INTERACTIONS BETWEEN HIPPO, FGF, AND TGFβ SIGNALING REGULATE MORPHOGENESIS OF EARLY MOUSE EMBRYOS By Robin E. Kruger A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Cell and Molecular Biology – Doctor of Philosophy 2023 ABSTRACT Intercellular molecular signaling regulates every aspect of development, from the very first cell fate decisions to communication between complex organ tissues. Much of what we know about molecular regulation of development in mammals comes from studies of mouse embryos. In mice, the first cell fate decision, which separates the outer, multipotent trophectoderm cells from the pluripotent inner cell mass, is regulated by the Hippo signaling pathway. It is known that silenced Hippo signaling in the outer cells maintains the cells’ polarity; however, my studies show that Hippo signaling does not direct the initiation of cell polarization in mouse embryos. The second cell fate decision in mouse embryos differentiates the inner cell mass into pluripotent epiblast and multipotent primitive endoderm. The Bone Morphogenic Protein (BMP) signaling pathway has been suggested to regulate preimplantation cell fate at this stage; however, I show that preimplantation lineage specification appears normal in mouse embryos that lack maternal and zygotic Smad4, an essential effector of canonical BMP signaling. Rather, my findings point to a previously unrecognized role for SMAD4 during early post-implantation stages, in which SMAD signaling restricts FGF signaling to promote epiblast growth and morphogenesis. In addition to these novel regulatory mechanisms in early cell fate, I present three technological advances in the study of early development, using recombinant fluorescent markers, more specific markers of pluripotency in reprogrammed stem cells, and ultra-low-input CUT&RUN to identify transcription factor binding sites in embryos. Altogether, these studies uncover novel mechanisms of cross-talk between regulatory pathways in development and present new strategies to investigate future questions in developmental biology. ACKNOWLEDGEMENTS First of all, I want to thank every member of the Ralston Lab for making my time in graduate school such a wonderful experience. I really could not have asked for a more supportive, fun, and smart group of people to spend five years with. Amy, you’ve taught me more than I thought possible. With your mentorship and guidance I’ve learned what it takes to be a scientist, but also what it takes to be a true mentor and a leader. Thank you for always being my advocate and for helping mold me into a well-rounded scientist. I hope I make you proud! Jenn, Alex, Tristan, and Dan, thank you for being my mentors and for teaching me the skills I needed to succeed in the lab. Barb, thank you for always being friendly and patient while fixing all my lab crises. And finally, my fellow labmates Tayler, Farina, Marcelio, Shannon, Ian, and Alyssa: you guys have been amazing friends through all of this. Every one of you have made me laugh, made me smile, helped me with experiments, debated science with me, and have helped me grow as a mentor, as a scientist, and as a person. I can’t wait to see where all your successes take you in life! I also want to thank the people who have made my professional development in graduate school a success. Thank you to the members of my guidance committee, Dr. Aitor Aguirre, Dr. Ingo Braasch, Dr. Chen Chen, and Dr. Almudena Veiga-Lopez, for your invaluable advice and support. Thank you to the Reproductive and Developmental Science Training Program (NIH T32 HD087166) to the directors, Dr. Keith Latham and Dr. Asgi Fazleabas, and to the coordinator Laurie Felton, for supporting my scientific training and giving me so many opportunities to share my work and improve as a iii developmental biologist. A special thank you to the Cell and Molecular Biology program, my graduate director Dr. Peggy Petroff, and academic coordinator Alaina Burghardt. Thank you so much for your support, for handling all my crises, and for saying yes to all my crazy ideas. You do an incredible job making your students feel like they matter, and that is so critical in helping them succeed. I really felt like I found a home in CMB, and I wish all the best for everyone in the program! I want to say a special thank you to the organizations I’ve been a part of during my time in graduate school for their support to my professional development: Graduate Women In Science Mid-Michigan, Cell and Molecular Biology Graduate Student Organization, and the Student Faculty Staff Hearing Board. Each one of these groups has taught me about what it takes to be a leader, to relate to people, to communicate effectively, and to take responsibility for a project from start to finish. Last but not least, I need to thank my amazing friends and family who helped me get to this point in my life. I cannot tell you how much your love and support have meant to me. Mom and Dad, thank you so much for always believing that I could do anything. You are the world’s best parents and I love you so much! Grandma and Grandpa, thank you so much for your love and support for everything I do. You’ve been a huge inspiration for me going into science, and I really appreciate how much you care about my work. Megan, you’re the best sister anyone could ask for, thank you for all the things you do for me! My frieNDs, Bella, Cristina, Danielle, Emma, Laura, and Mara: you guys have been an amazing support system for me ever since we met, and the fact that we iv are all still so close means the world to me. Skimble and Leia, thanks for sticking by me through everything and being my very best listeners. And finally, my wonderful husband, Adam Kruger. I truly would not have been able to do this if it were not for you. Thank you for everything, from moving with me to Lansing, to putting up with my unpredictable hours and weird nerdy science stuff, to just knowing how to cheer me up when things were tough. I hope you’re ready to be married to a doctor! To everyone mentioned and anyone else who has helped me through this journey, I can’t thank you enough for your guidance, love, and support. v TABLE OF CONTENTS LIST OF ABBREVIATIONS……………….……………………………………………….….vii CHAPTER 1. MOLECULAR SIGNALING MECHANISMS REGULATING MURINE PRE- AND POST-IMPLANTATION EMBRYONIC CELL FATE DECISIONS……………………1 CHAPTER 2. SMAD4 IS ESSENTIAL FOR EPIBLAST SCALING AND MORPHOGENESIS AFTER IMPLANTATION, BUT NONESSENTIAL PRIOR TO IMPLANTATION IN THE MOUSE……………………………………………………………38 CHAPTER 3. ESSENTIAL ROLES FOR HIPPO SIGNALING IN MAMMALIAN REPRODUCTION……………………………………………………………………….…….90 CHAPTER 4. HIPPO SIGNALING EFFECTORS MAINTAIN BUT DO NOT REGULATE INITIATION OF CELL POLARIZATION IN MOUSE BLASTOMERES…….……………115 CHAPTER 5. EFFICIENT GENERATION OF ENDOGENOUS PROTEIN REPORTERS FOR MOUSE DEVELOPMENT………………………………………………………….…130 CHAPTER 6. FLUORESCENT REPORTERS DISTINGUISH STEM CELL COLONY SUBTYPES DURING SOMATIC CELL REPROGRAMMING…………...……………...163 CHAPTER 7. IDENTIFICATION OF OCT4 BINDING SITES IN ICM CELLS BY uliCUT&RUN…….........................................................................................................189 CONCLUSION..…………………………………………………..……………………….…207 REFERENCES..……………………….……………………………………………………..209 vi AP AVE BMP BRE BSA CTB DVE E EMT EPI ESC EVT ExE FBS FGF hCG HuF ICM IFNT iPSC iXEN LDN LIST OF ABBREVIATIONS Alkaline phosphatase Anterior Visceral Endoderm Bone Morphogenic Protein BMP Response Element Bovine Serum Albumin Cytotrophoblast Distal Visceral Endoderm Embryonic Day Epithelial-to-Mesenchymal Transition Epiblast Embryonic Stem Cell Extravillous Trophoblast Extraembryonic Ectoderm Fetal Bovine Serum Fibroblast Growth Factor Human Chorionic Gonadotrophin Human Stromal Fibroblasts Inner Cell Mass Interferon Tau Induced Pluripotent Stem Cell Induced Extraembryonic Endoderm LDN-193189, ALK3/6 inhibitor vii m null MenSC mz null PE pERK PGC PrE PS pSMAD PV PVA r-SMAD RTK SSC STB T TE TGFβ VE XEN Maternal null Menstrual Stem Cell Maternal-zygotic null Parietal Endoderm Phosphorylated ERK protein(s) Primordial Germ Cell Primitive Endoderm Primitive Syncytium Phosphorylated SMAD protein(s) Placental Villi Polyvinyl alcohol Receptor-associated SMAD protein Receptor Tyrosine Kinase Spermatogonial Stem Cell Syncytiotrophoblast Brachyury Trophectoderm Transforming Growth Factor beta Visceral Endoderm Extraembryonic Endoderm viii CHAPTER 1. MOLECULAR SIGNALING MECHANISMS REGULATING MURINE PRE- AND POST- IMPLANTATION EMBRYONIC CELL FATE DECISIONS This study was supported by the National Institutes of Health R35 GM131759 and T32 HD087166. 1 Section 1.1. Abstract The study of embryonic development is critical to understanding the basic functions of stem cells and to determine the factors which promote the birth of healthy offspring. Most current knowledge of mammalian embryogenesis has come from decades of study into the most widely-used model: the mouse embryo. In this chapter, I introduce the morphological and molecular changes that occur during the first week of embryonic development in mice, from fertilization to gastrulation. I focus on the regulation of these early developmental processes by molecular signaling pathways, with an emphasis on the roles of Hippo, Fibroblast Growth Factor, and Bone Morphogenic Protein signaling. Section 1.2. Overview of mouse embryogenesis and early cell fate specification 1.2.1. Introduction to mouse development The study of development is the study of life itself. All multi-cellular organisms must go through developmental processes to reach reproductive maturity and pass their genetic material to their progeny. For complex organisms such as mammals, this process requires a single cell, the zygote, to divide and differentiate to derive hundreds of specialized cell types. These cell types not only need to specify properly during development, but also communicate and organize with one another as they expand to generate highly organized tissues, organs, and organ systems. By studying development, researchers can begin to understand the etiology of developmental abnormalities such as congenital disorders, pre-term birth, miscarriage, and infertility. These can be used to improve strategies for assisted reproductive therapies. Similarly, information gleaned from studies of natural development can be used to improve stem 2 cell models of embryos, tissues, and organs for uses in research, stem cell therapies, and personalized medicine (Rosner et al., 2021; Wallingford, 2019). Laboratory mice are the most common research models for the study of mammalian development. In humans, the process of embryonic development takes approximately 40 weeks; for mice, development from fertilization to birth takes 18-22 days (Murray et al., 2010). Despite the differences in timing, embryonic development in mice and humans is remarkably similar, going through similar processes in the same order and importantly, specifying many of the same cell types (Gupta et al., 2021; Taft, 2008). Developmental timepoints in mice are described by the number of days since fertilization, denoted by Embryonic Day (E). E0 is the moment of fertilization, which in nocturnal animals such as mice, typically occurs at night. After mating, male mice leave a waxy plug in the vaginal opening, allowing researchers to know when fertilization has occurred and to precisely determine the developmental stage of embryos developing internally. As a convention, mating is assumed to occur at midnight in laboratory tests, designating noon of the day the plug is detected as E0.5. This notation will be used throughout this work to designate developmental staging in mouse embryos. In this chapter, I describe the developmental processes that occur during embryogenesis in mice (Fig. 1.1). In particular, I focus on the stem cell progenitor types which emerge in the earliest cell fate decisions of embryogenesis and the populations they give rise to during early post-implantation development. I also discuss the molecular signaling mechanisms that regulate stem cell fate specification and 3 differentiation, with an emphasis on Hippo, Fibroblast Growth Factor (FGF), and Bone Morphogenic Protein (BMP) signaling. Though I focus on mouse development, many of the cell types and processes are similar to human development, showing a high degree of evolutionary conservation between these two species in their developmental progression. 1.2.2. Molecular mechanisms regulating developmental stages and processes in murine preimplantation development E0-E2.5: Cleavage divisions and zygotic genome activation An overview of mouse preimplantation development is important not only because the processes are very similar in human embryos, but also to provide the necessary context for the discussions of molecular signals regulating development presented in Chapters 2 and 4 of this work. The developmental processes in mouse embryos before the embryo implants into the uterine wall (E0-E4.5) are very well-characterized. Preimplantation development begins at fertilization, when the gametes of sperm and egg fuse to create a diploid zygote. The murine zygote is surrounded by a protective glycoprotein layer called the zona pellucida, which persists around the outside of the embryo throughout preimplantation development (Rankin et al., 2000). Similar to other species, the genome of the murine zygote is initially silenced, and embryonic development relies on maternally-derived mRNA transcripts and proteins deposited into the oocyte for early cellular activity (Cockburn & Rossant, 2010). Zygotic genome activation (ZGA) occurs at E1.5 after the first cleavage division into a 2-cell embryo, and >90% of maternal transcripts are degraded at this point (L. Li et al., 2010). However, 4 maternally-derived proteins may persist to the blastocyst stage and the effects of maternal proteins, such as chromatin modifications, may direct development even after the proteins themselves have been degraded (F. Chen et al., 2022; Israel et al., 2019; L. Li et al., 2010; D. Wu & Dean, 2020; H. Zhang et al., 2023). This is important because it poses an added layer of redundancy for researchers to consider when examining gene function in early embryos; even if both copies of a gene are knocked out in the zygotic genome, early phenotypes may be masked by compensation from maternally-derived transcripts and proteins. To completely eliminate a gene at this stage, it is necessary to create a maternal and zygotic loss-of-function model (Cockburn & Rossant, 2010). One strategy to create such a model is described in Chapter 2. E2.5-E3.0: Compaction, polarization, and the first cell fate decision regulated by Hippo signaling After fertilization, successive cell divisions in the embryo lead to morphological changes and corresponding decreases in developmental potential. The single-celled zygote is totipotent, meaning it has the potential to give rise to all cells in the embryo, both embryonic and extraembryonic (Chazaud & Yamanaka, 2016). In mice, totipotency persists through three embryonic cleavage events until the embryo consists of 8 cells, or blastomeres. At that point, around E2.75, the 8-cell embryo undergoes a significant morphological change known as compaction. During compaction, cell-cell contacts between blastomeres increase dramatically. E-cadherin becomes enriched at adherens junctions between cells to form the basis of a basolateral domain toward the center of the embryo (Sozen et al., 2014). Accumulation of E-cadherin at the cell-cell contacts 5 displaces actin filaments, which accumulate at the cell-free surfaces on the outside of the embryo (Lim & Plachta, 2021). Simultaneously, proteins such as Par3, Par6, and aPKCζ are upregulated and targeted to the cell-free membrane, forming an outer apical domain around the embryo. The apical domain interacts with the accumulated actin to form an actomyosin ring at the cell-free surface of each blastomere, which contracts to pull the edges of the cells toward one another (Lim & Plachta, 2021). This causes the cells to visibly flatten against one another until the boundaries between blastomeres are no longer distinct. Blastomere polarization and compaction, including regulation by Hippo signaling, are further discussed in Chapter 4. The process of compaction and cell polarization is the first developmental event that leads to symmetry breaking and cell differentiation in the embryo. Up until compaction, totipotent 8-cell blastomeres retain the ability to each give rise to a new blastocyst if separated (Tarkowski & Wróblewska, 1967). However, this potential is lost during the first cell division after compaction. During the 8-to-16-cell division, some compacted blastomeres divide asymmetrically so that not all daughter cells inherit the outer apical domain evenly. Those cells that lose the apical domain become apolar and move into the center of the embryo (Chazaud & Yamanaka, 2016). Apolar inner cells and polarized outer cells form the basis of the first cell fate decision in mouse embryos, which are driven to become inner cell mass (ICM) and trophectoderm (TE) cells, respectively (Chazaud & Yamanaka, 2016). Neither of these newly-formed cell populations retains their former totipotent state. The ICM cells are considered pluripotent, meaning they can still give rise to many, but not all, later cell types, while the TE cells are multipotent, 6 meaning that despite still being stem cell progenitors, their developmental potential is much more restricted (Rossant, 2001). ICM will eventually give rise to the fetus and extraembryonic tissues (discussed further below), while the TE lineage gives rise to embryonic placental cells (Chazaud & Yamanaka, 2016). An extensive body of work suggests that Hippo signaling plays an integral role in regulating the ICM-TE cell fate decision in mouse embryos (Alarcon, 2010; Anani et al., 2014; Cao et al., 2015; Cockburn et al., 2013; Frum et al., 2018, 2019; Hirate et al., 2013, 2015; Home et al., 2012; Leung & Zernicka-Goetz, 2013; Lorthongpanich et al., 2013; Mihajlović & Bruce, 2016; Nishioka et al., 2008, 2009; Plusa et al., 2005; Ralston et al., 2010; Ralston & Rossant, 2008; Rayon et al., 2014; Wicklow et al., 2014; Yagi et al., 2007; Yamamura et al., 2020; M. Zhu et al., 2017, 2020). These studies and others are the subject of several reviews, which I recommend for further information on this topic (Alarcon & Marikawa, 2018; Hirate & Sasaki, 2014; Karasek et al., 2020; Lorthongpanich & Issaragrisil, 2015; Sasaki, 2015, 2017; J. Sharma et al., 2021; Z. Wu & Guan, 2021; Yildirim et al., 2021). The collective model from these studies suggests that Hippo signaling is active in the apolar, inner cells of the morula which leads to an ICM fate, while the polarized outer cells silence Hippo signaling to induce a TE cell fate (Fig. 1.2). In the inner, apolar cells, the scaffolding protein Angiomotin (AMOT) localizes to adherens junctions on the cell membrane, and from there interacts with upstream Hippo family kinases LATS1/2 (Hirate et al., 2013). This interaction allows LATS1/2 to phosphorylate downstream 7 transcription factors YAP1 and WWTR1. Phosphorylation of YAP1 and WWTR1 prevents these factors from entering the nucleus and instead promotes their association with the 14-3-3 complex for degradation; therefore, phosphorylation of YAP1/WWTR1 downregulates their transcriptional activity (C. Y. Liu et al., 2010). In the TE of the preimplantation mouse embryo, YAP1/WWTR1 repress transcription of Sox2, the earliest marker of pluripotency (Frum et al., 2018, 2019). YAP1/WWTR1 degradation in the inner cells therefore upregulates Sox2 and promotes a pluripotent, inner cell mass fate. By contrast, in outer, polarized cells, AMOT is restricted to the apical domain through interaction with the aPKC-Par system (Hirate et al., 2013). AMOT is unable to promote phosphorylation of LATS1/2 as effectively from the apical domain, and in turn inactive LATS1/2 does not phosphorylate YAP1/WWTR1. Unphosphorylated YAP1/WWTR1 are translocated into the nucleus where they partner with TEAD4 to promote transcription of trophectoderm-specific transcription factors Cdx2 and Gata3 (Nishioka et al., 2008; Ralston et al., 2010) and repress Sox2 (Frum et al., 2018; Wicklow et al., 2014). Therefore, in mice, repression of Hippo signaling in the outer cells both actively promotes a TE cell fate and represses pluripotency. E3.25-E3.75: Blastocyst formation and the second cell fate decision regulated by FGF signaling This first cell fate decision coincides with the transition to the morula stage of development, which lasts from the 16-cell stage (at ~E3.0) to the 32-cell stage (at ~E3.25). At E3.25, tight junctions between outer TE cells allow the formation of a fluid- filled cavity within the embryo known as a blastocoel (Cockburn & Rossant, 2010). Once 8 the blastocoel is fully formed, around E3.5, the embryo is called a blastocyst. Inside the blastocyst, the apolar ICM cells are gathered to one side of the blastocoel, while the TE comprises an epithelial layer surrounding the outside. Soon after, at E3.75, the ICM undergoes a second cell fate decision to differentiate into pluripotent epiblast (EPI) and multipotent primitive endoderm (PrE) populations (Chazaud & Yamanaka, 2016). EPI cells will give rise to most of the eventual fetus as well as contribute to extraembryonic fetal membranes such as the chorion, amnion, and allantois. PrE gives rise primarily to the yolk sac endoderm, with some contribution to the definitive endoderm of the fetus (Kwon et al., 2008; Lu et al., 2001). Regulation by FGF signaling plays a key role in the EPI-PrE cell fate decision. As this is the result of a large body of work (Arman et al., 1998; Azami et al., 2019; Bessonnard et al., 2017; Chazaud et al., 2006; Feldman et al., 1995; Frankenberg et al., 2011; Frum et al., 2013; Grabarek et al., 2012; Kang et al., 2013, 2017; Krawchuk et al., 2013; Le Bin et al., 2014; Messerschmidt & Kemler, 2010; Mistri et al., 2018; Molotkov et al., 2017; Nichols et al., 2009; Ohnishi et al., 2014; Plusa et al., 2008; Rappolee et al., 1994; Wicklow et al., 2014; Wigger et al., 2017; Yamanaka et al., 2010) and has been recently reviewed (Brewer et al., 2016; Dorey & Amaya, 2010; Lanner & Rossant, 2010; Soszyńska et al., 2019), the main points are summarized here (Fig. 1.3). FGF activity is detected in the ICM of the blastocyst as early as E3.25, and is gradually upregulated in PrE cells as they become specified in the mid-blastocyst stage (Azami et al., 2019; Morgani et al., 2018). During the early blastocyst stage, the ICM cells co-express the transcription factors Nanog and Gata6, regulators of EPI and PrE, respectively. Nanog 9 induces expression of the FGF ligand Fgf4 (Frankenberg et al., 2011). Some bipotent ICM cells express higher levels of Fgf4 than others, making Fgf4 the first bimodally expressed gene among the ICM cells (Ohnishi et al., 2014). It is currently still unclear whether this asymmetry in Fgf4 expression arises stochastically or if it is regulated in some way. Neighboring ICM cells receive this signal through the receptor FGFR1 (Molotkov et al., 2017), which stimulates the MEK/ERK pathway in the receiving cells. ERK activity downregulates expression of Nanog, which relieves repression and upregulates Gata6 expression (Chazaud et al., 2006). Gata6-high cells upregulate expression of FGFR2, increasing the receptivity to FGF and reinforcing ERK activity and Gata6 upregulation in these cells. By the mid-blastocyst stage Gata6-high cells have also upregulated Pdgfrα, Sox17, and Gata4 to adopt a true PrE fate (Artus et al., 2012; Niakan et al., 2010). Meanwhile, upregulated Nanog expression in the Fgf4-high cells downregulates Gata6 to solidify an EPI fate. By E3.75, Gata6 and Nanog expression are mutually exclusive (Chazaud et al., 2006), although studies have shown that some ICM cells retain the potential to switch between EPI and PrE fates until E4.5 (Grabarek et al., 2012; Plusa et al., 2008; Wigger et al., 2017). In this manner, EPI and PrE cells arise in a “salt-and-pepper” distribution throughout the ICM, directed by localized high and low levels of FGF signaling (Fig. 1.3). FGF signaling is necessary and sufficient to direct specification of PrE over EPI cell fate in the mouse ICM. The ability to modulate cell fate using exogenous factors that modulate FGF signaling has enabled the in vitro testing of models of ICM differentiation. Morula-stage embryos cultured with exogenous FGF4 and heparin to blastocyst stage 10 develop ICMs comprised entirely of Gata6-positive cells (Yamanaka et al., 2010). Similarly, treating embryos with a combination of FGFR and MEK inhibitors (PD173074 and PD0325901, respectively) for the same developmental duration directs the entire ICM to express Nanog (Nichols et al., 2009; Yamanaka et al., 2010). However, ICM cells become unresponsive to FGF4 around E4.0 (Yamanaka et al., 2010). This suggests that modulations in FGF are sufficient to direct PrE specification in the early ICM even in EPI-biased cells, but that it is not sufficient to change ICM cell fate once an EPI or PrE program is fully established. Whether other signaling pathways interact with FGF to reinforce PrE fate or regulate the timing of FGF sensitivity in ICM cells remains an open question. The potential of BMP signaling to interfere with PrE fate induction by FGF signaling will be discussed further in Chapter 2. E4.0-E4.5: Embryo hatching and implantation After the second cell fate decision, the embryo undergoes only a few more morphological changes before implantation. EPI and PrE cells initially arise in a “salt- and-pepper” distribution throughout the ICM, but by E4.0 they sort into separate populations with the PrE forming an epithelial layer between the EPI and the blastocoel (Cockburn & Rossant, 2010). Around this time, the embryo hatches out of the zona pellucida, allowing the exposed TE cells to attach to the mother’s uterine wall (Ma et al., 2024). Embryo attachment facilitates a massive proliferation process in the uterine stromal cells known as decidualization, which provides the embryo a protected place to grow and establishes a maternal-fetal interface (Rinkenberger et al., 1997). During this 11 period, TE cells begin to invade the decidua, forming the basis of the future placenta. Shortly after E4.5, the embryo is considered to be implanted (Rossant & Tam, 2009). 