytes (Abcc9+, Kcnj8+, Rgs5+) circumscribed capillaries (Figure 3.30g) and smooth muscle cell progenitors (SMCPs) (Ifitm1+, Nr4a1+, Thy1+) were found associated with larger blood vessels (Figure 3.30h). A closer analysis of these vascular-associated cell types revealed commonalities and distinctions in transcriptome signatures for each cluster (Figure 3.31a-b). The VSMC cluster expressed bona fide VSMC genes such as Acta2, Myh11, and Synm288 (Figure 3.31c), but also shared genetic features with SMCPs for example: Cyr61, Eln, and Id3 (Figure 3.31e). These common genes have been previously identified in VSMCs288. Other than Abcc9+, Kcnj8+, and Rgs5+, which are typically pericyte specific markers, the pericyte cluster expressed unique genes such as Prex2 and Trpc3 (Figure 3.31d) which have recently become relevant as pericyte identifiers in scRNA-seq 98 datasets288. Similar to VSMCs, pericytes also shared genetic features with SMCPs, Cxcl12, Steap4, and Vtn are examples. The significant overlap between SMCPs and pericytes/VSMCs strengthens their identification as progenitors. Indeed, Thy1 has recently been described as a marker (among others) of smooth muscle and pericyte progenitor cells located in the adventitia of larger vessels289. Finally, among the vascular-associated cell types, Hic1 was found to be exclusive to pericytes and SMCPs (Figure 3.31d), further reinforcing their progenitor status. 99 Figure 3.30 Embryonic Hic1+ MPs give rise to distinct populations of adult stromal cells. (a) Schematic representation of strategy employed for lineage tracing embryonic Hic1+ MPs into adulthood. (b) UMAP colored by cluster of scRNA-seq data from sorted tdTomato+ cells isolated from the TA muscle of lineage-traced Hic1CreERT2; R26tdTomato mice. (c) Heatmap of scRNA-seq data showing enriched genes within each cluster. (d-h) Representative immunofluorescence images and violin plots of each cell population identified through lineage tracing performed from E10 until P56 on Hic1CreERT2; R26tdTomato mice. TdTomato+ populations persist as cells from the following lineages: fibroblasts (d); tenocytes and myotenocytes (e); pericytes (f), vascular smooth muscle cells (g); and smooth muscle cell progenitors (h). White arrowheads highlight cells of interest. White dotted boxes highlight regions that are magnified in larger panels (d’ and g’). Scale bars, 50µm. (i-k) Violin plots of select transcripts. 100 Figure 3.31 Characterization of mural cell fates. (a) UMAP colored by cluster of scRNA-seq data from sorted tdTomato+ cells isolated from the TA muscle of lineage-traced Hic1CreERT2; R26tdTomato mice. Dotted line encircles endothelium-associated clusters (VSMCs, pericytes, and SMCPs). (b) Heatmap of differentially expressed genes between VSMCs, pericytes, and SMCPs clusters. (c-e) Violin plots for expression of representative genes from each cluster indicated in (a). 101 Similar to previous findings45, Hic1 was expressed solely in fibroblasts, pericytes and SMCPs within adult SkM (Figure 3.30i). Pdgfrb was expressed to varying extents in all Hic1 lineage cell populations including Ly6a fibroblasts (Figure 3.30j,k). Perineural and endoneurial cells (Fzd2+, Ptch1+, Wnt6+) clustered separately in transcriptomic space and were observed in histological nerve samples (Figure 3.32a). Utilization of a Hic1CreERT2; R26tdTomato; Ptch1LacZ mouse line allowed for the in situ identification of Ptch1/tdTomato double-positive cells in the perineural compartment of limb nerves (Figure 3.32b). It is worth mentioning that other tdTomato+ connective tissue cell types were also present in histological samples of adult lineage-traced limbs; however, since only SkM was analyzed at the single-cell RNA-seq level, these cells were not described in the scRNA-seq analysis. These connective tissue cell types consisted in a variety of cell fates and in several anatomical locations such as cortical osteocytes in the humeral shaft (Figure 3.33a), BMSCs within the distal epiphysis of the humerus (Figure 3.33b), subcutaneous white fat throughout the limb hypodermis (Figure 3.33c), DFs were distributed throughout the reticular and papillary dermal layers (Figure 3.33d), and mineralized fibrocartilage within the tendon attachment site at the proximal head of the humerus (Figure 3.33e). Together, these analyses reveal that E10-labelled Hic1+ MPs contribute to select post-natal mesenchymal lineages across multiple tissues. 102 Figure 3.32 Histological confirmation of perineural fate for Hic1-lineage MPs. (a) Representative immunofluorescence image of tdTomato+ perineural and endoneurial cells identified through lineage tracing performed from E10 until P56 on Hic1CreERT2; R26tdTomato mice. Scale bars, 50µm (b) Expression of Ptch1 in tdTomato+ perineural fibroblasts lineage traced during adulthood in Hic1CreERT2; R26tdTomato; Ptch1Lacz mice. Scale bars, 20µm. White arrowheads highlight cells of interest. (c-e) Violin plots for selected genes representative of perineural cells. 103 Figure 3.33 Embryonic Hic1+ MPs contribute to diverse cell types that persist into adulthood. (a-e) Representative immunofluorescence images of E10-P56 limb tissues demonstrating contribution of Hic1+ MPs to the following lineages: osteocytes (a); BMSCs (b), subcutaneous adipocytes (c), DFs (d); and fibrocartilage (e). White arrowheads indicate cells of interest. White dotted-boxes in (b-f) highlights regions of interest that are magnified in adjoining panels. Scale bars, 20µm. BM, bone marrow; CB, cortical bone; TB, trabecular bone. 104 3.2.6 Embryonic Hic1+ fibroblasts persist into adulthood and have roles in SkM regeneration To determine contribution of E10-labelled Hic1+ MPs to post-natal SkM homeostasis and regeneration, scRNA-seq profiling and NTX-induced SkM injuries were performed in adult pulse (TAM at P56) (Figure 3.34a) and embryonic pulse (TAM at E10) (Figure 3.34b), chase experimental models. Single-cell transcriptomic data from adult pulsed (P56 – collect P70)45 and embryo pulsed (E10 – collect P70) SkM resident tdTomato+ cells were aggregated and clusters were annotated based on marker gene expression (Figure 3.34c-d). This experimental approach would allow for a direct comparison of the transcriptomic profiles between embryonic Hic1+ cells and adult Hic1+ cells in SkM under homeostasis. Both the adult and embryonic lineage-traced populations contained a large cluster of Ly6a+, Pdgfra+ fibroblasts (Figure 3.34e). Significant clusters consisting of Scx+, Tnmd+ tenocytes and Abcc9+, Kcnj8+ pericytes were also prevalent in both time courses (Figure 3.34f-g). SMCPs were most prevalent (66%) in cells labelled during embryogenesis (E10 – P70) and appeared to link the perycitic and VSMC populations in the 2D UMAP (Figure 3.34i). Interestingly, VSMCs were almost exclusively (94%) observed in the embryonic-pulse adult chase (E10 - P70) experiment (Figure 3.34h and 3.34k) and the absence of Hic1 expression in this tdTomato+ population reflects their more differentiated state (Figure 3.34l). A small population of perineurial fibroblasts was also found to be composed of >80% cells from the embryonic pulse experiment (Figure 3.34j). Hic1+ fibroblasts are defined by the markers Ly6a and Cd3440 and emerge at ~E16.5. Their origins and scope of contribution to embryonic development are unclear, however, studies have certainly 105 revealed a fundamental role for this population in SkM development38,55 as well as post-natal SkM homeostasis and regeneration42. Labelling at E10 yields comparable numbers of tdTomato+ cells (SkM-associated fibroblasts) to labelling at P56 (Figure 3.34k), revealing that Hic1+ MPs are a major source of post-natal fibroblasts including the two universal fibroblast populations196, Dpt+ Pi16+ and Dpt+ Col15a1+. Furthermore, consistent with prior findings40,41,45,48, NTX-induced SkM injury of embryonic-pulse (E10 – P70) Hic1CreERT2;R26tdTomato mice leads to a marked expansion of fibroblasts (Figure 3.35 a-b), which is comparable to that of Hic1+ cells labelled during adulthood (Figure 3.35c-d). Overall, the labelled cells display a similar post-damage proliferative phase followed by a return to baseline numbers (Figure 3.35e). These findings indicate that embryonic Hic1+ MPs give rise to fibroblasts which persist into adulthood and contribute to muscle repair and regeneration (Figure 3.35f). 106 Figure 3.34 Lineage tracing of embryonic Hic1+ MPs into adulthood reveals development-specific cell types. (a) Schematic of experimental timeline for lineage tracing of adult MPs. (b) Schematic of experimental timeline employed for post-natal lineage tracing of Hic1CreERT2; R26tdTomato embryos. (c) UMAP of aggregated scRNA-seq data from adult chase and embryonic chase tdTomato+ MPs, colored according to cluster ID. PnCs, perineural cells; SMCPs, smooth muscle cell progenitors; VSMCs, vascular smooth muscle cells. (d) UMAP of aggregated scRNA-seq data from adult chase and embryonic chase tdTomato+ MPs, colored according to TAM dose timepoint. (e-j) UMAP plots for representative genes used to identify each cluster. (k) Stacked bar graph illustrating the composition of each cluster with respect to time of TAM dose. (l) UMAP plots for Hic1 and tdTomato transcripts within the aggregated scRNA-seq data. 107 Figure 3.35 Embryonic Hic1-lineage FAPs persist postnatally and support muscle regeneration. (a) Schematic of experimental design for lineage tracing of embryonic Hic1+ progenitors into adulthood and subsequent muscle injury. (b) Representative tibialis anterior (TA) muscle sections from embryonic lineage-traced (TAM at E10) Hic1CreERT2; R26tdTomato mice at 3- and 14-days post NTX-induced injury. Contralateral TA shown as control. Scale bars, 100µm, 20µm inset. (c) Schematic of experimental design for lineage tracing of adult Hic1+ progenitors and subsequent muscle injury. (d) Representative tibialis anterior (TA) muscle sections from adult lineage-traced (TAM at P56) Hic1CreERT2; R26tdTomato mice at 3- and 14-days post NTX-induced injury. Contralateral TA shown as control. Scale bars, 100µm, 20 µm inset. (e) Quantification of tdTomato+ cell abundance in embryonic and adult lineage tracing models. (f) Graphical summary of the experimental results. Embryonic Hic1+ cells generate adult SkM-associated fibroblasts which become activated upon injury, cell number returns to pre-injury levels following injury resolution. 108 Chapter 4: Hic1 identifies a population of embryonic chondro-tenoprogenitors responsible for secondary bone morphogenesis 4.1 Introduction The developing limb relies on a multitude of cell types, signaling events, and cellular interactions converging to form a functional appendage. Nowhere is this interplay more evident than in the development of the limb musculoskeletal system where muscles, bones, and tendons must develop cooperatively and in tandem. Studies in birds and mice have shown that muscle and skeletal progenitors originate from differing embryonic sources262,263,290,291. Whereas bone and tendon progenitors derive from LPM, myogenic progenitors are somite-derived and migrate into the early limb primordium from hypaxial dermomyotomes. Timing is also an essential factor, myogenic progenitors initiate migration into the forelimb bud as early as E10278. At the same time, tenogenic progenitors (Scx+) become apparent in the somites and by E10.5 in the limb buds as well2. The myogenic and tenogenic domains of the somite (myotome and syndetome, respectively), while in close contact, do not overlap. Similarly, once within the limb bud, Scx+ and Myod1+ progenitors appear to be juxtaposed throughout the limb bud mesenchyme but always remain as separate progenitor populations2. Nevertheless, during musculoskeletogenesis, LPM-derived tenocytes must converge with somite-derived myocytes to form the MTJ and on the opposite end, tendons connect to bones at tendon-to-bone attachment sites which will later mature into functional entheses. In this way, musculoskeletogenesis effectively establishes a gradient of 109 material properties required for stress dissipation and efficient energy transfer from force-generating muscles through force-transmitting tendons to the load-bearing skeletal framework. Enthesis development has only recently began to be studied closely. In early limb skeletogenesis, LPM-derived mesenchymal cells condense and differentiate into chondrocytes to form the cartilage anlagen of what will eventually become bone. At this stage, all chondrocytes express the transcription factor Sox926–28. Lineage tracing experiments utilizing a Sox9-Cre mouse line have shown that Sox9+ mesenchymal cells are not only chondrocyte progenitors but also give rise to osteoblasts and tenocytes in the developing mouse limb292,293. Enthesis formation begins at this stage, when the primary cartilaginous template is morphologically modified by a distinct population of LPM-derived Scx+ Sox9+ progenitors. These cells give the premature cylindrical template a mature form via the addition of superstructures (eminences) to which tendons will attach25. This method of bone morphogenesis has been termed the ‘modular’ model since after a primary morphogenesis event in which Sox9+ progenitors form the initial featureless cartilaginous template of a bone, a secondary pool of Sox9+ Scx+ progenitors ‘add on’ the eminences to the cartilage anlage25. The molecular intricacies of enthesis morphogenesis are still being elucidated. TGFb signaling has been suggested to be required for the specification of attachment-unit progenitors and their subsequent differentiation has been linked to a SCX/BMP4 pathway25. Indeed, interfering with either Scx or Bmp4 expression has been shown to arrest the formation of tendon-bone-attachment units in the limb29. There is also a reliance on mechanical stimuli for enthesis maintenance and post-natal maturation19,21. 110 On the opposite end of the tendon, at the muscle attachment site, myotendinous junctions (MTJs) begin to develop during embryogenesis in mice and mature post-natally294. Studies in chick and zebrafish have elucidated the series of cellular events that entail MTJ formation10,294. The first indications of MTJ development are close interactions between myogenic cells and fibroblasts, followed by an accumulation of filamentous ECM proteins between muscle fibers and tendon cells, this protein secretion gives rise to a basement membrane at the interface between myotubes and tendon10. This basement membrane is composed mainly of laminins and collagen type IV, and it appears first at the prospective MTJ and extends to the entirety of the muscle cell thereafter294 . Initially, the deposited ECM proteins are sparce and have no common orientations. However, during the late embryonic and early neonatal periods, the fibrils will organize into mono-oriented fibril bundles in response to the contractile force of the developing muscle295. A fully-developed MTJ consists of finger-like extensions and invaginations of the muscle sarcolemma where actin filaments of the muscle cell cytoskeleton indirectly interact with extracellular components through transmembrane protein complexes, namely laminins and integrins296. The finger-like extensions serve to increase surface contact area between the myocytes and tendon in order to resist the muscle contraction forces10. Interestingly, these interdigitations between the muscle sarcolemma and tendon matrix shorten with age, often resulting in muscular atrophy297. Although MTJs are crucial components of the musculoskeletal system, little is known about the molecular mechanisms involved in MTJ formation. However, recent publications have suggested an important role for BMP-signaling and BMP-induced trans differentiation of muscle connective tissue (MCT) fibroblasts into myocytes in the establishment of functional MTJs during development11,12. Further work is required to fully unravel the molecular intricacies and cellular interplay that underlie the formation of such a unique biomechanical tissue. 111 Although maturation of both MTJs and entheses takes place postnatally, the early molecular and cellular events that initiate MTJ and enthesis development take place within the embryonic limb primordium. At this stage, the limb bud consists of a thin ectodermal sheath surrounding a pool of seemingly-homogenous mesenchyme. This perceived homogeneity at the macroscopic level is quickly revealed to be more complex when analyzed at a deeper level with cutting-edge techniques such as scRNA-seq, as the early limb mesenchyme contains a diverse agglomeration of progenitor cells such as myogenic, tenogenic, chondrogenic progenitors, among others. The development of a functional limb skeleton requires interactions between such muscle, tendon, and cartilage progenitors. It is unclear how Hic1+ progenitors figure within these embryonic cell populations and in the context of skeletal development. In this chapter, I show that Hic1+ MPs identified through the Hic1CrERT2; R26RtdTomato mouse line represent a cell population in the embryonic limb bud containing chondro- and teno-progenitors that directly contribute to the formation of tendon-bone attachment units and MTJs of the developing murine appendicular skeleton. I further demonstrate that perturbation of Hic1+ progenitor cells via both conventional and conditional Hic1 knockout mouse models, results in severe defects in secondary bone morphogenesis culminating in skeletal abnormalities. 4.2 Results 4.2.1 Hic1 identifies a subpopulation of ScxGFP+ MPs in the developing limb bud As mentioned in the previous chapter, in E10.75 embryos, Hic1 appears to be confined to the somitic mesoderm within the epaxial and hypaxial sclerotome compartments underlying the 112 dermomyotomes166. In this chapter, a closer look will be given to the expression of Hic1 in the embryonic forelimb exclusively. It is not until E11.5 that Hic1 first becomes evident within the limb bud. Hic1 initially exhibits limited and regionalized expression that is restricted to the proximal limb mesenchyme. At this stage, LacZ+ cells are well-established within the forelimb bud and begin to become apparent within the proximal hindlimb (Figure 4.1). Patterns formed by clusters of LacZ+ cells in the forelimb bud consist in dorsal and ventral streams along with a central core (Figure 4.1 and Figure 4.2a), this pattern persists until E12.5 with a continued aggregation of LacZ+ cells (Figure 4.1 and Figure 4.2b). At later stages, Hic1+ cells were visualized throughout the limb mesenchyme and circumscribed various chondrogenic anlagen (Figure 4.2c). Consistent with the interstitial distribution of the Hic1+ MP population in adult muscle, Hic1+ cells could also be detected within the emerging ventral and dorsal muscle masses (Figure 4.2d). Although LacZ+ cells were prevalent within the embryonic limb mesenchyme and surround the cartilage anlagen and muscle masses, no LacZ+ chondrocytes were observed (Figure 4.2d) indicating that expression of Hic1 is lost upon acquiring the chondrogenic fate. Indeed, micromass cultures of Hic1nLac/+ forelimbs showed an absence of LacZ+ cells within cartilage nodules; however, LacZ+ cells were abundant in the internodular regions (Figure 4.3a). Furthermore, treatment with pro-chondrogenic agents such as a retinoic acid receptor (RAR) pan-antagonist (AGN194310) or BMP4228,230 increased cartilage nodule formation and abrogated Hic1 expression (Figure 4.3a). In contrast, treatment with an RAR agonist (EC23) inhibited cartilage nodule formation. It should be noted that RAR agonist treatment markedly increased Hic1 expression as indicated by magenta-gal staining (Figure 4.3a). Similar micromass experiments performed with lineage traced Hic1CreERT2; R26tdTomato limb buds showed tdTomato+ cells within the cartilage nodules (Figure 4.3b). 113 Figure 4.1 Hic1nLacZ embryos show strips of LacZ+ cells extending from the hypaxial somites into the forelimb bud. Wholemount X-gal stained Hic1nLacZ/+ embryos show the gross distribution of Hic1+ cells in E11.5 and E12.5 limb buds. LacZ+ cells form dorsal and ventral clusters with a central core becoming apparent by E12.5. Red arrowheads indicate Hic1-expression region within the limb bud. Di, distal; Do, dorsal; P, proximal; V, ventral. Scale bars, 1mm.114 Figure 4.2 Characterization of Hic1 expression patterns in the embryonic limb bud. (a-c) Images of wholemount and transverse sections from Hic1nLacZ/+ embryonic forelimbs at early timepoints stained for nuclear LacZ expression. Sections were counterstained with NFR. Black arrowheads highlight two groups of LacZ+ cells. Scale bars, 1mm wholemount; 100 µm sections. (d) Representative image of LacZ-immunofluorescence co-stained E13.5 forelimb counterstained with DAPI. Scale bar 100µm. Ac, autopod pre-cartilage primordium; Hc, pre-cartilage primordium of humerus; URc, radio-ulnar pre-cartilage primordium; Dmm, dorsal muscle mass; Vmm, ventral muscle mass. 115 Figure 4.3 Hic1+ MPs localize to the internodular mesenchyme in limb micromass cultures. (a) Representative images of micromass cultures from E11.5 Hic1nLacZ/+ limb buds treated with EC23 (100 nM), AGN194310 (100 nM), or recombinant BMP4 (20 ng/ml) and co-stained with magenta-gal for detection of b-galactosidase activity and Alcian blue for visualization of cartilage nodules. Scale bar, 100 µm. (b) Representative confocal maximum intensity projections of micromass cultures from E11.5 Hic1CreERT2; R26tdTomato limb buds treated with compounds described in (a) and stained with antibodies against type II collagen. Scale bar, 100 µm. 116 Interestingly, this fate was not modified by treatment with chondrogenic agents (Figure 4.3b). Treatment of tdTomato+ cell-containing cultures with an RAR agonist did not yield an increased number of tdTomato+ cells indicating that CreERT2-induced recombination occurred prior to culture initiation. In summary, these findings indicate that Hic1 identifies a distinct cell population within the embryonic limb mesenchyme, a subset of which possess chondrogenic potential. Experiments described in chapter 3 demonstrated that Hic1+ MPs give rise to cartilage and tendon in the developing limb. The interesting distribution of Hic1nLacZ MPs in micromass cultures and the sensitivity of Hic1 expression to chondrogenic agents provoked interest in further exploring the events underlying Hic1+ MP differentiation into the chondrogenic and tenogenic fates during limb development. To explore the relationship between Hic1+ and tendon progenitor cells in the developing limb, a lineage tracing strategy was employed involving the Hic1CreERT2; R26tdTomato; ScxGFP mouse line with a focus on the embryonic forelimb. TAM administration at E10 allowed for the identification of rare tdTomato+ cells that were intermingled with ScxGFP+ cells at E11.75 (Figure 4.4a). FCM analysis at E11.5 and E13.5, demonstrated that tdTomato+ cells represent 1.0 ± 0.1% and 10.2 ± 1.3% of the ScxGFP+ cell population, respectively (Figure 4.4b-c). At the axial level at E11.5, tdTomato+ cells were concentrated in repeating segments along the antero-posterior axis of the embryo spanning from forelimb to hindlimb (Figure 4.5a) and overlapping with the ScxGFP+ somitic syndetomes but not with myotomes (Figure 4.5b). Micromass cultures of Hic1CreERT2; R26tdTomato forelimbs at E11.5 include myogenic progenitors that are present within the forelimb bud at that timepoint. As previously mentioned, in these cultures, tdTomato+ cells are located within the internodular regions and also circumscribing 117 cartilage nodules. Similarly, after ~2 days in culture, myocytes become apparent in these internodular regions (Figure 4.6). Figure 4.4 Characterization and quantification of tdTomato/ScxGFP double-positive cells in Hic1CreERT2; R26tdTomato; ScxGFP forelimbs. (a) Maximum intensity projection of wholemount E11.75 Hic1CreERT2; R26tdTomato; ScxGFP embryo. White dotted-line outlines the forelimb; red arrowhead highlights tdTomato+ cells within the forelimb. Scale bar, 500 µm. (b-c) FCM frequency analysis of dissociated Hic1CreERT2; R26tdTomato; ScxGFP forelimbs at E11.5 and E13.5. Data in (c) represent mean ± SD. 118 Figure 4.5 Spatial characterization of tdTomato+/ScxGFP+ cells in E11.5 Hic1CreERT2; R26tdTomato; ScxGFP embryos. (a) Wholemount fluorescence microscopy images of an E11.5 Hic1CreERT2; R26tdTomato; ScxGFP embryo. White box highlights forelimb region magnified in bottom (a’) panels. Scale bars, 1mm; 100µm inset. (b) Fluorescence microscopy images of Hic1CreERT2; R26tdTomato; ScxGFP embryo sagittal sections immunostained to identify myocytes. White box illustrates inset region in (b’). Scale bars, 100µm; 20µm inset.119 Figure 4.6 TdTomato+ MPs and myocytes localize to internodular regions in PLM cultures. (a) Representative fluorescent images of micromass cultures from E11.5 Hic1CreERT2; R26tdTomato limb buds stained with anti-myosin heavy chain. (b) Representative fluorescent images of micromass cultures from E11.5 Hic1CreERT2; R26tdTomato limb buds stained with anti-myosin heavy chain and treated with BMP4. Scale bars, 100µm. In congruence with tdTomato-cell distribution in the embryonic trunk, tdTomato+ cells in micromass cultures are in close proximity with myocytes yet do not contribute to the myogenic compartment (Figure 4.6). TdTomato+ MPs first appear at the proximal forelimb bud at E11.5. At this time, ScxGFP/tdTomato double-positive cells can be observed in histological sections of Hic1CreERT2; R26tdTomato; ScxGFP embryos (Figure 4.7a) and constitute approximately two-thirds (66%) of the tdTomato+ subpopulation (Figure 4.7b-c). One day later, at E12.5, clusters of double-positive cells are found associated with the chondrogenic anlagen (Figure 4.7d) and by E13.5, 85% of the tdTomato+ cells also express ScxGFP (Figure 4.7e-f). Collectively, these results indicate that tdTomato+ MPs entering the forelimb bud at E11.5 express the tenogenic master regulatory factor Scx (at a 2:1 ratio), display a modest contribution to the cartilage nodules in ex vivo limb cultures and are not myogenic progenitors. As development proceeds, ScxGFP/tdTomato double-120 positive cells become more abundant and are located in close proximity to the chondrogenic anlagen. To further characterize Hic1+ tenogenic progenitors in the embryonic forelimb, scRNA-seq was performed on FACS-purified tdTomato+ cells from Hic1CreERT2; R26tdTomato forelimbs at two early developmental timepoints (Figure 4.8a). The resulting single-cell transcriptomes from E11.5 (n = 3,208) and E12.5 (n = 4,722) samples were aggregated and queried for gene expression. Visualization of the data set in a 2D UMAP reveals that the cells from E11.5 and 12.5 coalesce to form a single main partition (Figure 4.8b). Louvain-Jaccard clustering categorized the dataset into 9 clusters mainly due to differences in expression of ECM-associated proteins (Col3a1, Lum, Dcn, Matn4), cellular signaling (Rspo3, Igf1, Ptn, Nbl1), and transcription factors associated with various developmental processes (Meis2, Ebf2, Tbx3, Osr2) (Figure 4.8c-d). However, all clusters expressed genes typical of embryonic mesenchyme such as Id1, Id2, Nnat, and Snai1 (Figure 4.8e). In contrast, genes indicative of LPM (Lhx9, Msx2) or myogenesis (Myod1, Pax3) were largely absent (Figure 4.8f-g). Gene markers of adult (Hic1, Pdgfra) and embryonic (Osr1, Tcf7l2) MPs were also broadly expressed (Figure 4.8h). Interestingly, chondrogenic (Col2a1, Sox5, Sox6, Sox9) and tenogenic (Egr1, Mkx, Scx, Tgfb2) markers largely overlapped and were present in both E11.5 and E12.5 cells (Figure 4.8i-j). Together, these results suggest that Hic1 identifies a subset of MPs within the embryonic limb mesenchyme that express determinants of both chondrogenic and tenogenic differentiation. 121 4.2.2 Hic1+ MP derivatives populate musculoskeletal attachment units Transcriptomic analysis of E11.5 and E12.5 tdTomato+ MPs revealed a possible contribution to the chondrogenic and tenogenic lineages. Unexpectedly, within this compartment, visualization of tdTomato+ cells at later timepoints (E14.5, E16.5, E18.5) revealed that they were limited to attachment units between muscle-tendon and tendon-bone, and not the tendon bodies nor primary chondrogenic anlagen. To further examine the unusual distribution of tdTomato+ cells in the developing appendicular skeleton, a lineage tracing strategy involving the Hic1CreERT2; R26tdTomato; ScxGFP was used. This strategy consisted in labelling Hic1+ cells at E10 and harvesting forelimbs at E18.5 to analyze the distribution of tdTomato+ cells using fluorescence microscopy (Figure 4.9a). To provide anatomical context for interpreting the contribution of tdTomato+ cells to the developing forelimb, muscles and bones of interest were highlighted as 3D-renders generated using the mouse limb anatomy atlas298 (Figure 4.9b). The distal epiphysis of the humerus, which contains multiple tendon-bone attachment units, exhibited an abundance of tdTomato+ chondrocytes (Figure 4.9c). In this region however, tdTomato+ chondrocytes were limited to the medial epicondyle and formed a clear demarcation between the tuberosity and the LPM-derived primary humeral cartilage (Figure 4.9c). Tendons emanating from this attachment site were double-positive for ScxGFP and tdTomato (Figure 4.9c, arrowheads). Hic1-lineage MPs were also found to form a major portion of the olecranon. Comparable to the medial epicondyle, tdTomato+ chondrocytes in this region were limited to the tuberosity and were especially concentrated in the regions in close contact with the nascent attachment unit (Figure 4.9d). Similarly, the tendons and fibrocartilage immediate to the olecranon were largely composed of tdTomato+ cells (Figure 4.9d). The distal-most skeletal elements (autopod) also contained several 122 Figure 4.7 Quantification of ScxGFP+ cells within the tdTomato+ MP population. (a) Representative images of Hic1CreERT2; R26tdTomato; ScxGFP forelimbs at E11.5. Scale bars, 100 µm; 20 µm inset. White box represents inset regions. (b) Representative plot from FCM analysis of E11.5 forelimbs gated for the quantification of ScxGFP+/tdTomato+ cells relative to the tdTomato+ population. (c) Pie chart generated from the FCM-based average cell counts of 3 independent E11.5 Hic1CreERT2; R26tdTomato; ScxGFP litters. (d) Representative images of Hic1CreERT2; R26tdTomato; ScxGFP forelimbs at E12.5. Scale bars, 100 µm; 20 µm inset. White box represents inset regions. (e) Representative plot from FCM analysis of E12.5 forelimbs gated for the quantification of ScxGFP+/tdTomato+ cells relative to the tdTomato+ population. (f) Pie chart generated from the FCM-based average cell counts of 3 independent E13.5 Hic1CreERT2; R26tdTomato; ScxGFP litters. White arrowheads in (a) and (d) indicate tdTomato+, ScxGFP+ cells; yellow arrowheads indicate tdTomato+ cells. 123 Figure 4.8 Early forelimb mesenchyme contains Hic1+ chondro-tenogenic MPs. (a) Graphical representation of experimental strategy used for scRNA-seq of FACS-purified tdTomato+ MPs from E11.5 and E12.5 forelimbs. (b) UMAP plot showing aggregated populations of tdTomato+ MPs isolated at E11.5 and E12.5. (c) Dot plot illustrating the highest-expressed genes per cluster from tdTomato+ MPs isolated from E11.5 and E12.5 Hic1CreERT2; R26tdTomato; ScxGFP forelimbs. (d) UMAP visualization of clusters for the E11.5 and E12.5 aggregated dataset. (e) Violin plots showing expression level of MP-specific genes per cluster. (f) UMAP visualization of LPM-specific genes. (g) UMAP visualization of muscle-specific genes. (h) UMAP visualization of genes expressed in embryonic MPs. (i) UMAP plots of genes involved in chondrogenesis. (j) UMAP plots of genes involved in tenogenesis. 124 clusters of tdTomato+ chondrocytes which were prevalent in tendon-bone attachment units of interphalangeal joints (Figure 4.9e). An analogous distribution of tdTomato+ cells was observed in the tendon compartment. In the olecranon, a majority of cells at the triceps brachii insertion were also tdTomato+ (Figure 4.9f). Likewise, the attachment sites of the tendons connecting the biceps brachii to the radial tuberosity and the supraspinatus muscle to the greater tubercle of the humerus were appreciably composed of tdTomato+ cells (Figure 4.9g-h). Similar to the examples illustrated above, Hic1-lineage MPs also contributed substantially to other tendon attachment sites of the forelimb but were largely absent from the tendon body (Figure 4.10a). To appreciate the contribution of tdTomato+ cells to the entire humeral cartilage, wholemount clearing and confocal microscopy was carried out on E10-18.5 humeri. As expected, tdTomato+ chondrocytes were highly concentrated at the medial and lateral epicondyles (Figure 4.10b). It is worth mentioning that the opposing side of the humerus, the humeral head which articulates with the scapula and forms the shoulder joint, is entirely tomato-negative (Figure 4.10b, dotted lines). These results highlight the contribution of tdTomato+ cells to tendon attachment sites (MTJ and tendon-bone) and their associated cartilage. The appendicular skeleton comprises two major functional units, the skeletal element (cartilage and bone) and fibrous connective tissues (tendons and ligaments). The primary skeletal anlagen are derived from the LPM, as are the majority of the tendons and ligaments299. The structures at the interface of bone and tendon represent the attachment units300, and mature to form the entheses and the associated bone protuberances and ridges. These eminences are formed by a distinct pool of Scx+ Sox9+ progenitor cells that generate a secondary skeletogenic field adjacent to the primary skeletal elements25,301,302. The Scx+ progenitors that generate the appendicular 125 tendon bodies originate from the LPM at E10.5300, prior to the emergence of the Hic1+ cells. The specific spatial localization of tdTomato+ chondrocytes observed throughout the developing forelimb suggested that a subset of Hic1-lineage MPs represent this unique Scx+ Sox9+ progenitor pool. To evaluate the relationship of tdTomato+ cells to Scx+ cells in the early forelimb, Hic1CreERT2; R26tdTomato; ScxGFP embryos were examined. Consistent with FCM experiments, histological analysis of E11.5 limb buds revealed clusters of ScxGFP+, SOX9+, tdTomato+ cells (Figure 4.11a). At E12.5, these triple-positive cells can be found localized to prospective attachment units (Figure 4.11b). This time line agrees with findings from Blitz et al.25, where Scx/Sox9 double-positive MPs were identified at E12.5 at the cartilage-tendon interface, and subsequently give rise to entheseal cartilages. Importantly, I identify these Scx+/Sox9+ MPs a day earlier (E11.5) before entry into the forelimb proper. Interestingly, at E12.5, gradients of differential GFP, tdTomato, and SOX9 expression can be observed in the areas of prospective tendon-bone attachment unit establishment. Chondrocytes located within the cartilage anlagen are SOX9+, at tendon-bone attachment sites tdTomato+/Sox9+ chondrocytes begin to appear. As the cartilage and fibrocartilage contribute to the tendon proper, tdTomato+/ScxGFP+ tenocytes become apparent and slowly decrease in abundance, until eventually only ScxGFP+ tenocytes form part of the distal tendons (Figure 4.10a). Together, these results indicate that tdTomato+ progenitors give rise to cartilage and tendon compartments of the tendon-bone attachment units during limb development. 