1.2.3. Molecular mechanisms regulating developmental stages and processes of murine post-implantation development through gastrulation E4.75-E5.5: Egg cylinder formation and cell differentiation Developmental and regulative processes become more complex after implantation as the embryo undergoes rapid proliferation and morphological changes as well as cell fate changes. The initial three cell types of EPI, PrE, and TE each begin to mature and differentiate as the embryo elongates into a cup-shaped structure known as the egg cylinder (reviewed in Arnold & Robertson, 2009; Filimonow & de la Fuente, 2022; Stower & Srinivas, 2018). These newly-differentiated cell types provide critical signals for the reorganization of the embryo and increase in complexity during gastrulation, and as such, their proper derivation and organization is crucial for further embryonic development. Several of these cell types and signaling pathways are discussed further in Chapter 2. Many morphological and molecular changes in cell fate between embryonic implantation and gastrulation have been observed. The polar TE cells, adjacent to the EPI in the blastocyst, proliferate and expand to form the most proximal end of the egg cylinder. These differentiate into more mature cell types including the extra-embryonic ectoderm (ExE), which eventually gives rise to the main cell types that form the embryonic portions of the placenta. Just as importantly, the ExE plays an essential role in pre- 12 gastrulation embryonic patterning, particularly as the source of BMP4 that is necessary for mesoderm induction and primordial germ cell formation (Ben-Haim et al., 2006; Winnier et al., 1995, discussed further in Section 1.3). The PrE also expands and differentiates after implantation, giving rise to the visceral endoderm (VE), which forms a single cell layer on the outside surface of the egg cylinder. These cells eventually give rise to the yolk sac endoderm (Filimonow & de la Fuente, 2022), as well as contribute to gut tube formation in the fetus (Kwon et al., 2008). Critically, the VE is the source of several essential extraembryonic signals to regulate embryonic axial patterning and gastrulation, including Nodal, BMP, and WNT (reviewed in S. M. Morgani & Hadjantonakis, 2020). EPI, although not referred to as a new cell type, undergoes significant changes during egg cylinder formation. In the blastocyst, expression of known pluripotency-promoting transcription factors such as Nanog, Sox2, and Oct4 keep epiblast cells in a “naïve” state of pluripotency (Nichols & Smith, 2009). After implantation, EPI downregulates Nanog, Klf4, Stella, and other naïve markers to mature into a “primed” state of pluripotency. Possibly as a result of this molecular change, the EPI changes morphologically after implantation from round and apolar cells in the blastocyst to columnar, epithelialized cells by E5.0. The epithelized EPI organizes into a polarized rosette at the distal end of the growing egg cylinder (Bedzhov & Zernicka-Goetz, 2014). The VE cells overlying the EPI deposit a basement membrane between the VE and EPI, inducing a basolateral domain on the outer edge of the EPI rosette and an apical domain in the center (Bedzhov & Zernicka-Goetz, 2014; S. Li et al., 2003; Niakan et al., 2010; Shahbazi et al., 2017). Soon after EPI 13 polarization, a lumen opens in the rosette’s center called the proamniotic cavity, forming the EPI into a single layer epithelium. Importantly for my studies, there is some debate in the literature as to what induces epiblast cavitation at early post-implantation stages. Early studies in embryoid bodies (spherical structures grown in vitro from EPI-derived embryonic stem cells) proposed that the central EPI cells receive a cell death signal from the maturing VE which causes them to undergo apoptosis (Coucouvanis & Martin, 1995, 1999). This is supported by a recent study which detected cells undergoing apoptosis in the developing lumen of E5.5 embryos (Halimi et al., 2022). However, other studies have suggested that this mechanism is not sufficient to explain epiblast cavitation. An alternative mechanism was proposed by Bedzhov and Zernicka-Goetz in which EPI polarization is the driving force behind cavitation. According to this study, the central apical domain secretes anti- adhesion proteins like podocalyxin, which help force the apical ends of the cells away from one another (Bedzhov & Zernicka-Goetz, 2014). This mechanism is supported by studies which show that embryos which lack essential components of cell polarity like integrin and laminin also fail to cavitate (Smyth et al., 1999; Stephens et al., 1995). A model for how these two mechanisms may be working simultaneously to promote cavitation is discussed in Chapter 2. E5.5-E7.5: Axial patterning and gastrulation After the establishment of the egg cylinder at E5.5 and associated differentiation of cell types, the complexity of the embryo begins to increase dramatically, with various 14 signaling gradients directing cell differentiation and cell movement. These regulate the process of gastrulation, or the formation of embryonic germ layers. Importantly, prior to gastrulation the post-implantation embryo is considered morphologically and molecularly symmetrical, with the only apparent axis being the proximal-distal axis (Arnold & Robertson, 2009; Takaoka & Hamada, 2012). Anterior-posterior and dorsal- ventral patterning occur concomitantly with the onset of gastrulation, and these processes are tightly linked to one another. The morphological and molecular changes that occur during axial patterning and gastrulation, and the signaling pathways that regulate them (including Nodal, BMP, WNT, and FGF signaling), have been extensively reviewed, and I recommend these for further information on the subject (Arnold & Robertson, 2009; Beddington & Robertson, 1999; Filimonow & de la Fuente, 2022; Matsuo & Hiramatsu, 2017; Morgani & Hadjantonakis, 2020; Perea-Gomez et al., 2001; Rossant & Tam, 2009; Stower & Srinivas, 2018; Takaoka & Hamada, 2012). The process of gastrulation and its regulation by BMP signaling will be discussed more thoroughly in Section 1.3. By E7.5, embryonic mesoderm is fully established, and more advanced structures begin to emerge, with the formation of the heart tube, head folds, amnion, and chorion. At this point, gastrulation is considered to be complete, and the dorsal-ventral and anterior-posterior axes are established. With the dorsal-ventral and anterior-posterior axes specified, the left-right axis becomes apparent, although the embryo does not show any overt left-right asymmetry until Nodal accumulates at the left side of the node ~E8.0 (Shiratori & Hamada, 2006). Thus, by the end of gastrulation, the embryo has specified all three germ layers and patterned the body axes, setting up the 15 more complex morphological and cell fate changes which occur in further stages of development. Section 1.3. The roles for Bone Morphogenic Protein (BMP) signaling in early mammalian embryonic development are not fully understood Bone Morphogenic Protein (BMP) signaling is essential for early mouse embryonic development and involved in many processes discussed in Section 1.2 (see highlighted processes in Fig. 1.1). BMP signaling is a highly conserved molecular signaling pathway, with homologs in all animals (Huminiecki et al., 2009), and its role in development has been studied for decades. The pathway is so named because it was first discovered as a factor which induced ectopic bone formation (E. A. Wang et al., 1988), but studies have since uncovered many other essential roles for BMP signaling regulating cell fate, cell survival, proliferation, and morphogenesis in both embryonic development and adult homeostasis (Reddi, 2003). Unsurprisingly, dysregulation of BMP signaling has also been implicated in a number of diseases, particularly congenital bone and heart defects (R. N. Wang et al., 2014). This section will focus on the roles of BMP signaling in pre- and peri-implantation mouse embryonic development, but many reviews discuss other important roles for BMP signaling (Bragdon et al., 2011; D. Chen et al., 2004; Reddi, 2003). Interestingly, studies of BMP signaling in mouse development disagree on the pathway’s earliest role in embryogenesis. This is essential knowledge for the studies in Chapter 2, which endeavor to resolve this conflict within the literature. 16 1.3.1. Biochemical interactions of the Bone Morphogenic Protein Signaling Pathway The biochemical interactions of BMP signaling have been well-described (Fig. 1.4). BMP is a subfamily of the TGFβ superfamily of signaling pathways, which also includes Activin, Nodal, and TGFβ signaling as well as Growth and Differentiation Factors, myostatin, and anti-Mullerian hormone (Reddi, 2003). BMPs may signal canonically through SMAD proteins or non-canonically through interaction with other pathways such as MEK/ERK, JNK/p38, or PI3K/Akt (Y. E. Zhang, 2009). Canonical BMP signaling is transduced through receptors known as activin-like kinases (ALKs) and internal effectors called SMAD proteins. BMPs, which are secreted proteins, form homo- or hetero-dimers to become biologically active. Dimerized ligands bind to membrane- bound ALK receptors on receiving cells. BMP signaling, as with other TGFβ pathways, requires two classes of protein to form an active receptor complex called Type I and Type II receptors. To induce a robust response, BMPs bind to heterodimers comprised of two Type I and two Type II receptor proteins. Both classes of protein are serine- threonine kinases, and upon assembly of the receptor complex the constitutively-active Type II receptors phosphorylate the Type I receptors to activate their kinase activity (Hari Reddi, 2003). Active Type I receptors can phosphorylate internal effector proteins called receptor-associated SMADs (r-SMADs). Many BMP receptors also transduce signals from Activin and Nodal signaling; however, Activin, Nodal, and TGFβ signals are transduced through the r-SMADs SMAD2/3 while BMP signals through SMAD1/5/9 (SMAD9 is also known as SMAD8) (Reddi, 2003). Phosphorylated r-SMADs can then bind a co-factor, SMAD4, which acts as a transcription factor. The completed SMAD 17 complex accumulates in the nucleus to bind DNA and affect gene expression. SMAD4 on its own has very weak binding affinity to DNA, and often requires co-factors such as p300 to initiate high-affinity and high-specificity DNA binding (Hill, 2016). It is important to note that SMAD4 is a shared co-factor for r-SMADs SMAD1/5/9 and SMAD2/3, making it an essential effector of both BMP and TGFβ/Activin/Nodal signaling (Weiss & Attisano, 2013). Also important is that while mammals express many different BMP ligands, receptors, and r-SMADs which may serve redundant or compensatory functions, SMAD4 is the only co-factor SMAD expressed in mammals (Huminiecki et al., 2009). Nearly every level of this pathway is carefully regulated by agonists, co-factors, and antagonists (Reddi, 2003). Noggin, Chordin, and Gremlin are three well-studied antagonists that bind BMPs and prevent their interaction with receptors (Walsh et al., 2010). BAMBI is another BMP antagonist which acts as a non-functional pseudoreceptor (Reddi, 2003). SMAD6 and SMAD7 are inhibitory SMADs which compete with SMAD1/5/9, and their expression is often induced by BMP signaling as a form of negative feedback. These regulators play essential roles in embryonic development; in fact, loss of Nog (Noggin), Grem1 (Gremlin), Chd (Chordin) and Smad7 cause embryonic lethality (Bragdon et al., 2011; Q. Chen et al., 2009). Artificial small- molecule inhibitors of BMP signaling have also been produced, such as dorsomorphin (and derivatives), LDN (and derivatives), and VU5350 (Lowery et al., 2016). These generally inhibit the kinase activity of BMP receptors, with varying degrees of specificity for each ALK. As discussed in Chapter 2, these have often been employed to 18 investigate BMP loss-of-function phenotypes, though the possibility of off-target inhibition of other kinases may confound findings. Notably, despite the vast array of possibilities for regulation of individual TGFβ family signaling pathways in vivo and in vitro, SMAD4 presents as a convenient target to assess the requirement for all canonical TGFβ signaling. 1.3.2. BMP signaling is required after implantation in extra-embryonic tissues for gastrulation and mesoderm formation in mammals Maternally-regulated BMP signaling gradients drive axis formation in non- mammalian models BMP signaling has been extensively studied in the embryonic development of non- mammalian model systems for its role as a morphogen governing body axis development. The role of BMP signaling in amphibians is especially famous due to the classic developmental biology experiments performed by Spemann, Mangold, and colleagues (Spemann & Mangold, 1923). The Spemann-Mangold group discovered that a bisected blastula-stage frog embryo would only develop dorsal structures if it included part of the “gray crescent” organizer structure. Later, other groups identified that the dorsalizing components of the gray crescent were antagonists to BMP signaling, including Noggin and Chordin (Sasai et al., 1994; Smith & Harland, 1992). Dorsal expression of these inhibitors creates a gradient of BMP signaling across early frog embryos, with the highest BMP activity at the ventral side of the embryo (Plouhinec et al., 2013). Addition of BMP ligands or overexpression of downstream SMADs can induce formation of ventral tissues in frog embryos, demonstrating that BMP activity is 19 sufficient to induce ventral fate and suppress dorsal fate (Suzuki et al., 1997). In flies, dpp (a homolog of BMP2/4) expression is enriched on the future dorsal side of the blastula-stage embryo forming a dorsal-ventral gradient of BMP activity. Injecting embryos with dpp mRNA dorsalizes the embryo, while dpp mutant embryos are ventralized (Dorfman & Shilo, 2001; O’Connor et al., 2006). In zebrafish, the BMP antagonist Chordin is expressed specifically on the eventual dorsal side of the embryo, forming a gradient of BMP activity which is stronger on the ventral side (Ramel & Hill, 2013). All of these symmetry-breaking events occur very early in development in these model systems, occurring prior to, or coinciding with, the maternal-to-zygotic transition (Lee et al., 2014). Furthermore, the BMP activity gradient in these model systems is partially established by maternally-derived factors which are asymmetrically distributed in the oocyte and embryo prior to zygotic genome activation. In flies, maternal expression of dorsal in the dorsal region is required for asymmetric expression of dpp (Rushlow et al., 1987). In zebrafish, the dorsalized localization of zygotic Chordin is induced by asymmetric localization of maternal β-catenin (Ramel & Hill, 2013). Similarly, ventralized expression of bmp2b and bmp4 in zebrafish is induced by asymmetric localization of maternally-derived radar and pou2 transcripts (Reim & Brand, 2006; Sidi et al., 2003). In addition to directing BMP gradient formation, maternally-derived components of the BMP pathway have also been shown to have direct effects on embryonic development in these species (Das et al., 1998; Faure et al., 2000; Kramer et al., 2002; F. Zhang et al., 2020). Therefore, body axis formation in non-mammalian species is strongly regulated by both maternal and zygotic control of BMP signaling activity. This has prompted many groups to investigate if such a gradient of BMP activity 20 also exists to direct axis specification in early mammalian embryos, and whether BMP activity in mammalian embryos may be driven by maternally-derived factors. BMP signaling is required for peri-implantation cell fate specification in mammals In contrast to other model systems, regulation of axial patterning is not the earliest role for BMP signaling in mammalian embryogenesis. A key piece of evidence for this conclusion is the fact that maternal factors regulating the establishment of a BMP morphogenic gradient is not conserved in mammals. In fact, no intrinsic pattern of maternally-derived molecular components, either mRNA or protein, has been demonstrated in mammalian oocytes or zygotes. Although many studies have described molecular asymmetries in early embryos (Jedrusik et al., 2008; Lim et al., 2020; Piotrowska-Nitsche et al., 2005; Piotrowska et al., 2001; Plachta et al., 2011; Torres- Padilla et al., 2007), no such asymmetries been shown to be deterministic of cell fate during embryonic development (Kurotaki et al., 2007; Motosugi et al., 2005; Rossant & Tam, 2009; Takaoka & Hamada, 2012). Additionally, the mammalian embryo does not specify body axes until relatively late in development compared to non-mammalian model species (Arnold & Robertson, 2009). Instead, the first cell fate decisions of mammalian embryos are dedicated to setting up the extraembryonic structures that the embryo will need to implant into the uterine wall and develop in an internal environment: the placenta, yolk sac, amnion, and chorion. Since components of BMP signaling are expressed prior to body axes specification in mammals, it suggests that it plays a role in these earlier cell fate decisions. 21 Genetic studies of BMP signaling in mammals have shown that BMP signaling is absolutely required for embryonic development. Zygotic knockout of many BMP pathway members in mouse models results in embryonic lethality before mid-gestation. Several of these knockouts, including those for the genes encoding the ligand Bmp4, the Type I receptor Bmpr1a, the Type II receptor Bmpr2, the cofactor Smad4, and downstream target gene Tlx2, all show a similar phenotype where the embryos fail to undergo gastrulation and arrest at early egg cylinder stages (Beppu et al., 2000; Mishina et al., 1995; Sirard et al., 1998; Tang et al., 1998; Winnier et al., 1995; X. Yang et al., 1998). Other BMP knockouts, including those for the genes encoding the receptor-associated SMADs Smad1 and Smad5, and the ligand Bmp2, are able to complete gastrulation and undergo turning, but die at early somite stages due to defects in mesoderm, heart development, and axial patterning (H. Chang et al., 1999; Hayashi et al., 2002; Tremblay et al., 2001; H. Zhang & Bradley, 1996). Additionally, many BMP knockout embryos display an obvious growth restriction at early egg cylinder stages and lack morphological distinction between the embryonic and extraembryonic parts of the embryo (Beppu et al., 2000; G. C. Chu et al., 2004; Di-Gregorio et al., 2007; Mishina et al., 1995; Sirard et al., 1998; Yamamoto et al., 2009; X. Yang et al., 1998; Ying et al., 2000). These phenotypes appear to result from a severe proliferation defect rather than an effect on cell survival, as several BMP knockout embryos show decreased BrdU incorporation or PCNA staining (Mishina et al., 1995; Sirard et al., 1998; X. Yang et al., 1998) and Smad4-null embryos did not display increased apoptosis as assessed by TUNEL staining (Sirard et al., 1998). This shows that BMP signaling is required for cell proliferation and cell fate specification after implantation. 22 BMP signaling regulates gastrulation and mesoderm formation in mouse embryos There is abundant evidence to show that BMP signaling is required prior to body axis specification in mammals to initiate mesoderm formation and gastrulation. Gastrulation, or germ layer formation, usually occurs between E6.25-E7.5 in mice and can be visualized by the appearance of a morphologically distinct tissue known as the primitive streak, which arises from the proximal epiblast (Arnold & Robertson, 2009; Morgani & Hadjantonakis, 2020; Takaoka & Hamada, 2012). The primitive streak represents one of first symmetry-breaking structures in the mouse embryo proper and is essential to establishing the anterior-posterior and dorsal-ventral axes. EPI cells which delaminate from the egg cylinder epithelium and pass through the primitive streak give rise to extraembryonic mesoderm (which contributes to the allantois and part of the amnion and chorion), axial mesoderm and definitive endoderm, and to primordial germ cells (PGCs) (Morgani & Hadjantonakis, 2020; Senft et al., 2019). Knockouts of Bmp4, Bmpr1a, Bmpr2, Acvr1a, and Smad4 completely fail to initiate mesoderm formation (Beppu et al., 2000; G. C. Chu et al., 2004; Di-Gregorio et al., 2007; Z. Gu et al., 1999; Mishina et al., 1995; Sirard et al., 1998; Winnier et al., 1995; X. Yang et al., 1998).These knockouts lack a visible primitive streak at E7.0 and do not express the mesoderm marker Brachyury (T). Other BMP-pathway knockouts, such as Bmp2, Bmp8b, Smad1, Smad5, and Tlx2 are able to initially form mesoderm and a primitive streak, but these structures fail to differentiate properly (Bosman et al., 2006; Tang et al., 1998; Tremblay et al., 2001; Ying et al., 2000; H. Zhang & Bradley, 1996). Some of these mutants initiate T expression, but expression is either decreased or fails to expand distally to designate a fully functional primitive streak (Tang et al., 1998; Tremblay et al., 2001). 23 Crucially, in BMP mutants which are able to initiate gastrulation, many mesoderm- derived structures such as the amnion and allantois are visibly smaller than normal, mispatterned, and fail to properly fuse to create the amniotic cavity and exocoelom (Bosman et al., 2006; H. Chang et al., 1999; Hayashi et al., 2002; Tang et al., 1998; Tremblay et al., 2001; Ying et al., 2000; Ying & Zhao, 2001; H. Zhang & Bradley, 1996). BMP mutants also fail to properly specify primordial germ cells (Arnold et al., 2006; Chuva De Sousa Lopes et al., 2004; Hayashi et al., 2002; Lawson et al., 1999; Senft et al., 2019; Tremblay et al., 2001; Ying et al., 2000; Ying & Zhao, 2001), which are normally derived from a common precursor with mesoderm (Saitou & Yamaji, 2012). In some cases, BMP signaling has been shown to be functionally sufficient to induce PGC formation, as distal epiblast explants from E6.25 embryos (which do not normally specify PGCs) cultured with exogenous BMP8B and BMP4 produced recognizable PGCs (Ying et al., 2001). Altogether, this shows that SMAD-mediated BMP signaling is required for successful gastrulation and mesoderm progenitor specification in early post-implantation mouse embryos. BMP signaling regulates axial patterning in mammals BMP does play a role in the axial patterning in mammals. However, as previously noted, this is not from a maternally-deposited morphogen gradient unlike other model species. Instead, the regulation of axial patterning by BMP is secondary to the role of BMP in mesoderm specification. Anterior-posterior axis formation in mammals is induced by anterior factors secreted from the anterior visceral endoderm (AVE), which forms opposite the primitive streak. Therefore the distal expansion of the primitive streak 24 across from the AVE helps specify posterior fate (Morgani & Hadjantonakis, 2020). In BMP mutants, the primitive streak is often not specified or fails to elongate, which in turn leads to defects in anterior-posterior axis formation (Arnold et al., 2006; Beppu et al., 2000; Fujiwara et al., 2002; Lawson et al., 1999; Sirard et al., 1998). This is shown by a failure of posterior markers such as Cripto and Fgf8 to show asymmetric expression in the epiblast by E7.5 in BMP knockouts (Beppu et al., 2000; Fujiwara et al., 2002). As a second mechanism of BMP regulation of axial patterning, BMP activity is involved in specifying the dorsal-ventral axis, which mimics the proximal-distal axis of the early egg cylinder. BMP patterns the primitive streak by cross-inhibiting Nodal, possibly through competition for the shared downstream cofactor SMAD4 (Morgani & Hadjantonakis, 2020; Senft et al., 2019). As the primitive streak elongates, the most distal streak cells move farther away from the BMP signal in the ExE and inhibition of Nodal signaling is released. Nodal in the more distal (anterior) primitive streak induces expression of BMP inhibitors Noggin and Chordin, reinforcing the decreased BMP signaling in the distal streak. This establishes differential signaling environments in the distal and proximal epiblast which helps characterize the dorsal-ventral axis (Morgani & Hadjantonakis, 2020; Senft et al., 2019) Finally, BMP may also be involved in specifying the left-right axis in mammals. Nodal is a right-side marker, since it is inhibited by Lefty1/2 on the left side of the embryo, and in E8.5 Bmp4-null and Smad1+/-; Smad5+/- embryos expression of Nodal is symmetrical, suggesting a defect in left-right patterning (Arnold et al., 2006; Fujiwara et al., 2002). 25 Possibly in support of this, several BMP mutants which survive to early somite stages display heart looping defects (Arnold et al., 2006; Bosman et al., 2006; H. Chang et al., 1999; Fujiwara et al., 2002; H. Zhang & Bradley, 1996). However, BMP signaling is known to be necessary for heart formation (Uchimura et al., 2009); therefore, it is difficult to determine whether the heart-looping failure is due to a left-right patterning defect or to a direct role for BMP in the heart (Fujiwara et al., 2002). As BMP mutants largely die in utero before any other left-right asymmetries are specified, it may be difficult to test this hypothesis further. However, we can conclude that any role for BMP signaling in left-right patterning is secondary to its earlier roles in gastrulation and embryo growth. BMP regulation of epiblast morphogenesis is non-cell-autonomous Although BMP is required for proliferation and mesoderm specification in the epiblast, several lines of evidence suggest BMP signaling is primarily active in the visceral endoderm. BMP activity has been visualized by several methods, including immunofluorescence or immunohistochemistry for pSMAD1/5/9 and transfection of reporter constructs which incorporate one or several “BMP response element” (BRE) motifs (Bosman et al., 2006; Chuva De Sousa Lopes et al., 2004; Di-Gregorio et al., 2007; Fernandes et al., 2016; Hayashi et al., 2002; Reyes de Mochel et al., 2015; Senft et al., 2019; Yamamoto et al., 2009). It is important to note that not all these reporters agree with one another on the extent of BMP activity in developing mouse embryos. Various studies report conflicting data on the earliest developmental timepoint at which BMP activity can be visualized, with some studies reporting BMP activity as early as the 26 4-cell stage (Reyes de Mochel et al., 2015) while others do not detect BMP activity until E6.5 (Chuva De Sousa Lopes et al., 2004). Similarly, studies do not agree on the cell type specificity of BMP signaling, with some reports claiming BMP activity is present nearly ubiquitously throughout the EPI and VE at early egg cylinder stages (Di-Gregorio et al., 2007) while others report it is restricted to the proximal EPI and extraembryonic VE (Chuva De Sousa Lopes et al., 2004) or to the proximal VE alone (Senft et al., 2019). Therefore, a consensus time course of BMP signaling in mouse embryos has not yet been constructed to date (I report one in Chapter 2). Nevertheless, several studies agree that BMP signaling is active in the VE, particularly around E6.5 when gastrulation is beginning (Chuva De Sousa Lopes et al., 2004; Di-Gregorio et al., 2007; Hayashi et al., 2002; Senft et al., 2019; Yamamoto et al., 2009). Additionally, BMP signaling is suggested to regulate VE maturation or patterning, as several BMP mutant models show disruption of VE morphology or molecular markers (Arnold et al., 2006; G. C. Chu et al., 2004; Z. Gu et al., 1999; Sirard et al., 1998; Tremblay et al., 2001; Yamamoto et al., 2009). Gain-of-function evidence is also available from studies of eXtraEmbryonic Endoderm (XEN) cells, a stem cell line derived from the preimplantation PrE (Kunath et al., 2005). These show that treating XEN cells with exogenous BMP4 upregulates differentiation markers for VE (Artus et al., 2012; Paca et al., 2012). This suggests an important role for BMP signaling in the VE. Interestingly, there is considerable evidence to suggest that BMP signaling in the VE plays an important non-cell-autonomous role to regulate development of the epiblast. Several groups have shown that embryos which lacked expression of BMP pathway 27 members only in the extraembryonic tissues resemble full-body BMP knockouts, suggesting that BMP signaling in the EPI is unable to rescue the proliferation or mesoderm defects. (Z. Gu et al., 1999; Lawson et al., 1999; C. Li et al., 2010; Tremblay et al., 2001; Yamamoto et al., 2009). Similarly, wild-type E5.5 epiblasts stripped of extraembryonic tissues were unable to form PGCs in culture, even when supplemented with exogenous BMP4 in the media (de Sousa Lopes et al., 2004). On the other hand, embryo models where BMP signaling is intact in extraembryonic tissues but deleted in EPI generally developed further than those with full-body BMP knockouts or BMP-null extraembryonic cells (G. C. Chu et al., 2004; Fujiwara et al., 2001; Z. Gu et al., 1999; C. Li et al., 2010; Sirard et al., 1998). Some of these models with wild-type extraembryonic cells actually completed gastrulation and underwent turning, formed mesoderm-derived structures such as allantois, and specified PGCs. This suggests that BMP activity in extraembryonic tissues is sufficient for proper mesoderm formation in the epiblast. However, it is important to note that these embryos, which lack BMP signaling in EPI, still died in utero between E9.5-E10.5 (G. C. Chu et al., 2004; Z. Gu et al., 1999; Sirard et al., 1998). These observations suggest that while BMP signaling in the peri- implantation VE alone is sufficient for gastrulation, mesoderm formation, and PGC specification, cell-autonomous BMP signaling in EPI-derived tissues is still required for later embryonic development. 