126 Figure 4.9 Hic1+ MPs contribute specifically to the bone-tendon attachment units but not the tendon body nor primary cartilage anlagen. (a) Overview of Hic1CreERT2; R26tdTomato; ScxGFP lineage tracing strategy employed for analysis of E18.5 skeletal elements. (b) 3D model of mouse forelimb bones and relevant muscles generated using the mouse limb anatomy atlas online tool to aid with orientation of embryonic sections in (c-h). (c-h) Representative images of Hic1CreERT2; R26tdTomato; ScxGFP forelimb sections highlighting tendon-bone attachment sites and contributions of tdTomato+ progenitors to the entheses. Yellow boxes/lines on 3D limb models represent plane-of-view of the corresponding histology panels. White boxes identify inset regions shown in the right of each panel. Scale bars, 100µm. 127 Figure 4.10 Wholemount visualization of forelimbs reveals localization of tdTomato+ cartilage and tendons. (a) Whole mount E18.5 image of Hic1CreERT2; R26tdTomato; ScxGFP forelimb cleared with CUBIC. Scale bar, 500 µm. White arrowheads indicate areas of overlapping tdTomato and GFP signal. D, distal; Pr, proximal. (b) Whole mount image of E18.5 Hic1CreERT2; R26tdTomato humerus. White box represents inset region highlighting tdTomato+ chondrocytes. Dotted ellipse encircles the humeral head. Scale bar, 500 µm; 100 µm inset. 128 Akin to tendon-bone attachment sites, on the opposite end of tendons the MTJ attaches muscle to the tendon body. The Underhill group has previously identified and characterized the myotenocyte, a cell that resides at the interface between muscle and tendon45. Myotenocytes possess a unique morphology and are located at the ends of myofibers immediately adjacent to the tendon ECM, these cells can conveniently be identified by expression of Col22a1. Collagen XXII has been identified as a marker of tissue junctions, and found at high concentrations in the MTJs of skeletal and cardiac muscle303. In the context of Hic1+ cells, scRNA-seq data showed that Col22a1 was not detected in limb tdTomato cells until E16.5, albeit in a small number of tenocyte-associated cells (Figure 4.12a). COL22A1 could not be detected at the MTJs of fetal forelimbs via immunofluorescence at E16.5 nor at E18.5. However, in Hic1CreERT2; R26tdTomato; ScxGFP embryos lineage traced from E10 into adulthood, COL22A1+/tdTomato+ myotenocytes were abundant throughout the forelimb musculature (Figure 4.12b). It is important to note that in adult Hic1+ MP lineage tracing studies, myotenocytes derivatives were only observed following muscle injury and therefore do not express Hic1 under homeostatic conditions45. Instead, a Hic1+ progenitor population(s) appear to reside within the epitenon and gives rise to these cells upon organ perturbation. Taken together, these findings indicate that a subset of Hic1-lineage MPs contribute to the formation of the MTJ in the limb musculoskeletal system and represent a source of Scx/Sox9 double-positive progenitors that constitute the tendon-bone attachment units. 129 4.2.3 Chondrocytes and tenocytes emerge from an embryonic subpopulation of bipotent Hic1+ MPs The transcriptome of cells that give rise to tendon-bone attachment sites has recently been investigated at the single cell level304. In this study, attachment cells were determined to be identifiable by E13.5 as bipotent progenitors that express a mixture of tenocyte and chondrocyte genes. This mixed expression profile is believed to allow the cells at the tendon-bone attachment site to act as a transitional tissue. The bipotency of this population is further reinforced by studies that identified these cells as Scx/Sox9 double-positive progenitors25,301 and by lineage tracing using a Sox9CreERT2 mouse model that demonstrated the tenogenic properties of Sox9+ cells292. As mentioned in the previous section of this thesis, tdTomato+ cells express chondrogenic and tenogenic genes and directly contribute to the development of the tendon-bone attachment unit. To determine whether Hic1+ MPs in the developing limb contained a subpopulation of tendon/cartilage bipotent progenitors, the scRNA-seq and scATAC-seq data acquired from Hic1CreERT2;R26tdTomato embryos were reanalyzed with a specific focus on the tenocyte and chondrocyte fates. URD provided an excellent overview of the diverse differentiation trajectories that Hic1-lineage MPs follow from E11.5 to E16.5. To complement these analyses and to further dissect the transcriptomic changes that MPs undergo during chondrogenic and tenogenic differentiation, the ‘Harmony’254 and ‘Palantir’255 python packages were used. ScRNA-seq analysis of 32,225 cells and visualization using force-directed layouts (FDLs) revealed four general fates of Hic1+ MPs up to E16.5 (Figure 4.13a-b). These consisted of chondrocytes (Col2a1+, Sox9+) (Figure 4.13c), tenocytes (Scx+, Thbs4+) (Figure 4.13d), fibroblasts (Dpt+, Pi16+) (Figure 4.13e), and DFs (Crabp1+, Twist2+) (Figure 4.13f). The DF fate contained two 130 Figure 4.11 A subset of tdTomato+ MPs are ScxGFP/SOX9 double-positive. (a-b) Representative immunofluorescence micrographs of Hic1CreERT2; R26tdTomato; ScxGFP forelimbs showing overlapping expression of tdTomato, ScxGFP, and SOX9 at E11.5 (a) and E12.5 (b). Arrowheads indicate triple-positive cells. Dotted boxes highlight inset regions in a’ and b’. Scale bars, 100 µm, 20µm inset. 131 Figure 4.12 TdTomato+ MPs contribute to the myotenocyte compartment during development of the MTJ. (a) Gene expression heatmap of Col22a1 overlayed on URD-generated dendrogram. Fibro1, fibroblast 1; Fibro2, fibroblast 2, RDFPs, reticular dermal fibroblast progenitors; PDFPs, papillary dermal fibroblast progenitors. (b) Fluorescence microscopy images of embryonic linage-traced adult muscle. Red arrow heads highlight tdTomato+ cells at the MTJ. Scale bars, 20um. 132 subpopulations identified by separate points in the FDL (Figure 4.13f, arrowheads), these consist of the previously described RDFPs and PDFPs. The connective tissue fates, specifically fascia and perichondral cells were interspersed among the fibroblast populations and cell fates that had a small number of cells, such as pericytes and perineural cells were not sufficiently resolved using the Harmony analysis tool. To focus specifically on skeletogenic fates, a subset of the scRNA-seq data was created containing only chondrogenic and tenogenic cells (Figure 4.14a). In the previous chapter, Id1 and Snai1 were identified as markers of E11.5 MPs. Indeed, Id1+, Snai1+ MPs were abundant at E11.5 and were therefore used to determine the pseudotime start point (Figure 4.14b-c). Interestingly, within this progenitor population, a subset of cells co-expressed Scx and Sox9 beyond E12.5 and up to E16.5 (Figure 4.14d). On the other hand, markers specific to differentiated cartilage and tendon, Acan and Tnmd respectively, were exclusive to E16.5 terminal cells (Figure 4.14e). To analyze the transcriptional changes that MPs undergo during chondrogenesis and tenogenesis, gene expression trends were plotted for genes of interest. As expected, transcripts associated with an undifferentiated state were highly expressed in early pseudotime cells, but drastically declined as pseudotime progressed in both the chondrogenic and tenogenic trajectories (Figure 4.14f). Interestingly, even though Scx and Sox9 are widely expressed at E11.5 and E12.5, the gene expression trends diverge with Sox9 decreasing in the tenogenic trajectory and increasing in the chondrogenic curve (Figure 4.14g). On the other hand, Scx expression increased in the tenogenic trajectory and decreased in the chondrogenic one as pseudotime progressed. Transcripts for Acan and Tnmd were non-existent at early timepoints and appear only in their respective trajectories (chondrogenic and tenogenic fates) late in the pseudotime values (Figure 4.14h). 133 Similar to scRNA-seq, epigenetic profiling revealed a latent potential for Hic1+ MPs towards cartilage and tendon with OCRs in gene loci specific for these fates (Figure 4.15a-b). To further explore the chromatin landscapes that underlie the chondrogenic and tenogenic fates, cell classification labels were transferred from the scRNA-seq dataset using an anchor-based method for cross-modality integration259 and individual pseudotime trajectories were constructed by subsetting the scATAC-seq dataset (Figure 4.15c-d). Differential accessibility analysis in pseudotime revealed that genes associated with the chondrogenic program remain open in the chondrocyte cluster but become progressively closed in the tenocyte cluster (Figure 4.15e-g; Acan, Col2a1 and Col9a3). Similarly, tenogenic gene loci are opened in the tendon cluster at late pseudotime values (Figure 4.15h-j; Tnmd, Thbs4 and Kera). Additionally, prospective unique lineage-associated regulatory elements were identified within chondrogenic and tenogenic gene regions (Figure 4.15k) which underlie the progressive lineage restriction observed in the scRNA- seq gene expression trends. Taken together, these results suggest that Hic1 identifies a population of bipotent chondro-tenogenic progenitors in the developing forelimb. 134 Figure 4.13 Analysis of the Hic1CreERT2; R26tdTomato embryonic forelimb scRNA-seq dataset with Harmony identifies four general fates. (a) Force-directed layouts displaying developmental timepoints of Hic1-lineage MPs. (b) Force-directed layout displaying pseudotime for an Id1hi/Snai1hi start cell to terminal states. (c-f) Force-directed layouts displaying expression levels of representative cell type markers. (c) cartilage, (d) tendon, (e) fibroblasts, (f) DFs, arrowheads indicate DF subpopulations. 135 Figure 4.14 ScRNA-seq analyses reveal differentiation trajectories for chondrogenic and tenogenic progenitors. (a) Force-directed layouts displaying developmental timepoints of Hic1-lineage chondro-tenoprogenitor subcluster. (b) Force-directed layout displaying pseudotime for an Id1hi/Snai1hi start cell and Acanhi or Tnmdhi terminal states. (c-e) Force-directed layouts displaying expression levels of representative cell type markers. (f-h) Expression trends of marker genes with respect to pseudotime. 136 Figure 4.15 A subpopulation of Hic1+ MPs are bipotent chondro-tenoprogenitors. (a-b) Genome browser tracks from scATAC-seq analyses displaying the promoter sum signal near the indicated gene loci. Grey boxes highlight clusters exhibiting promoter sum signal relevant to the indicated cell function or fate. (c) UMAP plot of scATAC-seq chondrocyte and MP cluster subset colored by embryonic time point (left) and pseudotime values (right). (d) UMAP plot of scATAC-seq tenocyte and MP cluster subset colored by embryonic time point (left) and pseudotime values (right). (e-j) Chromatin differential accessibility graphs for representative genes in chondrocyte (e, f, g) and tenocyte (h, i, j) subclusters. (k) Coverage plots for MP, chondrocyte, and tenocyte clusters focused on the indicated gene loci highlighting cis-co-accessible network links. 137 4.2.4 Hic1 deletion negatively impacts skeletal morphogenesis Previous studies have shown that Hic1 deletion is embryonic lethal and is associated with a spectrum of defects including limb abnormalities161. To further explore the nature of these defects and, in particular, to assess the impact of Hic1 deletion on enthesis morphogenesis, Hic1 knockout animals were examined. Skeletal preparations of Hic1-/- (Hic1-null) embryos displayed a significantly reduced crown-rump measurement of 14 ± 1mm compared to 22 ± 0.5mm for wildtype littermates (Figure 4.16a). There were no differences in size between heterozygous and wild type fetuses (Figure 4.16a). Indeed, Hic1+/- mice were viable, fertile, and did not display any apparent abnormalities. Interestingly, Hic1-null forelimbs contained multiple defects including reductions and malformation of the deltoid tuberosity and olecranon (Figure 4.16b). Further analysis of the impact of Hic1 deletion on the later stages of appendicular skeletal development was limited by embryonic lethality. This impediment was circumvented with a strategy whereby the conditional deletion of Hic1f/f alleles was achieved specifically in the limb bud mesenchyme using a transgenic Prrx1-Cre line213. In contrast to the conventional Hic1 KO (cKO), Prrx1-Cre; Hic1f/f mice were viable and generally healthy (Figure 4.17a) other than locomotion abnormalities. To study the effects of Hic1 knockout on appendicular skeletogenesis, bones from adult Prrx1-Cre; Hic1f/f mice were analyzed using micro computed tomography (µCT) imaging. In contrast to Hic1-null limbs, Prrx1-Cre; Hic1f/f bones were similar in length to wildtype littermates (Figure 4.17b). Strikingly however, the humerus, radius and ulna from the conditional Hic1 KOs displayed severe anomalies in the placement, morphology, and overall presence of bone protuberances, eminences, and ridges (Figure 4.17c-d and 4.18). A comprehensive list of deformities observed in the µCT images of forelimb long bones are summarized in Table 4.1. In particular, the ulnar 138 tuberosity was absent (Figure 4.17d, red panel), the olecranon was noticeably thinner and shorter, and the radial tuberosity, radial and ulnar ridges were all absent (Figure 4.17d, blue panel). Consequently, Hic1 KO skeletal elements, such as the radius and ulna, presented with a featureless cylindrical morphology (Figure 4.17, green panel). Furthermore, the region containing the interosseous ligament was notably-rich for tdTomato+ cells (Figure 4.18a) and, likely as a result, the interosseous volume was increased in the Hic1 KO mice compared to wild-type littermates. The elbow joint also presented faulty morphogenesis of the olecranon process, medial epicondyle, and olecranon fossa (Figure 4.17d, yellow panel). The proximal humerus also presented with abnormalities affecting the positioning and mass of the greater and lesser tubercles (Figure 4.18b-c). Similarly, on the distal end of the humerus, the supinator crest appeared narrow with a reduced curvature (Figure 4.18d). The coronoid process of the ulna which acts as a bony pillar to prevent dislocation of the elbow joint, was severely reduced (Figure 4.18e). Also, within the elbow joint, the proximal end of the radius did not articulate appropriately with the radial fossa, resulting in an increased interosseus space (Figure 4.18f). This incorrect articulation could be a result of changes in the morphology of the radial neck (Figure 4.18g). Taken together, Hic1 is necessary for enthesis formation and the perturbation of Hic1+ MPs is sufficient to cause a spectrum of skeletal malformations that involves deficiencies in bone morphogenesis. 139 Figure 4.16 Characterization of Hic1nLacZ/nLacZ skeletal deformities. (a) Bar graph showing crown-rump measurements of Hic1 wild-type, heterozygous and knockout embryos. Data represent individual sample measurements +/- SD, unpaired t test, *P<0.05, ***P<0.005. (b) Alcian blue and Alizarin red whole-mount skeletal preparations of E17.5 Hic1 knockout embryos (Hic1nLacZ/nLacZ) and wildtype littermates. Complete embryo shown in the left panel and corresponding forelimb shown in the right panel. Scale bars, 5 mm for whole embryo images and 1 mm for forelimb panels. Arrowheads highlight the deltoid tuberosity (black) and olecranon (blue). 140 141 Figure 4.17 Hic1 deletion severely impacts secondary skeletal morphogenesis. (a) Gross anatomical dorsal view of wildtype and Prrx1-Cre; Hic1f/f adult mice (P56). (b) Bar graphs of bone-length measurements taken from forelimb long bones of wildtype and Prrx1-Cre; Hic1f/f adult mice. Data represent individual bones measured +/- SD. (c-d) µCT volume-rendered reconstructions of adult Prrx1-Cre; Hic1f/f and wildtype forelimbs. Colored boxes in (c) represent regions of interest highlighted in (d). White lines in green panel represent cross-sections shown beneath. H, humerus; O, olecranon; R, radius; U, ulna; red arrowhead, ulnar tuberosity; blue arrow, radial tuberosity; grey arrowhead, olecranon process; white arrowhead, medial epicondyle; star, olecranon fossa. Scale bars, (c) 5mm; (c) 1mm. 142 (Figure legend on next page) 143 Figure 4.18 Consequences of Hic1 deletion on forelimb long bone morphology. (a) Representative fluorescence micrographs of E18.5 Hic1CreERT2; R26tdTomato showing distribution of tdTomato+ cells throughout the forearm interosseus space and at ligament attachment sites. R, radius; U, ulna. Dotted lines denote the bone surfaces, arrowheads highlight tdTomato-rich areas within the interosseus space. Scale bars, 100µm. (b-g) µCT volume-rendered reconstructions of adult wildtype and Prrx1-Cre; Hic1f/f forelimbs highlighting tendon-bone attachment regions. Each panel contains an overview of the complete bone at the appropriate viewing angle adjacent to a magnified region of interest (white boxes). Top to bottom: greater tubercle, lesser tubercle, supinator crest, coronoid process, deltoid tuberosity, extra growth on radial neck are highlighted (white arrowheads). Red arrowheads indicate the articular surface for sesamoid bone on distal humerus. Scale bars, 1mm. 144 Table 4.1 List of observed morphological deficiencies in long bones of Prrx1-Cre; Hic1f/f mice. Long Bone Affected Structure Associated Muscles or Ligaments Skeletal Defect Description Humerus Deltoid tuberosity Deltoid muscle Reduced area Greater tubercle Supraspinatus, infraspinatus and teres minor muscles Positioning and decreased size Lateral epicondyle Extensor muscles and radial collateral ligament Size reduction Lesser tubercle Subscapularis muscle Positioning and decreased mass Medial epicondyle Flexor muscles and radial collateral ligament Size reduction Olecranon fossa N/A Change in morphology Supinator crest N/A Reduced curvature, narrow Radius Distal radius N/A Incorrect articulation with humerus 145 Radial neck N/A Extra bony growth Radial ridge Interosseous membrane Lacking Radial tuberosity Biceps brachii muscles Absent Ulna Coronoid Process Brachialis muscle and ulnar collateral ligament Severely reduced Olecranon Triceps brachii muscles Reduced Ulnar ridge Interosseous membrane Absent Ulnar tuberosity Brachialis muscle Absent Interosseous space Interosseous membrane Increased volume 146 Chapter 5: Conclusion 5.1 Overview In adult SkM, Hic1 identifies three major MP sub-populations including, fibroblasts, pericytes and tenogenic progenitors45. Herein, Hic1-based genetic lineage tracing tools have been used to develop an atlas of Hic1+ MP derivatives starting from the onset of Hic1 expression in limb mesenchyme through to P56. In this regard, Hic1 defines a rare sub-population of cells in the early limb mesenchyme. While barely evident at E11.5, Hic1-lineage traced cells subsequently undergo appreciable expansion culminating in embryonic and post-natal progeny that reflect the diversity of fibroblasts, other specialized MPs and pericytes. A number of other markers have been used to identify various subpopulations within these lineages including Grem1106, Lepr110, Osr155 and Tcf7l239, but Hic1 appears to identify an MP compartment situated at the apex of this lineage tree thereby capturing the totality of their fates. With the exception of VSMCs, there is excellent concordance between the embryonic lineage-traced derivatives and those identified in the adult. Furthermore, again with the exception of VSMCs, the embryo-traced Hic1+ derivatives express Hic1 in the adult indicating that their progenitor status is likely maintained. In the URD-generated mesenchymal lineage tree, Hic1 is expressed in the trunk and branch populations and is lost in specific tip populations associated with the emergence of differentiated phenotypes, i.e., chondrocytes and tenocytes. Osr1 was previously shown to define embryonic FAPs and its expression parallels that of Hic1, except it is downregulated in the emerging pericyte lineage as well as in adult SkM pericytes. Similarly, Pdgfra is a useful marker for identifying 147 various MPs, however, it is also absent in pericytes and is accordingly down-regulated in the pericyte branch. Dpt is expressed much later in embryonic development along with Ly6a and defines the emergence of the Fibro1 and 2 populations. Hic1 effectively labels these two major emerging mesenchymal cell branches, fibro-skeletogenic and pericytic within the developing limb. Collectively, these findings appear to indicate two separate but major branches defining the embryonic limb mesenchyme, a fibro-skeletogenic branch and a distinct pericyte branch. Furthermore, integrated scRNA-seq and scATAC-seq analyses revealed that Hic1+ progenitors encompass a population of bipotent chondro-tenoprogenitors that give rise to differentiated cartilage, tendon, and the transitional cells found in between these two types of tissues. Other specialized cell types were similarly identified as Hic1+ MP progeny with a combined histological and transcriptomic approach (Figure 5.1). Myotenocytes, the cells that represent the interface between muscle and tendon, were characterized by expression of Col22a1 and their unique morphology and tissue localization. DFs were subcategorized as PDFPs or RDFPs based on gene expression and spatial localization within the dermis. On the other hand, it is important to note that Hic1+ MPs did not give rise to any epithelial, endothelial, SkM, or nervous cell types in the embryonic or adult limb. The Hic1f/f and Prrx1-Cre; Hic1f/f mouse models allowed for the exploration of the functional consequences that Hic1 knockout would have on embryonic development and limb skeletogenesis. Although the embryonic knockout of Hic1 has been previously characterized161, its effects on limb development and skeletogenesis have been overshadowed by more severe phenotypes such as acrania, exencephaly, cleft palate, and omphalocele which lead to embryonic lethality. As described in previous sections of this text, lineage tracing experiments performed on 148 Hic1CreERT2; R26tdTomato embryos revealed a significant contribution of Hic1-lineage MPs to the appendicular chondrogenic elements. Furthermore, Hic1 has been identified as an RAR target gene271 and it is well established that retinoids regulate chondrogenesis during limb development230,305. These findings suggest a need to study the phenotypic properties of the Hic1 knockout mouse model with respect to skeletogenesis and other aspects of the musculoskeletal development. Therefore, the experiments described in this thesis are focused on the skeletogenic repercussions of Hic1 deletion on both the conventional and limb mesenchyme-specific knockout mouse models. Overall, the cell fate and dynamics if Hic1+ MPs has been interrogated to an extent that will allow for the elucidation of the embryonic origin, fate and function of these cells in the context of limb development and skeletogenesis. Experimental strategies have been employed to investigate Hic1 MP biology from the onset of Hic1 expression at E10.5 to postnatal timepoints that are well into adulthood. In this way, a continuum of Hic1+ MP functions can be linked to diverse processes such as embryonic organogenesis, skeletal morphogenesis, and tissue repair and regeneration. 149 Figure 5.1 Fate of Hic1+ MPs in limb development. Schematic of the diverse mesenchymal limb lineages captured by labelling Hic1CreERT2; R26tdTomato embryos from development into adulthood. Genes displayed in red indicate cell types that were not identified in scRNA-seq. 150 5.2 On the Embryonic Origin of Hic1+ MPs The generation of a functional limb during embryonic development relies on several migratory events. Myogenic progenitors, melanocytes, endothelial cells, nerve axons and glial cells all migrate from various regions of the embryo into the developing limb to establish their respective tissues and organs. In contrast, appendicular chondrocytes and tenocytes originate from LPM-derived limb mesenchyme. The findings described herein propose the possibility that a specialized subset of tenogenic and chondrogenic progenitors arise from the somitic mesoderm and not the LPM. Specifically, the connecting cells located at either end of the tendon body: the MTJ cells and those of the enthesis. Although this concept may initially seem as contradicting the current dogma of musculoskeletogenesis, an argument exists for a case in which this cellular mechanism may be rationalized alongside the accepted tenets of tendon evolution, development, and function. Firstly, it is important to mention the technical limitations of early lineage tracing methodologies, mainly quail-chick chimeras, that were employed during the late 20th century with the goal of determining the embryonic origin of limb muscle progenitors. This inter-species transplantation model was initially used to trace the migratory routes of neural crest cells306, it was also crucial in determining the somitic origin of muscle progenitors307. Although quail-chick chimeras represented an extremely effective and elegant lineage tracing model at the time, it was not without certain weaknesses that may have resulted in overlooking of dynamic rarer migratory populations. This surgical technique required an extensive amount of dexterity and specialization on part of the technician performing the experiments. It was not uncommon for experiments to be tallied as different surgeries would result in varying results13. Indeed, some early findings utilizing this method published contradicting claims with respect to contributions of paraxial and LPM to 151 limb tissues308,309. Furthermore, the emphasis of most early lineage tracing experiments was strictly on myogenic and neural crest progenitors as the chondrogenic and tenogenic populations were believed to be predominantly LPM-derived. Tendon-focused studies relied mainly on tissue cross-sections without a method of focusing on MTJs or tendon-bone attachment sites290. Quail-chick chimeras, along with the discovery of Scleraxis as a tendon progenitor marker, allowed for the identification of the syndetome2. However, these initial Scx-based studies focused on the axial skeleton and the contribution of somitic mesenchyme to axial tendons with the exception of sternum-associated tendons which were found to be LPM derived, this conclusion was then extrapolated to limb tendons. I would like to emphasize that the results described in this thesis do not suggest that all limb tendons are somite derived. Rather, the data seem to indicate that the specialized attachment cells located at either end of the tendon, areas that have been historically neglected from close analyses, may be somite-derived. It is also important to mention the evolutionary differences that exist between avian and mammalian limb development. At the molecular level, gene regulatory networks involved in early limb specification and outgrowth are highly conserved among vertebrates310. This is the case for mouse-chicken comparative transcriptomics but, as expected, there is a divergence into species-specific spatiotemporal gene expression at later stages of limb development311. At the cellular level, chick and mice embryos display discrepancies in migratory patterns of limb-bound progenitor populations. Specifically, mouse nerve axons extend into the limb bud concurrent with migrating myogenic progenitors312,313; in chicken, the nerves undergo a “waiting” period of ~24 hours within the torso mesenchyme and only penetrate the limb field once primary myotube formation has commenced314,315. Furthermore, migration of myogenic progenitors from the somite 152 in rodents is confounded in comparison to birds as the same region that gives rise to the upper arm is responsible for migration into the developing diaphragm, a mammal-specific organ247,313. Examination at the tissue level also presents significant differences; for instance, avian limb tendons have been known to calcify under normal conditions316, mammalian tendons, on the other hand, do not share this property. It is not clearly understood how appendicular tendon calcification may have been an evolutionary asset317; however, mineralized tendons have been shown to reduce strain and provide greater load bearing capacity318. The aforementioned divergences in mouse and chick development are expected, as the last common ancestor that linked humans and birds lived ~300 million years ago compared to ~80 million for humans and mice319. Evidently, rodent embryology will more closely resemble that of humans; however, non-mammalian model organisms present unique advantages for the study of developmental processes such as the use of in ovo manipulation techniques in the case of the chick. Therefore, the impactful contributions of non-mammalian model organisms to the fields of cell and developmental biology must not be understated. Nonetheless, as novel technologies emerge that facilitate the interrogation of cell fate and ontogeny in mammals, the formulation of theories and conclusions therefrom should maintain awareness and acknowledge the gaps in evolutionary history that exist between species used in previous research. The single cell genomics data presented here suggest that in E11.5 mouse embryos, a population of MPs distinct from the LPM are present in the forelimb bud and gives rise to diverse cell types. Histological findings indicate that a subset of these cells may derive from paraxial mesoderm. However, it remains unclear whether all the fates described for this population (DFs, fibroblasts, pericytes, chondrocytes, tenocytes, among others) could therefore be derived from the 153 same embryonic source and whether these fates share a common multipotent progenitor(s). Regarding the former issue of a somitic origin for Hic1+ MP cell derivatives, subsets of pericytes and VSMCs have been previously shown to be somite-derived89. As previously mentioned, limb chondrocytes and tenocytes are LPM-derived. However, Hic1-lineage chondrocytes and tenocytes are specialized cells found predominantly at tissue transition sites and represent separate populations with respect to the general cartilage anlagen25 and possibly the tendon body cells as well, since tendon elongation has been shown to require recruitment of new tenogenic progenitors after tendon has attached to skeletal and muscular targets320. These specialized functions may justify a distinct origin for Hic1-lineage chondrocytes and tenocytes in the limb. A somitic origin for DFs and connective tissues is more controversial, with the methodology described in these studies it is not possible to exclude the possibility of dual somite-LPM origins for labelled cells. In a similar fashion, the multipotency of Hic1+ MPs is a potential point of contention. Although scATAC-seq analyses at E12.5 showed a generally homogenous chromatin state throughout Hic1+ MP descendants, certain cellular populations that are difficult to capture or less abundant at this stage of limb development may be underrepresented. Similarly FCM experiments revealed that a minority (33%) of labelled tdTomato cells in E11.5 Hic1CreERT2; R26tdTomato; ScxGFP are ScxGFP-negative, this suggests a degree of heterogeneity within this population. Furthermore, transcriptomic analysis of tdTomato+ cells in E11.5 forelimbs displayed modest differences in gene expression between clusters with an overall shared expression of MP markers such as Id1, Id2, Nnat, and Snai1. Clonal lineage tracing experiments performed in chick limb buds of a similar timepoint revealed that there are no lineage-restricted cells at this stage and that single progenitor cells can give rise to multiple cell types including clones of cartilage, tendon, and dermal DFs125, suggesting the possibility of Hic1+ MPs representing multilineage progenitors or a heterogeneous 154 population of uni- and/or multi-lineage progenitors. In summary, while the limited complexity observed in the single cell-omics analyses of tdTomato+ cells at E11.5 and E12.5 are consistent with a common embryonic origin for the Hic1+ MPs, this does not preclude the possibility of multiple overlapping populations with more restricted lineage trajectories. Alternatively, Hic1 may represent a functional marker that is expressed in disparate mesenchymal lineages, indicative of a shared regulatory program. 5.3 Hic1+ MPs and “classic” mesenchymal cell types Mesenchymal cells are, in part, functionally defined by their ability to contribute to the skeletogenic lineages; including, chondrogenic, osteogenic and/or tenogenic. Strong transcriptomic signatures reflective of tenogenic and chondro/osteogenic fates are evident in the URD plots and were validated with histological analyses. In addition, within the tenogenic branch, a Col22a1-containing tenogenic signature could be identified reflective of myotenocytes within the MTJ302,303. Within embryonic and adult limb bones, lineage-traced osteocytes as well as presumptive BMSCs could be identified based on their morphology. BMSCs and osteocytes were located in cortical and trabecular bone of the humerus and corresponding bone marrow, respectively. Neither of these populations could be reliably detected in the embryonic URD plots. An osteogenic signature is embedded within the tip of the chondrogenic branch and this could reflect a bona fide osteogenic lineage and/or chondrocyte maturation and the appearance of hypertrophic chondrocytes (HC), which express many osteoblastic markers. Consistent with this, the tip population is also positive for the HC-expressed gene Col10a1. Alternatively, Hic1+ progeny make a limited contribution to embryonic bone formation and thus, the detection of this 155 population would be compromised by its rarity and/or difficulty in liberating tdTomato+ cells from bone. In this regard, this population is remarkably similar to the Lepr+ bone mesenchymal stromal cell114. In the post-natal bone marrow, Lepr identifies the majority of the mesenchymal stromal population. While Lepr+ cells can be identified in the embryonic bone marrow, this population makes a negligible contribution to osteocytes therein110. In contrast, Lepr+ cells contribute substantially to new osteocytes generated with normal adult bone turnover and following fracture110,114. Very few tdTomato+ cells can be identified in the embryonic cortical bone and thus, it appears that Hic1 progeny make a limited contribution to the embryonic skeleton. Multi-potent MPs are, in part, defined by their ability to generate white adipocytes. Transcripts indicative of adipogenic lineage potential (Cebpa, Cebpb, Cebpd, Pparg, Zfp423) were embedded in multiple branches of the URD lineage tree (including fibro 1, fibro 2, perineurium, DFs, and fascia), however, they were most prevalent within the fibroblast 1 and 2 branches. Similarly, gene loci corresponding to the aforementioned adipogenic markers were accessible in all scATAC-seq cell clusters. Subcutaneous white fat first becomes apparent in later stages of embryogenesis (~E18.5) and expands postnatally, when the need for energy storage increases321. No tdTomato+ fat cells were observed in E18.5 Hic1CreERT2; R26tdTomato forelimb dermis, although the transcription factor Zfp423 which identifies preadipogenic fibroblasts321 was widely expressed, suggesting that cells with adipogenic potential are present within the tdTomato+ fraction of embryonic forelimb MPs. When lineage tracing embryonic Hic1+ cells into adulthood, a large quantity of tdTomato+ PLIN1+ adipocytes were detected histologically in the limb; however, this population was not detected in scRNA-seq. This likely reflects the technical challenges in sorting large fragile adipocytes and their subsequent inefficient capture into droplets. Nevertheless, the 156 adipogenic potential of adult Hic1+ MPs has been corroborated in vitro and in vivo through morphological, immunofluorescence, and histochemical approaches45,203,204. 