28 1.3.3. Proposed roles for BMP signaling in preimplantation mammalian development do not agree with post-implantation roles The existing literature does not agree on the earliest role for BMP signaling in mammalian development. Most genetic BMP knockout studies did not report any phenotypes earlier than E5.5-E6.0, at which point Bmp4, Bmp2, Bmp8b, Smad4, Acvr1a, and Bmpr1a mutants appear morphologically normal (Di-Gregorio et al., 2007; Z. Gu et al., 1999; Mishina et al., 1995; Winnier et al., 1995; X. Yang et al., 1998; Ying et al., 2000; H. Zhang & Bradley, 1996). None of these studies conducted any molecular characterization of BMP knockout embryos at these peri-implantation stages; however, a few studies examined Smad4-null and Bmpr2-null blastocysts at E3.5 and reported that the knockout did not cause an apparent phenotype (Beppu et al., 2000; Sirard et al., 1998; X. Yang et al., 1998). Functionally, Bmpr1a-null and Smad4-null blastocysts were able to form recognizable EPI, PrE, and TE-derived cell types which were indistinguishable from controls (Mishina et al., 1995; Sirard et al., 1998; X. Yang et al., 1998). This suggests that without active BMP signaling, preimplantation embryos are able to specify all three cell lineages properly. In contrast to these observations, more recent studies which focused specifically on preimplantation development have proposed that BMP signaling plays an important developmental role at these stages (Graham et al., 2014; Reyes de Mochel et al., 2015; Stuart et al., 2019). These studies detected pSMAD1/5/9 as early as the 4-cell stage in wild-type embryos and X-gal staining in BRE-LacZ reporter embryos at the morula stage (Graham et al., 2014; Reyes de Mochel et al., 2015), suggesting that BMP 29 signaling is active at preimplantation stages. Importantly, this contrasts with a finding published later that GFP is undetectable in BRE:gfp reporter embryos at E3.5 (Fernandes et al., 2016). To test the requirement for BMP signaling in preimplantation development, wild-type embryos were treated with a variety of inhibitors to block BMP receptor activity (Dorsomorphin, Noggin, (5Z)-7-Oxozeaenol, DMH2, and LDN-193189). As a complementary assay, these studies also knocked down BMP signaling by microinjecting wild-type embryos with mRNA to overexpress dominant-negative forms of BMPR1A, BMPR2, and SMAD4. Both dominant-negative receptors and all inhibitor treatments resulted in similar outcomes in both studies. BMP loss-of-function in preimplantation embryos decreased the total amount of TE and PrE cells at the blastocyst stage, resulting in embryos with significantly decreased total cell numbers (Graham et al., 2014; Reyes de Mochel et al., 2015; Stuart et al., 2019). These studies also reported that embryos treated with BMP inhibitors displayed changes in cell cycle time specifically in TE and PrE, which may account for the decreased amount of extraembryonic cells (Graham et al., 2014; Reyes de Mochel et al., 2015). Despite some differences in findings, these studies paint a fairly cohesive picture of a requirement for BMP signaling in preimplantation development, possibly for specification or maintenance of extraembryonic cell types. Notably, gain-of-function studies failed to show that exogenous BMP4 treatment during preimplantation development caused any change in extraembryonic cell fate specification, though they did report a slight decrease in EPI cells (Goissis et al., 2023). This study was also unable to replicate the earlier results and did not report any changes in cell number or cell fate when preimplantation embryos were treated with inhibitors to BMP signaling. 30 This questions the reproducibility of these results and means that the exact role for BMP signaling in preimplantation development has yet to be fully resolved. It is unknown why none of the preimplantation defects described with BMP inhibitor treatment were observed in genetic knockouts of BMP signaling, though there are several possibilities which may explain this discrepancy. On the one hand, as with any small molecule treatment, it is possible that the “BMP” inhibitors have off-target effects which adversely affect preimplantation development. Indeed, many of these inhibitors have been shown to cause considerable off-target effects in vitro; dorsomorphin in particular can inhibit at least 30 other kinases in addition to BMPR1A at the concentrations used in these studies (Lowery et al., 2016). Overexpression of proteins at non-physiological levels and microinjection may also be detrimental to preimplantation embryos (Joris et al., 1998). This would suggest that the genetic knockout studies are a more reliable assay for BMP loss-of-function. However, as preimplantation embryos are known to contain maternally-derived transcripts for many BMP pathway components (Xie et al., 2010), it is possible that these could compensate for loss of the zygotic genes and mask potential preimplantation phenotypes in the genetic knockout studies. Furthermore, BMP4 is known to be produced in the maternal decidua and BMP signaling is active in this tissue during embryo implantation (Monsivais et al., 2021), meaning that essential downstream factors which are normally produced by BMP signaling in the embryo could be supplemented by the uterus. Thus, currently published studies do not clearly specify what role, if any, BMP signaling plays 31 in preimplantation mouse development. My studies in Chapter 2 address this gap in the literature. Section 1.4. Conclusions In summary, molecular signaling pathways dictate morphogenic and cell fate changes in many aspects of mouse embryonic development. Hippo signaling is crucial for regulating the first cell fate decision and polarization of blastomeres in preimplantation embryos. FGF signaling regulates the second cell fate decision between EPI and PrE cells within the preimplantation ICM. BMP signaling has well-described roles after implantation in regulating gastrulation, mesoderm formation, PGC specification, cell proliferation, and anterior-posterior patterning through development of the primitive streak. However, the role of BMP signaling at preimplantation stages is still unclear, as studies using BMP signaling inhibitors have proposed that BMP signaling regulates TE and PrE cell specification and cell cycle rate, but inconsistencies with genetic knockout studies calls these results into question. Thus, despite the depth of knowledge about molecular signaling pathways in mouse embryos, there are still many unanswered questions about their precise roles in development and how each of these pathways may interact with one another to direct proper embryogenesis. 32 FIGURES Figure 1.1. Key events in mouse development from zygote to gastrulation. 33 Figure 1.1 (cont’d). The first week of embryonic development in mice can be divided into pre- and post- implantation stages. During pre-implantation, two cell fate decisions give rise to three important stem cell progenitor types: epiblast, primitive endoderm, and trophectoderm. After implantation, these cell types mature and differentiate to begin to pattern the embryo in preparation for germ layer formation and body axis specification during gastrulation. E = Embryonic Day. Blue boxes denote processes which are proposed to be regulated by BMP signaling. 34 Figure 1.2. Hippo signaling regulates the first cell fate decision in mouse embryos. Beginning at the 16-cell stage in mouse embryonic development, the outer, polarized cells sequester AMOT at the apical domain and prevent its interaction with LATS1/2. This prevents LATS1/2 from phosphorylating YAP1/WWTR1, which are able to enter the nucleus and promote the transcription of TE genes. Inner cells do not have an apical domain, so LATS1/2 are able to phosphorylate YAP1/WWTR1 and prevent their translocation to the nucleus. Without YAP1/WWTR1 transcriptional activity, pluripotency genes are upregulated to establish an ICM cell fate. 35 Figure 1.3. Fibroblast Growth Factor signaling regulates the second cell fate decision in mouse embryos. Within the ICM, cells with higher levels of Fgf4 expression secrete the FGF4 ligand to neighboring cells. Active FGF signaling through MAPK and ERK promotes transcription of Gata6, promoting a PrE fate. Decreased signaling in the Fgf4-producing cells increases transcription of Nanog, promoting an EPI fate. GATA6 and NANOG mutually repress one another, creating feedback loops which solidify the cell fates. 36 Figure 1.4. Biochemical interactions of BMP signaling in mammals. Dimerized BMP ligands bind heterotetramers of Type I and Type II serine-threonine kinase receptors. Activated receptor complexes phosphorylate internal receptor-associated SMAD proteins, which allows them to form complexes with the co-factor SMAD4. The completed SMAD complexes accumulate in the nucleus and regulate transcription, most often in conjunction with other transcription factors. 37 CHAPTER 2. SMAD4 IS ESSENTIAL FOR EPIBLAST SCALING AND MORPHOGENESIS AFTER IMPLANTATION, BUT NONESSENTIAL PRIOR TO IMPLANTATION IN THE MOUSE Robin E. Kruger1,2, Tristan Frum3,4, Stephanie L. Hickey5, A. Sophie Brumm6, Kathy K. Niakan6,7,8,9, Farina Aziz1, Marcelio A. Shammami2,10, Jada G. Roberts11, and Amy Ralston2,3 1) Cell and Molecular Biology Ph.D. Program, Michigan State University, East Lansing, MI, 48824, USA 2) Reproductive and Developmental Sciences Training Program, Michigan State University, East Lansing, MI, 48824, USA 3) Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA 4) Current address: Department of Internal Medicine, University of Michigan Medical School, Ann Arbor, MI, 48109, USA 5) Research Technology Support Facility, Michigan State University, East Lansing, MI, 48824, USA 6) Human Embryo and Stem Cell Laboratory, The Francis Crick Institute; London, NW1 1AT, UK 7) The Centre for Trophoblast Research, Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge, CB2 3EG, UK 38 8) Wellcome Trust – Medical Research Council Stem Cell Institute, University of Cambridge, Jeffrey Cheah Biomedical Centre, Puddicombe Way, Cambridge, CB2 0AW, UK 9) Epigenetics Programme, Babraham Institute, Cambridge, CB22 3AT, UK 10) Genetics and Genome Sciences Ph.D. Program, Michigan State University, East Lansing, MI 48824, USA 11) Molecular, Cellular, and Integrative Physiology Ph.D. Program, Michigan State University, East Lansing, MI 48824, USA This chapter is modified from Kruger et al., submitted to Development. Figures and text pertaining to Figure 2.3, Supplemental Figure 2.3F-G, Supplemental Figure 2.5, and Supplemental Figure 2.6 are not included in the submitted manuscript. This study was supported by the National Institutes of Health R35 GM131759 and T32 HD087166. Work in the laboratory of KKN was supported by the Wellcome (221856/Z/20/Z) and by the Francis Crick Institute (FC001120). 39 Section 2.1. Abstract Bone Morphogenic Protein (BMP) signaling plays an essential and highly conserved role in axial patterning in embryos of many externally developing animal species. However, in mammalian embryos, which develop inside the mother, early development includes an additional stage known as preimplantation. During preimplantation, the epiblast lineage is segregated from the extraembryonic lineages that enable implantation and development in utero. Yet, the requirement for BMP signaling in mouse preimplantation is imprecisely defined. We show that, in contrast to prior reports, BMP signaling (as reported by SMAD1/5/9 phosphorylation) is not detectable until implantation, when it is detected in the primitive endoderm – an extraembryonic lineage. Moreover, preimplantation development appears normal following deletion of maternal and zygotic Smad4, an essential effector of BMP signaling. In fact, mice lacking maternal Smad4 are viable. Finally, we newly uncover a requirement for zygotic Smad4 in epiblast scaling and cavitation immediately after implantation, via a mechanism involving FGFR/ERK attenuation. Altogether, our results demonstrate no role for BMP4/SMAD4 in the first lineage decisions during mouse development. Rather, multi-pathway signaling among embryonic and extraembryonic cell types drives epiblast morphogenesis post-implantation. Section 2.2. Introduction In animal embryos, including mice, frogs, fish, and flies, the Bone Morphogenic Protein (BMP) signaling pathway oversees critical patterning events early in development. In non-mammalian species, BMP signaling is critical for specification of the dorsal/ventral 40 axis of the early embryo (De Robertis & Sasai, 1996; O’Connor et al., 2006; Zinski et al., 2018). However, the mammalian embryo has an additional developmental task immediately following fertilization: specification of the extraembryonic lineages that will give rise to placenta and yolk sac and enable development within the mother. Published studies support roles for BMP signaling in both extraembryonic lineage specification, prior to implantation, and subsequent axial patterning, which occurs after implantation (Graham et al., 2014; Reyes de Mochel et al., 2015; Senft et al., 2019; Yamamoto et al., 2009). However, differences in technical approaches used, as well as challenges intrinsic to mouse such as small embryo size and internal development, have limited origination of a universally accepted model of the role of BMP signaling in mouse embryos throughout pre- and post-implantation stages. BMP is one of several related and highly conserved molecular signaling pathways belonging to the Transforming Growth Factor beta (TGFβ) superfamily of cytokines. The molecular mechanisms of TGFβ signaling have been carefully studied (C. Chang, 2016; Massagué & Sheppard, 2023). BMP proteins, like other members of the TGFβ pathway, are secreted ligands that elicit cellular responses by binding to heterodimeric, transmembrane serine-threonine kinase receptors. The activated receptor complex then phosphorylates members of a family of intracellular effectors known as receptor- associated SMADs (r-SMADs). Phosphorylation of r-SMADs allows their association with a co-factor SMAD and accumulation in the nucleus, where they impact chromatin and transcription (Hill, 2016). In mammals, r-SMAD activity is encoded by several Smad paralogues, with SMAD1, SMAD5, and SMAD9 (also known as SMAD8) primarily 41 transducing BMP signals and SMAD2 and SMAD3 primarily transducing Nodal, Activin, and TGFβ. Notably, the mammalian genome encodes a single co-factor Smad, SMAD4, which is shared by BMP, Nodal, Activin, and TGFβ signaling pathways. Across species, BMP signaling has been visualized in embryos using antibodies that specifically recognize the phosphorylated form of the BMP-responsive r-SMAD(s). This approach has been used to observe gradients of BMP signaling activity that correspond with the dorsal/ventral axis in fly, fish, and frog embryos (Dorfman & Shilo, 2001; Plouhinec & De Robertis, 2009; Schohl & Fagotto, 2002; Tucker et al., 2008). In mouse, no graded pSMAD1/5/9 pattern has been reported. Prior to implantation, pSMAD1/5/9 is reportedly detected in all cell types of the embryo at multiple stages (Graham et al., 2014; Reyes de Mochel et al., 2015). After implantation, pSMAD1/5/9 is detected within a subdomain of extraembryonic cells, and not within the embryo itself until it is detected in emerging mesoderm during gastrulation (Senft et al., 2019). These observations suggest fundamental differences in the roles of BMP signaling between mammalian and non-mammalian animal embryos, but raise the need for additional, functional lines of evidence. In mice, individual members of the BMP signaling pathway appear to be dispensable prior to embryonic day 6.5 (E6.5). Knockout of genes encoding the predominant ligand Bmp4 (Lawson et al., 1999; Winnier et al., 1995), the receptors Bmpr2 (Beppu et al., 2000), Bmpr1a (Mishina et al., 1995), Actr1a (Z. Gu et al., 1999), the r-SMADs encoded by Smad1 (Tremblay et al., 2001) and Smad5 (H. Chang et al., 1999), and the co-Smad 42 Smad4 (Sirard et al., 1998; X. Yang et al., 1998, 2002) all point to essential roles for BMP signaling in extraembryonic mesoderm, extraembryonic endoderm, and germ cell development. Mechanistically, BMP also interacts with Nodal to pattern the visceral endoderm and to specify distal, and then anterior visceral endoderm, structures required to spatially pattern the embryo and specify the primitive streak (Robertson, 2014; Waldrip et al., 1998; Yamamoto et al., 2009). These events define gastrulation and anterior/posterior axial patterning in mouse, processes which therefore rely on BMP signaling. None of these studies reported that BMP signaling loss-of-function had any effect on development prior to E5.5. However, maternal gene products provided within the oocyte could complicate interpretation of knockout phenotypes resulting from zygotic gene deletion only. Indeed, evidence exists that BMP pathway members are maternally supplied and functional in embryos of other animal species (Das et al., 1998; Faure et al., 2000; Kramer et al., 2002; Miyanaga et al., 2002; F. Zhang et al., 2020). Finally, mouse embryos are particularly challenging to recover between E4.5 and E6.5 and we lack an in vitro protocol that robustly recapitulates in vivo development during these stages, presenting a barrier to the facile testing of a possible role for BMP signaling during the peri-implantation period. By contrast, preimplantation embryos are relatively easy to isolate and culture in vitro. Accordingly, several studies have suggested a role for BMP signaling in preimplantation development. Culturing preimplantation embryos in the presence of small-molecule BMP inhibitors led to decreased numbers and cell cycle rate of extraembryonic trophectoderm (TE) and primitive endoderm (PrE) cells, as well as changes in 43 expression of lineage-specific transcription factors, including markers of PrE (SOX17, GATA6), TE (CDX2), and inner cell mass (ICM, OCT4) (Graham et al., 2014; Reyes de Mochel et al., 2015; Stuart et al., 2019). Some of these observations were recapitulated following microinjection of siRNA against Bmp4 or overexpression of dominant-negative forms of Bmpr2 (Graham et al., 2014). Overexpression of dominant-negative Smad4 reportedly phenocopied loss of the upstream signaling components. In principle, these approaches could interfere with the activities of both maternally and zygotically expressed signaling components and thereby achieve more extreme loss of function. However, pSMAD1/5/9 was not examined in these manipulated embryos, so the extent to which these manipulations disrupted BMP signaling is unclear. Moreover, inhibitors are prone to off-target effects, which could further confound interpretation of results (Lowery et al., 2016). In the present study, we visualize pSMAD1/5/9 in wild-type and embryos in which Bmp4 has been maternally and zygotically deleted, as well as lineage specification and morphogenesis in embryos lacking maternal and zygotic Bmp4 or Smad4 throughout preimplantation, peri- implantation, and early post-implantation stages. We report that, in contrast to previous studies, BMP signaling is apparently dispensable during mouse preimplantation development. However, we observed that SMAD4-mediated signaling is essential for peri-implantation epiblast morphogenesis shortly after implantation, when it helps attenuate FGF/ERK signaling to enable the timely execution of epiblast morphogenetic events. 44 Section 2.3. Materials and Methods scRNA-seq Analysis Single-cell RNA-seq data generated by Nowotschin et al. was used to analyze the expression of TGFβ genes in mouse E3.5, E4.5, E5.5, and E6.5 blastocysts (Nowotschin et al., 2019). The analysis was completed using R v4.1.0 with tools from Seurat v4.3.0 (R-core Team 2021) (Hao et al., 2021). We normalized the UMI counts using SCTransform and cells were visualized in 2D space using UMAP performed on the first 30 principal components (Choudhary & Satija, 2022; Hafemeister & Satija, 2019). After excluding TGFβ genes expressed in <10 cells, we used Seurat’s FindAllMarkers function with the Wilcoxon rank-sum test to identify TGFβ genes enriched in each cell type versus all other cells. The p-values were corrected for multiple comparisons using the Bonferroni method. Genes with p-adj<0.01 and average log2 fold change<0.25 were considered cluster enriched. Heatmaps were generated using the pheatmap (v 1.0.12) after averaging the normalized expression for each gene in each cell type. Mouse Strains and Genotyping All animal research was conducted in accordance with the guidelines of the Michigan State University Institutional Animal Care and Use Committee under approved protocol 202300108. Wild type embryos were derived from CD-1 mice (Charles River). The following alleles were used in this study and maintained in a CD-1 background: Bmp4tm1Jfm/J (W. Liu et al., 2004); Smad4tm2.1Cxd/J (X. Yang et al., 2002); Tg(Zp3- cre)93Knw (De Vries et al., 2000). Null alleles were generated by breeding dams 45 carrying homozygous floxed alleles and the Zp3Cre allele to CD-1 males. Mouse genotypes were determined by PCR using genomic DNA extracted using the REDExtract-N-Amp kit (Sigma XNAT) according to the manufacturer’s protocol. Embryo genomic DNA was extracted using the same kit scaled to 10 µL total volume. Genomic extracts (1–2 µL) were then subjected to PCR using allele-specific primers (see Table 2.3). Embryo Collection and Culture Mice were maintained on a 12 hour light/dark cycle. Preimplantation (E2.5-E4.5) embryos were collected by flushing the oviduct or uterus with M2 medium (Sigma M7167). Post-implantation (E4.75-E6.5) embryos were collected by dissecting the embryos from the decidua in ice-cold PBS containing 1% FBS (HyClone SH30396.02) or Bovine Serum Albumin (BSA, Sigma A7888). During embryo collection, dissected embryos were held in warm M2 media. For embryo culture, KSOM medium (Millipore MR-121-D) was equilibrated overnight prior to embryo collection. Where indicated, the following were included in the culture medium: 1 µM or 0.25 µM LDN-193189 in DMSO (Stemgent 04-0074-02); 1 µg/mL recombinant FGF4 in PBS with 0.1% BSA (R&D 235- F4); 1 µg/mL heparin (Sigma H3149); 100 ng/mL recombinant BMP4 in 4 mM HCl (R&D 314-BP); 1 µM PD173074 in DMSO (Selleckchem S1264); 5 µM PD0325901 in DMSO (Stemgent 04-0006); or 0.2% DMSO (New England BioLabs B0515A) as control. Embryos were cultured at 37°C in a 5% CO2 incubator under light mineral oil (Millipore ES-005-C). For serum-free culture conditions (KSOM+PVA-BSA), embryos were cultured as described in Frum & Ralston, 2020. 46 Real-time PCR of Oocytes Smad4 expression levels in oocytes were assessed by real-time PCR as previously described (Blij et al., 2012). Smad4 levels were assessed in oocytes from three wild- type and three Smad4 maternal null females. Oocytes collected from each female were pooled for mRNA extraction and cDNA synthesis. RT-PCR was performed in quadruplicate technical replicates for each cDNA sample. Primers were (5′-3′): Actb, CTGAACCCTAAGGCCAACC and CCAGAGGCATACAGGGACAG; Smad4 (wild-type allele) CGCGGTCTTTGTACAGAGTTA and ACACTGCCGCAGATCAAAG; Smad4 (deleted allele), CACAGGACAGAAGCGATTGA and CCAAACGTCACCTTCACCTT. Immunofluorescence and Confocal Microscopy Preimplantation embryos (E2.5-E4.75) were fixed with 4% formaldehyde (Polysciences 04018) for 10 minutes, permeabilized with 0.5% Triton X-100 (Sigma Aldrich X100) for 30 minutes, and then blocked with blocking solution (10% Fetal Bovine Serum (HyClone SH30396.02), 0.1% Triton X-100) overnight at 4°C. Embryos were incubated with primary antibody overnight at 4°C. The next day, embryos were washed in blocking solution for 30 minutes, incubated in secondary antibody diluted in blocking solution for 1 hour, washed in blocking solution for 30 minutes, then stained with nuclear stain diluted in block for 10 minutes or overnight. Post-implantation embryos (E5.0-E6.5) were fixed with 4% formaldehyde for 1 hour, washed 3 times in 0.1% Tween-20 (Sigma Aldrich P9416), permeabilized for 4 hours in 0.5% Triton X-100, and then blocked with blocking solution (3% BSA (Sigma Aldrich 47 A7888); 0.3% Triton X-100 in PBS) overnight at 4°C. Embryos were incubated with primary antibody overnight at 4°C. The next day, embryos were washed three times in 0.1% Tween-20 for 5 minutes, then incubated in secondary antibody diluted in blocking solution overnight. The following day embryos were washed three times in 0.1% Tween- 20 for 5 minutes, then stained with nuclear stain diluted in block for 10 minutes or overnight. All embryos (preimplantation or post-implantation) which used antibodies against pSMAD1/5/9 were fixed with 4% formaldehyde for 1 hour, methanol dehydration- rehydration series (25%, 50%, 75%, 100%) for 5 minutes each, washed three times in freshly-made 1% Triton X-100 for 10 minutes, washed 20 minutes in ice-cold acetone at -20°C, washed three times in freshly-made 1% Triton X-100 for 10 minutes, then then blocked with blocking solution (10% Fetal Bovine Serum, 0.1% Triton X-100 in PBS) overnight at 4°C. Embryos were incubated with primary antibody overnight at 4°C. The next day, embryos were washed three times in freshly-made 0.1% Triton X-100 for 10 minutes, incubated in secondary antibody diluted in blocking solution for 2 hours, washed three times in freshly-made 0.1% Triton X-100 for 10 minutes, then stained with nuclear stain diluted in blocking solution for 10 minutes or overnight. All embryos (preimplantation or post-implantation) which used antibodies against pERK were fixed with 4% formaldehyde for 1 hour, washed three times for 5 minutes in PBS, washed 20 minutes in ice-cold methanol at -20°C, permeabilized 30 minutes in 0.1% Tween-20, then blocked with blocking solution (3% Bovine Serum Albumin; 0.3% Triton 48 X-100 in PBS) overnight at 4°C. Embryos were incubated with primary antibody overnight at 4°C. The next day, embryos were washed three times in PBS for 5 minutes, incubated in secondary antibody diluted in blocking solution for 2 hours, washed three times in PBS for 5 minutes, then stained with nuclear stain diluted in block for 10 minutes or overnight. All solutions contained HALT protease inhibitor (Thermo Scientific 78430) and PhosSTOP phosphatase inhibitor (Roche 04906837001) diluted 1:500. Antibodies used are listed in Table 2. Embryos were imaged using an Olympus FluoView FV1000 Confocal Laser Scanning Microscope system with 60X PlanApoN oil (NA 1.42) objective. For each embryo, z-stacks were collected, with 5 µm intervals between optical sections. All embryos were imaged prior to knowledge of their genotypes. Embryo Analysis For each embryo, z-stacks were analyzed using Fiji (ImageJ), which enabled the labeling, based on DNA stain, of all individual cell nuclei. Using this label to identify individual cells, each cell in each embryo was then assigned to relevant phenotypic categories, without knowledge of embryo genotype. Phenotypic categories included marker expression (e.g., OCT4 positive or negative) and marker localization (e.g., pSMAD1/5/9 nuclear, absent, or unlocalized). Statistical analysis was performed using GraphPad Prism (v. 9.5.1). Figure images were assembled using Adobe Illustrator. 49 Section 2.4. Results Phosphorylated SMAD1/5/9 is first detectable in peri-implantation embryos To determine when BMP signaling becomes active in the mouse embryo, we first developed a method to examine the localization of transcription factors SMAD1, 5, and 9, which are phosphorylated in response to ligand/receptor binding (Dijke & Hill, 2004). To achieve this, we used immunofluorescence and an antibody that recognizes phosphorylated SMAD1/5/9 (pSMAD1/5/9, Fig. 2.1) (Senft et al., 2019; Xu et al., 2019; Yuan et al., 2015). We did not detect pSMAD1/5/9 in preimplantation embryos flushed from uteri between E3.75-E4.25 (Fig. 2.1A, Supp. Fig. 2.1A). We first observed pSMAD1/5/9 in E4.5 peri-implantation embryos (Fig. 2.1A), when it was detected in nuclei of a few inner cell mass cells in 29% of embryos examined (Fig. 2.1B-C). By E4.75, when embryos have undergone implantation, we observed pSMAD1/5/9-positive cells in 87.5% of the embryos evaluated (Fig. 2.1A-C). Starting at E5.0, we observed pSMAD1/5/9- positive cells within 100% of embryos examined (Fig. 2.1A-C, Supp. Fig. 1A). The observed pSMAD1/5/9 overlapped with a sub-set of GATA6-expressing primitive endoderm (E4.5-E4.75) and visceral endoderm (E5.5-E5.75) cells (Fig. 2.1A). To determine whether the observed pSMAD1/5/9 signal was specific, we treated E5.5 wild-type embryos with LDN-193189 (LDN hereafter) which has been used to disrupt BMP signaling in mouse embryos (Graham et al., 2014; Reyes de Mochel et al., 2015). A concentration of 1 µM LDN was reported as sufficient to inhibit BMP signaling in preimplantation mouse embryos (Reyes de Mochel et al., 2015). However, we found that treatment with 1 µM LDN was highly toxic to embryos (Supp. Fig. 2.1B). 