5.4 Hic1+ MP derivatives contribute to fibroblast and peripheral nerve mesenchymal components during development A recent study has defined two “universal” fibroblast populations that are shared across tissues, defined by the transcripts Dpt, Col15a1 (Fibro1) and Dpt, Pi16 (Fibro2)196. The URD plots identify profiles reflective of these two populations and while Col15a1 is not expressed in the Fibro1 compartment at E16.5, other genes associated with this sub-population (Cxcl12, Fbln1, Penk) were found to be enriched in this branch. A recent study that employed scRNA-seq to characterize EGFP+ cells from a PdgfrbEGFP knock-in, further refined the classification of SkM fibroblast types and their contribution to the muscle connective tissues322. They identified presumptive signatures reflective of endomysial cells, perimysial and epimysial cells and a new population they refer to as paramysial cells which are situated between the endomysium and perimysium. Interestingly, the Dpt+ Pi16+ (Pdgfra+ Hic1+) sub-population was enriched in the endomysium, which is a thin connective tissue layer that circumscribes individual myofibers. This population selectively expresses transcripts of numerous trophic factors (i.e., Wnt2, Wnt5a, Sema3c), which is consistent both with the proximity of the endomysium to the satellite cell niche and its function in regulating this compartment. A previous study identified multiple Hic1+ MP types within the adult dermis, including: hair follicle fibroblast progenitors (Sox2+, Itga8+, Mgp+, Acan+) which repopulate the dermal 157 sheath (Itga8+, Itga5+, Acan+, Myh10+, Myl9+, Cd200+) and dermal papillae (Sox2+, Lef1+, Crabp1+, Rspo3+, Corin+, Vcan+) fibroblast compartments, papillary fibroblasts (Dpp4+, Dlk1-, Blimp1+, Lrig1+, Ly6a-, Trps1+) and reticular fibroblasts (Dpp4-, Dlk1+, Ly6a-). During embryogenesis these populations emerge from a common (Pdgfra+, Dlk1+, Lrig1+) fibroblast progenitor59. Pdgfra and Dlk1 are broadly expressed throughout many of the branches, whereas Lrig1+ cells are restricted to the presumptive DF lineage and this parallels the appearance of En1, another DF progenitor marker. At 14.5, the DF lineage bifurcates into PDFP and RDFP branches reflected by the differential expression of Blimp1, Lef1, Vcam1 and Trps1. At this branch point, Sox2 transcripts are detectable and restricted to the PDFP branch and expression can be further confirmed by anti-SOX2 staining at E18.5. Consistent with previous reports, the PDFP branch will give rise to all of the specialized fibroblasts within the skin and signatures reflective of this potential are embedded in this branch tip including: Cd200, Col11a1, Lef1, Myl9 and Rspo3. Furthermore, the point of RDFP and PDFP bifurcation is associated with differential expression of multiple Irx genes, with Irx1, 3 and 5 enriched in the RF branch. In addition to Sox2, other Sox transcripts are preferentially expressed in the PDFPs, such as Sox11 and Sox18. In comparison to all of the other lineages, the DF lineage exhibits a prominent canonical WNT signature that is associated with Axin2, Lef1 and Nkd2 expression immediately following the emergence of this lineage at E12.5. Collectively, Hic1 effectively labels the common fibroblast progenitor in the E12.5 limb mesenchyme and continues to be expressed in these lineages, identifying enduring progenitor subsets which support skin homeostasis and regeneration into and throughout adulthood203. 158 In a simple ex vivo culture experiment, Bunge et al. showed that the perineurium appeared to derive from fibroblasts323. Hic1 progeny give rise to all of the nerve-associated fibroblasts and in the adult, these cells continue to express Hic1, indicative of their progenitor status. In the URD lineage tree presented here, cells reflective of the perineurium branch off from the common Dpt+ fibroblast branch and markers reflective of a perineurial signature (Wnt6, Ptch1, Cldn1) can be detected at E16.5. Hic1+ cells traced at E10 – P56 clearly show contribution to the various mesenchymal components of the sciatic nerve, including the epineurium, perineurium and endoneurium. The signatures of the perineurium (Wnt6, Cldn1, Ptch1) and endoneurium (Sox9, Cd34, Fzd2) align well with the recently published scRNA-seq sciatic nerve dataset (SNAT)136. However, the epineurial cluster within this data aligns with the Dpt+ fibroblast cluster and as such, is likely embedded within the Hic1+ fibroblast clusters. 5.5 Embryonic Hic1+ MPs represent a source of mural cells Pericytes form a distinct lineage within the URD tree and this is reflected by their expression of a core group of transcripts which exhibit limited overlap with fibroblasts. Within the time points captured, pericytes appear after the branch point at E12.5 and are defined by the transcripts Rgs5, Kcnj8, Abcc9, Myh11 with Myocd appearing weakly in the tip cells. Interestingly there are a number of transcripts broadly expressed at the top of the tree that are quickly extinguished in the pericyte arm including Osr1, Pdgfra, Scx, Tcf7l2, Sox11, Snai1 and Crabp2. The transcriptional program underlying the specification of the pericytic lineage has yet to be identified75. However, these findings would indicate that the acquisition of a pericytic fate may be, in part, related to the decreased expression of transcription factors and/or signaling pathways 159 associated with alternative fates. Unexpectedly, the E10-P56 chase revealed a major tdTomato+ Hic1- population of VSMCs (Cnn1+, Tagln+, Myocd+, Myh11+, and Pdlim3+) and tdTomato+ VSMCs (CNN1+) were detected in the tunica media of arterioles within SkM. A distinct hybrid cluster expressing transcripts reflective of VSMCs and pericytes was also detected. However, in addition to many VSMC and pericytic markers, this cluster also expresses several unique transcripts including Dlk1, Adra2a, Rgs16 and is abundant for Cd90. Dlk1 is typically associated with stem and progenitor cells and a similar Dlk1+ Thy1+ population has been described in human subcutaneous fat, located within the vessel adventitia and on capillaries324. Furthermore, Dlk1 expression has been defined in a subset of brain pericytes325. Thus, the Hic1+ Dlk+ population appears to represent a distinct pericyte-like progenitor along a pericyte to VSMC lineage continuum288,322. Overall, Hic1+ MPs represent a distinct mesenchymal population which emerge later in limb development, after the initiation of the LPM-derived primary skeletal elements. This MP sub-population appears coincident with migrating somite-derived myogenic precursors along with infiltrating neuronal processes. Nonetheless, Hic1+ MPs generate a unique cellular lineage hierarchy which culminates in embryonic and post-natal progeny that reflect the diversity of specialized functions of MPs. As previously mentioned, the embryonic origin of many of these mesenchymal lineages is unclear. Early chick-quail transplant experiments have indicated a somitic origin for VSMCs89, whereas most of the other mesenchymal lineages appear to derive from the LPM. The potentially distinct lineage trajectories of fibrogenic/skeletogenic arms versus the pericyte/VSMC branch in part reflect a dichotomous origin. 160 5.6 Embryonic Hic1+ MP progeny in enthesis and MTJ development The appendicular and axial skeletons are derived from different embryonic sources. The LPM gives rise to all the bones within the limbs and the somitic sclerotome contributes to the formation of most of the ribs and the entire vertebral column. Similarly, tendons in the limb have been shown to be LPM-derived and those associated with the rib cage and vertebrae originate from the syndetome, a Scx+ compartment of tenogenic progenitors in the somite. In this work, a population of teno-chondroprogenitors identified by Hic1 expression are revealed in the somite and embryonic limb. Within the early limb bud, these progenitors proliferate and localize to the developing dorsal and ventral muscle masses on the periphery of the chondrogenic anlagen. As the limb matures, Hic1+ MP derivatives contribute to the tendon and cartilage compartments of the developing skeleton. Transcriptomic analyses indicate that the gene expression signatures of Hic1+ MPs differ from that of surrounding LPM cells and Hic1+ MPs represent a relatively rare population in the E11.5 and E12.5 limb bud. Furthermore, Scx/Sox9 double-positive cells are evident at these timepoints, although these cells clearly constitute a subset as there are clusters of cells that don’t express either master regulatory factor. This could indicate a heterogeneous population as discussed previously. These comments notwithstanding, Scx/Sox9 double-positive cells represent teno-chondroprogenitors which have been shown to populate tendon-bone attachment units during early development25,301,304. Within the developing appendicular skeleton, Hic1+ MPs do not contribute appreciably to the primary cartilaginous anlagen, but exhibit specialized skeletogenic fates that generate many of the bone markings. Consistent with this, conditional deletion of Hic1 leads to featureless bones, 161 with minimal impact on the primary skeletogenic components. Collectively, these observations indicate that Hic1+ MPs are distinct from LPM-derived mesenchyme that is essential for formation of the primary anlagen. In contrast, Hic1+ MP derivatives contribute to a “secondary” wave of skeletogenesis that sculpts bone morphology, generating various bone markings essential for function; termed herein, secondary skeletal morphogenesis. In this regard, Hic1+ MPs contribute exclusively to events that underpin the assembly of functional connections between force-generating muscle tissue and the skeleton. Accordingly, the development of the interfaces between bone and tendon (enthesis) and tendon and muscle (MTJ) both require a particular Scx+ MP population which is derived from a common Hic1+ progenitor. Single cell transcriptomics of lineage-traced Hic1+ MPs across several developmental timepoints revealed four major differentiation trajectories including fibroblasts, DFs, chondrocytes, and tenocytes. As early as E11.5, Scx/Sox9 double-positive progenitors can be found within the limb bud mesenchyme, extending the chronological depth of previous studies which identified these progenitors beginning at E12.525. Furthermore, scATAC-seq showed that Hic1+ MPs have latent chondrogenic and tenogenic potential at E12.5, which becomes increasingly restricted as cells progress through each pseudotime trajectory ultimately culminating in a decrease of chondro-tenogenic bipotency by E16.5. The emergence of non-skeletogenic cell fates (i.e. fibroblasts and DFs) from Hic1+ MPs and their relation to Scx+/Sox9+ progenitors requires further investigation. Experimental approaches with higher sensitivity and resolution may be required to elucidate whether Hic1+ MPs represent a common progenitor for connective tissue cells in the developing limb or constitute a pool of heterogeneous progenitor Hic1+ subpopulations. Interestingly, gene markers that have been used to identify and lineage trace embryonic and post-162 natal fibroblasts (Osr1, Tcf7l2) and DFs (Dlk1, Pdgfra) largely overlap with the Scx+/Sox9+ cells in transcriptomic space of sorted Hic1CreERT2; R26tdTomato cells, suggesting a common Hic1+ fibroblastic progenitor. The importance of Hic1+ MPs in adult tissue renewal and regeneration has previously been demonstrated in muscle, heart, and skin. In this work, I show that Hic1+ MPs are also crucial in appendicular skeletogenesis, specifically at the entheses, MTJs and tendons, all critical components of the appendicular musculoskeletal system. Future studies will involve determining whether Hic1+ MPs play a similar role in adult skeletal repair with respect to injuries, such as bone fractures or tendon ruptures, and identifying the gene regulatory networks and molecular mechanisms involved therein. In this way, therapeutic strategies that harness Hic1+ MP’s developmental potential may be used to recapitulate embryonic skeletogenesis and enhance resolution of skeletal injuries or post-operative prognoses. 5.7 Embryonic Hic1+ MP progeny in adult tissue repair and regeneration Adult tissues harbor a variety of resident MP populations that serve vital functions in tissue homeostasis, growth, renewal and regeneration. The origin of many of these populations has remained nebulous. Tracing of E10-labelled Hic1+ MPs demonstrated a contribution to numerous terminally differentiated embryonic mesenchymal cell types, but is also sufficient to capture many of these post-natal MP populations. In particular, post-natal SkM interstitial fibroblasts are efficiently labelled at E10, and congruent with known adult fibroblast/FAP activity40,42,45,48, undergo a notable expansion following injury and during regeneration. Adult Hic1+ MPs figure 163 prominently in these processes40,45,48, indicating that embryonic Hic1+ MPs not only support limb ontogenesis, but also persist and serve as a reservoir for post-natal tissue maintenance and restoration. 5.8 Final remarks Hic1+ MPs represent a distinct mesenchymal population which emerge later in limb development, after the initiation of the LPM-derived primary skeletal elements. This MP sub-population uniquely derives from the hypaxial somite and appear coincident with migrating somite-derived myogenic precursors along with infiltrating neuronal processes. This multi-potent progenitor generates a cellular lineage hierarchy which culminates in progeny that reflect the diversity of specialized functions of MPs, such as supporting the development and maintenance of skin (PDFPs, RDFPs) and muscle (FAPs), providing HSC niche support (BMSPs), vascularization and blood vessel homeostasis (pericytes). Furthermore, within the developing appendicular skeleton, Hic1+ MPs contribute exclusively to what is referred to herein as secondary skeletal morphogenesis, events that underpin the assembly of functional connections between force-generating muscle tissue and the skeleton. In this manner, the development of the interfaces between bone and tendon (enthesis) and tendon and muscle (myotendinous junction) both require a particular Scx+ MP population which is derived from a common Hic1+ progenitor. Moreover, adult Hic1+ MPs participate in regeneration of the MTJ3, indicating that embryonic Hic1+ MPs not only support MTJ ontogenesis, but also persist and serve as a reservoir for post-natal tissue growth, renewal and regeneration. 164 In conclusion, this body of work presents a comprehensive understanding of the origin, fate, and function of Hic1+ MPs in the developing forelimbs. With the use of cell lineage tracing, single cell-omics, histology, knockout mouse models, and in vitro and ex vivo techniques, the results suggest that Hic1+ MPs represent a migratory population that originates from somitic mesenchyme and migrates into the developing limb bud where differentiation occurs into several mesenchymal fates. Some of these cell fates persist well beyond embryogenesis into adulthood and have roles in SkM repair and regeneration, and likely other homeostatic and restorative activities. The importance of Hic1 function in limb skeletogenesis was also addressed. Proper function of Hic1+ MPs is essential for the establishment of bone eminences and protuberances to which tendons attach revealing a specialized subpopulation of Hic1+ chondro-tenogenic progenitors that are the cornerstone of secondary bone morphogenesis. Further studies are required to determine the molecular and cellular mechanisms of secondary bone morphogenesis in which Hic1 is involved. Similarly, future studies should investigate whether Hic1+ MPs play a similar role in axial and craniofacial skeletogenesis, as they do in the case of the appendicular skeleton. Lastly, Hic1+ MPs are found in all vascularized tissues to certain extents, it would be interesting to determine the embryonic origin, fate and function of Hic1+ MPs in the development of organs in which Hic1+ MPs figure prominently during repair or regeneration such as dorsal skin, heart, liver, and lungs. The studies that follow will build on the present work that has laid down the groundwork for understanding Hic1+ MP biology in the context of embryonic development and skeletal morphogenesis. 165 References 1. Benjamin, M. & Ralphs, J. R. The cell and developmental biology of tendons and ligaments. Int. Rev. Cytol. 196, 85–130 (2000). 2. Brent, A. E., Schweitzer, R. & Tabin, C. J. A Somitic Compartment of Tendon Progenitors. Cell 113, 235–248 (2003). 3. Bénazet, J.-D. & Zeller, R. Vertebrate limb development: moving from classical morphogen gradients to an integrated 4-dimensional patterning system. Cold Spring Harb. Perspect. Biol. 1, a001339 (2009). 4. Hecht, M. Evolutionary Biology. (1986). 5. Hall, B. K. & Miyake, T. All for one and one for all: condensations and the initiation of skeletal development. BioEssays News Rev. Mol. Cell. Dev. Biol. 22, 138–147 (2000). 6. Helmbacher, F. & Stricker, S. Tissue cross talks governing limb muscle development and regeneration. Semin. Cell Dev. Biol. 104, 14–30 (2020). 7. Kardon, G. Muscle and tendon morphogenesis in the avian hind limb. Dev. Camb. Engl. 125, 4019–4032 (1998). 8. Kardon, G., Harfe, B. D. & Tabin, C. J. A Tcf4-Positive Mesodermal Population Provides a Prepattern for Vertebrate Limb Muscle Patterning. Dev. Cell 5, 937–944 (2003). 9. Besse, L. et al. Individual Limb Muscle Bundles Are Formed through Progressive Steps Orchestrated by Adjacent Connective Tissue Cells during Primary Myogenesis. Cell Rep. 30, 3552-3565.e6 (2020). 10. Tidball, J. G. & Lin, C. Structural changes at the myogenic cell surface during the formation of myotendinous junctions. Cell Tissue Res. 257, 77–84 (1989). 166 11. Esteves de Lima, J. et al. Unexpected contribution of fibroblasts to muscle lineage as a mechanism for limb muscle patterning. Nat. Commun. 12, 3851 (2021). 12. Yaseen, W. et al. Fibroblast fusion to the muscle fiber regulates myotendinous junction formation. Nat. Commun. 12, 3852 (2021). 13. Chevallier, A., Kieny, M. & Mauger, A. Limb-somite relationship: origin of the limb musculature. J. Embryol. Exp. Morphol. 41, 245–258 (1977). 14. Christ, B., Jacob, H. J. & Jacob, M. Experimental analysis of the origin of the wing musculature in avian embryos. Anat. Embryol. (Berl.) 150, 171–186 (1977). 15. Gaut, L. & Duprez, D. Tendon development and diseases. Wiley Interdiscip. Rev. Dev. Biol. 5, 5–23 (2016). 16. Schweitzer, R. et al. Analysis of the tendon cell fate using Scleraxis, a specific marker for tendons and ligaments. Dev. Camb. Engl. 128, 3855–3866 (2001). 17. Tozer, S. & Duprez, D. Tendon and ligament: development, repair and disease. Birth Defects Res. Part C Embryo Today Rev. 75, 226–236 (2005). 18. Chen, J. W. & Galloway, J. L. The development of zebrafish tendon and ligament progenitors. Dev. Camb. Engl. 141, 2035–2045 (2014). 19. Thomopoulos, S. et al. Decreased muscle loading delays maturation of the tendon enthesis during postnatal development. J. Orthop. Res. Off. Publ. Orthop. Res. Soc. 25, 1154–1163 (2007). 20. Kahn, J. et al. Muscle contraction is necessary to maintain joint progenitor cell fate. Dev. Cell 16, 734–743 (2009). 167 21. Schwartz, A. G., Lipner, J. H., Pasteris, J. D., Genin, G. M. & Thomopoulos, S. Muscle loading is necessary for the formation of a functional tendon enthesis. Bone 55, 44–51 (2013). 22. Benjamin, M. et al. The skeletal attachment of tendons--tendon ‘entheses’. Comp. Biochem. Physiol. A. Mol. Integr. Physiol. 133, 931–945 (2002). 23. Lu, H. H. & Thomopoulos, S. Functional attachment of soft tissues to bone: development, healing, and tissue engineering. Annu. Rev. Biomed. Eng. 15, 201–226 (2013). 24. Zelzer, E., Blitz, E., Killian, M. L. & Thomopoulos, S. Tendon‐to‐bone attachment: From development to maturity. Birth Defects Res. Part C Embryo Today Rev. 102, 101–112 (2014). 25. Blitz, E., Sharir, A., Akiyama, H. & Zelzer, E. Tendon-bone attachment unit is formed modularly by a distinct pool of Scx- and Sox9-positive progenitors. Dev. Camb. Engl. 140, 2680–2690 (2013). 26. Bi, W., Deng, J. M., Zhang, Z., Behringer, R. R. & de Crombrugghe, B. Sox9 is required for cartilage formation. Nat. Genet. 22, 85–89 (1999). 27. Akiyama, H., Chaboissier, M.-C., Martin, J. F., Schedl, A. & de Crombrugghe, B. The transcription factor Sox9 has essential roles in successive steps of the chondrocyte differentiation pathway and is required for expression of Sox5 and Sox6. Genes Dev. 16, 2813–2828 (2002). 28. Hattori, T. et al. SOX9 is a major negative regulator of cartilage vascularization, bone marrow formation and endochondral ossification. Dev. Camb. Engl. 137, 901–911 (2010). 29. Blitz, E. et al. Bone ridge patterning during musculoskeletal assembly is mediated through SCX regulation of Bmp4 at the tendon-skeleton junction. Dev. Cell 17, 861–873 (2009). 168 30. Frantz, C., Stewart, K. M. & Weaver, V. M. The extracellular matrix at a glance. J. Cell Sci. 123, 4195–4200 (2010). 31. Järveläinen, H., Sainio, A., Koulu, M., Wight, T. N. & Penttinen, R. Extracellular matrix molecules: potential targets in pharmacotherapy. Pharmacol. Rev. 61, 198–223 (2009). 32. Nagalingam, R. S., Al-Hattab, D. S. & Czubryt, M. P. What’s in a name? On fibroblast phenotype and nomenclature 1. Can. J. Physiol. Pharmacol. 97, 493–497 (2019). 33. Tallquist, M. D. & Molkentin, J. D. Redefining the identity of cardiac fibroblasts. Nat. Rev. Cardiol. 14, 484–491 (2017). 34. Kalluri, R. & Zeisberg, M. Fibroblasts in cancer. Nat. Rev. Cancer 6, 392–401 (2006). 35. Sefton, E. M. & Kardon, G. Connecting muscle development, birth defects, and evolution: An essential role for muscle connective tissue. Curr. Top. Dev. Biol. 132, 137–176 (2019). 36. Soliman, H. et al. Multipotent stromal cells: One name, multiple identities. Cell Stem Cell 28, 1690–1707 (2021). 37. Ritso, M., Tung, L. W. & Rossi, F. M. V. Emerging skeletal muscle stromal cell diversity: Functional divergence in fibro/adipogenic progenitor and mural cell populations. Exp. Cell Res. 410, 112947 (2022). 38. Mathew, S. J. et al. Connective tissue fibroblasts and Tcf4 regulate myogenesis. Dev. Camb. Engl. 138, 371–384 (2011). 39. Murphy, M. M., Lawson, J. A., Mathew, S. J., Hutcheson, D. A. & Kardon, G. Satellite cells, connective tissue fibroblasts and their interactions are crucial for muscle regeneration. Dev. Camb. Engl. 138, 3625–3637 (2011). 40. Joe, A. W. B. et al. Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nat. Cell Biol. 12, 153–163 (2010). 169 41. Uezumi, A., Fukada, S., Yamamoto, N., Takeda, S. & Tsuchida, K. Mesenchymal progenitors distinct from satellite cells contribute to ectopic fat cell formation in skeletal muscle. Nat. Cell Biol. 12, 143–152 (2010). 42. Wosczyna, M. N. et al. Mesenchymal Stromal Cells Are Required for Regeneration and Homeostatic Maintenance of Skeletal Muscle. Cell Rep. 27, 2029-2035.e5 (2019). 43. Dulauroy, S., Di Carlo, S. E., Langa, F., Eberl, G. & Peduto, L. Lineage tracing and genetic ablation of ADAM12(+) perivascular cells identify a major source of profibrotic cells during acute tissue injury. Nat. Med. 18, 1262–1270 (2012). 44. Mashinchian, O., Pisconti, A., Le Moal, E. & Bentzinger, C. F. The Muscle Stem Cell Niche in Health and Disease. Curr. Top. Dev. Biol. 126, 23–65 (2018). 45. Scott, R. W., Arostegui, M., Schweitzer, R., Rossi, F. M. V. & Underhill, T. M. Hic1 Defines Quiescent Mesenchymal Progenitor Subpopulations with Distinct Functions and Fates in Skeletal Muscle Regeneration. Cell Stem Cell 25, 797-813.e9 (2019). 46. Heredia, J. E. et al. Type 2 innate signals stimulate fibro/adipogenic progenitors to facilitate muscle regeneration. Cell 153, 376–388 (2013). 47. Uezumi, A. et al. Mesenchymal Bmp3b expression maintains skeletal muscle integrity and decreases in age-related sarcopenia. J. Clin. Invest. 131, 139617 (2021). 48. Lemos, D. R. et al. Nilotinib reduces muscle fibrosis in chronic muscle injury by promoting TNF-mediated apoptosis of fibro/adipogenic progenitors. Nat. Med. 21, 786–794 (2015). 49. Juban, G. et al. AMPK Activation Regulates LTBP4-Dependent TGF-β1 Secretion by Pro-inflammatory Macrophages and Controls Fibrosis in Duchenne Muscular Dystrophy. Cell Rep. 25, 2163-2176.e6 (2018). 170 50. Mann, C. J. et al. Aberrant repair and fibrosis development in skeletal muscle. Skelet. Muscle 1, 21 (2011). 51. Dadgar, S. et al. Asynchronous remodeling is a driver of failed regeneration in Duchenne muscular dystrophy. J. Cell Biol. 207, 139–158 (2014). 52. Contreras, O., Rossi, F. M. & Brandan, E. Adherent muscle connective tissue fibroblasts are phenotypically and biochemically equivalent to stromal fibro/adipogenic progenitors. Matrix Biol. Plus 2, 100006 (2019). 53. Uezumi, A., Ikemoto-Uezumi, M. & Tsuchida, K. Roles of nonmyogenic mesenchymal progenitors in pathogenesis and regeneration of skeletal muscle. Front. Physiol. 5, 68 (2014). 54. Lemos, D. R. et al. Functionally convergent white adipogenic progenitors of different lineages participate in a diffused system supporting tissue regeneration. Stem Cells Dayt. Ohio 30, 1152–1162 (2012). 55. Vallecillo-García, P. et al. Odd skipped-related 1 identifies a population of embryonic fibro-adipogenic progenitors regulating myogenesis during limb development. Nat. Commun. 8, 1218 (2017). 56. Driskell, R. R. & Watt, F. M. Understanding fibroblast heterogeneity in the skin. Trends Cell Biol. 25, 92–99 (2015). 57. Jinno, H. et al. Convergent genesis of an adult neural crest-like dermal stem cell from distinct developmental origins. Stem Cells Dayt. Ohio 28, 2027–2040 (2010). 58. Fernandes, K. J. L. et al. A dermal niche for multipotent adult skin-derived precursor cells. Nat. Cell Biol. 6, 1082–1093 (2004). 171 59. Driskell, R. R. et al. Distinct fibroblast lineages determine dermal architecture in skin development and repair. Nature 504, 277–281 (2013). 60. Sorrell, J. M. & Caplan, A. I. Fibroblast heterogeneity: more than skin deep. J. Cell Sci. 117, 667–675 (2004). 61. Philippeos, C. et al. Spatial and Single-Cell Transcriptional Profiling Identifies Functionally Distinct Human Dermal Fibroblast Subpopulations. J. Invest. Dermatol. 138, 811–825 (2018). 62. Biernaskie, J. et al. SKPs derive from hair follicle precursors and exhibit properties of adult dermal stem cells. Cell Stem Cell 5, 610–623 (2009). 63. Blanpain, C., Lowry, W. E., Geoghegan, A., Polak, L. & Fuchs, E. Self-renewal, multipotency, and the existence of two cell populations within an epithelial stem cell niche. Cell 118, 635–648 (2004). 64. Cotsarelis, G., Sun, T. T. & Lavker, R. M. Label-retaining cells reside in the bulge area of pilosebaceous unit: implications for follicular stem cells, hair cycle, and skin carcinogenesis. Cell 61, 1329–1337 (1990). 65. Rahmani, W. et al. Hair follicle dermal stem cells regenerate the dermal sheath, repopulate the dermal papilla, and modulate hair type. Dev. Cell 31, 543–558 (2014). 66. Rompolas, P. et al. Live imaging of stem cell and progeny behaviour in physiological hair-follicle regeneration. Nature 487, 496–499 (2012). 67. Festa, E. et al. Adipocyte lineage cells contribute to the skin stem cell niche to drive hair cycling. Cell 146, 761–771 (2011). 68. Wynn, T. A. Cellular and molecular mechanisms of fibrosis. J. Pathol. 214, 199–210 (2008). 172 69. Rinkevich, Y. et al. Skin fibrosis. Identification and isolation of a dermal lineage with intrinsic fibrogenic potential. Science 348, aaa2151 (2015). 70. Walmsley, G. G. et al. Scarless wound healing: chasing the holy grail. Plast. Reconstr. Surg. 135, 907–917 (2015). 71. Rognoni, E. et al. Inhibition of β-catenin signalling in dermal fibroblasts enhances hair follicle regeneration during wound healing. Dev. Camb. Engl. 143, 2522–2535 (2016). 72. Phan, Q. M. et al. Lef1 expression in fibroblasts maintains developmental potential in adult skin to regenerate wounds. eLife 9, e60066 (2020). 73. Mitchell, T. S., Bradley, J., Robinson, G. S., Shima, D. T. & Ng, Y.-S. RGS5 expression is a quantitative measure of pericyte coverage of blood vessels. Angiogenesis 11, 141–151 (2008). 74. Gerhardt, H. & Betsholtz, C. Endothelial-pericyte interactions in angiogenesis. Cell Tissue Res. 314, 15–23 (2003). 75. Armulik, A., Genové, G. & Betsholtz, C. Pericytes: developmental, physiological, and pathological perspectives, problems, and promises. Dev. Cell 21, 193–215 (2011). 76. Sims, D. E. & Westfall, J. A. Analysis of relationships between pericytes and gas exchange capillaries in neonatal and mature bovine lungs. Microvasc. Res. 25, 333–342 (1983). 77. Nehls, V. & Drenckhahn, D. Heterogeneity of microvascular pericytes for smooth muscle type alpha-actin. J. Cell Biol. 113, 147–154 (1991). 78. Ozerdem, U., Grako, K. A., Dahlin-Huppe, K., Monosov, E. & Stallcup, W. B. NG2 proteoglycan is expressed exclusively by mural cells during vascular morphogenesis. Dev. Dyn. Off. Publ. Am. Assoc. Anat. 222, 218–227 (2001). 173 79. Lindahl, P., Johansson, B. R., Levéen, P. & Betsholtz, C. Pericyte loss and microaneurysm formation in PDGF-B-deficient mice. Science 277, 242–245 (1997). 80. Bondjers, C. et al. Transcription profiling of platelet-derived growth factor-B-deficient mouse embryos identifies RGS5 as a novel marker for pericytes and vascular smooth muscle cells. Am. J. Pathol. 162, 721–729 (2003). 81. Daneman, R., Zhou, L., Kebede, A. A. & Barres, B. A. Pericytes are required for blood-brain barrier integrity during embryogenesis. Nature 468, 562–566 (2010). 82. Armulik, A. et al. Pericytes regulate the blood-brain barrier. Nature 468, 557–561 (2010). 83. Stratman, A. N., Malotte, K. M., Mahan, R. D., Davis, M. J. & Davis, G. E. Pericyte recruitment during vasculogenic tube assembly stimulates endothelial basement membrane matrix formation. Blood 114, 5091–5101 (2009). 84. Yemisci, M. et al. Pericyte contraction induced by oxidative-nitrative stress impairs capillary reflow despite successful opening of an occluded cerebral artery. Nat. Med. 15, 1031–1037 (2009). 85. Paquet-Fifield, S. et al. A role for pericytes as microenvironmental regulators of human skin tissue regeneration. J. Clin. Invest. 119, 2795–2806 (2009). 86. Bergwerff, M., Verberne, M. E., DeRuiter, M. C., Poelmann, R. E. & Gittenberger-de Groot, A. C. Neural crest cell contribution to the developing circulatory system: implications for vascular morphology? Circ. Res. 82, 221–231 (1998). 87. Etchevers, H. C., Vincent, C., Le Douarin, N. M. & Couly, G. F. The cephalic neural crest provides pericytes and smooth muscle cells to all blood vessels of the face and forebrain. Dev. Camb. Engl. 128, 1059–1068 (2001). 174 88. Dias Moura Prazeres, P. H. et al. Pericytes are heterogeneous in their origin within the same tissue. Dev. Biol. 427, 6–11 (2017). 89. Pouget, C., Pottin, K. & Jaffredo, T. Sclerotomal origin of vascular smooth muscle cells and pericytes in the embryo. Dev. Biol. 315, 437–447 (2008). 90. Yamaguchi, T. P., Dumont, D. J., Conlon, R. A., Breitman, M. L. & Rossant, J. flk-1, an flt-related receptor tyrosine kinase is an early marker for endothelial cell precursors. Dev. Camb. Engl. 118, 489–498 (1993). 91. Yamashita, J. et al. Flk1-positive cells derived from embryonic stem cells serve as vascular progenitors. Nature 408, 92–96 (2000). 92. Darland, D. C. & D’Amore, P. A. Blood vessel maturation: vascular development comes of age. J. Clin. Invest. 103, 157–158 (1999). 93. Asahina, K., Zhou, B., Pu, W. T. & Tsukamoto, H. Septum transversum-derived mesothelium gives rise to hepatic stellate cells and perivascular mesenchymal cells in developing mouse liver. Hepatol. Baltim. Md 53, 983–995 (2011). 94. Eroschenko, V. P. & Fiore, M. S. H. di. DiFiore’s atlas of histology with functional correlations. (Wolters Kluwer Health/Lippincott Williams & Wilkins, 2008). 95. Comazzetto, S., Shen, B. & Morrison, S. J. Niches that regulate stem cells and hematopoiesis in adult bone marrow. Dev. Cell 56, 1848–1860 (2021). 96. Liggett, L. A. & Sankaran, V. G. Unraveling Hematopoiesis through the Lens of Genomics. Cell 182, 1384–1400 (2020). 97. Morrison, S. J. & Spradling, A. C. Stem cells and niches: mechanisms that promote stem cell maintenance throughout life. Cell 132, 598–611 (2008). 175 98. Kiel, M. J. et al. SLAM family receptors distinguish hematopoietic stem and progenitor cells and reveal endothelial niches for stem cells. Cell 121, 1109–1121 (2005). 99. Nombela-Arrieta, C. et al. Quantitative imaging of haematopoietic stem and progenitor cell localization and hypoxic status in the bone marrow microenvironment. Nat. Cell Biol. 15, 533–543 (2013). 100. Acar, M. et al. Deep imaging of bone marrow shows non-dividing stem cells are mainly perisinusoidal. Nature 526, 126–130 (2015). 101. Sugiyama, T., Kohara, H., Noda, M. & Nagasawa, T. Maintenance of the hematopoietic stem cell pool by CXCL12-CXCR4 chemokine signaling in bone marrow stromal cell niches. Immunity 25, 977–988 (2006). 102. Méndez-Ferrer, S. et al. Mesenchymal and haematopoietic stem cells form a unique bone marrow niche. Nature 466, 829–834 (2010). 103. Ding, L., Saunders, T. L., Enikolopov, G. & Morrison, S. J. Endothelial and perivascular cells maintain haematopoietic stem cells. Nature 481, 457–462 (2012). 104. Greenbaum, A. et al. CXCL12 in early mesenchymal progenitors is required for haematopoietic stem-cell maintenance. Nature 495, 227–230 (2013). 105. Park, D. et al. Endogenous bone marrow MSCs are dynamic, fate-restricted participants in bone maintenance and regeneration. Cell Stem Cell 10, 259–272 (2012). 106. Worthley, D. L. et al. Gremlin 1 identifies a skeletal stem cell with bone, cartilage, and reticular stromal potential. Cell 160, 269–284 (2015). 107. Seike, M., Omatsu, Y., Watanabe, H., Kondoh, G. & Nagasawa, T. Stem cell niche-specific Ebf3 maintains the bone marrow cavity. Genes Dev. 32, 359–372 (2018). 176 108. Ding, L. & Morrison, S. J. Haematopoietic stem cells and early lymphoid progenitors occupy distinct bone marrow niches. Nature 495, 231–235 (2013). 