50 Nevertheless, treatment with 0.25 µM LDN led to complete loss of pSMAD1/5/9 signal in E5.5 embryos (Supp. Fig. 2.1B). Altogether, these observations suggest that BMP signaling becomes active around the time of embryo implantation but is not active during preimplantation stages. BMP pathway members are present, but largely inactive, prior to implantation A prior report showed that BMP4 is sufficient to influence gene expression in preimplantation mouse embryos (Goissis et al., 2023), suggesting that preimplantation embryos can respond to exogenous BMP signals. We therefore examined expression dynamics of genes encoding BMP pathway members during preimplantation stages. We analyzed published single-cell RNA-seq data from mouse embryos at stages E3.5- E6.5 (Nowotschin et al., 2019). At E3.5, many core components of canonical BMP signaling were detectable, including the ligand Bmp4, Type I receptor Bmpr1a, Type II receptors Bmpr2 and Acvr2b, receptor-associated SMAD Smad5, and co-factor SMAD Smad4 (Fig. 2.1D and Supp. Fig. 2.2). Next, we investigated whether pSMAD1/5/9 could be induced in preimplantation embryos treated with exogenous BMP. We cultured compacted 8-cell stage embryos (E2.75) in 300 ng/ml BMP4 for 36 hours to the blastocyst stage (equivalent in cell number to E3.75, as confirmed by cell counts). Although we did not observe pSMAD1/5/9 in any control embryos cultured in unsupplemented medium, we observed low, but detectable levels of pSMAD1/5/9 in 82% (n=14/17) of embryos treated with exogenous BMP4, further supporting the faithful detection of pSMAD1/5/9 by 51 immunofluorescence analysis (Fig. 2.1E-F). Notably, pSMAD1/5/9 was detected only in the ICM but did not preferentially colocalize with either SOX2-positive epiblast (EPI) or SOX2-negative PrE cells. Therefore, we conclude that BMP signaling is not highly active during preimplantation development, but ICM cells are competent to respond to exogenous BMP signals at these stages, consistent with published investigations (Graham et al., 2014; Reyes de Mochel et al., 2015). We next analyzed the pSMAD1/5/9 in embryos shortly after implantation. Consistent with prior reports (Senft et al., 2019; Yamamoto et al., 2009), we detected pSMAD1/5/9 within a zone of the visceral endoderm (VE) that flanks the extraembryonic ectoderm (ExE) at E5.5 and E5.75 (Fig. 2.1A, Supp. Fig. 2.1A). This observation is also consistent with evidence that several key components, including Bmp2, Smad1, Smad5, and Bmpr2, are substantially upregulated around the time of implantation (E4.5-E5.5), particularly in the PrE/VE lineage (Fig. 2.1D, Supp. Fig. 2.1A- B). Notably, culturing E5.5 embryos in the presence of exogenous BMP4 for 6 hours was sufficient to expand the zone of pSMAD1/5/9 within the VE in a dose-dependent manner (Supp. Fig. 2.1C). Thus, the availability of ligand could limit the extent of pathway activation, during both pre- and post-implantation stages. Finally, we evaluated pSMAD1/5/9 in Bmp4-null embryos at E5.5. We were unable to detect pSMAD1/5/9in Bmp4-null embryos, although it was observed at wild-type levels and localization in homozygous wild-type littermate controls (Fig. 2.1G-H). In Bmp4 heterozygous embryos, we observed an intermediate phenotype where some 52 pSMAD1/5/9 was detectable but trended toward lower numbers of pSMAD1/5/9-positive cells than wild-type (Fig. 2.1H). This suggests that at E5.5, BMP4 plays a major role in initiating BMP signaling activity in the mouse, and that this function of BMP4 is dose- dependent. Maternal Bmp4 and Smad4 are not required for development Previous knockout studies of BMP signaling components did not report preimplantation phenotypes (Beppu et al., 2000; Mishina et al., 1995; Sirard et al., 1998; Winnier et al., 1995). However, other groups reported defects in preimplantation lineage specification using pathway inhibitors or microinjection of RNAi or mRNA for dominant-negative overexpression (Graham et al., 2014; Reyes de Mochel et al., 2015; Stuart et al., 2019). One way to reconcile these disparate findings is to invoke a model in which some components of the BMP pathway are maternally imparted to the oocyte and participate in preimplantation development to compensate for previously reported zygotic null mutations. To further investigate this possibility, we examined cell fate specification in embryos lacking both maternal (m) and zygotic (z) Bmp4 or Smad4 using the female germ line-expressed Zp3-Cre (De Vries et al., 2000) in combination with floxed alleles of either Bmp4 or Smad4 (see Supp. Fig. 2.3A for breeding scheme). RT-qPCR analysis confirmed the absence of detectable Smad4 transcript in Smad4 mz null embryos (Supp. Fig. 2.3B), as we have observed for many other loci deleted using this approach (Blij et al., 2012; Frum et al., 2013, 2018; Wicklow et al., 2014). 53 Remarkably, we were able to recover either Bmp4 or Smad4 mz null blastocysts at predicted rates, indicating no requirement for maternal Bmp4 or Smad4 on fertilization or embryo development. Moreover, both Bmp4 and Smad4 mz null embryos exhibited normal morphology, total cell number, and ratio of trophectoderm and ICM cells (Fig. 2.2, Supp. Fig. 2.3C-E). In addition, the ICM marker Oct4 was detected at normal levels within the ICM, and CDH1 localization strongly suggested that the TE was properly polarized (Supp. Fig. 2.4). Finally, the expression of EPI and PrE cell fate markers at E3.75, E4.25, and E4.5 (Fig. 2.2, Supp. Fig. 2.3C-E) was unaffected in either Bmp4 or Smad4 mz null embryos at these stages, consistent with normal ICM differentiation. Our observations support the conclusion that canonical BMP signaling does not play a major role in preimplantation development. As the embryos in these studies were fixed immediately after collection from the uterus, it is possible that uterine-derived signals compensated for loss of BMP4 in embryos. To eliminate potential exogenous BMP4 from the uterus, we collected Bmp4 mz null embryos at E1.5 and cultured for 60 hours to blastocyst stage. As serum may contain BMP agonists or antagonists that can affect embryo development in culture, we conducted this experiment in serum-free conditions by supplementing KSOM media with polyvinyl alcohol without bovine serum albumin (Frum & Ralston, 2020). The cultured Bmp4 mz null embryos again displayed normal morphology, and we did not observe any difference in total cell number or EPI:PrE ratio when compared to wild-type controls (Supp. Fig. 2.3F-G). In a parallel set of experiments, we allowed Smad4 m null embryos to develop to term. Mice lacking m Smad4 were born and developed 54 apparently normally to 4 months old (10/10 mice, two litters). We conclude that maternal Bmp4 and Smad4 are dispensable for development and that neither zygotic gene plays a predominant role prior to implantation. Exogenous BMP4 is capable of inducing cell fate changes in preimplantation embryos Because we were able to induce pSMAD1/5/9 expression in preimplantation embryos cultured with exogenous BMP4, we wanted to test the functional consequences of that induced signaling. BMP signaling has been suggested to affect preimplantation cell fate (Goissis et al., 2023; Graham et al., 2014; Reyes de Mochel et al., 2015). To test whether our induced BMP signaling could affect preimplantation cell fate, we cultured wild-type E2.75 embryos in 100 ng/ml or 300 ng/mL BMP4 for 36 hours to blastocyst stage. Examination of cell fate markers in these embryos by immunofluorescence did not reveal any significant differences in total cell number or the amount of EPI, PrE or TE cells (Fig. 2.3A-B, Supp. Fig. 2.5). This suggests that BMP4 treatment alone is not sufficient to induce changes in cell fate in preimplantation embryos. To test this further, we cultured embryos with 100 ng/mL BMP4 in the presence of 1 ug/mL exogenous FGF4 and heparin, a treatment known to cause changes in preimplantation cell fate (Yamanaka et al., 2010). Treatment with FGF4/heparin alone directed the entire inner cell mass of treated embryos toward a PrE cell fate, as expected (Fig. 2.3A-B, Supp. Fig. 2.5C). Embryos treated with both exogenous BMP4 and exogenous FGF4/heparin did not rescue EPI specification, but instead showed a 55 significant increase in the amount of inner cell mass cells which did not express markers for either EPI or PrE cell fate (Fig. 2.3C, Supp. Fig. 2.5C). Importantly, this effect was not seen in Smad4 mz null embryos cultured under the same conditions (Fig. 2.3C, Supp. Fig. 2.5C), suggesting that the increase in unspecified ICM cells relies on BMP signaling capability. Overall, these results suggest that although BMP signaling does not normally function in preimplantation cell fate specification, it has a capability to do so under specific culture conditions. Smad4 is required for epiblast cavitation at E5.5 in a Bmp4-independent manner Prior studies mainly focused on characterization of BMP signaling loss of function phenotypes at later stages (>E5.5) (Sirard et al., 1998; Winnier et al., 1995; X. Yang et al., 1998). However, we first observed BMP signaling activity in most embryos just after implantation at E4.75, prompting us to examine embryos lacking Bmp4 or Smad4 beginning at E4.75. At E4.75, Smad4 null embryos were grossly morphologically normal (Fig. 2.4A). However, upon close examination, Smad4 null embryos displayed a significant decrease in total cell number (Fig. 2.4B). We quantified the number of EPI, PrE, and TE cells in these embryos and discovered that the decreased cell number was most pronounced in EPI cells at this stage (Fig. 2.4B-C). By E5.5, Smad4 null embryos were visibly reduced in size and all displayed disorganization in EPI, VE, and ExE compartments, as expected from studies performed at E6.5 (Sirard et al., 1998; X. Yang et al., 1998) (Fig. 2.4D). Strikingly, the epiblast was greatly reduced in cell number (Fig. 2.4D-F) relative to controls, and had 56 not yet cavitated, as the proamniotic cavity was not present among most (>80%, n=7/9) Smad4 null embryos examined (Fig. 2.4D, 2.4L). Notably, relative to extraembryonic lineages, the amount of EPI cells was disproportionately decreased in Smad4 null embryos since the epiblast length was decreased even when normalized to proximal- distal embryo length (Fig. 2.4G-I, Supp. Fig. 2.6A). By contrast, the size of the ExE was appropriately scaled to the reduced size of Smad4 null embryos at E5.5 (Fig. 2.4J-K). This suggests that Smad4 is not only required for general embryonic growth, but also specifically required for epiblast growth relative to total embryo size. Notably, these phenotypes were not observed in E5.5 Bmp4-null embryos, which did not differ from wild-type in morphology, embryo size, or EPI or PrE cell number (Supp. Fig. 2.6B-E), suggesting that signaling pathways other than BMP signaling feed into SMAD4- regulation of epiblast growth and morphogenesis at this stage. Defects in Smad4-null embryos are more widespread across cell types at E6.5 We next examined the epiblasts of Smad4 mz null embryos at E6.5, which in other studies had been reported to display primarily extra-embryonic defects with relatively unaffected epiblasts (Sirard et al., 1998; X. Yang et al., 1998). Many of the defects we observed in E5.5 Smad4-null embryos were still observed at E6.5 (Supp. Fig. 2.7A). Smad4-null embryos showed a more significant reduction in total embryo size at this stage (Supp. Fig. 2.7B), as well as a highly disorganized and reduced EPI compartment (Supp. Fig. 2.7C). However, the difference in epiblast length as a percentage of total embryo length was no longer statistically significant between Smad4-null embryos and controls (Supp. Fig. 2.7D). Additionally, 60% of the observed E6.5 embryos had 57 cavitated (N=3/5) suggesting that loss of Smad4 delays initiation of cavitation but does not prevent cavitation entirely. This is in agreement with previous studies, which did not report a specific defect in epiblast size or cavitation at E6.5 (Sirard et al., 1998; X. Yang et al., 1998). This observation suggests that by E6.5 the overall growth restriction of the entire embryo is so severe that the specific effect on the growth of the EPI is no longer apparent. None of the phenotypes observed in E6.5 Smad4-null embryos were as severe in E6.5 Bmp4-null embryos, although there was a slight significant decrease in total embryo length in E6.5 Bmp4-null embryos (Supp. Fig. 2.7E). The observed difference in total embryo size was not specific to the EPI (Supp. Fig. 2.7F-G). This suggests that there is a requirement for Bmp4 at E6.5, but that the more severe effect of Smad4 does not rely on Bmp4. Epiblast cavitation requires SMAD4-dependent inhibition of FGF/ERK signaling Having discovered that Smad4 is required for epiblast cavitation and growth at E5.5, we began to investigate the mechanism underlying this role. We were struck by the observation that the ExE appeared disproportionately large, relative to the size of the EPI in E5.5 Smad4 null embryos. To confirm the identity of the ExE cells, we examined markers of ExE, including phosphorylated ERK (pERK), which is elevated within the ExE (Corson et al., 2003). In Smad4 null embryos, we observed pERK throughout putative ExE, consistent with their identity as ExE cells. However, we observed dramatically elevated levels of pERK within this region (Fig. 2.5A). 58 Since pERK in the ExE is known to be dependent on signaling by the Fibroblast Growth Factor (FGF) pathway (Corson et al., 2003), we therefore hypothesized that increased pERK could be due to elevated FGF signaling in Smad4 null embryos. To test this hypothesis, we used a previously-published protocol to inhibit FGF signaling in embryos (Yamanaka et al., 2010), which effectively eliminated pERK in control embryos (Supp. 2.8A). Notably, inhibition of FGF signaling partially rescued epiblast defects in E5.5 Smad4 null embryos (Fig. 2.5A). We observed a significant increase in epiblast scaling, as well as increased rates of cavitation (Fig. 2.5B-D). However, FGF inhibitor treatment did not rescue the overall growth restriction of Smad4-null embryos (Fig. 2.5E), nor was the number of OCT4-positive cells restored (Fig. 2.5F). These observations are consistent with SMAD4 repressing FGF signaling in the ExE during early post- implantation stages as a critical regulator of epiblast morphogenesis. Our observations also suggest that additional pathways regulate embryo growth downstream of SMAD4. To determine whether elevated FGF signaling is sufficient to antagonize cavitation, we treated embryos with exogenous FGF4. In wild-type E5.5 embryos treated with exogenous 1 µg/mL FGF4, we observed elevated levels of pERK within the ExE and ectoplacental cone (Supp. Fig. 2.8A). However, we observed no impact on epiblast cavitation or epiblast size following this treatment (Supp. Fig. 2.8A-D). Altogether, these data suggest that pERK/FGF signaling antagonizes EPI cavitation, but upregulation of pERK alone is insufficient to induce cavitation defects in wild-type embryos, at least under the conditions tested here. 59 Section 2.5. Discussion Despite being a highly conserved developmental signaling pathway, the role of BMP signaling in regulating peri-implantation mammalian development has not been fully elucidated. In part, this may be due to the technical challenges surrounding studying mammalian embryos at pre- and peri-implantation stages of development. Our study undertook a detailed examination of molecular markers of BMP signaling activity and the phenotypes of maternal-zygotic genetic knockout models of BMP signaling to identify the earliest role of BMP signaling in development. Although we failed to detect a role for BMP signaling in preimplantation embryos, we newly identified a role for Smad4 in epiblast organization and cavitation at early post-implantation stages. We focused our initial studies on defining the role of BMP signaling in preimplantation mouse development because there was not a consensus on this role in the existing literature. Previous studies of Bmp4 and Smad4 genetic knockouts in mice have largely focused on the more overt phenotypes which occur after implantation (G. C. Chu et al., 2004; C. Li et al., 2010; Senft et al., 2019; Sirard et al., 1998; Winnier et al., 1995; X. Yang et al., 1998). However, some studies have reported deleterious effects of BMP knockdown on in vitro cultured preimplantation embryos (Graham et al., 2014; Reyes de Mochel et al., 2015; Stuart et al., 2019). These studies found a range of effects, but largely reported a specific deficiency in extra-embryonic cell types, either by decreased cell numbers (Graham et al., 2014) or slower cell cycle (Reyes de Mochel et al., 2015). In contrast, our study did not uncover any defect in Bmp4- or Smad4-null preimplantation embryos, which did not differ from wild-type in morphology, cell type 60 specification, or expression of key developmental transcription factors even in Bmp4- null embryos cultured in defined media without BMP4 from E1.5 to blastocyst. These data suggest that the normal development of flushed embryos lacking BMP signaling factors is not due to replacement factors supplied by the uterus. Furthermore, in contrast to previous studies (Graham et al., 2014; Reyes de Mochel et al., 2015), we were unable to detect any pSMAD1/5/9 signal at preimplantation stages in flushed embryos, suggesting a lack of endogenous BMP signaling activity. Overall, our data strongly suggest that BMP signaling (or indeed, any signaling pathway mediated through SMAD4) does not play a role in preimplantation development or stem cell progenitor specification in mice. Several differences exist between our current study and previous studies of BMP signaling in preimplantation development which may explain our failure to replicate the previous results. The earlier studies primarily examined preimplantation embryos in culture with serum. This may explain why they detected pSMAD1/5/9, as it is possible that the reported pSMAD1/5/9 signal was an artifact of culture conditions in undefined media. In support of this hypothesis, we were able to induce pSMAD1/5/9 when preimplantation embryos were cultured with exogenous BMP4. Differences in the embryo fixation or permeabilization protocols, the specificity of the antibodies used, or the genetic background of the embryos may also contribute to the differences in pSMAD1/5/9 expression or developmental phenotypes. Indeed, genetic background has already been shown to affect the severity of phenotypes in Bmp4 knockout models (Winnier et al., 1995). Additionally, these earlier studies primarily modulated BMP 61 signaling with either pharmacological inhibitors or overexpression of dominant-negative receptors. As overexpressing proteins at above-physiological levels and off-target effects of chemical inhibitors have both been shown to induce detrimental phenotypes, it is also possible that the previously reported changes in preimplantation development were the result of non-specific effects. Several studies have reported that commonly- used BMP inhibitors such as dorsomorphin, LDN-193189, and DMH2 have the capability to inhibit dozens of non-specific kinases (Boergermann et al., 2010; Lowery et al., 2016; J. Vogt et al., 2011). We report a novel requirement for Smad4 in post-implantation development. Previous descriptions of Smad4-null embryos report growth restriction and disorganized VE beginning at E5.5, with embryonic lethality by E8.5 (G. C. Chu et al., 2004; Sirard et al., 1998; X. Yang et al., 1998). Our results are consistent with these findings, but also uncover a previously unappreciated defect specific to the epiblast. The proportion of EPI cells is decreased starting at E4.75, suggesting that in the absence of Smad4 the number of EPI cells does not scale with embryo growth. The structure of the epiblast is also disrupted in these embryos, as Smad4-null embryos had largely disorganized epiblasts which failed to form a proamniotic cavity. It is unlikely that this defect is cell- autonomous, as embryos with an epiblast-specific knockout of Smad4 do not display an observable defect until after gastrulation (G. C. Chu et al., 2004). Rather, it is likely that Smad4 is required in the visceral endoderm to produce a signal that promotes EPI development non-cell-autonomously (Fig. 2.6A). This model is supported by the finding that VE-specific Smad4-knockout embryos fail to gastrulate properly and more closely 62 resemble whole-body Smad4-null embryos (C. Li et al., 2010). However, there was no defect reported in the VE-specific Smad4-KO embryos earlier than E6.5. It is possible that this discrepancy comes from the VE-specific Cre driver used to generate the knockout in this study, which is not active until E5.75. Since our study first observed an epiblast defect at E4.75, an earlier knockout of Smad4 specific to the primitive endoderm lineage would be needed to test that the epiblast defect is in fact a non-cell-autonomous effect from the visceral endoderm. It is also interesting to note that a cavitation defect was not reported in previous Smad4 knockout mouse models which targeted the same exon 8 (Sirard et al., 1998; X. Yang et al., 1998). It is possible that the phenotypic differences are due to differences in genetic background, as both of these studies were completed in C57/B6 mice, as opposed to the CD-1 background used in the current study. However, we also observed that some Smad4 null embryos were cavitated at E6.5, so it is possible that the phenotypic discrepancies can be explained by a difference in developmental staging. Our observations are consistent with prior observation that BMP signaling is necessary and sufficient for cavitation of embryoid bodies (Coucouvanis & Martin, 1999). In this context, BMP2 and BMP4 may be functionally redundant, and could explain why we observed normal cavitation in E5.5 Bmp4-null embryos. Alternatively, SMAD4 could regulate epiblast morphogenesis through another TGF-β pathway. Knockout of Nodal has been shown to decrease embryo size and expression of Oct4 mRNA at early post- implantation stages (Brennan et al., 2001; Mesnard et al., 2006), but Nodal-null embryos cavitate normally and expression of OCT4 protein is apparently unaffected at E5.5 (Senft et al., 2019). As the cavitation defect and loss of OCT4-positive cells are 63 more severe in Smad4-null embryos than either Bmp4- or Nodal-null models alone, it suggests that Smad4 may regulate epiblast morphogenesis through a combination of TGFβ pathways. Our data also suggests that SMAD promotes cavitation by attenuating pERK levels. Inhibition of FGF/MAPK signaling rescued cavitation in Smad4-null embryos, even though the epiblast remained disproportionately small. This observation allows us to propose that cavitation is not dependent on epiblast size, but rather the embryo signaling environment. Our data suggest that ectopic upregulation of pERK resulting from loss of Smad4 is detrimental to epiblast cavitation; however, the mechanisms of how these two pathways interact with one another and regulate cavitation remain unclear. SMAD4 could regulate several processes associated with epiblast cavitation, including epiblast cell apoptosis or epiblast cell polarization and lumenogenesis (Bedzhov & Zernicka-Goetz, 2014; Coucouvanis & Martin, 1995, 1999; Halimi et al., 2022) (Fig. 2.6B-C). In this model, Smad4 is sufficient for initiation of cavitation, possibly through regulation of apoptosis by BMP signaling (Fig. 2.6A). Separately, Smad4 also restricts MAPK signaling through ERK phosphorylation, which expands distally in the absence of Smad4. Meanwhile, FGF/MAPK partially regulates rosette formation, possibly through regulation of basement membrane formation by the emVE. In the presence of Smad4, when FGF/MAPK is restricted, rosette formation occurs properly, and cavitation is successful. In the absence of Smad4, the epiblast does not receive the signal for programmed cell death and does not initiate lumen formation. FGF/MAPK is also ectopically expanded, and rosette formation fails (Fig. 2.6B). By inhibiting ectopic 64 FGF/MAPK in Smad4-null embryos at the critical timepoint just before cavitation, basement membrane deposition occurs and rosette formation is allowed to proceed, forming a small proamniotic cavity. However, since the apoptotic signal from Smad4 is still missing, cavitation is not completely rescued (Fig. 2.6C). This model suggests that the reason pERK upregulation was insufficient to compromise EPI cavitation in wild-type embryos is that by the time of treatment at E5.25, EPI polarization was already complete. In this way, in vitro modulation of FGF/MAPK signaling only affects cavitation in the absence of Smad4. Although the above proposed mechanisms are consistent with our data, it is not proven from the current study that Smad4 regulates cavitation through apoptosis or basement membrane deposition. Alternatively, SMAD4 could promote epiblast maturation, defined here as the transition from a naïve to primed state which normally occurs in epiblast cells between preimplantation and post-implantation stages (Boroviak et al., 2015; Nichols & Smith, 2009). Epiblast maturation has been shown to be critical in formation of proamniotic cavity (Carbognin et al., 2023; Shahbazi et al., 2017). Other mechanisms are also proposed to play a role in embryonic cavitation such as apical domain maintenance (Meng et al., 2017), tight junction formation (C. J. Chan et al., 2019), cellular adhesion through ECM interactions (Liang et al., 2005; Sakai et al., 2003), and establishment of an osmotic gradient (Dumortier et al., 2019). SMAD4 and pERK may regulate any or a combination of these factors. Further studies will be needed to interrogate the interaction of these pathways further, to identify the precise regulatory 65 mechanisms governing post-implantation epiblast morphogenesis downstream of SMAD4 and pERK. Section 2.6. Acknowledgements We thank the lab of Dr. David Arnosti for their generous loan of laboratory equipment. We also thank Barbara Makela and Ella Markley for technical support. Section 2.7. Diversity and Inclusion Statement The authors wholeheartedly support all efforts to increase the inclusion of scientists from underrepresented backgrounds (including, but not limited to, women, LGBTQIA+, people of color, Indigenous people, neurodivergent people, people with disabilities, and people from disadvantaged backgrounds) in developmental biology and related careers. This manuscript reflects the efforts of authors who identify as members of several of these groups. The authors believe that diverse perspectives are essential for scientific excellence and innovation, yet we acknowledge the continued existence of systemic barriers to success for scientists of underrepresented and marginalized communities. We support the development of initiatives to address disparities and biases in scientific publishing and encourage further efforts to implement inclusive practices. 66 TABLES Allele CD-1 Common Allele Name CD-1 Tg(Zp3- cre)93Knw/J Zp3Cre Bmp4tm1Jfm/J Bmp4fl Smad4tm2.1Cx d/J Smad4fl Strain, (species) Source or Reference CD-1 (Mus musculus) Mixed background (M. musculus) Charles River Laboratories De Vries et al, 2000 (PMID: 10686600) Mixed background (M. musculus) Mixed background (M. musculus) Liu et al, 2004 (PMID: 15070745; PMCID: PMC384774 ) Yang et al, 2002 (PMID: 11857783) Identifier(s) Additional Information Wild-type Presence of this allele assessed by genotyping for Cre and Zp3Cre Bmp4del or Bmp4 null refers to recombined allele Crl:CD1(IC R), Strain code: 022 RRID:IMSR _JAX:00365 1 MGI:J:6790 3 RRID:IMSR _JAX:01687 8 MGI:J:8923 7 RRID:IMSR _JAX:01746 2 MGI:J:7514 0 Smad4del or Smad4 null refers to recombined allele Table 2.1. Animal Resources Table for Chapter 2. 67 Antibody/Stain Source or Identifiers Dilution Additional Information Goat-anti- hGATA6 Goat-anti- GATA4 Goat-anti- SOX17 Mouse-anti- CDX2 Rabbit-anti- CDX2 Mouse-anti- OCT4 AF1700 1:100 sc1237 1:2000 AF1924 1:2000 CDX-88 sc-5279 1:500 1:200 1:100 reference(s) R&D Systems Santa Cruz Biotechnolog y R&D Systems Abcam Santa Cruz Biotechnolog y Reprocell Abcam ab76541 Sigma Cell Signaling Rabbit-anti- NANOG Goat-anti-SOX2 Neuromics Rat-anti-CDH1 Rabbit-anti- Phospho- Smad1 (Ser463/465)/ Smad5 (Ser463/465)/ Smad9 (Ser465/467) Rabbit-anti- Phospho-p44/42 MAPK (Erk1/2) (Thr202/Tyr204) Donkey-anti- goat IgG Donkey-anti- mouse IgG Donkey-anti- rabbit IgG Goat-anti-rabbit IgG Goat-anti- mouse IgG DRAQ5 Cell Signaling Jackson Laboratories Jackson Laboratories Invitrogen Invitrogen Invitrogen Cell Signaling RCAB002P-F 1:400 GT15098 U3254 13820 1:2000 1:500 1:50 9101 1:100 A-11055 1:400 Alexa488 715-165-150 1:400 711-165-152 1:400 A-11034 A-11029 4084 1:400 1:400 1:400 Cy3 Cy3 Alexa488 Alexa488 Table 2.2. Antibody Table for Chapter 2. 68 Allele Bmp4WT, Bmp4fl Bmp4del Smad4WT, Smad4fl Smad4del Cre Zp3Cre Forward Primer GAGCTAAGTTTTGCTGG TTTGC GCTAAGTTTTGCTGGTT TGC TAAGAGCCACAGGGTCA AGC TAAGAGCCACAGGGTCA AGC CTAGGCCACAGAATTGA AAGATCT CGAGATTGAGGGAAGC AGAG Reverse Primer GCCCATGAGCTTTTCTG AGA TGACTAGGGGAGGAGT AGAAGGTG TTCCAGGAAAAACAGGG CTA GACCCAAACGTCACCTT CAG GTAGGTGGAAATTCTAG CATCATCC CAGGTTCTTGCGAACCT CAT Table 2.3. Allele-specific primers for PCR genotyping for Chapter 2. 69 FIGURES Figure 2.1. BMP signaling becomes active in primitive endoderm at implantation. 70 Figure 2.1 (cont’d). A) SMAD1/5/9 phosphorylation (pSMAD) in wild-type CD-1 embryos at E3.75, E4.5, E4.75, and E5.5. In all cases, positive pSMAD signal co-localizes with GATA6 as a marker of primitive endoderm and visceral endoderm. B) Quantification of total number of pSMAD-positive cells in wild-type embryos in A and Supplemental Figure 2.1A. C) Quantification of the percentage of embryos from A and Supplemental Figure 2.1A which display any pSMAD-positive cells versus no pSMAD-positive cells. D) Heat map of the mean normalized expression of BMP pathway genes from scRNA-seq data from Nowotschin et al., 2019. E) pSMAD in wild-type embryos collected at E2.75 and cultured for 36 hours in media containing 300 ng/mL exogenous BMP4. F) Quantification of the total number of pSMAD-positive cells in embryos from E revealed significantly more pSMAD-positive cells in BMP4-treated embryos. G) pSMAD staining is absent in Bmp4 z null embryos at E5.5. H) Quantification of total number of pSMAD- positive cells in wild-type and Bmp4-null embryos at E5.5 revealed significantly fewer pSMAD-positive cells in Bmp4-null embryos. All pairwise comparisons were assessed by analysis of variance (ANOVA) with Tukey’s post-hoc test. White arrowheads indicate positive pSMAD signal. Red arrowhead indicates a GATA6-positive cell which does not express pSMAD. Scale bars represent 10 μm. 71 Figure 2.2. Maternal and zygotic Smad4 and Bmp4 are dispensable for blastocyst formation and preimplantation cell fate specification. A) Immunofluorescence for SOX17 and NANOG as respective markers of primitive endoderm (PrE) and epiblast (EPI) in flushed E3.75 wild-type CD-1 embryos and embryos lacking maternal and zygotic Bmp4 (mz null). Quantification did not reveal any significant difference in cell number or cell fate between Bmp4 mz null embryos and controls. 72 Figure 2.2 (cont’d). “Mixed” indicates co-expression of SOX17 and NANOG. B) Immunofluorescence for SOX17 and NANOG in flushed E4.25 embryos lacking maternal Bmp4 only (m null) and Bmp4 mz null embryos. Quantification did not reveal any significant difference in cell number or cell fate between Bmp4 mz null embryos and controls. “Mixed” indicates co- expression of SOX17 and NANOG. C) Immunofluorescence for SOX17 and NANOG in flushed E3.75 Smad4 m null and Smad4 mz null embryos. Quantification did not reveal any significant difference in cell number or cell fate between Smad4 mz null embryos and controls. D) Immunofluorescence for SOX17 and NANOG in flushed E4.25 Smad4 m null and Smad4 mz null embryos. Quantification did not reveal any significant difference in cell number or cell fate between Smad4 mz null embryos and controls. All pairwise comparisons were assessed by Student’s t-test. Scale bars represent 10 μm. 73 Figure 2.3. Exogenous BMP4 is capable of inducing cell fate changes in preimplantation embryos. A) Wild-type embryos collected at E2.75 and cultured for 36 hours in untreated control media, 100 ng/mL BMP4, 300 ng/mL BMP4, 1 ug/mL FGF4 + heparin, or both 100 ng/mL BMP4 and 1 ug/mL FGF4 + heparin. Smad4 mz null embryos collected at E2.75 and cultured for 36 hours in both 100 ng/mL BMP4 and 1 ug/mL FGF4 + heparin. Embryos were stained by immunofluorescence for NANOG and SOX17 as markers of EPI and PE, respectively. B) Quantification of the amount of EPI, PE, mixed, and no expression cells as a percentage of total ICM cells in embryos from A. EPI=NANOG-positive only, PE=SOX17-positive only, mixed=NANOG/SOX17 double- positive, no expression=NANOG/SOX17 double-negative. C) Quantification of NANOG/SOX17 double-negative cells as a percentage of ICM cells in embryos from A. Comparisons were assessed by analysis of variance (ANOVA) with Tukey's post-hoc test. 74 Figure 2.4. BMP-independent function of Smad4 is required for post-implantation epiblast organization and maintenance. A) E4.75 Smad4 mz null embryos stained by immunofluorescence for SOX17 and NANOG. B) Quantification of EPI, PrE, and TE cell numbers from embryos in A revealed a significant decrease in EPI cells in Smad4 mz null embryos when compared to controls. C) Quantification of the EPI, PrE, and TE cells as a percentage of total cell number from embryos in A revealed a significant decrease in EPI percentage in Smad4 mz null embryos. D) E5.5 Smad4-/- embryos stained by immunofluorescence for OCT4 and GATA6 as markers of EPI and VE, respectively. 75 Figure 2.4 (cont’d). Smad4-/- refers to combined Smad4 z null and Smad4 mz null embryos. E) Quantification of the number of OCT4-positive cells in wild-type, Smad4+/-, and Smad4-/- embryos. F) Quantification of EPI and PrE cell numbers from Smad4+/- and Smad4-/- embryos at E5.5 revealed a specific, significant decrease in epiblast cell number in Smad4 mz null embryos when compared to controls (p<0.05 by Student’s t-test). The difference in VE cell numbers was not significant (p>0.05). G) Quantification of the proximal-distal length wild-type, Smad4+/-, and Smad4-/- embryos at E5.5. H) Quantification of the proximal-distal length of the EPI of wild-type, Smad4+/-, and Smad4-/- embryos at E5.5. I) Quantification of the proximal-distal length of the EPI as a percentage of total length of wild-type, Smad4+/-, and Smad4-/- embryos at E5.5. J) Quantification of the proximal-distal length of the ExE of wild-type, Smad4+/-, and Smad4-/- embryos at E5.5. K) Quantification of the proximal-distal length of the ExE as a percentage of total length of wild-type, Smad4+/-, and Smad4-/- embryos at E5.5. L) Quantification of the proportion of Smad4+/- and Smad4-/- embryos with a proamniotic cavity at E5.5. Comparisons in B,C, F were assessed by Student’s t-test. Comparisons in E, G-K were assessed by analysis of variance (ANOVA) with Tukey's post-hoc test. 76 Figure 2.5. Inhibition of FGF signaling partially rescues epiblast cavitation in E5.5 Smad4 null embryos. A) Wild-type and Smad4 z null embryos collected at E5.5 and cultured for 6 hours after dissection in control media or media containing FGFR/MEK inhibitors (see Methods), then stained by immunofluorescence for OCT4 and phosphorylated ERK (pERK). Dashed line in enlargement notates the proamniotic cavity. B) Quantification of the proportion of treated and untreated Smad4-/- embryos with a proamniotic cavity at E5.5. C) Quantification of proximal-distal length of the EPI, D) proximal-distal length of the EPI as a proportion of total length, E) total proximal- distal length, and F) OCT4-positive cell number in treated and untreated E5.5 Smad4-/- embryos. 77 Figure 2.6. Possible model of FGF inhibition rescuing rosette formation but not embryo growth in Smad4-null embryos. A) In wild-type embryos, SMAD4 in the visceral endoderm promotes lumen formation in the epiblast by regulating a pro- apoptotic signal. Separately, SMAD4 also inhibits ERK phosphorylation in the ExE, which allows for polarization and rosette formation in the epiblast. B) In Smad4-null embryos, pERK is upregulated, causing an ectopic increase in pERK and preventing epiblast polarization. The pro-apoptotic signal is also lost. The combination of these two factors lead to epiblast disorganization and a failure to cavitate. C) Treatment with FGF inhibitors prevents upregulation of pERK in Smad4-null embryos. Repressed pERK levels allow epiblast polarization to proceed, resulting in a small proamniotic cavity. 78 Supplemental Figure 2.1. SMAD1/5/9 phosphorylation in post-implantation embryos can be modulated. A) Immunofluorescence for GATA6 and SMAD1/5/9 phosphorylation (pSMAD) in wild-type E4.25, E5.0, and E5.75 embryos. 79 Supplemental Figure 2.1 (cont’d). B) Immunofluorescence for GATA6 and pSMAD in wild-type E5.5 embryos after 6 hours of culture in control media or BMP inhibitor treatments. Note the lack of pSMAD with 0.25 µM LDN-193189 treatment. 1 µM LDN-193189 resulted in severe toxicity to the treated embryos. Graph represents quantification of the proportion of embryos displaying unique SMAD1/5/9 phosphorylation patterns. C) Immunofluorescence for GATA6 and pSMAD in wild-type E5.5 embryos after 6 hours of culture in indicated concentrations of exogenous BMP4 treatment. pSMAD showed a dose-dependent increase as BMP4 concentration increased. Graph represents quantification of proportion of embryos displaying unique SMAD1/5/9 phosphorylation patterns. “Proximal VE only” and “Proximal VE and DVE” refer to pSMAD signal in some cells in those tissues, while “Entire VE” refers to pSMAD signal in every observed VE cell. pSMAD signal in proximal VE is indicated by white arrowheads. pSMAD signal in DVE is indicated by yellow arrowheads. pSMAD signal in proximal EPI is indicated by red arrowhead. 80 Supplemental Figure 2.2. BMP pathway genes are upregulated after implantation. A) Single-cell RNA-seq data generated by Nowotschin et al. reclustered using Seurat and identified by cell type. B) BMP ligand genes enriched in identified cell types and stages. Genes with p-adj<0.01 and average log2 fold change <0.25 were considered cluster enriched (*). 81 Supplemental Figure 2.3. Maternal and zygotic Smad4 and Bmp4 are dispensable for blastocyst formation and preimplantation cell fate specification. A) Breeding scheme for the generation of Smad4 maternal-zygotic (mz) null embryos using the Zp3Cre allele described in de Vries et al, 2000. The same strategy was applied to generate Bmp4 mz null embryos. B) Level of Smad4 mRNA detected in wild-type and Zp3Cre; Smad4fl/fl oocytes by qPCR, normalized to Actin mRNA level. 82 Supplemental Figure 2.3 (cont’d). C) Quantification of the ratio of EPI and PrE cells in the inner cell mass of embryos from D. D) Immunofluorescence for SOX17 and NANOG in dissected E4.5 Smad4 m null and Smad4 mz null embryos. E) Quantification of the number of cells occupying the inner cell mass or trophectoderm and total cell number in embryos from D. F) Immunofluorescence for SOX17 and NANOG in wild type, Bmp4 zygotic (z) null and Bmp4 mz null embryos cultured in KSOM +PVA -BSA (see Methods) from E1.5 to E3.75. G) Quantification of EPI and PrE cell numbers from F did not reveal any significant difference in ICM composition between Bmp4 mz null embryos and controls. “Mixed” indicates co-expression of SOX17 and NANOG. Pairwise comparisons were evaluated by Student’s t-test. 83 Supplemental Figure 2.4. Maternal and zygotic Smad4 are dispensible for OCT4 expression in preimplantation mouse embryos. 84 Supplemental Figure 2.4 (cont’d). A) Immunofluorescence for OCT4 and CDH1 in E2.75 Smad4 m null and Smad4 mz null embryos. B) Quantification of the number of OCT4-positive cells as a percentage of total cells in embryos from A. C) Immunofluorescence for OCT4 and CDH1 in E3.75 Smad4 m null and Smad4 mz null embryos. D) Quantification of the number of OCT4- positive cells as a percentage of total cells in embryos from B. E) Immunofluorescence for OCT4 and CDH1 in E4.25 Smad4 m null and Smad4 mz null embryos. F) Quantification of the number of OCT4-positive cells as a percentage of total cells in embryos from E. All comparisons were assessed by analysis of variance (ANOVA). 85 Supplemental Figure 2.5. Additional data for the effect of BMP4 treatment on preimplantation cell fate. A) Quantification of total cell numbers for embryos from Fig. 2.3A. B) Quantification of TE cell numbers for embryos from Fig. 2.3A. C) Percentage of ICM cells with EPI, PE, mixed, and no expression from Fig. 2.6B represented as individual data points. EPI=NANOG-positive only, PE=SOX17-positive only, mixed=NANOG/SOX17 double-positive, no expression=NANOG/SOX17 double- negative. All comparisons were assessed by analysis of variance (ANOVA) with Tukey's post-hoc test. 86 Supplemental Figure 2.6. Bmp4 is dispensable for embryo growth and organization at E5.5. A) Correlation between the number of OCT4+ cells and proximal-distal length of the EPI in wild-type, Smad4+/-, and Smad4-/- embryos at E5.5 from Fig. 2.4 and Fig. 2.5. Correlation assessed by Pearson’s coefficient. B) Immunofluorescence for GATA6 and CDX2 as respective markers of visceral endoderm and extra-embryonic ectoderm in E5.5 wild-type, Bmp4+/-, and Bmp4-/- embryos. C) Quantification of the proximal-distal length of embryos from B. D) Quantification of the proximal-distal length of the EPI in embryos from B. E) Quantification of the proximal- distal length of the EPI as a percentage of total length in embryos from B. 87 Supplemental Figure 2.7. Defects in Smad4-null embryos are more widespread across cell types at E6.5. A) E6.5 Smad4 mz null and Bmp4 z null embryos stained by immunofluorescence for CDX2 and GATA6. Epiblast length was inferred from the unstained cells distal to the CDX2-positive compartment in these embryos. B) Quantification of the proximal-distal length of wild-type, Smad4 m null, and Smad4 mz null embryos at E6.5. C) Quantification of the proximal-distal length of the EPI of wild- type, Smad4 m null, and Smad4 mz null embryos at E6.5. D) Quantification of the proximal-distal length of the EPI as a percentage of total length of wild-type, Smad4 m null, and Smad4 mz null embryos at E6.5. E) Quantification of the proximal-distal length of wild-type, Bmp4+/-, and Bmp4-/- embryos at E6.5. F) Quantification of the proximal- distal length of the EPI in wild-type, Bmp4+/-, and Bmp4-/- embryos at E6.5. G) Quantification of the proximal-distal length of the EPI as a percentage of total length in wild-type, Bmp4+/-, and Bmp4-/- embryos at E6.5. All comparisons were assessed by analysis of variance (ANOVA) with Tukey's post-hoc test. 88 Supplemental Figure 2.8. ERK phosphorylation is responsive to modulation of FGF signaling. A) Wild-type embryos collected at E5.5 and cultured for 6 hours in control media, 1 µg/mL FGF4 + heparin, or media containing FGFR/MEK inhibitors (see Methods), then stained by immunofluorescence for OCT4 and phosphorylated ERK (pERK). Note decreased pERK in inhibitor-treated embryos and increased pERK in FGF4-treated embryos. B) Quantification of proximal-distal length of all embryos from Figure 2.5A and Supplemental Figure 2.8A. C) Quantification of proximal-distal length of the EPI in all embryos from Figure 2.5A and Supplemental Figure 2.8A. D) Quantification of proximal-distal length of the EPI as a percentage of total length in all embryos from Figure 2.5A and Supplemental Figure 2.8A. All comparisons were assessed by analysis of variance (ANOVA) with Tukey's post-hoc test. 89 CHAPTER 3. ESSENTIAL ROLES FOR HIPPO SIGNALING IN MAMMALIAN REPRODUCTION Robin E. Kruger1,2, Farina Aziz1, and Amy Ralston2,3 1) Cell and Molecular Biology Ph.D. Program, Michigan State University, East Lansing, MI, 48824, USA 2) Reproductive and Developmental Sciences Training Program, Michigan State University, East Lansing, MI, 48824, USA 3) Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA This chapter is a completed manuscript that will be submitted for publication in Reproduction. This study was supported by the National Institutes of Health R35 GM131759 and T32 HD087166. 90 Section 3.1. Abstract Hippo signaling, a highly conserved molecular signaling pathway with well-known functions in development, has long been known to be essential for mammalian fertility. In this review, we discuss recent advances which have uncovered the functions of Hippo signaling in key reproductive organs such as the testes, ovaries, uterus, and placenta. We also discuss human reproductive disorders whose etiology may be related to dysregulation of Hippo signaling and possible therapies which have been proposed to correct this dysregulation. Finally, we highlight current gaps in knowledge where further studies of Hippo signaling may uncover new insights into molecular regulation of reproduction and fertility. Section 3.2. Introduction to the Hippo signaling pathway The Hippo signaling pathway is a highly conserved pathway which is known to play essential roles in both development and adult homeostasis. The biochemical interactions of this pathway have been well-described in several model systems (M. Fu et al., 2022). The pathway is so named because loss-of-function mutations caused massive organ overgrowth in fruit flies, causing a bloated appearance reminiscent of a hippopotamus (Harvey et al., 2003). Subsequent studies using biochemical and genetic approaches uncovered core components of the Hippo pathway, including inputs and outputs. Unlike many signaling pathways, Hippo signaling relies on a large variety of upstream signals to initiate its activity. In addition to extracellular ligands, initiation of this pathway is often regulated by mechanical cues from changes in the extra-cellular matrix 91 or cytoskeleton. Hippo signaling may also be modulated by soluble factors such as steroids, or in response to stress (reviewed in Fu et al., 2022). The main output of Hippo signaling is modulation of the transcriptional activity of homologous transcription factors YAP1 and WWTR1. In the absence of Hippo signaling, these proteins complex with other transcription factors (particularly TEAD factors) to bind DNA and effect changes in gene expression. Active Hippo signaling initiates a phosphorylation cascade in which upstream kinases MST1/2 phosphorylate and activate downstream kinases LATS1/2. Activated LATS proteins form a complex with the scaffolding protein Angiomotin (AMOT), which facilitates their ability to phosphorylate YAP1 and WWTR1. Phosphorylation prevents YAP1/WWTR1 from entering the nucleus, and instead targets these proteins for degradation. In this way, active Hippo signaling prevents the transcriptional activity of YAP1/WWTR1 (see Fig. 1.2). Several excellent review articles have described Hippo roles in homeostasis, disease, development, and regeneration, in model organisms and humans (Davis & Tapon, 2019; M. Fu et al., 2022; Misra & Irvine, 2018; Moya & Halder, 2016; Zheng & Pan, 2019). Yet, the roles of Hippo signaling in reproduction have not been comprehensively reviewed. Relying heavily on studies in mice, we summarize known roles for Hippo signaling in reproduction, beginning with the adult testis, ovary, and uterus, which then sets the stage for understanding Hippo signaling during early development and formation of the extraembryonic tissues. 92 Section 3.3. Hippo signaling in male reproduction Male reproduction processes and cell types In male mammalian reproduction, Hippo signaling is greatly involved in the regulation of spermatogenesis. Spermatogenesis takes place within the seminiferous tubules of the testes, where immature, self-renewing spermatogonial stem cells (SSCs) reside on the inner edge of the tubule. These germ cells gradually mature as they move inward toward the tubule lumen. SSCs first differentiate into spermatogonia, which can divide by mitosis to produce meiotically-capable spermatocytes. After completing meiosis, haploid spermatocytes differentiate further into spermatids, where they undergo the morphological changes to become mature sperm in the process of spermiogenesis. During spermiogenesis, the spermatids develop an acrosomal head, intermediate mid- piece, and flagellar tail. Finally, the mature spermatid enters the lumen of the seminiferous tubule as a fully-developed spermatozoon. Spermatozoa travel from the seminiferous tubule to the epididymis, where they undergo final maturation before ejaculation. The somatic cells of the testis, Sertoli cells, Leydig cells, and structural interstitial cells, are essential to the process of spermatogenesis. Leydig cells function primarily to produce testosterone, which among several other targets is received by Sertoli cells. Sertoli cells act as “nurse” cells for the developing germ cells within the seminiferous tubules, providing nutrients, structural support, and aiding in waste removal. Tight junctions between Sertoli cells help form the blood-testes barrier, providing a favorable microenvironment for spermatogenesis. Finally, Sertoli cells produce anti-Mullerian 93 hormone and inhibin, both key molecules in supporting male sexual development. Proper function of Sertoli and Leydig cells is essential for spermatogenesis and male fertility. Hippo signaling maintains testicular somatic cell identity and promotes spermatogenesis non-cell-autonomously Hippo signaling is essential to male fertility, as demonstrated by the decreased fertility in Hippo genetic knockout models in mice (Hossain et al., 2007; St John et al., 1999). Several knockouts, including those of Lats1, Lats2, and Yap1;Wwtr1 report decreased testis size (Abou Nader et al., 2022; Levasseur et al., 2017; St John et al., 1999). However, although YAP1 and WWTR1 protein are expressed and localized to the nucleus in germ cells after puberty (Levasseur et al., 2017), the direct, cell-autonomous effect of Hippo signaling on spermatogenesis appears to be limited. Conditional knockout of Yap1 in mouse germ cells had no apparent effect on the expression of germ cell markers, spermatogonial stem cell formation, or sperm count in vivo (Abou Nader et al., 2019). Neither knockdown of fat, expanded, hippo, salvador, or warts, nor overexpression of yorkie, had any discernible effect on germ cell cyst formation in Drosophila (S. Sun et al., 2008). However, in sheep, Mst2, Lats1/2, and Yap1 mRNA and protein are all strongly upregulated in mature, ejaculated sperm as opposed to immature spermatozoa recovered from the cauda epididymis, suggesting that Hippo signaling may play a role in adult sperm function even if it is not required cell- autonomously in spermatogenesis (G.-M. Zhang et al., 2019). 94 Although Hippo signaling is apparently dispensable in germ cells, it is essential to maintain the cell identities of Sertoli and Leydig cells. Conditional double knockout of Lats1 and Lats2 in murine testicular somatic cells (using a Nr5a1-Cre strain, which drives Cre expression in a common progenitor cell population for Sertoli cells and fetal Leydig cells as well as adrenal precursors), led to smaller and disorganized testes as early as embryonic day E14.5 (Abou Nader et al., 2022). These knockouts showed decreased expression of many Sertoli cell markers including Sox9. More strikingly, testes in these animals showed severe dysgenesis, marked by a progressive increase in spindle-shaped, fibroblast-like cells and corresponding decrease in Sertoli and Leydig cells (Abou Nader et al., 2022). Further analysis revealed a loss of polarity markers in Sertoli cells, suggesting that without Lats1/2 these cells undergo an epithelial-to- mesenchymal transition resulting in fibrosis of the testis interstitium (Abou Nader et al., 2022). On the other hand, conditional double knockout of Yap1 and Wwtr1 in Sertoli cells decreased expression of male-specific genes such as Dhh, Dmrt1, Sox9, and Wt1 at pre-pubertal stages (Levasseur et al., 2017). The fact that loss of Lats1/2 resulted in a more detrimental phenotype than loss of Yap/Wwtr is consistent with a requirement for active Hippo signaling in Sertoli cell differentiation. Similar results were reported with loss of cell identity and transdifferentiation in Leydig cells in the Lats1/2 conditional knockout model, showing that Hippo signaling is necessary for maintenance of cell identity in both of these cell types. It is, of course, possible that the two cell identities are not independent from one another; for example, if loss of Hippo signaling prevented Leydig cell differentiation, decreased 95 steroidogenesis could indirectly compromise Sertoli cell identity. However, knockdown of Yap1/Wwtr1 in Sertoli cells did not impact testosterone levels, suggesting that YAP1/WWTR1 transcriptional activity in Sertoli cells does not compromise Leydig cell function (Levasseur et al., 2017). It is important to note that the loss of cell identity in Sertoli and Leydig cells does not result in sex reversal, as neither Yap1/Wwtr1 knockout or Lats1/2 knockout resulted in upregulation of granulosa cell genes in vivo (Abou Nader et al., 2022; Levasseur et al., 2017). Altogether, these data suggest that active Hippo signaling is required for maintenance of somatic testicular cell types. The requirement for Hippo signaling in somatic cell identity leads to an essential, non- cell-autonomous role in spermatogenesis. This role was first observed in Drosophila, where deletion of expanded (a hippo co-factor) in germ cells themselves did not have an effect on germ cell cyst formation; however, expanded deletion in the surrounding somatic cells led to overproliferation of spermatogonia (S. Sun et al., 2008). Later, several studies showed that disruption of Hippo signaling in Sertoli cells disrupts spermatogenesis (Abou Nader et al., 2022; Ni et al., 2019; S. Sen Sharma & Majumdar, 2017). Conditional double knockout of Lats1 and Lats2 in mouse Sertoli cells results in small, disorganized testes with very few observable seminiferous tubules, and germ cells in this model were mostly apoptotic by E17.5 (Abou Nader et al., 2022). Similarly, knockout of Yap1 and Wwtr1 in Sertoli cells does not impair early spermatogenesis (Levasseur et al., 2017). This suggests that active Hippo signaling, or silenced YAP1/WWRT1 activity, is specifically required in Sertoli cells for early spermatogenesis. 96 The specific transcriptional targets of YAP1/WWTR1 and exact mechanisms by which Hippo signaling regulates Sertoli cell fate remain to be identified. One study suggested that Hippo signaling may be required for stabilization of cilia, as kidney cells with increased nuclear localization of YAP1 also showed increased expression of AURKA, a cilia disassociation protein (W. H. Shi et al., 2023). In support of this hypothesis, Wwtr1 knockout mice which survive to birth also display cilia malformations in renal cells (Hossain et al., 2007). This possibility is intriguing as patients with autosomal dominant polycystic kidney disease, which affects cilia formation, also commonly display male infertility and sperm motility defects (W. H. Shi et al., 2023). Another study suggested that Yap1 regulates the response of Sertoli cells to cAMP (S. Sen Sharma & Majumdar, 2017). As cAMP signaling in mature Sertoli cells regulates follicle stimulating hormone (FSH) response, this suggests a mechanism for Yap1 regulation of Sertoli cell fate (S. Sen Sharma et al., 2019). It was also shown that FSH treatment could upregulate YAP1 and increase YAP1 nuclear localization in Sertoli cells, suggesting a regulatory feedback loop in which increased FSH induces YAP1 nuclear localization, and then transcriptional targets of YAP1 increase cAMP signaling, which in turn allows for a stronger response to FSH. Excess YAP1 is kept controlled by increased expression of Hippo kinases and increased YAP1 phosphorylation (S. Sen Sharma et al., 2019). This model positions Hippo signaling and YAP1 as master regulators in Sertoli cell maturation and function, making them essential factors for male reproduction. 97 Section 3.4. Hippo signaling in the ovary Ovarian structure and cell types Many processes critical to female mammalian reproduction, including oogenesis, ovulation, and production of female sex hormones such as estrogens and progesterone occur in the ovaries. The ovary is a non-ductal structure composed of two layers of tissue. The internal medulla is a highly vascularized collagenous support matrix for the outer cortex, which itself contains the ovarian functional units known as follicles. Follicles contain all the essential pieces necessary for ovulation. Each follicle has a single germ cell at its center (which becomes a mature oocyte during ovulation) and several surrounding layers of somatic cells including theca cells and granulosa cells. These cells grow and differentiate through several stages of follicular maturity. Follicle formation commences when pre-granulosa cells surround the early germ cell to form the primordial follicle. As the follicle matures, the granulosa cells differentiate into cumulus cells, which provide nutrients and growth factors for the maturing oocyte, and mural granulosa cells, which are responsible for estrogen production. Following ovulation, granulosa cells luteinize and form the bulk of the corpus luteum, a temporary endocrine structure which produces progesterone during early pregnancy. In addition to their hormone-producing capabilities, granulosa cells are also non-vascularized and separated from surrounding cells by a basal lamina, creating a blood-follicle barrier between the oocyte and potential toxins. 