109. Himburg, H. A. et al. Distinct Bone Marrow Sources of Pleiotrophin Control Hematopoietic Stem Cell Maintenance and Regeneration. Cell Stem Cell 23, 370-381.e5 (2018). 110. Zhou, B. O., Yue, R., Murphy, M. M., Peyer, J. G. & Morrison, S. J. Leptin-receptor-expressing mesenchymal stromal cells represent the main source of bone formed by adult bone marrow. Cell Stem Cell 15, 154–168 (2014). 111. Tikhonova, A. N. et al. The bone marrow microenvironment at single-cell resolution. Nature 569, 222–228 (2019). 112. Baryawno, N. et al. A Cellular Taxonomy of the Bone Marrow Stroma in Homeostasis and Leukemia. Cell 177, 1915-1932.e16 (2019). 113. Baccin, C. et al. Combined single-cell and spatial transcriptomics reveal the molecular, cellular and spatial bone marrow niche organization. Nat. Cell Biol. 22, 38–48 (2020). 114. Zhou, B. O. et al. Bone marrow adipocytes promote the regeneration of stem cells and haematopoiesis by secreting SCF. Nat. Cell Biol. 19, 891–903 (2017). 115. Matsushita, Y. et al. A Wnt-mediated transformation of the bone marrow stromal cell identity orchestrates skeletal regeneration. Nat. Commun. 11, 332 (2020). 116. Shen, B. et al. A mechanosensitive peri-arteriolar niche for osteogenesis and lymphopoiesis. Nature 591, 438–444 (2021). 117. Ono, N., Balani, D. H. & Kronenberg, H. M. Stem and progenitor cells in skeletal development. Curr. Top. Dev. Biol. 133, 1–24 (2019). 177 118. Bianco, P. et al. The meaning, the sense and the significance: translating the science of mesenchymal stem cells into medicine. Nat. Med. 19, 35–42 (2013). 119. Maes, C. et al. Osteoblast precursors, but not mature osteoblasts, move into developing and fractured bones along with invading blood vessels. Dev. Cell 19, 329–344 (2010). 120. Sheng, G. The developmental basis of mesenchymal stem/stromal cells (MSCs). BMC Dev. Biol. 15, 44 (2015). 121. Grcevic, D. et al. In vivo fate mapping identifies mesenchymal progenitor cells. Stem Cells Dayt. Ohio 30, 187–196 (2012). 122. Shin, M., Nagai, H. & Sheng, G. Notch mediates Wnt and BMP signals in the early separation of smooth muscle progenitors and blood/endothelial common progenitors. Dev. Camb. Engl. 136, 595–603 (2009). 123. Isenmann, S. et al. TWIST family of basic helix-loop-helix transcription factors mediate human mesenchymal stem cell growth and commitment. Stem Cells Dayt. Ohio 27, 2457–2468 (2009). 124. Füchtbauer, E. M. Expression of M-twist during postimplantation development of the mouse. Dev. Dyn. Off. Publ. Am. Assoc. Anat. 204, 316–322 (1995). 125. Pearse, R. V., Scherz, P. J., Campbell, J. K. & Tabin, C. J. A cellular lineage analysis of the chick limb bud. Dev. Biol. 310, 388–400 (2007). 126. Nagoshi, N. et al. Ontogeny and multipotency of neural crest-derived stem cells in mouse bone marrow, dorsal root ganglia, and whisker pad. Cell Stem Cell 2, 392–403 (2008). 127. Glejzer, A. et al. Wnt1 and BMP2: two factors recruiting multipotent neural crest progenitors isolated from adult bone marrow. Cell. Mol. Life Sci. CMLS 68, 2101–2114 (2011). 178 128. Komada, Y. et al. Origins and properties of dental, thymic, and bone marrow mesenchymal cells and their stem cells. PloS One 7, e46436 (2012). 129. Morikawa, S. et al. Development of mesenchymal stem cells partially originate from the neural crest. Biochem. Biophys. Res. Commun. 379, 1114–1119 (2009). 130. Isern, J. et al. The neural crest is a source of mesenchymal stem cells with specialized hematopoietic stem cell niche function. eLife https://elifesciences.org/articles/03696 (2014) doi:10.7554/eLife.03696. 131. Stassart, R. M., Möbius, W., Nave, K.-A. & Edgar, J. M. The Axon-Myelin Unit in Development and Degenerative Disease. Front. Neurosci. 12, 467 (2018). 132. Carr, M. J. et al. Mesenchymal Precursor Cells in Adult Nerves Contribute to Mammalian Tissue Repair and Regeneration. Cell Stem Cell 24, 240-256.e9 (2019). 133. Stierli, S. et al. The regulation of the homeostasis and regeneration of peripheral nerve is distinct from the CNS and independent of a stem cell population. Dev. Camb. Engl. 145, dev170316 (2018). 134. Hidmark, A. S., Nawroth, P. P. & Fleming, T. Analysis of Immune Cells in Single Sciatic Nerves and Dorsal Root Ganglion from a Single Mouse Using Flow Cytometry. J. Vis. Exp. JoVE (2017) doi:10.3791/56538. 135. Joseph, N. M. et al. Neural crest stem cells undergo multilineage differentiation in developing peripheral nerves to generate endoneurial fibroblasts in addition to Schwann cells. Dev. Camb. Engl. 131, 5599–5612 (2004). 136. Gerber, D. et al. Transcriptional profiling of mouse peripheral nerves to the single-cell level to build a sciatic nerve ATlas (SNAT). eLife 10, e58591 (2021). 137. Gartner, L. P. & Hiatt, J. L. Color textbook of histology. (W.B. Saunders, 2001). 179 138. Junqueira, L. C. U. & Carneiro, J. Basic histology: text & atlas. (McGraw-Hill, 2005). 139. Morris, R. J. & Beech, J. N. Differential expression of Thy-1 on the various components of connective tissue of rat nerve during postnatal development. Dev. Biol. 102, 32–42 (1984). 140. Ma, D. et al. A Subpopulation of Foxj1-Expressing, Nonmyelinating Schwann Cells of the Peripheral Nervous System Contribute to Schwann Cell Remyelination in the Central Nervous System. J. Neurosci. Off. J. Soc. Neurosci. 38, 9228–9239 (2018). 141. Kania, A., Johnson, R. L. & Jessell, T. M. Coordinate roles for LIM homeobox genes in directing the dorsoventral trajectory of motor axons in the vertebrate limb. Cell 102, 161–173 (2000). 142. Huber, A. B., Kolodkin, A. L., Ginty, D. D. & Cloutier, J.-F. Signaling at the growth cone: ligand-receptor complexes and the control of axon growth and guidance. Annu. Rev. Neurosci. 26, 509–563 (2003). 143. Huettl, R.-E., Haehl, T. & Huber, A. B. Fasciculation and guidance of spinal motor axons in the absence of FGFR2 signaling. PloS One 7, e41095 (2012). 144. Bronner, M. E. & Simões-Costa, M. The Neural Crest Migrating into the Twenty-First Century. Curr. Top. Dev. Biol. 116, 115 (2016). 145. Brockes, J. P., Fields, K. L. & Raff, M. C. Studies on cultured rat Schwann cells. I. Establishment of purified populations from cultures of peripheral nerve. Brain Res. 165, 105–118 (1979). 146. Assouline, J. G., Bosch, E. P. & Lim, R. Purification of rat Schwann cells from cultures of peripheral nerve: an immunoselective method using surfaces coated with anti-immunoglobulin antibodies. Brain Res. 277, 389–392 (1983). 180 147. Richard, L., Védrenne, N., Vallat, J.-M. & Funalot, B. Characterization of Endoneurial Fibroblast-like Cells from Human and Rat Peripheral Nerves. J. Histochem. Cytochem. Off. J. Histochem. Soc. 62, 424–435 (2014). 148. Kumar, A., Godwin, J. W., Gates, P. B., Garza-Garcia, A. A. & Brockes, J. P. Molecular basis for the nerve dependence of limb regeneration in an adult vertebrate. Science 318, 772–777 (2007). 149. Kumar, A. & Brockes, J. P. Nerve dependence in tissue, organ, and appendage regeneration. Trends Neurosci. 35, 691–699 (2012). 150. Takeo, M. et al. Wnt activation in nail epithelium couples nail growth to digit regeneration. Nature 499, 228–232 (2013). 151. Rinkevich, Y. et al. Clonal analysis reveals nerve-dependent and independent roles on mammalian hind limb tissue maintenance and regeneration. Proc. Natl. Acad. Sci. U. S. A. 111, 9846–9851 (2014). 152. Johnston, A. P. W. et al. Dedifferentiated Schwann Cell Precursors Secreting Paracrine Factors Are Required for Regeneration of the Mammalian Digit Tip. Cell Stem Cell 19, 433–448 (2016). 153. Clements, M. P. et al. The Wound Microenvironment Reprograms Schwann Cells to Invasive Mesenchymal-like Cells to Drive Peripheral Nerve Regeneration. Neuron 96, 98-114.e7 (2017). 154. Cattin, A.-L. et al. Macrophage-Induced Blood Vessels Guide Schwann Cell-Mediated Regeneration of Peripheral Nerves. Cell 162, 1127–1139 (2015). 155. Cattin, A.-L. & Lloyd, A. C. The multicellular complexity of peripheral nerve regeneration. Curr. Opin. Neurobiol. 39, 38–46 (2016). 181 156. Fujii, H. et al. Methylation of the HIC-1 candidate tumor suppressor gene in human breast cancer. Oncogene 16, 2159–2164 (1998). 157. Chen, W. et al. Epigenetic and genetic loss of Hic1 function accentuates the role of p53 in tumorigenesis. Cancer Cell 6, 387–398 (2004). 158. Hao, M. et al. HIC1 loss promotes prostate cancer metastasis by triggering epithelial-mesenchymal transition. J. Pathol. 242, 409–420 (2017). 159. Konishi, H. et al. Detailed deletion mapping suggests the involvement of a tumor suppressor gene at 17p13.3, distal to p53, in the pathogenesis of lung cancers. Oncogene 17, 2095–2100 (1998). 160. Wales, M. M. et al. p53 activates expression of HIC-1, a new candidate tumour suppressor gene on 17p13.3. Nat. Med. 1, 570–577 (1995). 161. Carter, M. G. et al. Mice deficient in the candidate tumor suppressor gene Hic1 exhibit developmental defects of structures affected in the Miller-Dieker syndrome. Hum. Mol. Genet. 9, 413–419 (2000). 162. Chen, W. Y. et al. Heterozygous disruption of Hic1 predisposes mice to a gender-dependent spectrum of malignant tumors. Nat. Genet. 33, 197–202 (2003). 163. Dehennaut, V., Loison, I., Boulay, G., Van Rechem, C. & Leprince, D. Identification of p21 (CIP1/WAF1) as a direct target gene of HIC1 (Hypermethylated In Cancer 1). Biochem. Biophys. Res. Commun. 430, 49–53 (2013). 164. Van Rechem, C. et al. Differential regulation of HIC1 target genes by CtBP and NuRD, via an acetylation/SUMOylation switch, in quiescent versus proliferating cells. Mol. Cell. Biol. 30, 4045–4059 (2010). 182 165. Sherr, C. J. & Roberts, J. M. CDK inhibitors: positive and negative regulators of G1-phase progression. Genes Dev. 13, 1501–1512 (1999). 166. Grimm, C. et al. Isolation and embryonic expression of the novel mouse gene Hic1, the homologue of HIC1, a candidate gene for the Miller-Dieker syndrome. Hum. Mol. Genet. 8, 697–710 (1999). 167. Deltour, S., Guerardel, C. & Leprince, D. Recruitment of SMRT/N-CoR-mSin3A-HDAC-repressing complexes is not a general mechanism for BTB/POZ transcriptional repressors: the case of HIC-1 and gammaFBP-B. Proc. Natl. Acad. Sci. U. S. A. 96, 14831–14836 (1999). 168. Deltour, S., Pinte, S., Guerardel, C., Wasylyk, B. & Leprince, D. The human candidate tumor suppressor gene HIC1 recruits CtBP through a degenerate GLDLSKK motif. Mol. Cell. Biol. 22, 4890–4901 (2002). 169. Pinte, S. et al. The tumor suppressor gene HIC1 (hypermethylated in cancer 1) is a sequence-specific transcriptional repressor: definition of its consensus binding sequence and analysis of its DNA binding and repressive properties. J. Biol. Chem. 279, 38313–38324 (2004). 170. Chen, W. Y. et al. Tumor suppressor HIC1 directly regulates SIRT1 to modulate p53-dependent DNA-damage responses. Cell 123, 437–448 (2005). 171. Briones, V. R., Chen, S., Riegel, A. T. & Lechleider, R. J. Mechanism of fibroblast growth factor-binding protein 1 repression by TGF-beta. Biochem. Biophys. Res. Commun. 345, 595–601 (2006). 172. Briggs, K. J. et al. Cooperation between the Hic1 and Ptch1 tumor suppressors in medulloblastoma. Genes Dev. 22, 770–785 (2008). 183 173. Valenta, T., Lukas, J., Doubravska, L., Fafilek, B. & Korinek, V. HIC1 attenuates Wnt signaling by recruitment of TCF-4 and beta-catenin to the nuclear bodies. EMBO J. 25, 2326–2337 (2006). 174. Friedenstein, A. J., Chailakhjan, R. K. & Lalykina, K. S. The development of fibroblast colonies in monolayer cultures of guinea-pig bone marrow and spleen cells. Cell Tissue Kinet. 3, 393–403 (1970). 175. Friedenstein, A. & Kuralesova, A. I. Osteogenic precursor cells of bone marrow in radiation chimeras. Transplantation 12, 99–108 (1971). 176. Friedenstein, A. J. et al. Precursors for fibroblasts in different populations of hematopoietic cells as detected by the in vitro colony assay method. Exp. Hematol. 2, 83–92 (1974). 177. Heersche J.N.M. & Kanis J.A. Bone and Mineral Research. (Elsevier, 1990). 178. Howlett, C. R. et al. Mineralization in in vitro cultures of rabbit marrow stromal cells. Clin. Orthop. 251–263 (1986). 179. Friedenstein, A. J., Latzinik, N. W., Grosheva, A. G. & Gorskaya, U. F. Marrow microenvironment transfer by heterotopic transplantation of freshly isolated and cultured cells in porous sponges. Exp. Hematol. 10, 217–227 (1982). 180. Schofield, R., Lorimore, S. A. & Wright, E. G. Ectopic implantation studies using Sl/Sld marrow and recipients. Exp. Hematol. 15, 217–220 (1987). 181. Caplan, A. I. Mesenchymal stem cells. J. Orthop. Res. Off. Publ. Orthop. Res. Soc. 9, 641–650 (1991). 182. Caplan, A. I. The mesengenic process. Clin. Plast. Surg. 21, 429–435 (1994). 183. Pittenger, M. F. et al. Multilineage potential of adult human mesenchymal stem cells. Science 284, 143–147 (1999). 184 184. Caplan, A. I. Mesenchymal Stem Cells: Time to Change the Name! Stem Cells Transl. Med. 6, 1445–1451 (2017). 185. Sacchetti, B. et al. Self-renewing osteoprogenitors in bone marrow sinusoids can organize a hematopoietic microenvironment. Cell 131, 324–336 (2007). 186. da Silva Meirelles, L., Chagastelles, P. C. & Nardi, N. B. Mesenchymal stem cells reside in virtually all post-natal organs and tissues. J. Cell Sci. 119, 2204–2213 (2006). 187. Caplan, A. I. All MSCs are pericytes? Cell Stem Cell 3, 229–230 (2008). 188. Crisan, M. et al. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell 3, 301–313 (2008). 189. Bianco, P. & Robey, P. G. Skeletal stem cells. Dev. Camb. Engl. 142, 1023–1027 (2015). 190. Sacchetti, B. et al. No Identical ‘Mesenchymal Stem Cells’ at Different Times and Sites: Human Committed Progenitors of Distinct Origin and Differentiation Potential Are Incorporated as Adventitial Cells in Microvessels. Stem Cell Rep. 6, 897–913 (2016). 191. Sipp, D., Robey, P. G. & Turner, L. Clear up this stem-cell mess. Nature 561, 455–457 (2018). 192. Caplan, A. I. There Is No ‘Stem Cell Mess’. Tissue Eng. Part B Rev. 25, 291–293 (2019). 193. Plikus, M. V. et al. Fibroblasts: Origins, definitions, and functions in health and disease. Cell 184, 3852–3872 (2021). 194. Pradhan, R. N., Krishnamurty, A. T., Fletcher, A. L., Turley, S. J. & Müller, S. A bird’s eye view of fibroblast heterogeneity: A pan-disease, pan-cancer perspective. Immunol. Rev. 302, 299–320 (2021). 195. Giordani, L. et al. High-Dimensional Single-Cell Cartography Reveals Novel Skeletal Muscle-Resident Cell Populations. Mol. Cell 74, 609-621.e6 (2019). 185 196. Buechler, M. B. et al. Cross-tissue organization of the fibroblast lineage. Nature 593, 575–579 (2021). 197. Kunisaki, Y. et al. Arteriolar niches maintain haematopoietic stem cell quiescence. Nature 502, 637–643 (2013). 198. Ding, L., Saunders, T. L., Enikolopov, G. & Morrison, S. J. Endothelial and perivascular cells maintain haematopoietic stem cells. Nature 481, 457–462 (2012). 199. Quante, M. et al. Bone marrow-derived myofibroblasts contribute to the mesenchymal stem cell niche and promote tumor growth. Cancer Cell 19, 257–272 (2011). 200. Kramann, R. et al. Perivascular Gli1+ progenitors are key contributors to injury-induced organ fibrosis. Cell Stem Cell 16, 51–66 (2015). 201. Bonyadi, M. et al. Mesenchymal progenitor self-renewal deficiency leads to age-dependent osteoporosis in Sca-1/Ly-6A null mice. Proc. Natl. Acad. Sci. U. S. A. 100, 5840–5845 (2003). 202. Houlihan, D. D. et al. Isolation of mouse mesenchymal stem cells on the basis of expression of Sca-1 and PDGFR-α. Nat. Protoc. 7, 2103–2111 (2012). 203. Abbasi, S. et al. Distinct Regulatory Programs Control the Latent Regenerative Potential of Dermal Fibroblasts during Wound Healing. Cell Stem Cell 27, 396-412.e6 (2020). 204. Soliman, H. et al. Pathogenic Potential of Hic1-Expressing Cardiac Stromal Progenitors. Cell Stem Cell 26, 205-220.e8 (2020). 205. Sulston, J. E., Schierenberg, E., White, J. G. & Thomson, J. N. The embryonic cell lineage of the nematode Caenorhabditis elegans. Dev. Biol. 100, 64–119 (1983). 206. Kimmel, C. B., Warga, R. M. & Schilling, T. F. Origin and organization of the zebrafish fate map. Dev. Camb. Engl. 108, 581–594 (1990). 186 207. Axelrod, D. Carbocyanine dye orientation in red cell membrane studied by microscopic fluorescence polarization. Biophys. J. 26, 557–573 (1979). 208. Serbedzija, G. N., Bronner-Fraser, M. & Fraser, S. E. A vital dye analysis of the timing and pathways of avian trunk neural crest cell migration. Dev. Camb. Engl. 106, 809–816 (1989). 209. Eagleson, G. W. & Harris, W. A. Mapping of the presumptive brain regions in the neural plate of Xenopus laevis. J. Neurobiol. 21, 427–440 (1990). 210. Lemischka, I. R., Raulet, D. H. & Mulligan, R. C. Developmental potential and dynamic behavior of hematopoietic stem cells. Cell 45, 917–927 (1986). 211. Price, J. Retroviruses and the study of cell lineage. Dev. Camb. Engl. 101, 409–419 (1987). 212. Le Douarin, N. & Kalcheim, C. The neural crest. (Cambridge University Press, 1999). 213. Logan, M. et al. Expression of Cre Recombinase in the developing mouse limb bud driven by a Prxl enhancer. Genes. N. Y. N 2000 33, 77–80 (2002). 214. Lang, D. et al. Pax3 functions at a nodal point in melanocyte stem cell differentiation. Nature 433, 884–887 (2005). 215. Atit, R. et al. Beta-catenin activation is necessary and sufficient to specify the dorsal dermal fate in the mouse. Dev. Biol. 296, 164–176 (2006). 216. Sebo, Z. L., Jeffery, E., Holtrup, B. & Rodeheffer, M. S. A mesodermal fate map for adipose tissue. Dev. Camb. Engl. 145, dev166801 (2018). 217. Buckingham, M. E. & Meilhac, S. M. Tracing cells for tracking cell lineage and clonal behavior. Dev. Cell 21, 394–409 (2011). 218. Blanpain, C. & Simons, B. D. Unravelling stem cell dynamics by lineage tracing. Nat. Rev. Mol. Cell Biol. 14, 489–502 (2013). 