98 Theca cells become associated with the follicle at the primary follicle stage, forming the final vascularized layer outside of the basal lamina. The primary role of the theca cells is to produce androgens, which are secreted to the aromatase-expressing granulosa cells to be converted into estrogens. After ovulation, the theca cells also contribute to the corpus luteum as theca lutein cells and continue to play a steroidogenic role until the corpus luteum is resorbed. These somatic cells are essential to the reproductive process; loss of any of these cell types will result in oocyte death, failed ovulation, or pregnancy loss. Hippo signaling is essential for proper ovarian function The Hippo pathway has been shown to play an essential role in female fertility, particularly through regulation of ovarian cell growth, follicle maturation, and key ovarian functions such as ovulation and steroidogenesis. The first evidence of a Hippo requirement in female reproduction was produced by St John et al., 1999, who showed that 60% of female mice with a full-body knockout of Lats1 were completely sterile and did not progress through the estrus cycle. Further studies have shown that Hippo signaling activity through LATS1/2 is necessary in granulosa cells for key ovarian processes such as follicle maturation (L. L. Hu et al., 2019; St John et al., 1999; S. Sun et al., 2015; Ye et al., 2017), ovulation (Ji et al., 2017; Plewes et al., 2019; T. Sun & Diaz, 2019), steroidogenesis (M. Fu et al., 2022; Lv et al., 2020; Plewes et al., 2019; Tsoi et al., 2019), maintenance of granulosa cell fate (Lv et al., 2020; Tsoi et al., 2019), and granulosa cell proliferation (M. Fu et al., 2022; Lv et al., 2020; Plewes et al., 2019; T. Sun & Diaz, 2019). Similarly, changes in Hippo signaling have been associated with 99 defects in ovarian function such as polycystic ovarian syndrome (Cheng et al., 2015; Jiang et al., 2017; Kawamura et al., 2013; T. Li et al., 2012; Maas et al., 2018), primary ovarian insufficiency (Ye et al., 2017), and simple age-related ovarian functional decline (J. Li et al., 2015; Xiang et al., 2015). Further information may be found in several recent reviews on this topic (Clark et al., 2022; Hsueh et al., 2015; Maas et al., 2018; Xia & Du, 2022; M. Zhu et al., 2023). A point we want to emphasize is that collective evidence supports a model in which a balance of Hippo signaling activity is needed for proper ovarian function. Studies show that both YAP1 inactivation (by Yap1 knockdown) and YAP1 hyperactivation (by Lats1 knockout) both resulted in decreased follicle formation (L. L. Hu et al., 2019; St John et al., 1999; S. Sun et al., 2015; Ye et al., 2017). Similar studies have reported ovulation defects and decreased steroidogenesis both when Hippo signaling is overactive and when it is underactive (D. Fu et al., 2014; Lv et al., 2020; Plewes et al., 2019; Tsoi et al., 2019). Furthermore, a need for balanced Hippo signaling in granulosa cells has been demonstrated, which is important because studies have demonstrated that Hippo signaling in the ovary is primarily required in granulosa cells rather than in the oocyte itself (Abbassi et al., 2016; Yu et al., 2016). Too little Hippo activity in granulosa cells, as in the loss of Lats1/2, causes them to transdifferentiate into a variety of cell types, leading to loss of steroidogenesis function and disruption of the follicular structure (Tsoi et al., 2019). This in turn leads to a deficiency in follicle abundance, ovulation defects, and a severe decrease in fertility. Interestingly, all previous studies of Lats KO in the ovary reported growth of ovarian tumor- or cyst-like structures (St John et al., 1999; S. 100 Sun et al., 2015). It is possible that these phenomena may have actually been transdifferentating cell populations which had not been fully characterized, though more work in this area is needed to test this hypothesis. Too much Hippo activity, as in the case of Yap1 knockdown, prevents proper maturation and proliferation of granulosa cells to result in a decrease in mature follicles and compromised fertility. Further research is required to elucidate how this balance of Hippo activity is maintained. Model for Differential Regulation of Hippo in Follicle Maturation Altogether, the evidence suggests that there is an essential but dynamic role for Hippo signaling in follicle maturation. The requirements for Hippo activity change over the lifespan of a follicle. Early follicles have higher expression of Hippo kinases (Mst1/2, Lats1/2), decreased nuclear localization of YAP1, and more Hippo activity in early granulosa cells which helps to prevent premature follicle activation (De Roo et al., 2020; L. L. Hu et al., 2019; Lv et al., 2019; S. Sun et al., 2015; Xiang et al., 2015). Active Hippo signaling also prevents unrestricted granulosa cell proliferation and maintains granulosa cell identity (S. Sun et al., 2015; Tsoi et al., 2019). However, as follicles begin to mature they downregulate Hippo signaling to allow YAP1 to induce granulosa cell proliferation and maturation (J. Li et al., 2015). As further evidence of a stage- dependent role for Hippo signaling in granulosa cells, functional studies of YAP1 were conducted in cell-specific Yap1 knockouts in both proliferating and mature granulosa cells (Lv et al., 2019). In mature cells, YAP1 localized in the cytoplasm and loss of Yap1 in these cells produced no functional or morphological differences. In proliferating granulosa cells, however, loss of Yap1 led to small ovaries and significantly fewer 101 follicles. It is possible that this differential regulation of YAP1 activity is partially regulated by mechanical signals, as the stiffer ECM of the outer ovarian cortex may induce Hippo activity in primordial follicles. As the maturing follicles move inward to the less stiff ovarian medulla, the mechanical signal may relax to silence Hippo signaling and allow YAP1 nuclear entry (Hsueh et al., 2015). In support of this hypothesis, strips of ovarian tissue have been shown to have less pYAP and more advanced follicles near the edges of the strip, where the ECM has been disrupted (Grosbois & Demeestere, 2018). After ovulation, Hippo turns back on (possibly in response to endocrine signals such as LH or hCG) to limit proliferation and induce corpus luteum formation (Ji et al., 2017; T. Sun & Diaz, 2019). Therefore, Hippo signaling serves as a possible master regulator for the changes in granulosa cells through the whole lifetime of a follicle. Remaining questions about Hippo signaling in the ovary Despite the abundant reported studies, there remain some unanswered questions which are needed to put together a full picture of Hippo signaling in the ovary. Though the requirement for and effects of Hippo signaling in granulosa cells are clear, it is still unknown what regulates Hippo signaling activity in granulosa cells. It would be especially interesting to discover the mechanism by which granulosa cells modulate Hippo signaling throughout follicle maturation. Similarly, many of the direct genetic targets of YAP1/WWTR1 which regulate granulosa cell fate, proliferation, and follicle maturation have not been reported. 102 Much remains to be discovered on the role of Hippo signaling in the oocyte as well. It is known that both Yap1 and Wwtr1 mRNA are maternally loaded in the oocyte (Xie et al., 2010). However, YAP1 protein localization is strictly cytoplasmic and Yap1 is apparently dispensable for follicle formation, follicular maturation, and ovulation (Abbassi et al., 2016; L. L. Hu et al., 2019; S. Sun et al., 2015; Yu et al., 2016). WWTR1, by contrast, appears to be strongly nuclear in the oocyte (S. Sun et al., 2015). This functional localization of WWTR1 suggests a difference between Yap1 and Wwtr1 in regard to their respective roles in the oocyte. Since maternal Wwtr1 KO embryos develop into adult mice when fertilized by wild-type sperm (although these offspring are subfertile), it appears that its function in the oocyte may be dispensable (Hossain et al., 2007). However, all developmental studies with Wwtr1 have been done in whole-body knockouts; no oocyte-specific knockouts of Wwtr1 have been examined to date. Oocyte-specific knockout models of Lats or other Hippo kinases have not been examined either. Though full-body knockout of Lats1 resulted in high rates of oocyte apoptosis, it is possible that this is caused by loss of granulosa cells (and thereby loss of growth factors and sex hormones essential for oocyte development), rather than any direct impact on the oocyte itself (S. Sun et al., 2015). More cell-type-specific knockout models are essential to determine the direct effect of active Hippo signaling in ovarian germ cells. 103 Section 3.5. Hippo signaling in the uterus Uterine tissues and processes Hippo signaling is known to have a profound effect on the uterus, particularly in the process of decidualization. The uterus is comprised of three tissue layers, with an outer perimetrium, middle, muscular myometrium, and inner endometrium. The endometrium is the site of embryo attachment during pregnancy, and as such, this layer has been the focus of most studies regarding Hippo signaling in reproduction. During early pregnancy, the endometrium undergoes drastic morphological and functional remodeling to accommodate the incoming blastocyst for the duration of the pregnancy; this process is known decidualization. Decidualization takes place around four days after fertilization in mouse while in human it begins during the secretory phase of the menstrual cycle (Okada et al., 2018). One of the hallmarks of decidualization is the differentiation of elongated, fibroblast-like endometrial stromal cells to rounded, epithelial-like decidual cells. These cells secrete hormones such as prolactin that are fundamental for embryo implantation and development of the invading trophoblasts to form the placenta. Thus, decidualization is critical for the success of a pregnancy and impairment to this process leads to multitude of pregnancy disorders including infertility and miscarriage. Hippo signaling is required for uterine decidualization The components of the Hippo signaling pathway have been implicated in modulating the decidualization process in mammals. Several groups have reported changes in the expression of Hippo pathway members during decidualization. Strakova et al. reported 104 a decrease in the expression of nuclear WWTR1 after inducing in vitro decidualization in human stromal fibroblast cells (Strakova et al., 2010). Increased phosphorylation of LATS1 and WWTR1 have been reported in human endometrial stromal cells cocultured in conditioned media from menstrual stem cells (H. Zhu et al., 2019). Consistent with the upregulation of LATS kinase activity during decidualization, there was decreased expression of WWTR1 in the uterine stroma during the secretory phase of the menstrual cycle in humans (Strakova et al., 2010). Interestingly, the ability of endometrial stromal cells to undergo decidualization was not compromised with the loss of Lats1 and Lats2 which might suggest the dispensability of Hippo signaling in this context (St-Jean et al., 2019). It could also mean that Lats1/2 become dispensable once the decidual cell fate has been established. Taken together, these suggest that Hippo signaling is active during decidualization in uterine stromal cells. The mechanism by which Hippo regulates decidualization is still under investigation. Interestingly, treatment with a commonly used decidualization inducer, cAMP, did not affect WWTR1 protein level in human stromal fibroblast cells, which suggests that the expression of WWTR1 in decidualization is controlled more by hormones (progesterone/PGR signaling) than by cAMP/PKA signaling (Strakova et al., 2010). In contrast to Wwtr1, both mRNA and protein level of Yap1 is upregulated in human decidual tissue compared to the non-pregnant endometrial stromal cells. Consistent with that, an increase in both YAP1 and TEAD1 mRNA and protein levels has been reported during decidualization in cultured human endometrial cells (H. Chen et al., 2017). Knockdown of Yap1 using shRNA in these cells showed lack of typical decidual 105 morphology along with decreased expression of decidua-specific markers and TEAD1, indicating that YAP1 might regulate decidualization in endometrial stromal cells through TEAD1 (H. Chen et al., 2017). The contrasting role of YAP1 and WWTR1 in this context suggests that these homologous proteins play distinctive roles in regulating decidualization. Hippo disruption may lead to development of endometrial disorders In addition to their role in regulating decidualization, dysregulation of YAP1/WWTR1 levels has been reported to be involved in the development of endometriosis and endometrial fibrosis (Pei et al., 2019, 2022; Song et al., 2016; H. Zhu et al., 2019). YAP1 mRNA and protein level is significantly higher in endometrial stem cells in women with endometriosis compared to women without endometriosis (Song et al., 2016). YAP1- TEAD1 complex causes increased proliferation and decreased autophagy of endometrial stromal cells from endometriosis via interaction with the negative regulator of autophagy, mTOR (Pei et al., 2019, 2022). This suggests that disrupted Hippo signaling in endometriosis leads to YAP1 overactivation and overproliferation of endometrial cells. Moreover, phosphorylated WWTR1 causes downregulation of the expression of fibrotic genes in endometrial fibrosis, suggesting that disruption of Hippo signaling may also be a driving factor in the etiology of this disease (H. Zhu et al., 2019). These data indicate a connection between the Hippo pathway and the development of endometrial disorders, though more research is needed to thoroughly define this relationship. Taken together, published studies indicate that Hippo signaling 106 is involved in the dynamic changes occurring in the uterine endometrium during the reproductive cycle and is essential to establishing a healthy pregnancy. Section 3.6. Hippo signaling in the trophoblast and placenta lineage Cell types and mechanisms of placental development After a pregnancy is established, Hippo signaling plays a vital role in the specification and development of the placental lineage. The placenta is a large reproductive organ which acts as an exchange barrier between the mother and the developing fetus. It is comprised of both maternal and fetal components, with the key functional cell type being the trophoblast cells (Rossant & Cross, 2001). The maternal components of the placenta arise from the uterine decidual cells and maternal vasculature, while chorion- derived epithelial cells and allantois-derived vascular cells generate the fetal vascular compartment (Rossant & Cross, 2001). Placentation is characterized by invasion of multipotent, fetal trophoblasts into the maternal endometrium and establishing direct contact between trophoblasts and maternal blood (Cross et al., 1994; Simmons et al., 2007). Placental villi, the functional unit of the human placenta, are generated when rows of fetal trophectoderm-derived cytotrophoblast (CTB) cells break though the expanding primitive syncytium. As this process continues, the villous CTBs continue to expand and differentiate into two principal trophoblast cell types: syncytiotrophoblasts (STBs) and extravillous trophoblasts (EVTs). STBs are responsible for secreting crucial pregnancy hormones, such as human chorionic gonadotrophin (hCG) to maintain the corpus luteum after ovulation (Costa, 2016). The invasive EVT lineage is involved in remodeling the uterine vasculature to enhance maternal blood supply to support the 107 growing fetus (Knöfler et al., 2019). EVTs also express nonclassical human leukocyte antigen-G to promote immune tolerance to the fetus (PrabhuDas et al., 2015). Any change in this developmental program of the placenta can cause substantial effects on the fetus and its ability to cope with the in utero environment (Perez-Garcia et al., 2018; Rossant & Cross, 2001). Therefore, proper development of the trophoblast lineage (i.e., the synchrony between self-renewal and differentiation ability of the trophoblast progenitors) is crucial for successful placentation. Hippo signaling regulates trophoblast specification and placentation The Hippo pathway is a critical regulator of many steps of placental development, beginning with an essential role in the differentiation of placental progenitors. As discussed in Chapter 1, the Hippo pathway is well known to regulate a global gene expression program to establish TE lineage in preimplantation mouse embryos (Nishioka et al., 2008; Ralston et al., 2010; Yagi et al., 2007). Interestingly, YAP1 and TEAD4 continues to support trophectoderm lineage development even after implantation. In early post implantation mouse embryo, it has been demonstrated that conditional deletion of Tead4 in trophoblast stem cell-like progenitor cells leads to impaired placentation together with the repression of trophoblast stem state specific genes, such as Esrrb, Tfap2C and Gata3 (Saha et al., 2020). Moreover, both mouse and human TSCs undergo differentiation and display loss of stem state colony morphology with Tead4-knockdown, signifying a requirement for TEAD4 to maintain self-renewal ability and stemness in mouse TSCs (Saha et al., 2020). Another study showed that depletion of WWTR1 in CTB-derived human TSCs led to loss of stem-state 108 colony morphology together with a strong reduction in cell proliferation (Ray et al., 2022). Thus, the transcriptional activity of Hippo effectors is critical to maintain the progenitor state of early trophoblasts. After implantation, upregulation of Hippo signaling activity and silencing of YAP1/WWTR1 is essential to allow differentiation of more mature trophoblast lineages. YAP1 protein expression level is higher in first-trimester human placentae compared to those of the full-term group, showing that Yap1 is downregulated after placentation is complete. (Sudol et al., 1995; M. Sun et al., 2018). Furthermore, loss of TEAD4 in human TSCs promotes expression of differentiated STB specific genes such as CGA, CGB isoforms and PSGs. Similarly, YAP1-knockout trophoblastic JEG3 carcinoma cells (representing early pregnancy trophoblasts with TSC-like properties) showed decreased expression of stemness/proliferation associated genes such as cyclin A (Ccna), Cdk6, Cyr61, Itga6 and Tead4 whereas overexpression of YAP1 led to induced expression of these genes (Meinhardt et al., 2020). Consistently, overexpression of YAP1 caused downregulation of STB markers such as Cgβ, Gcm1, Ovol1, Endou and Gdf15. As a proposed mechanism to explain YAP1-mediated inhibition of STB differentiation, the YAP1/TEAD4 complex has been shown to interact with the genomic regions of cell cycle regulators such as Ccna2 and Cdk6 as well as the promoter regions of the STB specific genes such as CGB5 and CGB7 (Meinhardt et al., 2020; Saha et al., 2020). Depletion of Wwtr1 in human TSCs also resulted in strong upregulation of STB-specific genes and extended culture of this cells led to a STB fate in a culture medium that otherwise supported the maintenance of TSC stem state (Ray et al., 2022). This shows 109 that YAP1/TEAD4 and WWTR1 are key factors in promoting cell proliferation and in maintaining a self-renewing stem state in trophoblast cells, which must be downregulated for trophoblast differentiation to STBs. However, YAP1 and WWTR1 may not necessarily work together to complete these roles. Overexpression of YAP1 in villous CTBs led to decreased WWTR1 while YAP1 knockout led to increased WWTR1 expression (Meinhardt et al., 2020). This YAP1 mediated alteration of WWTR1 expression suggests a regulatory link between them and suggests that these proteins may work via independent mechanisms to regulate trophoblast differentiation. Taken together, the roles of YAP1, WWTR1 and TEAD4 maintain a balance between CTB self- renewal and differentiation establishes Hippo as a critical regulator of human trophoblast development. In addition to the roles in maintenance, self-renewal, and differentiation of trophoblasts, downstream mediators of the Hippo pathway have been implicated as critical regulators of placentation and embryo implantation (Bai et al., 2018; Kusama et al., 2016; Meinhardt et al., 2020; Ray et al., 2022; Saha et al., 2020). During placentation, YAP1/WWTR1 are essential for proliferation and invasion of trophoblasts. In support of this, TEAD4 and YAP1 shows strong overlapping expression in the nuclei of proliferative villous CTBs in the first trimester placenta, whereas WWTR1 is abundant in the nuclei of EVTs (Meinhardt et al., 2020; Ray et al., 2022; Saha et al., 2020). Overexpression of YAP1 protein in human trophoblast cell line HTR-8/SVneo led to increased invasive ability of the cells whereas knockdown of YAP1 yielded the opposite outcome (M. Sun et al., 2018). Together, this suggests that YAP1 enhances human trophoblast invasion. 110 Hippo signaling is also involved during conceptus attachment into the bovine endometrium. Interferon tau (IFNT) is the pregnancy recognition protein in ruminants and is secreted by trophoblast cells of peri-implantation conceptuses (Imakawa et al., 1987). IFNT exerts an anti-luteolytic activity to maintain the integrity of corpus luteum and secretion of progesterone, and thus essential for maintenance of pregnancy (Antoniazzi et al., 2013; Roberts et al., 1992). Bovine IFNT level peaks during peri- attachment period and the level decreases soon after the initiation of the conceptus attachment to the endometrium (Hansen et al., 1999; Kusama et al., 2016). Notably, this downregulation of IFNT in bovine conceptuses has been associated with a decrease in Yap1 mRNA expression level together with an increase in phosphorylated YAP1 level (Kusama et al., 2016). Consistent with this, nuclear localization of both TEAD2 and TEAD4 was lower during the conceptus attachment period compared to the peri- attachment period. Taken together, these studies suggest that activation of Hippo causes downregulation of IFNT to promote bovine conceptus attachment. Additionally, YAP1-TEAD complex downregulates bovine OVOL2, a transcription factor essential for mesenchymal-to-epithelial transition, concurrently with conceptus implantation. This downregulation of OVOL2 subsequently induces trophectoderm epithelial-to- mesenchymal progression and aids in the non-invasive type of trophoblast implantation in the endometrium (Bai et al., 2018). Therefore, Hippo signaling is crucial for early gestational processes beginning with trophoblast invasion to conceptus attachment among mammals. 111 Hippo dysregulation is implicated in complications of pregnancy The Hippo pathway has been implicated in several pregnancy associated complications, such as preeclampsia, intrauterine growth retardation, recurrent pregnancy loss, and preterm birth (Ray et al., 2022; Saha et al., 2020; St John et al., 1999; M. Sun et al., 2018). Abnormal placentation due to shallow trophoblast invasion is central to the pathogenesis of preeclampsia during pregnancy (Fisher, 2015; Goldman-Wohl & Yagel, 2002). It was shown that both YAP1 mRNA and protein expression level is decreased in preeclamptic human placentae compared to the control first trimester and full-term group (M. Sun et al., 2018). Since invasion ability of trophoblasts depends on the enhanced YAP1 expression level, the decreased YAP1 level in preeclamptic placentas might be associated with loss of invasion and thus leading to a disease state (M. Sun et al., 2018). This YAP1-mediated trophoblast invasion defect could be related to microRNA-21 (miR-21) mediated modulation of Hippo signaling. miR-21 is upregulated in preeclamptic human placentae compared to the matched normal placentae (M. Hu et al., 2022; Zhou et al., 2020). Transfection of human trophoblast line HTR-8/SV neo with miR-21 mimic resulted in significant inhibition of both the invasion and migration of the cells (M. Hu et al., 2022). It has been shown that PP2A Bβ, the regulatory subunit of the highly conserved serine threonine phosphatase enzyme PP2A, dephosphorylates LATS1 and leads to increased nuclear YAP1 (M. Hu et al., 2022). Consequently, abnormal level of miR-21 during placentation interferes with the PP2A Bβ activity which leads to Hippo pathway activation and subsequent sequestration of YAP1 in the cytoplasm. In addition, loss of TEAD4 has been associated with defective development of placental villi, particularly defective formation of CTB/STB layer, and thus leading to 112 idiopathic recurring pregnancy loss in patients (Saha et al., 2020). Finally, placentae from pregnancies complicated with extreme preterm birth showed drastically low level of WWTR1 expression in CTB layer indicating the dysregulation of Hippo effectors as a factor in poor placentation (Ray et al., 2022). Thus, Hippo pathway members orchestrate an intricate developmental program during early human placentation and improper activation of the pathway may result in poor placentation leading to adverse pregnancy outcomes. Section 3.7. Conclusions In conclusion, Hippo signaling is a critical regulator of many aspects of mammalian reproduction. In particular, the transcriptional activity of YAP1/WWTR1 appears to regulate many of the reproductive processes involving cellular proliferation, such as spermatogenesis, follicle growth, decidualization, and placentation. Similarly, repression of YAP1/WWTR1 through activation of Hippo signaling kinases is crucial to prevent overproliferation and allow differentiation of reproductive cell types. Thus, precise activation and repression of Hippo signaling is essential to promote fertility and successful pregnancy in mammals. Dysregulation of Hippo signaling can lead to many detrimental reproductive disorders, whether from Hippo hyperactivity causing poor placentation in preeclampsia to Hippo hypoactivity leading to overproliferation of uterine stromal cells in endometriosis. The position of Hippo signaling as such a key regulator in these disorders makes it an attractive therapeutic target; however, there are some drawbacks. The fact that most reproductive processes appear to require a balance of Hippo signaling activity, neither too much nor too little, means that strategies inhibiting 113 or stimulating the Hippo pathway itself are likely to cause severe side effects. Similarly, since Hippo signaling regulates similar processes of cell proliferation and differentiation in many other tissues, the therapeutic targeting to the reproductive system would need to be very precise. One way to address this would be to look more deeply into factors upstream and downstream of Hippo signaling in reproductive tissues, many of which have yet to be elucidated. These factors may be more specific to reproduction than the Hippo pathway itself and may present more attractive therapeutic targets. There is still much to be learned about Hippo signaling regulation in reproduction, and this field represents exciting opportunities for further discovery and abundant applications in human health and disease. 114 CHAPTER 4. HIPPO SIGNALING EFFECTORS MAINTAIN BUT DO NOT REGULATE INITIATION OF CELL POLARIZATION IN MOUSE BLASTOMERES Robin E. Kruger and Amy Ralston This study was supported by the National Institutes of Health R35 GM131759 and T32 HD087166. 