187 219. Indra, A. K. et al. Temporally-controlled site-specific mutagenesis in the basal layer of the epidermis: comparison of the recombinase activity of the tamoxifen-inducible Cre-ER(T) and Cre-ER(T2) recombinases. Nucleic Acids Res. 27, 4324–4327 (1999). 220. Jaisser, F. Inducible gene expression and gene modification in transgenic mice. J. Am. Soc. Nephrol. JASN 11 Suppl 16, S95–S100 (2000). 221. Kretzschmar, K. & Watt, F. M. Lineage tracing. Cell 148, 33–45 (2012). 222. Eisner, C. PDGFRα+ and HIC1+ mesenchymal progenitors in murine skeletal tissue ; roles in homeostasis and regeneration. (2020). 223. Uchida, A. et al. Development and function of smooth muscle cells is modulated by Hic1 in mouse testis. Dev. Camb. Engl. 147, dev185884 (2020). 224. Gilbert, S. F. Developmental biology. (Sinauer Associates, 2000). 225. White, A. & Wallis, G. Endochondral ossification: a delicate balance between growth and mineralisation. Curr. Biol. CB 11, R589-591 (2001). 226. Mackie, E. J., Ahmed, Y. A., Tatarczuch, L., Chen, K.-S. & Mirams, M. Endochondral ossification: how cartilage is converted into bone in the developing skeleton. Int. J. Biochem. Cell Biol. 40, 46–62 (2008). 227. Pizette, S. & Niswander, L. BMPs are required at two steps of limb chondrogenesis: formation of prechondrogenic condensations and their differentiation into chondrocytes. Dev. Biol. 219, 237–249 (2000). 228. Hoffman, L. M. et al. BMP action in skeletogenesis involves attenuation of retinoid signaling. J. Cell Biol. 174, 101–113 (2006). 188 229. Barna, M. & Niswander, L. Visualization of cartilage formation: insight into cellular properties of skeletal progenitors and chondrodysplasia syndromes. Dev. Cell 12, 931–941 (2007). 230. Dranse, H. J., Sampaio, A. V., Petkovich, M. & Underhill, T. M. Genetic deletion of Cyp26b1 negatively impacts limb skeletogenesis by inhibiting chondrogenesis. J. Cell Sci. 124, 2723–2734 (2011). 231. Ahrens, P. B., Solursh, M. & Reiter, R. S. Stage-related capacity for limb chondrogenesis in cell culture. Dev. Biol. 60, 69–82 (1977). 232. Solursh, M., Ahrens, P. B. & Reiter, R. S. A tissue culture analysis of the steps in limb chondrogenesis. In Vitro 14, 51–61 (1978). 233. Niswander, L. Methods in avian embryology experimental and molecular manipulation of the embryonic chick limb. Methods Cell Biol. 87, 135–152 (2008). 234. Daniels, K., Reiter, R. & Solursh, M. Micromass cultures of limb and other mesenchyme. Methods Cell Biol. 51, 237–247 (1996). 235. Underhill, T. M., Dranse, H. J. & Hoffman, L. M. Analysis of chondrogenesis using micromass cultures of limb mesenchyme. Methods Mol. Biol. Clifton NJ 1130, 251–265 (2014). 236. Bell, D. M. et al. SOX9 directly regulates the type-II collagen gene. Nat. Genet. 16, 174–178 (1997). 237. Meech, R., Edelman, D. B., Jones, F. S. & Makarenkova, H. P. The homeobox transcription factor Barx2 regulates chondrogenesis during limb development. Dev. Camb. Engl. 132, 2135–2146 (2005). 189 238. Lorda-Diez, C. I., Montero, J. A., Martinez-Cue, C., Garcia-Porrero, J. A. & Hurle, J. M. Transforming growth factors beta coordinate cartilage and tendon differentiation in the developing limb mesenchyme. J. Biol. Chem. 284, 29988–29996 (2009). 239. Bruce, S. J. et al. Inactivation of Patched1 in the mouse limb has novel inhibitory effects on the chondrogenic program. J. Biol. Chem. 285, 27967–27981 (2010). 240. Shimono, K. et al. Inhibition of ectopic bone formation by a selective retinoic acid receptor alpha-agonist: a new therapy for heterotopic ossification? J. Orthop. Res. Off. Publ. Orthop. Res. Soc. 28, 271–277 (2010). 241. Jensen, C. & Teng, Y. Is It Time to Start Transitioning From 2D to 3D Cell Culture? Front. Mol. Biosci. 7, 33 (2020). 242. Wolpert, L. Vertebrate limb development and malformations. Pediatr. Res. 46, 247–254 (1999). 243. Arostegui, M. & Underhill, T. M. Murine Limb Bud Organ Cultures for Studying Musculoskeletal Development. Methods Mol. Biol. Clifton NJ 2230, 115–137 (2021). 244. Smith, E., Kanczler, JM, & Oreffo, ROC. A new take on an old story: chick limb organ culture for skeletal niche development and regenerative medicine evaluation. Eur. Cell. Mater. 26, 91–106 (2013). 245. Diether Neubert & Hans-Jürgen Barrach. Techniques Applicable to Study Morphogenetic Differentiation of Limb Buds in Organ Culture. in Methods in Prenatal Toxicology 241–251 (Georg Thieme Publishers, 1977). 246. Ueda, H. R. et al. Tissue clearing and its applications in neuroscience. Nat. Rev. Neurosci. 21, 61–79 (2020). 190 247. Sefton, E. M., Gallardo, M. & Kardon, G. Developmental origin and morphogenesis of the diaphragm, an essential mammalian muscle. Dev. Biol. 440, 64–73 (2018). 248. Pryce, B. A., Brent, A. E., Murchison, N. D., Tabin, C. J. & Schweitzer, R. Generation of transgenic tendon reporters, ScxGFP and ScxAP, using regulatory elements of the scleraxis gene. Dev. Dyn. Off. Publ. Am. Assoc. Anat. 236, 1677–1682 (2007). 249. Tainaka, K. et al. Whole-body imaging with single-cell resolution by tissue decolorization. Cell 159, 911–924 (2014). 250. Hama, H. et al. Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain. Nat. Neurosci. 14, 1481–1488 (2011). 251. Cao, J. et al. The single-cell transcriptional landscape of mammalian organogenesis. Nature 566, 496–502 (2019). 252. Cao, C. et al. Comprehensive single-cell transcriptome lineages of a proto-vertebrate. Nature 571, 349–354 (2019). 253. Satija, R., Farrell, J. A., Gennert, D., Schier, A. F. & Regev, A. Spatial reconstruction of single-cell gene expression data. Nat. Biotechnol. 33, 495–502 (2015). 254. Nowotschin, S. et al. The emergent landscape of the mouse gut endoderm at single-cell resolution. Nature 569, 361–367 (2019). 255. Setty, M. et al. Characterization of cell fate probabilities in single-cell data with Palantir. Nat. Biotechnol. 37, 451–460 (2019). 256. van Dijk, D. et al. Recovering Gene Interactions from Single-Cell Data Using Data Diffusion. Cell 174, 716-729.e27 (2018). 257. Farrell, J. A. et al. Single-cell reconstruction of developmental trajectories during zebrafish embryogenesis. Science 360, eaar3131 (2018). 191 258. Pliner, H. A. et al. Cicero Predicts cis-Regulatory DNA Interactions from Single-Cell Chromatin Accessibility Data. Mol. Cell 71, 858-871.e8 (2018). 259. Stuart, T. et al. Comprehensive Integration of Single-Cell Data. Cell 177, 1888-1902.e21 (2019). 260. Scott, R. W. & Underhill, T. M. Methods and Strategies for Lineage Tracing of Mesenchymal Progenitor Cells. Methods Mol. Biol. Clifton NJ 1416, 171–203 (2016). 261. Rigueur, D. & Lyons, K. M. Whole-mount skeletal staining. Methods Mol. Biol. Clifton NJ 1130, 113–121 (2014). 262. Armand, O., Boutineau, A. M., Mauger, A., Pautou, M. P. & Kieny, M. Origin of satellite cells in avian skeletal muscles. Arch. Anat. Microsc. Morphol. Exp. 72, 163–181 (1983). 263. Lepper, C. & Fan, C.-M. Inducible lineage tracing of Pax7-descendant cells reveals embryonic origin of adult satellite cells. Genes. N. Y. N 2000 48, 424–436 (2010). 264. Seale, P. et al. Pax7 is required for the specification of myogenic satellite cells. Cell 102, 777–786 (2000). 265. Kuang, S., Kuroda, K., Le Grand, F. & Rudnicki, M. A. Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell 129, 999–1010 (2007). 266. Reid, J. M. et al. Pharmacokinetics of endoxifen and tamoxifen in female mice: implications for comparative in vivo activity studies. Cancer Chemother. Pharmacol. 74, 1271–1278 (2014). 267. Buckingham, M. Making muscle in mammals. Trends Genet. TIG 8, 144–148 (1992). 268. Hutcheson, D. A., Zhao, J., Merrell, A., Haldar, M. & Kardon, G. Embryonic and fetal limb myogenic cells are derived from developmentally distinct progenitors and have different requirements for beta-catenin. Genes Dev. 23, 997–1013 (2009). 192 269. Deutsch, U., Dressler, G. R. & Gruss, P. Pax 1, a member of a paired box homologous murine gene family, is expressed in segmented structures during development. Cell 53, 617–625 (1988). 270. Wallin, J. et al. The role of Pax-1 in axial skeleton development. Dev. Camb. Engl. 120, 1109–1121 (1994). 271. Hassan, H. M. et al. Regulation of Active DNA Demethylation through RAR-Mediated Recruitment of a TET/TDG Complex. Cell Rep. 19, 1685–1697 (2017). 272. Choi, S., Ferrari, G. & Tedesco, F. S. Cellular dynamics of myogenic cell migration: molecular mechanisms and implications for skeletal muscle cell therapies. EMBO Mol. Med. 12, e12357 (2020). 273. Barriga, E. H. & Theveneau, E. In vivo Neural Crest Cell Migration Is Controlled by ‘Mixotaxis’. Front. Physiol. 11, 586432 (2020). 274. Wilting, J. et al. Angiogenic potential of the avian somite. Dev. Dyn. Off. Publ. Am. Assoc. Anat. 202, 165–171 (1995). 275. Ambler, C. A., Nowicki, J. L., Burke, A. C. & Bautch, V. L. Assembly of trunk and limb blood vessels involves extensive migration and vasculogenesis of somite-derived angioblasts. Dev. Biol. 234, 352–364 (2001). 276. Relaix, F., Rocancourt, D., Mansouri, A. & Buckingham, M. Divergent functions of murine Pax3 and Pax7 in limb muscle development. Genes Dev. 18, 1088–1105 (2004). 277. Baggiolini, A. et al. Premigratory and migratory neural crest cells are multipotent in vivo. Cell Stem Cell 16, 314–322 (2015). 193 278. Yvernogeau, L., Auda-Boucher, G. & Fontaine-Perus, J. Limb bud colonization by somite-derived angioblasts is a crucial step for myoblast emigration. Dev. Camb. Engl. 139, 277–287 (2012). 279. Ling, F., Kang, B. & Sun, X.-H. Id proteins: small molecules, mighty regulators. Curr. Top. Dev. Biol. 110, 189–216 (2014). 280. Zhao, Q., Eberspaecher, H., Lefebvre, V. & De Crombrugghe, B. Parallel expression of Sox9 and Col2a1 in cells undergoing chondrogenesis. Dev. Dyn. Off. Publ. Am. Assoc. Anat. 209, 377–386 (1997). 281. Shukunami, C., Takimoto, A., Oro, M. & Hiraki, Y. Scleraxis positively regulates the expression of tenomodulin, a differentiation marker of tenocytes. Dev. Biol. 298, 234–247 (2006). 282. He, P. et al. The changing mouse embryo transcriptome at whole tissue and single-cell resolution. Nature 583, 760–767 (2020). 283. Okamoto, O. & Fujiwara, S. Dermatopontin, a novel player in the biology of the extracellular matrix. Connect. Tissue Res. 47, 177–189 (2006). 284. Driskell, R. R., Giangreco, A., Jensen, K. B., Mulder, K. W. & Watt, F. M. Sox2-positive dermal papilla cells specify hair follicle type in mammalian epidermis. Dev. Camb. Engl. 136, 2815–2823 (2009). 285. Hemavathy, K., Ashraf, S. I. & Ip, Y. T. Snail/slug family of repressors: slowly going into the fast lane of development and cancer. Gene 257, 1–12 (2000). 286. Sakai, L. Y., Keene, D. R., Morris, N. P. & Burgeson, R. E. Type VII collagen is a major structural component of anchoring fibrils. J. Cell Biol. 103, 1577–1586 (1986). 194 287. Houweling, A. C. et al. Gene and cluster-specific expression of the Iroquois family members during mouse development. Mech. Dev. 107, 169–174 (2001). 288. Vanlandewijck, M. et al. A molecular atlas of cell types and zonation in the brain vasculature. Nature 554, 475–480 (2018). 289. Rampin, A, Campagnolo, P., Mastrullo, V., Madeddu, P., & Spinetti, G. Adventitial and Skeletal Muscle Pericytes in Health and Ischemic Tissue Regeneration. in Biology of Pericytes – Recent Advances. vol. 68 (Humana, Cham, 2021). 290. Chevallier, A. Role of the somitic mesoderm in the development of the thorax in bird embryos. II. Origin of thoracic and appendicular musculature. J. Embryol. Exp. Morphol. 49, 73–88 (1979). 291. Gros, J., Manceau, M., Thomé, V. & Marcelle, C. A common somitic origin for embryonic muscle progenitors and satellite cells. Nature 435, 954–958 (2005). 292. Soeda, T. et al. Sox9-expressing precursors are the cellular origin of the cruciate ligament of the knee joint and the limb tendons. Genes. N. Y. N 2000 48, 635–644 (2010). 293. Akiyama, H. et al. Osteo-chondroprogenitor cells are derived from Sox9 expressing precursors. Proc. Natl. Acad. Sci. U. S. A. 102, 14665–14670 (2005). 294. Charvet, B., Ruggiero, F. & Le Guellec, D. The development of the myotendinous junction. A review. Muscles Ligaments Tendons J. 2, 53–63 (2012). 295. Kostrominova, T. Y., Calve, S., Arruda, E. M. & Larkin, L. M. Ultrastructure of myotendinous junctions in tendon-skeletal muscle constructs engineered in vitro. Histol. Histopathol. 24, 541–550 (2009). 296. Trotter, J. A., Samora, A. & Baca, J. Three-dimensional structure of the murine muscle-tendon junction. Anat. Rec. 213, 16–25 (1985). 195 297. Ciena, A. P. et al. Ultrastructure of the myotendinous junction of the medial pterygoid muscle of adult and aged Wistar rats. Micron Oxf. Engl. 1993 41, 1011–1014 (2010). 298. Delaurier, A. et al. The Mouse Limb Anatomy Atlas: an interactive 3D tool for studying embryonic limb patterning. BMC Dev. Biol. 8, 83 (2008). 299. Nassari, S., Duprez, D. & Fournier-Thibault, C. Non-myogenic Contribution to Muscle Development and Homeostasis: The Role of Connective Tissues. Front. Cell Dev. Biol. 5, 22 (2017). 300. Schweitzer, R., Zelzer, E. & Volk, T. Connecting muscles to tendons: tendons and musculoskeletal development in flies and vertebrates. Dev. Camb. Engl. 137, 2807–2817 (2010). 301. Sugimoto, Y. et al. Scx+/Sox9+ progenitors contribute to the establishment of the junction between cartilage and tendon/ligament. Dev. Camb. Engl. 140, 2280–2288 (2013). 302. Subramanian, A. & Schilling, T. F. Tendon development and musculoskeletal assembly: emerging roles for the extracellular matrix. Dev. Camb. Engl. 142, 4191–4204 (2015). 303. Koch, M. et al. A novel marker of tissue junctions, collagen XXII. J. Biol. Chem. 279, 22514–22521 (2004). 304. Kult, S. et al. Bi-fated tendon-to-bone attachment cells are regulated by shared enhancers and KLF transcription factors. eLife 10, e55361 (2021). 305. Cash, D. E., Bock, C. B., Schughart, K., Linney, E. & Underhill, T. M. Retinoic acid receptor alpha function in vertebrate limb skeletogenesis: a modulator of chondrogenesis. J. Cell Biol. 136, 445–457 (1997). 306. Ribatti, D. Nicole Le Douarin and the use of quail-chick chimeras to study the developmental fate of neural crest and hematopoietic cells. Mech. Dev. 158, 103557 (2019). 196 307. Beresford, B. Brachial muscles in the chick embryo: the fate of individual somites. J. Embryol. Exp. Morphol. 77, 99–116 (1983). 308. Dhouailly, D. & Kieny, M. [Participation of the somatopleural mesoderm of the side to the building of an additional limb in birds]. Comptes Rendus Hebd. Seances Acad. Sci. Ser. Sci. Nat. 271, 519–522 (1970). 309. Pinot, M. [The role of the somitic mesoderm in the early morphogenesis of the limbs in the fowl embryo]. J. Embryol. Exp. Morphol. 23, 109–151 (1970). 310. Tissières, V. et al. Gene Regulatory and Expression Differences between Mouse and Pig Limb Buds Provide Insights into the Evolutionary Emergence of Artiodactyl Traits. Cell Rep. 31, 107490 (2020). 311. Jhanwar, S. et al. Conserved and species-specific chromatin remodeling and regulatory dynamics during mouse and chicken limb bud development. Nat. Commun. 12, 5685 (2021). 312. Hurren, B., Collins, J. J. P., Duxson, M. J. & Deries, M. First Neuromuscular Contact Correlates with Onset of Primary Myogenesis in Rat and Mouse Limb Muscles. PLoS ONE 10, (2015). 313. Hirasawa, T. & Kuratani, S. Evolution of the muscular system in tetrapod limbs. Zool. Lett. 4, 27 (2018). 314. Tosney, K. W. & Landmesser, L. T. Development of the major pathways for neurite outgrowth in the chick hindlimb. Dev. Biol. 109, 193–214 (1985). 315. Wang, G. & Scott, S. A. The ‘waiting period’ of sensory and motor axons in early chick hindlimb: its role in axon pathfinding and neuronal maturation. J. Neurosci. Off. J. Soc. Neurosci. 20, 5358–5366 (2000). 197 316. Abdalla, O. Ossification and mineralization in the tendons of the chicken (Gallus domesticus). J. Anat. 129, 351–359 (1979). 317. Agabalyan, N. A., Evans, D. J. R. & Stanley, R. L. Investigating tendon mineralisation in the avian hindlimb: a model for tendon ageing, injury and disease. J. Anat. 223, 262–277 (2013). 318. Landis, W. J. & Silver, F. H. The structure and function of normally mineralizing avian tendons. Comp. Biochem. Physiol. A. Mol. Integr. Physiol. 133, 1135–1157 (2002). 319. Benton, M. J. Vertebrate palaeontology. (Wiley Blackwell, 2015). 320. Huang, A. H. et al. Requirement for scleraxis in the recruitment of mesenchymal progenitors during embryonic tendon elongation. Dev. Camb. Engl. 146, dev182782 (2019). 321. Gupta, R. K. et al. Transcriptional control of preadipocyte determination by Zfp423. Nature 464, 619–623 (2010). 322. Muhl, L. et al. Single-cell analysis uncovers fibroblast heterogeneity and criteria for fibroblast and mural cell identification and discrimination. Nat. Commun. 11, 3953 (2020). 323. Bunge, M. B., Wood, P. M., Tynan, L. B., Bates, M. L. & Sanes, J. R. Perineurium originates from fibroblasts: demonstration in vitro with a retroviral marker. Science 243, 229–231 (1989). 324. Zwierzina, M. E. et al. Characterization of DLK1(PREF1)+/CD34+ cells in vascular stroma of human white adipose tissue. Stem Cell Res. 15, 403–418 (2015). 325. Bondjers, C. et al. Microarray analysis of blood microvessels from PDGF-B and PDGF-Rbeta mutant mice identifies novel markers for brain pericytes. FASEB J. Off. Publ. Fed. Am. Soc. Exp. Biol. 20, 1703–1705 (2006).
Thesis/Dissertation
eng
Cell and Developmental Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Underhill, T M
Origin, fate, and function of embryonic Hic1-positive mesenchymal progenitors in the developing limb
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