115 Section 4.1. Abstract Hippo signaling effectors YAP1 and WWTR1 are key regulators of polarization in mammalian morula-stage embryos; however, it is unknown whether these factors regulate the initiation of embryo polarization or the maintenance of polarization. In this study, I examined the localization of apical domain markers aPKCζ and PARD6B in wild- type and Yap1;Wwtr1 maternal-zygotic null embryos at early compaction stages to assess whether Yap1 and Wwtr1 were required for the initiation of blastomere polarization. I found that polarization initiated at wild-type rates in the absence of Yap1 and Wwtr1. This suggests that Hippo effectors are required to maintain polarization in late morula and blastocyst-stage embryos after initial formation of the apical domain at the 8- to -16-cell stage. Section 4.2. Introduction Hippo signaling is a critical regulator of the first cell fate decision in mammalian embryogenesis. Active Hippo signaling in the inner cells of morula-stage embryos induces expression of inner cell mass (ICM) factors such as Sox2, while silenced Hippo in the outer cells allows transcription of key trophectoderm (TE) factors such as Cdx2 and Gata3 (see Chapter 1, Fig. 1.2). One of the key upstream factors governing differential Hippo activity in inside and outside cells of preimplantation embryos is the presence or absence of apical-basal cell polarity. Formation of the apical domain first arises during compaction of blastomeres at the late 8-cell stage (E2.75) (Vinot et al., 2005). The apical domain is characterized by 116 localization of PAR complex (composed of PAR3, PAR6, and aPKC proteins) and p- ERM to the cell membrane (Alarcon, 2010; Plusa et al., 2005; M. Zhu et al., 2017). These proteins help direct the formation of an apical F-actin “cap”, which further assists in blastomere compaction (Maître et al., 2016; Zenker et al., 2018). Recently, keratin proteins have also been suggested to accumulate at the apical domain and help direct F-actin assembly (Lim et al., 2020). Meanwhile, the nascent basolateral domain is characterized by upregulation of E-cadherin and EMK1 and forms adherens junctions between neighboring cells, which also help maintain apicobasal polarity (Shirayoshi et al., 1983; Stephenson et al., 2010; Vinot et al., 2005). Following compaction and polarization, asymmetric cell divisions cause some cells to inherit the apical domain proteins and remain outside cells, while those that do not inherit an apical domain lose their polarity and are directed to the inside of the embryo (Johnson & Ziomek, 1981; Korotkevich et al., 2017; Lim et al., 2020). This leads to transcriptional and morphological differences between inside and outside cells. The presence or absence of an apical domain heavily regulates the activity of Hippo signaling in blastomeres. Single, dissociated blastomeres lose apical domain expression and concomitantly display Hippo signaling activity (Lorthongpanich et al., 2013). Hippo signaling is also active in blastomeres cultured in media containing Ca2+, which prevents polarization and cell-cell contact (Anani et al., 2014). In embryos lacking maternal and zygotic E-cadherin, aPKCζ is ectopically localized to the membrane of inner cells, and in these embryos Hippo signaling is inactive (Stephenson et al., 2010). Finally, if a section of apical domain is added to an initially apolar blastomere, the cell 117 will move to the outside of the embryo and typically adopt a trophectoderm identity (Korotkevich et al., 2017). This shows that cell polarization upstream of Hippo signaling is necessary and sufficient for regulating ICM and TE cell fate at early morula stages. Interestingly, evidence suggests that polarization may also be downstream of Hippo signaling, forming a feedback mechanism to direct the first cell fate decision. At the blastocyst stage, dual loss of Hippo effector proteins YAP1 and WWTR1 leads to a loss of apical aPKC, PAR3, and ZO-1 (Frum et al., 2018). This suggests that Hippo signaling is partially responsible for ensuring the proper apical localization of these proteins, which is critical to embryonic development as loss of either aPKC or Pard6b causes blastomeres to adopt an inside position (Hirate et al., 2015; Plusa et al., 2005). However, it is unknown whether YAP1/WWTR1 are required for the initial localization of these factors at the apical domain or simply for their maintenance at that localization. To test the hypothesis that YAP1/WWTR1 are essential for initiating blastomere polarization, I examined apical domain markers aPKCζ and PARD6B at early morula stages in wild-type and Yap1;Wwtr1 mz null embryos. I found that, surprisingly, polarization appears normal in Yap1;Wwtr1 mz null embryos at the 16-cell stage, suggesting that Hippo signaling regulates the maintenance but not the initial localization of these apical domain factors. 118 Section 4.3. Materials and Methods Mouse Strains and Genotyping All animal research was conducted in accordance with the guidelines of the Michigan State University Institutional Animal Care and Use Committee under approved protocol 202300108. Wild type embryos were obtained from CD-1 mice (Charles River). The following alleles were used in this study and maintained in a CD-1 background: Yap1tm1.1Eno (Xin et al., 2011); Wwtr1tm1.1Eno (Xin et al., 2013); Tg(Zp3-cre)93Knw (De Vries et al., 2000). Null alleles were generated by breeding dams carrying homozygous floxed alleles and the Zp3Cre allele to wild-type CD-1 males. Mouse genotypes were determined by PCR using genomic DNA extracted using the REDExtract-N-Amp kit (Sigma XNAT) according to the manufacturer’s protocol. Embryo genomic DNA was extracted using the same kit scaled to 10 µL total volume. Genomic extracts (1–2 µL) were then subjected to PCR using allele-specific primers (see Table 4.1). Immunofluorescence and Confocal Microscopy Preimplantation embryos (E2.5-E3.0) were fixed with 4% formaldehyde (Polysciences 04018) for 10 minutes, permeabilized with 0.5% Triton X-100 (Sigma Aldrich X100) for 30 minutes, and then blocked with blocking solution (10% Fetal Bovine Serum (HyClone SH30396.02), 0.1% Triton X-100) overnight at 4°C. Embryos were incubated with primary antibody overnight at 4°C. The next day, embryos were washed in blocking solution for 30 minutes, incubated in secondary antibody diluted in blocking solution for 1 hour, washed in blocking solution for 30 minutes, then stained with nuclear stain diluted in block for 10 minutes or overnight. 119 Antibodies used are listed in Table 4.2. Embryos were imaged using an Olympus FluoView FV1000 Confocal Laser Scanning Microscope system with 60X PlanApoN oil (NA 1.42) objective. For each embryo, z-stacks were collected, with 5 mm intervals between optical sections. All embryos were imaged prior to knowledge of their genotypes. Embryo Analysis For each embryo, z-stacks were analyzed using Fiji (ImageJ), which enabled the labeling, based on DNA stain, of all individual cell nuclei. Statistical analysis was performed using GraphPad Prism (v. 9.5.1). Figure images were assembled using Adobe Illustrator. Section 4.4. Results Apical domain formation is first initiated at the 8-cell stage (Vinot et al., 2005). Therefore, I first investigated polarization in wild-type embryos at this stage. Apical aPKCζ was detectable in all 8-cell embryos, consistent with previous reports (Frum et al., 2018; Vinot et al., 2005). However, in most 8- to 10-cell embryos only ~60% of outer blastomeres expressed apical aPKCζ. Furthermore, most embryos still did not express apical PARD6B (Fig. 4.1A). This suggested that although outer blastomere polarization had initiated at the 8-cell stage, it was not yet complete. I therefore performed a time course to determine the first stage where polarization was complete in all embryos. At the 16- to 18-cell stage, I observed that nearly 100% of outer cells expressed apical aPKCζ and PARD6B, a significant upregulation from previous stages (Fig. 4.1B). The 120 percentage of polarized cells remained at nearly 100% after this stage (Fig. 4.1B). From these observations, I concluded that polarization of outer blastomeres (as defined by apical localization of aPKCζ and PARD6B) is normally complete at the 16-cell stage. Having identified the developmental stage at which apical domain formation is normally complete in mouse embryos, I then assessed whether the loss of Hippo effectors YAP1 and WWTR1 would result in decreased or delayed apical domain formation. Yap1 and Wwtr1 have been shown to have partially redundant roles in early mouse embryos (Nishioka et al., 2009), and maternally-derived transcripts of both are present and functional in oocytes and early embryos (Frum et al., 2018; Varelas et al., 2010; Xie et al., 2010; Yu et al., 2016). Therefore, to avoid potential compensation from any source of genetic material, I generated embryos lacking maternal and zygotic Yap1 and Wwtr1 (Yap1/Wwtr1 mz null, see breeding scheme in Fig. 4.2). If my hypothesis that YAP1/WWTR1 are required to initiate blastomere polarization was correct, I expected to see decreased initial apical localization of aPKCζ and PARD6B in Yap1/Wwtr1 mz null embryos. In contrast to this hypothesis, I detected apical localization of aPKCζ and PARD6B in 16-cell Yap1/Wwtr1 mz null embryos at a similar level to wild-type (Fig. 4.3A). The percentage of outer blastomeres which were polarized did not differ between Yap1/Wwtr1 mz null embryos and wild-type or single-knockout controls (Fig. 4.3B). This suggests that initial apical localization of aPKCζ and PARD6B does not depend on Yap1 and Wwtr1. 121 Section 4.5. Discussion Hippo signaling effectors YAP1 and WWTR1 have been shown to be necessary for outer cell polarization in mouse blastocysts (Frum et al., 2018); however, the mechanism of this requirement is still unknown. My data suggest that YAP1 and WWTR1 do not regulate the initial formation of an apical domain at the 8- to 16-cell stages, as marker proteins aPKCζ and PARD6B show normal apical localization in Yap1/Wwtr1 mz null embryos at this stage. Rather, it appears that YAP1 and WWTR1 are required for apical domain maintenance between the early morula and blastocyst stages. It is unknown how YAP1/WWTR1 may be regulating apical domain maintenance in early embryos. YAP1/WWTR1 generally induce molecular changes by binding to TEAD family transcription factors in order to regulate gene expression (Zheng & Pan, 2019). TEAD4 is canonically considered the main transcriptional binding partner for YAP1/WWTR1 in mouse embryos, as Tead4-null embryos largely recapitulate the phenotypes associated with Yap1;Wwtr1 loss-of-function and Lats1/2 gain-of-function (Nishioka et al, 2008; 2009; Frum et al 2018). Interestingly, however, Tead4 z null embryos retain normal apical localization of aPKCζ at late morula stage (exact timepoint not reported), indicating that polarization is maintained properly (Nishioka et al., 2008). This suggests that YAP1/WWTR1 may regulate apical domain maintenance through partnership with proteins other than TEAD4 (Frum et al 2018). 122 Likewise, the exact mechanism by which targets of YAP1/WWTR1 and their binding partner(s) may regulate apical domain maintenance is still unknown. However, there is evidence to suggest that regulation of the actin cytoskeleton may be a good candidate. Actin remodeling is an essential part of blastomere compaction at the late 8-cell stage, a process largely concomitant with cell polarization (M. Zhu & Zernicka-Goetz, 2020). During compaction, F-actin is excluded from cell-cell contacts and accumulates at the cell-free, outer membranes. Increased tension from unequal actomyosin contractility helps drive compaction of blastomeres (Maître et al., 2016). Shortly afterward, polarization initiates and accumulation of Par complex proteins helps remodel the actomyosin network into a characteristic apical ring (Zenker et al., 2018). Interestingly, initial localization of apical domain proteins is unperturbed when actomyosin contractility is decreased by inhibiting myosin II activity, suggesting initiation of polarization may be independent of actin dynamics (M. Zhu et al., 2017). However, polarization is not maintained in embryos where actin nucleation is inhibited (S. C. Sun et al., 2013). Significantly, there is abundant evidence that actin polymerization can regulate Hippo signaling activity in mammalian and fly systems (Hirate et al., 2013, reviewed in Matsui & Lai, 2013; Reddy et al., 2013), and this regulation is not one-sided; in flies, Hippo pathway kinase activation can also downregulate F-actin accumulation (Matsui & Lai, 2013). If F-actin accumulation is essential for the maintenance of blastomere polarization but not its initiation, it is possible that Yap1;Wwtr1 mz null embryos fail to maintain polarity by inappropriate downregulation of F-actin. More studies are needed to determine the validity of this model and define the factors which maintain blastomere polarity downstream of Hippo signaling. 123 Section 4.6. Acknowledgements I thank Drs. Tristan Frum and Jennifer Watts for their technical assistance in this study. 124 Allele Yap1WT, Yap1fl Yap1del Wwtr1WT, Wwtr1fl Wwtr1del Cre Zp3Cre TABLES Forward Primer Reverse Primer ACATGTAGGTCTGCATG CCAGAGGAGG AGGCTGAGACAGGAGG ATCTCTGTGAG ACATGTAGGTCTGCATG CCAGAGGAGG GGCTTGTGACAAAGAAC CTGGGGCTATCTGAG TGGTTGAGACAGCGTGC ACTATGGAG CCCACAGTTAAATGCTT CTCCCAAGACTGGG TGACAAAGAACCTGGGG CTA CTAGGCCACAGAATTGA AAGATCT CGAGATTGAGGGAAGCA GAG AACTGCTAACGTCTCCT GCC GTAGGTGGAAATTCTAG CATCATCC CAGGTTCTTGCGAACCT CAT Table 4.1. Allele-specific primers for PCR genotyping for Chapter 4. Antibody/Stain Source Mouse-anti- aPKCζ Rabbit-anti- PARD6B Goat-anti- mouse IgG Donkey-anti- rabbit IgG Santa Cruz Biotechnology Novus Biologicals Invitrogen Jackson Immuno Research Cell Signaling DRAQ5 Identifiers sc-17781 Dilution 1:100 Fluorophore NBP1-87337 1:100 A-11029 1:400 Alexa488 711-165-152 1:400 Cy3 4084 1:400 Table 4.2. Antibody Table for Chapter 4 125 FIGURES Figure 4.1. Polarization of outer blastomeres is complete at the 16-cell stage. A) Immunofluorescence for aPKCζ and PARD6B in wild-type preimplantation mouse embryos collected at designated timepoints. B) Quantification of the percentage of outside blastomeres in embryos from A which display apical localization of aPKCζ. Statistical analysis was performed using analysis of variance (ANOVA) with Tukey’s post-hoc test. 126 Figure 4.2. Breeding Scheme for Yap1;Wwtr1 maternal-zygotic (mz) null embryos. Yap1;Wwtr1 mz null embryos were generated using the Zp3Cre allele described in De Vries et al., 2000. Yap1;Wwtr1 mz null embryos lacked zygotic expression of both genes and did not have maternally-derived transcripts of either gene deposited in the oocyte before fertilization. 127 Figure 4.3. Initial polarization of outer blastomeres does not depend on Yap1/Wwtr1. 128 Figure 4.3 (cont’d). A) Immunofluorescence for aPKCζ and PARD6B in 16-cell mouse embryos lacking maternal Yap1;Wwtr1 and either heterozygous or null for zygotic Yap1;Wwtr1. B) Quantification of the percentage of outside blastomeres from embryos in A which display apical localization of aPKCζ. Statistical analysis was performed using analysis of variance (ANOVA) with Tukey’s post-hoc test. 129 CHAPTER 5. EFFICIENT GENERATION OF ENDOGENOUS PROTEIN REPORTERS FOR MOUSE DEVELOPMENT Daniel O’Hagan1, Robin E. Kruger2, Bin Gu3,4, and Amy Ralston1,2 1) Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824 2) Reproductive and Developmental Sciences Training Program, Michigan State University, East Lansing, MI, 48824 3) Department of Obstetrics, Gynecology and Reproductive Biology, Michigan State University, East Lansing, MI, 48824 4) Institute for Quantitative Health Science and Engineering, Michigan State University, East Lansing, MI, 48824 This chapter is modified from O’Hagan et al., published in Development on May 20, 2021. This study was supported by the National Institutes of Health R35 GM131759 and T32 HD087166. 130 Section 5.1. Abstract Fluorescent proteins and epitope tags can reveal protein localization in cells and animals, yet the large size of many tags hinders efficient genome targeting. Accordingly, many studies have relied on characterizing overexpressed proteins, which might not recapitulate endogenous protein activities. Here, we present two strategies for higher throughput production of endogenous protein reporters in mice, focusing on the blastocyst model of development. Our first strategy makes use of a split fluorescent protein mNeonGreen2 (mNG2). Knock-in of a small portion of the mNG2 gene, in frame with gene coding regions of interest was highly efficient in embryos, potentially obviating the need to establish mouse lines. When complemented by the larger portion of the mNG2 gene, fluorescence was reconstituted and endogenous protein localization faithfully reported in living embryos. Our second strategy achieves in-frame knock-in of a relatively small protein tag, which provides high efficiency and higher sensitivity protein reporting. Together, these two approaches provide complementary advantages and enable broad downstream applications. Section 5.2. Introduction Mouse models are essential tools for research to uncover human disease mechanisms. To produce new mouse lines, embryos are collected and genetically manipulated during the preimplantation stage, and are then transferred to surrogate mothers for gestation. Thus, preimplantation embryos are the starting point for many studies. Within preimplantation, the blastocyst stage of development is also an alluring model in its own right. This is in part because the blastocyst provides technical advantages, including 131 optical transparency, the capacity to develop ex vivo in a cell culture incubator, and the ease of collecting dozens of embryos at a time. These properties have enabled discovery of the molecular mechanisms of the first steps in mammalian development. Moreover, embryonic stem cells (ESC) are derived from blastocysts, providing additional models for basic and applied research. Thus, technological advances using the blastocyst can impact broader areas of biomedical research. One powerful approach to elucidating the molecular mechanisms of development and disease has been live imaging of fluorescent reporters in vivo, which enables time- resolved analysis of gene expression at the cellular level (Nowotschin & Hadjantonakis, 2014). Live imaging of gene expression in vivo is often achieved by knocking in genes encoding Green Fluorescent Protein (GFP) and other fluorescent proteins downstream of gene promoters, to create a gene reporter. This approach requires establishing and breeding new mouse lines. An alternative method of protein detection is to use antibodies to localize endogenous proteins, for example by immunofluorescence. However, immunofluorescence does not allow visualization of dynamic processes. Moreover, identification of reliable and specific antibodies can also be time-intensive and, for some antigens, may not exist. Our goal was to help overcome some of these challenges by developing an alternative, streamlined pipeline for the detection or screening of endogenous proteins in vivo. We focus on preimplantation mouse embryos, where we present two complementary approaches to enhance the efficiency of detecting endogenous proteins in vivo. We 132 provide guidelines for implementation of these approaches in broader experimental settings. Section 5.3. Materials and Methods Animal Use All animal research was conducted in accordance with the guidelines and approval of the Michigan State University Institutional Animal Care and Use Committee. Most experiments were performed using male or female CD-1 mice, at least 6-8 weeks of age, maintained on a 12-hour day/night cycle with food and water ad libitum. Plasmid construction The pR26-CAG-mNG2(∆11) targeting vector was cloned by insertion of a synthesized dsDNA fragment encoding mNG2(∆11) (Table 5.4) into the previously published vector pR26-AsisI/MluI (Addgene #74286) (V. T. Chu et al., 2016) via restriction/ligation with AsisI and MluI. After cloning, the Lox-Stop-Lox site was removed by exposure to recombinant Cre recombinase (NEB), using the NEB standard protocol. The in vitro transcription plasmids for Clta-mNG2(11), mNG2(∆11), and Cdx2-mNG2(11) were cloned by inserting a synthesized dsDNA fragment (Table 5.4) containing the respective coding sequence into a pcDNA3.1-poly(A)83 vector (K. Yamagata et al., 2005) downstream of the T7 promoter via restriction ligation with HindIII and NotI. pX459- sgRosa26-1 was generated by inserting the guide RNA sequence targeting Rosa26 (Table 5.3) into pSpCas9(BB)-2A-Puro (PX459) V2.0 (Addgene #62988) via restriction ligation with BbsI. 133 mRNA synthesis In vitro transcription (IVT) was performed using the T7 mMessage mMachine kit (Life Technologies). Each IVT construct was digested with XbaI, followed by ethanol precipitation, and was then used in the IVT reaction per manufacturer’s instructions. Resulting mRNA was purified using the MEGAclear kit (Ambion). mRNA quantity and quality were assessed by Nanodrop spectrophotometer and by agarose gel, respectively. Zygote and 2-cell embryo microinjection Target-specific crRNA and non-variable tracrRNA obtained from Integrated DNA Technologies were each suspended in injection buffer (1 mM Tris HCl, pH 7.5; 0.1 mM EDTA), mixed at a 1:1 molar ratio and annealed in a thermal cycler by ramping down from 95 ºC to 25 ºC at 0.1 ºC/second. The annealed RNAs were then mixed with recombinant Cas9 Protein at a 5:1 RNA:Cas9 molar ratio and allowed to form RNPs for 15 minutes at room temperature. RNPs were mixed with donor ssODN synthesized as Ultramers from Integrated DNA Technologies (Table 5.2) and diluted to working concentrations (Table 5.5) by addition of injection buffer. For mRNA injections, each mRNA was diluted in injection buffer to 350 ng/µl and injected either into one pronucleus of the mouse zygote or into one blastomere of the 2-cell mouse embryo. Injection mixes and mRNAs were aliquoted and stored at -80 ºC, avoiding freeze/thaw cycles. Zygotes were harvested from naturally mated pregnant mice on the day that copulatory plugs were detected. Oviducts were flushed with M2 medium (Millipore Sigma), and 134 then injection mix was delivered into one pronucleus or the nucleus of one blastomere via microinjection (Nagy et al., 2003). Injected zygotes were cultured in KSOM + amino acids (Millipore Sigma) for up to five days before being either fixed or imaged live. Only embryos that survived injection and appeared to have cavitated or to be attempting cavitation were included in the analysis. Generation of R26-mNG2(∆11) Mouse Line The R26-mNG2(∆11) mouse line was generated by zygote microinjection at the Michigan State University Transgenic and Genome Editing Facility. Rosa26 sgRNA was synthesized by in vitro transcription (IVT) of PCR-amplified region of PX459-sgRosa26- 1 using the Life Technologies MEGAshortscript T7 kit. Transcripts were subsequently purified using the MEGAclear kit. A mixture containing 5 ng/µl circular pR26-CAG- mNG(∆11) and 125 ng/µl Cas9 RNP complexed with sgRosa26 guide RNA was injected into one pronucleus of C57BL/6J mouse zygotes. Zygotes were then transferred to CD- 1 recipient mice. After birth, tail tips were screened by PCR for successful integration of the R26-CAG-mNG(∆11) allele. Immunofluorescence and Confocal Microscopy V5 was detected by mouse anti-V5 antibody (Thermo Fisher, R96025). Embryos were fixed with 4% formaldehyde (Polysciences) for 10 minutes, permeabilized with 0.5% Triton X-100 (Millipore Sigma) for 30 minutes, and then blocked in 10% Fetal Bovine Serum (Hyclone) with 0.1% Triton X-100 for 1 hour at room temperature. Embryos were then incubated in anti-V5 at a dilution of 1:400 in blocking solution at 4 ºC overnight. 135 The next day, embryos were stained with goat anti-mouse Alexa488 (Invitrogen, A- 11030) at a 1:400 dilution in blocking solution for 1 hour at room temperature. Embryos were then stained for 10 minutes at room temperature in 50 µM Hoechst nucleic acid stain (Thermo Fisher) or DRAQ5 (Cell Signaling, 4084S, 1:400 dilution). Rabbit-anti- Nanog, (Reprocell, RCAB002P-F) was used at 1:400 dilution, with Cy3-conjugated donkey-anti-rabbit IgG (Jackson Laboratories, 711-165-152) at 1:400 dilution. Split mNG2 embryos were imaged either fixed or live after Hoechst staining. Imaging was performed using an Olympus FluoView FV1000 Confocal Laser Scanning Microscope system with 20x UPlanFLN objective (0.5 NA) and 5x digital zoom or with 60x PlanApoN oil (NA 1.42) objective. For each embryo, z-stacks were collected with 3 µm intervals between optical sections. Optical sections are displayed as an intensity projection over the Z axis. Figures were prepared using FIJI, Adobe Photoshop, and Adobe Illustrator. Genotyping To genotype embryos, genomic DNA was extracted from single blastocysts by placing each blastocyst in a microtube containing 4.4 µl Extraction buffer (REDExtract-N-Amp Tissue PCR Kit, Millipore Sigma) mixed with 1.1 µl of Tissue Prep Buffer, and then incubating tubes at 56 ºC for 30 minutes, 24 ºC for 5 minutes, and 95 ºC for 5 minutes. After incubation, 5 µl Neutralization buffer was added to each tube. In subsequent reactions, 1 µl of embryo extract was used as PCR template, and locus-specific primers (Table 5.1). To genotype adult mice, genomic DNA was extracted from ear punch 136 biopsies using the Wizard SV Genomic DNA Purification System (Promega), and PCR was performed using Herculase II Polymerase (Agilent). Sequencing To confirm the identity of select PCR products, the products were directly cloned into pCR2.1 TOPO using the Invitrogen TOPO TA Cloning Kit (Invitrogen). Plasmids containing the PCR product were prepped with the Spin Miniprep Kit (Qiagen) and then sequenced by RTSF Sanger method at the Genomics Core at Michigan State University. mNG2(∆11) ES Cell Derivation R1/E ES cells (ATCC) were cultured on CF-1 feeder MEFs (Applied Stem Cell) in ES cell medium [DMEM supplemented with 1000 U/mL leukemia inhibitory factor (Millipore Sigma), 15% (v/v) fetal bovine serum (Hyclone), 2 mM L-Glutamax (Thermo Fisher), 0.1 mM beta-mercaptoethanol (Millipore Sigma), 0.1 MEM non-essential amino acids (Millipore Sigma), 1 mM Sodium Pyruvate (Millipore Sigma), and 1% (v/v) penicillin/streptomycin (Gibco)]. Passage 13 R1/E ES cells were cultured to approximately 70% confluence in a 10 cm dish, and then electroporated with pX459- sgRosa26-1 and pR26-CAG-mNG2(∆11) as follows: pelleted cells were resuspended in 800 µl Embryo-Max Electroporation Buffer (Millipore Sigma) containing 20 µg each plasmid, and cells were then electroporated in a 0.4 cm electrode gap electroporation cuvette (Bio-Rad) using Bio-Rad Gene Pulser XCell electoporator (250 V, 500 µF, infinite Ω). Subsequently, 400 µl electroporated cells were then diluted in 10 mL ES cell 137 medium, and then plated on a 10 cm dish on puromycin-resistant DR4 feeder MEFs (Applied Stem Cell). After 24 hours, selection was started with ES cell medium containing 1.25 µg/mL puromycin (Gibco). After 12 days, colonies were picked into 96- well plates and expanded over several more passages. Cell lines were genotyped by PCR using R26F3 and SAR primers to detect insertion of mNG2(∆11) in the Rosa26 locus (Table 5.1). Section 5.4. Results A mouse line to enable in vivo implementation of a split fluorescent protein Like GFP, the yellow-green, monomeric fluorescent protein mNeonGreen (mNG), derived originally from the marine invertebrate Branchiostoma lanceolatum, is an eleven-stranded beta-barrel, but is up to three times brighter than GFP (Shaner et al., 2013). The mNG derivative, mNG2 can be split into two separate coding units, mNG2(∆11), which lacks the eleventh beta-strand, and mNG2(11), which is the eleventh beta-strand (Feng et al., 2017). Individually, the two resulting proteins lack appreciable fluorescence. However, when the larger protein mNG2(∆11) is complemented by the 16- amino acid mNG2(11), fluorescence is reconstituted (Fig. 5.1A), and the two proteins are capable of self-assembly through non-covalent intermolecular interactions (Cabantous et al., 2005). We sought to make use of this fluorescence complementation strategy to evaluate localization of endogenous proteins in mouse embryos because we reasoned that tagging endogenous proteins with the smaller, 16-amino acid mNG2(11) coding region 138 would be more efficient than knocking in the full-length gene encoding the full-length, 236 amino acid fluorescent protein. Then, to provide the complementary protein, we aimed to establish a mouse line capable of constitutive expression of mNG2(∆11). Our goal was to introduce an expression construct including cytomegalovirus enhancer, chicken beta-actin promoter, rabbit beta-globin splice acceptor (CAG) sequences, and the mNG2(∆11) coding region into the Rosa26 (R26) locus by homologous recombination (Fig. 5.1B), which would enable constitutive, ubiquitous expression of mNG2(∆11) throughout mouse tissues and development (Friedrich & Soriano, 1991). However, prior to attempting knock-in in mouse zygotes, we first established an R26- mNG2(∆11) embryonic stem (ES) cell line using a Cas9/CRISPR-mediated knock-in strategy (V. T. Chu et al., 2016) (see Methods). These R26-mNG2(∆11) ES cells provided a renewable source of positive control genomic DNA for subsequent experiments. To produce a mouse line capable of expressing mNG2(∆11), we subsequently introduced the mNG2(∆11) expression construct into the Rosa26 (R26) locus in zygotes, following the strategy we had used in ES cells. Injected zygotes were transferred to recipient females, allowed to gestate, and then founder mice carrying mNG2(∆11) were identified by PCR genotyping (Fig. 5.1C-D) and genomic sequencing (not shown). A single founder mouse was then expanded, and bred to homozygosity to establish R26-mNG2(∆11)/R26-mNG2(∆11) mice. In principle, providing mNG2(11) in trans to R26-mNG2(∆11) would lead to reconstitution of the green fluorescent protein. For simplicity, we called this the GOGREEN system. 139 As an initial evaluation of the GOGREEN system, our first test was to determine if we could detect fluorescence complementation in embryos by epifluorescence microscopy (Fig. 5.2A). For this test, we generated mRNA encoding mNG2(11)-tagged Clathrin, light polypeptide (Clta). For negative controls, mRNA encoding either mNG2(∆11) or mNG2(11)-Clta were injected individually into wild type embryos (Fig. 5.2B). For a positive control, wild type zygotes were co-injected with mRNAs encoding both R26- mNG2(∆11) and mNG2(11)-Clta, and these exhibited greatly elevated fluorescence over both negative controls. Finally, R26-mNG2(∆11)/+ zygotes were injected with mRNA encoding mNG2(11)-Clta, which led to elevated fluorescence at the blastocyst stage, demonstrating functionality of the GOGREEN system in vivo using epifluorescence and an exogenous mNG2(11)-tagged protein. Fluorescence reconstitution by split fluorescent protein knock-in We next aimed to evaluate the performance of the GOGREEN system when mNG2(11) was endogenously expressed from several genomic loci. Our goal was to derive R26- mNG2(∆11)/+ zygotes and, in these, perform CRISPR/Cas9-mediated knock-in of mNG2(11) in frame with proteins of interest (Fig. 5.3A) to produce mNG2(11) fusion proteins capable of complementing mNG2(∆11) and reporting endogenous protein patterns. To achieve in-frame mNG2(11) knock-in, we designed targeting constructs encoding the 16 amino acid mNG2(11), plus a three-amino acid linker, flanked by genomic locus- specific homology arms of 30 nucleotides each (Fig. 5.3B). Resulting targeting 140 constructs ranged from 117-120 nucleotides in length, permitting their synthesis as a single stranded oligodeoxyribonucleotides (ssODN) by a commercial vendor (see Methods). For our first knock-in attempts, we targeted cytoskeletal proteins, including intermediate filaments and beta-actin, because their subcellular localizations in mouse preimplantation have long been known (Chisholm & Houliston, 1987; Coonen et al., 1993; Reima & Lehtonen, 1985). We designed CRISPR reagents to knock mNG2(11) in-frame with Keratins (Krt) Krt8 and Krt18, as well as Actin, beta (Actb). Following injection of the knock-in mixture into R26-mNG2(∆11)/+ zygotes, embryos were cultured to the blastocyst stage, and then imaged by confocal microscopy. For each knock-in, we observed the very unique fluorescent meshwork of cortical filamentous proteins expected, in accordance with published observations (Coonen et al., 1993; Lim et al., 2020; Ralston & Rossant, 2008; Reima & Lehtonen, 1985). These observations are indicative of faithful protein reporting. Individual embryos were then harvested, and gene targeting evaluated by PCR (Fig. 5.3D) and sequencing (not shown). In all cases, monoallelic targeting was highly efficient (Fig. 5.3E). These observations demonstrate the utility of the GOGREEN system for efficiently reporting localization of endogenous proteins in vivo. Given that the dynamics of cytoskeletal protein localization and turnover during preimplantation development are actively studied (Anani et al., 2014; Schwarz et al., 2015; Zenker et al., 2018), the GOGREEN system could provide new tools since these proteins are usually visualized either in fixed embryos or by injection of mRNAs encoding tagged proteins, both of which could introduce unwanted artifacts. 141 Fluorescence complementation in the nuclear compartment Thus far, we had evaluated the ability of the GOGREEN system to report endogenous cytoplasmic proteins in vivo. However, we were uncertain whether the GOGREEN system could effectively report the dynamics of endogenous nuclear protein expression, owing to the possibility that the two components of the GOGREEN system might end up separated by the nuclear membrane. To investigate the performance of the GOGREEN system in visualizing nuclear proteins in vivo, we evaluated fluorescence in embryos after targeting the genes Nucleophosmin (Npm1) and NOP58 ribonuclear protein (Nop58), which both encode nucleolar proteins. As for previous experiments, we targeted mNG2(11) in frame with target genes in the R26-mNG2(∆11)/+ genetic background. Remarkably, were able to detect fluorescence within the nuclear compartment (Fig. 5.4A) in embryonic cells following mNG2(11) knock-in (Fig. 5.4B, C). Npm1 fluorescence recapitulated the pattern reported by immunofluorescence (E. J. Vogt et al., 2012), while the observed Nop58 pattern is novel. These observations indicate that the nuclear envelope does not necessarily present a barrier to fluorescence complementation, in spite of fact that the GOGREEN components lack nuclear localization sequences. Robust detection of low abundance endogenous proteins Having observed that the GOGREEN system can detect both cytoplasmic and nuclear proteins, we next tested its performance in reporting transcription factor localization, since there is great interest in imaging transcription factor dynamics in living 142 preimplantation embryos (B. Gu et al., 2018; McDole et al., 2011; Posfai et al., 2017; Saiz et al., 2013, 2015). We next evaluated CDX2, YAP1, GATA6, and NANOG, four transcription factors with essential activities during preimplantation development (Frum et al., 2018; Mitsui et al., 2003; Schrode et al., 2014; Strumpf et al., 2005). However, we were unable to detect appreciable fluorescent signal in embryos of any of these four mNG2(11) knock-ins in the mNG2(∆11) background (Fig. 5.4D and data not shown). We therefore developed a second and alternative knock-in tagging strategy for detecting endogenous transcription factors. We selected the V5 epitope, a 14 amino acid protein derived from the simian virus 5 (SV5) paramyxovirus because its small size promised high knock-in efficiency and because of the existence of low background, commercially available, monoclonal anti- V5 antibody that could be used for immunofluorescent detection of V5-tagged proteins in embryos. We then designed V5-encoding targeting constructs for generating in-frame V5 fusion proteins (Fig. 5.5A,B). We targeted the V5 tag in frame with key transcription factors in zygotes, and then observed immunofluorescence patterns in blastocysts by confocal microscopy. We were able to detect V5 signals that were clear and specific after targeting the nuclear factors such as GATA3, CTCF, and NANOG (Fig. 5.5C-E). Importantly, the patterns of V5- tagged GATA3, NANOG, and CTCF recapitulated their reported expression patterns in blastocysts (Home et al., 2009; Marcho et al., 2015; Ralston & Rossant, 2008; Strumpf et al., 2005). In a parallel set of experiments, we harvested embryos prior to blastocyst 143 stage and then costained these with anti-V5 and anti-NANOG antibodies. In these embryos, we detected NANOG expression in every V5-expressing cell (Supp. Fig. 5.1), confirming the utility of the V5 knock-in approach for faithfully reporting gene expression, even at preimplantation stages prior to blastocyst. These observations highlight the utility of the V5-tagging system to evaluate the endogenous expression patterns of known or novel transcription factors. Having observed that V5 outperformed the GOGREEN system, in terms of transcription factor detection, we hypothesized that protein abundance could be the limiting factor for detection using the GOGREEN system. Consistent with this hypothesis, we were able to detect the transcription factor CDX2 using GOGREEN when Cdx2-mNG2(11) was overexpressed by mRNA injection (Fig. 5.6A,B). Finally, we evaluated the abundance of transcripts encoding proteins evaluated in this study, as measured by RNA sequencing of individual blastocysts (Aksoy et al., 2013). Remarkably, transcript abundance predicted protein detectability using the GOGREEN system (Fig. 5.6C). Moreover, proteins of extremely low abundance could still be detected using the V5 system. This analysis therefore provides a guideline for informing subsequent experimental design and in selecting the optimal protein tagging approach. Ultimately, the GOGREEN and V5 systems together enable detection of endogenous proteins across the range of protein expression levels, facilitating multiple downstream applications, and opening doors for new discoveries. 144 Section 5.5. Discussion Split GFP and V5 epitope tagging have been used for protein detection in cell lines and in some animal models (Hefel & Smolikove, 2019; Kamiyama et al., 2016; Kim et al., 2012; Leonetti et al., 2016; To et al., 2016; M. Yamagata & Sanes, 2012; H. Yang et al., 2013), but their use as knock-in mouse reporters has not been systematically compared across diverse genomic loci. Here, we presented a systematic comparison of their performance, sensitivity, and efficiency of endogenous protein reporting. We note that both approaches are similarly efficient, averaging ~60% knock-in efficiency across more than a dozen loci tested. This rate is much higher than targeting full-length fluorescent proteins by zygote injection. For example, we observed a 6% knock-in efficiency at the Rosa26 locus with the nearly full-length fluorescent protein mNG2(∆11). In fact, we observed upwards of 75-100% knock-in efficiency for multiple loci, which exceeds allele inheritance rates in most mating strategies. Moreover, the relatively short length of the ssODN enables higher efficiency targeting and ease of synthesis, bypassing traditional molecular cloning methods required for producing longer donors. Finally, tagging with GOGREEN and V5 enables efficient detection of endogenous proteins, thereby circumventing artifacts caused by imaging fluorescently tagged, overexpressed proteins. We note opportunities for applying biochemical and molecular techniques in vivo. V5 is commonly used for purifying proteins from cells and tissues for the downstream identification of protein or nucleotide interactions, including immunoprecipitation-western blotting or mass spectrometry, chromatin-immunoprecipitation or ribonucleotide 145 pulldown and sequencing (ChIP-seq, RIP-seq) or Cleavage Under Targets and Release Using Nuclease (CUT&RUN) (Hainer et al., 2019; Skene & Henikoff, 2017). We therefore envision that the approaches described here could be used to generate stable mouse lines that enable anti-V5 antibody-mediated discovery of protein localization patterns, protein and RNA binding partners and DNA binding sites throughout the genome. Both GOGREEN and V5 systems present exciting opportunities for biological investigation outside of preimplantation mouse development as well. For example, V5 or mNG2(11) knock-in embryos could be transferred to recipient females to allow for postimplantation development so that protein localization can be evaluated in later developmental processes or in adult tissues and organs. Additionally, both the GOGREEN and V5 systems could be adaptable to viral transduction (Yoon et al., 2018), which could extend applications to adult organs and tissues. Our studies thus provide guidelines, molecular reagents, and genotyping assays to enable these applications. In considering endogenous protein tagging applications, we identify several key considerations. First, care should be given to the design of the tagged protein, and whether the location and nature of the tag interfere with protein function. Validation for protein function and localization can be confirmed using appropriate strategies, including mouse genetics and, if possible, by confirming protein localization by immunofluorescence. Second, guide RNA design should follow best practices so as to minimize the chance of on/off-target indel alleles; targeting protein c-termini may help 146 avoid unwanted phenotypes caused by frame-shift mutations. Third, the genotyping strategy should confirm that the tag has been knocked in in-frame with the target protein at the sequence level. Related to this, strategies for identifying random ssODN insertions should be considered (Lanza et al., 2018). Finally, if microinjection is to be used as the delivery method, consultation with institutional transgenic facility with proper technical expertise should be sought, when available to ensure optimal experimental design. Finally, both the GOGREEN and V5 systems could also be used in embryos from species such as humans or other primates, where breeding to establish knock-in lines is either inappropriate or impractical. There would be additional advantages to applying either system to emerging mammalian models, such as marsupials, where protein- specific antibodies have not yet been developed. For live imaging, mNG2(∆11) could be provided by mRNA injection, while mNG2(11) would be knocked in frame into genes of interest. If fixed imaging of low abundance proteins is preferred, then V5 could be knocked in. Either system promises new opportunities for discovery of developmental principles in mouse as well as understudied mammalian species. Section 5.6. Acknowledgements We thank Dr Kristin Parent for comments on the manuscript; current and former members of the Ralston Laboratory, especially Dr Michael Halbisen and Dr Tristan Frum, for helpful discussions and training; Axel Schmitter for assistance in the 147 laboratory; and Drs Elena Demeriva and Huirong Xie at the MSU Transgenic and Genome Editing Facility for the R26 mouse knock-in line. 148 Allele R26- mNG2(∆11) Krt18- mNG2(11) Krt8- mNG2(11) mNG2(11)- Actb Npm1- mNG2(11) mNG2(11)- Nop58 Ctcf-V5 V5-Gata3 Nanog-V5 Cdx2- mNG2(11) TABLES Forward CTGCCCGAGCGGAAAC GCCACTGAC GGCTGTTTATAACTAAGG CTTGGTC TGTGGTTGTGAAGAAGA TTGAAACC CCAGCGTTTGCCTTTTAT GGTAATA GGCAACACTGGCCATAA AGTATTTA GATATTTTAAGGCCGTCT CTTTCCG CAGAATACAGGTGCAAT TGAGAACA CTTTTGCTAAACTATCCC GCAAAGA CCACTAGGGAAAGCCAT GCGCATTT GAGAGGAAAATCAAGAA GAAGCAGC Reverse CCTGGACTACTGCGCCC TACAGA GGACAGTCATATCTCCTA CTTCGTC ATACAACTGAATTGGGTT TGGATGG CACTCCCAAAGTAACAG GTCACTT CAAACACAGTAGGGAAA GTTCTCAC CAACAACTCCATCTCAC CTACCTTA CATCCTTGAAGTTTTCGT TCTCAGT TTGCCTTGACCATCGAT GTTAAAAA GGAAGAAGGAAGGAAC CTGGCTTTGC GAGGAATCTCTTCTGAG GATTCTCG Table 5.1. Genotyping primers used in Chapter 5. 149 Sequence 5’ to 3’ Allele Krt18-mNG2(11) T*TCCCAGGGGTTCCCTCCTTCTCTGCCTCACATCATATCGG TAAAGGCCTTTTGCCACTCCTTGAAGTTGAGCTCGGTGCCA GAGCCGTGC CTCAGAACTCTGGTGTCATTAGTCT*C Krt8-mNG2(11) G*TGTCCGAGTCTTCTGATGTCGTGTCCAAGGGCTCTGGCA CCGAGCTCAACTTCAAGGAGTGGCAAAAGGCCTTTACCGAT ATGATGTGAA TGGCCACTGAAGTCCTTGCCAGCCT*G mNG2(11)-Actb G*ACGACCAGCGCAGCGATATCGTCATCCATGCCACCTCCC ATCATATCGGTAAAGGCCTTTTGCCACTCCTTGAAGTTGAGC TCGGTCATG GCGAACTATCAAGACACAAAAGAAGGCT*A Ctcf-V5 mNG2(11)- Nop58 Npm1-mNG2(11) C*AAGATCTCTGGCAGTGGAGGAAATCTCTTGGCTCTGGCA CCGAGCTCAACTTCAAGGAGTGGCAAAAGGCCTTTACCGAT ATGATGTAAG AAAAGGGTTTAAACAGTTTGAAATA*T C*GCGTAGCGCCGCCCTGACCTGGTCTCATCATGACCGAGC TCAACTTCAAGGAGTGGCAAAAGGCCTTTACCGATATGATG GGAGGTGG CATGTTGGTCCTGTTTGAAACGTCCGTTGG*C C*CTGAGATGATCCTCAGCATGATGGACCGGGGCTCTGGCG GCAAGCCGATCCCTAACCCTCTGCTGGGCCTGGACAGCAC TTGATGCTG GGGCCTTGCTCGGCACCAGGA*C G*GGCGAGAGGGCGCGAGCACAGCCGAGGACATGGGCAA GCCGATCCCTAACCCTCTGCTGGGCCTGGACAGCACTGGA GGTGGCATGG AGGTGACTGCGGACCAGCCGCGCTG*G A*CTTTAAGCCCAGATGTTGCGTAAGTCTCAAGTGCTGTCCA GGCCCAGCAGAGGGTTAGGGATCGGCTTGCCGCCAGAGCC TATTTCACC TGGTGGAGTCACAGAGTAGT*T Nanog-V5 V5-Gata3 Cdx2-mNG2(11) C*GCCGCCGCTTCAGACCACGGGAGGGGTCACATCATATCG GTAAAGGCCTTTTGCCACTCCTTGAAGTTGAGCTCGGTGCC AGAGCCCT GGGTGACAGTGGAGTTTAAAACCCCTC*C Table 5.2. Synthesized ssODN sequences used in Chapter 5. Phosphorothioate bonds (indicated by *) were added during oligo synthesis to enhance oligo resistance to endogenous exonuclease degradation. 150 Allele R26-mNG(∆11) Krt18-mNG2(11) Krt8-mNG2(11) mNG2(11)-Actb Npm1-mNG2(11) mNG2(11)-Nop58 Ctcf-V5 V5-Gata3 Nanog-V5 Cdx2-mNG2(11) Guide Sequence (5’ to 3’) ACTCCAGTCTTTCTAGAAGATGG ACCAGAGTTCTGAGGCACTGAGG TGATGTCGTGTCCAAGTGAATGG TGTGTCTTGATAGTTCGCCATGG GAGGAAATCTCTTTAAGAAAAGG CTGACCTGGTCTCATCATGTTGG GAGCAAGGCCCCAGCATCACCGG GAGCACAGCCGAGGACATGGAGG CGTAAGTCTCATATTTCACCTGG CAGACCACGGGAGGGGTCACTGG Table 5.3. CRISPR Guides used in Chapter 5. Underlined sequence = Protospacer Adjacent Motif (PAM). 151 Purpose mNG2(D11) Synthesized dsDNA Fragment Clta-mNG2(11) Synthesized dsDNA Fragment for in vitro transcription (IVT) Sequence 5’ to 3’ gccagctaggccttgaccaaagttcctctggaattgcgatgcgatcgcggagaattct ccacgcgtcgccaccATGGTGAGCAAGGGTGAGGAGGATAACAT GGCCTCTCTCCCAGCGACTCATGAGTTACACATCTTTGGC TCCATCAACGGTGTGGACTTTGACATGGTGGGTCAGGGTA CCGGCAATCCAAATGATGGTTATGAGGAGTTAAACCTGAA GTCCACCAAGGGTGACCTCCAGTTCTCCCCCTGGATTCTG GTCCCTCATATCGGGTATGGCTTCCATCAGTACCTGCCCTA CCCTGACGGGATGTCGCCTTTCCAGGCCGCCATGGTAGAT GGCTCCGGATACCAAGTCCATCGCACAATGCAGTTTGAAG ATGGTGCCTCCCTTACTGTTAACTACCGCTACACCTACGAG GGAAGCCACATCAAAGGAGAGGCCCAGGTGATGGGGACT GGTTTCCCTGCTGACGGTCCTGTGATGACCAACACGCTGA CCGCTGCGGACTGGTGCATGTCGAAGAAGACTTACCCCA ACGACAAAACCATCATCAGTACCTTTAAGTGGAGTTACACC ACTGTAAATGGCAAACGCTACCGGAGCACTGCGCGGACC ACCTACACCTTTGCTAAGCCAATGGCGGCTAACTATCTGAA GAACCAGCCGATGTACGTGTTCCGTAAGACGGAGCTCAAG CACTCCATGGGAACAGGTGGCGGCGGAAGTTAGagcgatcg cactccaatgccctggatcgcacgcgtaaatgattgcagatccactagttctagag ctagttaagcttgctgtcgccaccATGACCGAGCTCAACTTCAAGGAG TGGCAAAAGGCCTTTACCGATATGATGGGAGGTGGCATGG CCGAGCTTGACCCTTTTGGAGCTCCAGCAGGCGCACCAG GTGGTCCTGCTCTGGGTAATGGTGTTGCCGGGGCAGGTG AGGAAGACCCTGCTGCAGCATTTCTTGCCCAGCAAGAATC TGAGATAGCTGGGATCGAAAATGACGAGGCATTTGCCATT CTCGACGGTGGAGCCCCCGGACCCCAACCTCATGGGGA GCCTCCTGGCGATGCAGTTGACGGGGTTATGAACGGAGA ATACTATCAGGAGTCTAACGGCCCCACAGATAGCTACGCC GCCATATCAGAGGTTGATAGACTTCAATCAGAGCCAGAGA GCATACGAAAATGGCGCGAAGAACAGACCGAGCGATTGG AAGCTCTTGATGCTAACAGTCGCAAGCAAGAAGCCGAATG GAAGGAAAAAGCCATCAAAGAACTGGAGGAATGGTATGCC AGACAGGATGAGCAGTTGCAAAAGACAAAGGCAAATAATA GAGCCGCAGAGGAGGCTTTTGTCAATGATATAGACGAGTC CAGCCCCGGAACCGAGTGGGAACGGGTCGCTAGACTCTG TGATTTCAATCCAAAATCCAGCAAGCAGGCCAAAGACGTG TCACGAATGCGGTCAGTGCTTATCTCATTGAAGCAAGCAC CACTGGTTCATTGAagagccgcggccgctcgact Table 5.4. dsDNA fragments synthesized for Chapter 5. Coding sequences are capitalized. 152 Purpose mNG2(D11) Synthesized dsDNA Fragment for IVT Cdx2-mNG2(11) Synthesized dsDNA Fragment for IVT Sequence 5’ to 3’ ctagttaagcttgctgtcgccaccATGGTGAGCAAGGGTGAGGAGGAT AACATGGCCTCTCTCCCAGCGACTCATGAGTTACACATCTT TGGCTCCATCAACGGTGTGGACTTTGACATGGTGGGTCAG GGTACCGGCAATCCAAATGATGGTTATGAGGAGTTAAACCT GAAGTCCACCAAGGGTGACCTCCAGTTCTCCCCCTGGATT CTGGTCCCTCATATCGGGTATGGCTTCCATCAGTACCT GCCCTACCCTGACGGGATGTCGCCTTTCCAGGCCGCCAT GGTAGATGGCTCCGGATACCAAGTCCATCGCACAATGCAG TTTGAAGATGGTGCCTCCCTTACTGTTAACTACCGCTACAC CTACGAGGGAAGCCACATCAAAGGAGAGGCCCAGGTGAT GGGGACTGGTTTCCCTGCTGACGGTCCTGTGATGACCAA CACGCTGACCGCTGCGGACTGGTGCATGTCGAAGAAGAC TTACCCCAACGACAAAACCATCATCAGTACCTTTAAGTGGA GTTACACCACTGTAAATGGCAAACGCTACCGGAGCACTGC GCGGACCACCTACACCTTTGCTAAGCCAATGGCGGCTAAC TATCTGAAGAACCAGCCGATGTACGTGTTCCGTAAGACGG AGCTCAAGCACTCCATGGGAACAGGTGGCGGCGGAAGTT AGagagccgcggccgctcgact ctagttaagcttgctgtcgccaccATGTACGTTTCATACTTGCTGGACA AGGACGTCAGTATGTATCCCAGTAGTGTTCGCCACTCAGG GGGATTGAATTTGGCCCCCCAAAATTTCGTTAGCCCCCCT CAATACCCTGACTATGGCGGCTATCATGTAGCCGCCGCAG CAGCCGCAACTGCTAACTTGGATAGTGCCCAATCTCCCGG GCCCTCCTGGCCAACAGCATACGGGGCTCCCCTCCGAGA AGATTGGAATGGGTATGCTCCTGGGGGTGCAGCAGCAGC AAATGCTGTGGCCCACGGTCTGAATGGGGGCAGTCCTGC TGCCGCTATGGGGTATTCTAGTCCTGCCGAATACCATGCC CATCACCACCCACATCACCATCCACACCATCCCGCCGCCT CTCCATCCTGTGCAAGCGGGCTGCTCCAAACCCTGAACCT CGGACCCCCAGGTCCTGCTGCCACTGCAGCTGCAGAACA ACTCTCACCAAGTGGTCAGCGGCGAAATCTGTGCGAATG GATGCGAAAACCAGCACAACAGTCACTCGGCTCTCAAGTC AAAACCCGCACCAAAGACAAGTACAGGGTCGTTTATACCG ATCATCAGCGGTTGGAACTGGAGAAAGAGTTCCATTTTAG CAGGTATATAACTATTCGACGCAAATCTGAGCTTGCTGCAA CTTTGGGGTTGTCAGAGCGGCAAGTTAAGATCTGGTTTCA AAACCGACGAGCTAAGGAAAGAAAAATTAAGAAAAAGCAA CAACAACAGCAGCAACAGCAACAGCAGCAACCCCCCCAA CCTCCTCCACAGCCATCCCAACCACAGCCAGGGGCATTG CGCAGTGTACCTGAGCCATTGAGCCCAGTGACCAGCCTC CAGGGTTCTGTACCAGGAAGCGTTCCCGGTGTGCTGGGG CCCGCTGGTGGCGTGCTTAACTCAACAGTGACACAGGGC TCTGGCACCGAGCTCAACTTCAAGGAGTGGCAAAAGGCC TTTACCGATATGATGTGAagagccgcggccgctcgact Table 5.4. (cont’d). 153 Gene Targeted Krt8 Actb Krt18 Nop58 Npm1 Ctcf Gata3 Nanog Tag mNG2(11) mNG2(11) mNG2(11) mNG2(11) mNG2(11) V5 V5 V5 Cas9 RNP (ng/µL) ssODN (ng/µL) 100 100 100 100 100 25 100 100 20 20 20 20 10 5 20 20 Table 5.5: Concentrations of Cas9 RNP and ssODN used for targeting each gene in Chapter 5. 154 FIGURES Figure 5.1. A mouse line for fluorescence complementation in vivo. (A) Deletion of mNG2(11), the 11th beta-strand of the fluorescent protein mNeonGreen2 (mNG2), eliminates its fluorescent properties. However, complementation by co-expression of mNG2(Δ11) and the 11th beta-strand mNG2(11) enables non-covalent association of the two proteins and reconstitution of the mNG2 fluorescent properties. (B) Strategy for CRISPR/Cas9 knock-in of the mNG2(Δ11) expression construct into the mouse Rosa26 (R26) locus. Sequence of single-guide RNA (sgRNA), location of genotyping primers (R26F3 and SAR) and predicted Cas9 cut site are shown. HA, homology arm; RNP, ribonucleoprotein. (C) PCR genotyping of tail tip biopsies from offspring born following zygote injection of CRISPR/Cas9 reagents to target the CAG-mNG2(Δ11) expression construct to the R26 locus in zygotes. 155 Figure 5.1 (cont’d). Successful homologous recombination suggested by PCR amplification of a 1.389 kb band from genomic DNA. B, C57BL/6 wild-type genomic DNA; L, DNA ladder; +, positive control (targeted embryonic stem cells); −, negative control (no DNA template). Numbers indicate individual mice screened by PCR; stars indicate potential founders (D) Summary of R26 targeting with mNG2(Δ11) expression construct. 156 Figure 5.2. Fluorescence complementation in vivo using the GOGREEN system. (A) Strategy for testing fluorescent complementation in early embryos. Zygotes carrying mNG2(Δ11) were harvested and then injected with mRNA encoding mNG2(11)-tagged clathrin (CLTA). Zygotes were subsequently cultured ex vivo to later stages and fluorescence examined in individual embryos. Individual embryo genotypes were determined by PCR. In control experiments, zygotes were produced harvested from wild-type parents. (B) Fluorescence reconstitution can be detected by epifluorescence. Negative controls, wild type embryos injected with mRNA encoding only mNG2(Δ11) or mNG2(11)-tagged Clta exhibit background levels of fluorescence (columns 1 and 2); positive control, wild-type embryos co-injected with mNG2(Δ11) and mNG2(11)-Clta exhibit reconstituted fluorescence (column 3); test of R26-mNG2(Δ11) mice, heterozygous knock-in embryos injected with mRNA encoding R26-mNG2(11)-Clta also exhibit reconstituted fluorescence above background (column 4). n, number of embryos evaluated in the experiment. Scale bars: 20 μm. 157 Figure 5.3. The GOGREEN system enables detection of endogenous proteins. (A) Experimental design: R26-mGN2(Δ11)/+ knock-in zygotes (generated per cross shown in Fig. 5.2A) are injected with CRISPR/Cas9 targeting reagents to knock mNG2(11) into loci of interest, in frame with target proteins. Embryos are then cultured ex vivo, imaged and genotyped to evaluate the efficiency of mNG2(11) knock-in. gRNA, guide RNA; ssODN, single-stranded oligodeoxyribonucleotide. (B) Overview of strategy for targeting mNG2(11) to genomic loci to produce fusion proteins. (C) The GOGREEN system enables detection of endogenous cytoskeletal proteins including intermediate filaments KRT18, KRT8 and ACTB. Note that 100% of embryos inherited R26-mNG2(Δ11). Scale bars: 20 μm. Sample sizes in E. (D) PCR genotyping of embryos, including those shown in C, to identify which embryos were successful mNG2(11) knock-ins.(E) Summary of mNG2(11) knock-in results observed for the three loci shown. *, one additional embryo presented the knock-in genotype, but not the expected fluorescent phenotype; **, one additional embryo presented the fluorescent phenotype, but not the expected genotype. 158 Figure 5.4. The GOGREEN system can report endogenous protein localization in the nucleus. (A) In the R26-mNG2(Δ11) background, knock-in of mNG2(11) in frame with the coding regions of two different nuclear proteins demonstrates fluorescent reconstitution of mNG2 and localization within nuclei. Scale bars: 20 μm. Sample sizes in C and D. (B) PCR genotyping to confirm knock-in of mNG2(11) into indicated loci for individual embryos, including those shown in A. (C) Efficient knock-in of nuclear proteins shown in A and B. *, one additional embryo presented the fluorescent phenotype but not the expected genotype. (D) Summary of mNG2(11) knock-in efficiencies, as determined by PCR genotyping, that were undetectable using the GOGREEN system. 159 Figure 5.5. A V5-based system for the detection of diverse transcription factors with a single antibody. (A) Strategy for knocking the V5-encoding gene into loci of interest in wild-type zygotes, to enable streamlined detection of diverse endogenous proteins with a monoclonal anti-V5 antibody. (B) Overview of V5 targeting strategy. (C) Examples of proteins detected as V5 fusion proteins, following knock-in as illustrated in A and B. Scale bars: 20 μm. Sample sizes in E. (D) PCR genotyping of embryos to confirm V5 knock-in at indicated genomic loci, including those shown in C. (E) Summary of V5 knock-in efficiency at indicated loci. *, one additional embryo presented the knock- in genotype, but not the expected fluorescent phenotype. 160 Figure 5.6. Complementary systems enable detection of endogenous proteins over a range of expression levels. (A) Overview of experimental design: mRNA injection into one of two blastomeres of the early mouse embryo, followed by imaging. (B) Overexpression of mRNAs encoding mNG2(Δ11) and Cdx2-mNG2(11) leads to reconstituted fluorescence in the nucleus of the injected blastomere. n, number of embryos presenting the phenotype shown. Scale bars: 20 μm. (C) Relative abundance of endogenous mRNAs encoding tagged proteins, as measured by RNA-seq (Aksoy et al., 2013), and detection results using GOGREEN or V5 systems. Each gray dot indicates a unique gene transcript. RPKM, reads per kilobase per million reads. For all genes shown, mNG2(11) or V5 knock-in was confirmed by PCR. 161 Supplemental Figure 5.1. NANOG-V5 is detected in NANOG-expressing cells. A) Experimental design. B) Immunofluorescent detection of V5 and NANOG in embryos prior to the blastocyst stage (n =3). 162 CHAPTER 6. FLUORESCENT REPORTERS DISTINGUISH STEM CELL COLONY SUBTYPES DURING SOMATIC CELL REPROGRAMMING Alexandra Moauro1,2, Robin E. Kruger3,4, Daniel O’Hagan5, and Amy Ralston4,5. 1) Molecular, Cellular and Integrative Physiology Ph.D. Program, Michigan State University, East Lansing, MI, 48824 2) D.O.-Ph.D. Program, Michigan State University, East Lansing, MI, 48824 3) Cell and Molecular Biology Ph.D. Program, Michigan State University, East Lansing, MI, 48824 4) Reproductive and Developmental Sciences Program, Michigan State University, East Lansing, MI, 48824 5) Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824 This chapter is modified from Moauro et al., published in Cellular Reprogramming on December 12, 2022. This study was supported by the National Institutes of Health R35 GM131759 and T32 HD087166. 163 Section 6.1. Abstract Somatic cell reprogramming was first developed to create induced pluripotent stem (iPS) cells. Since that time, the highly dynamic and heterogeneous nature of the reprogramming process has come to be appreciated. Remarkably, a distinct type of stem cell, called induced extraembryonic endoderm stem (iXEN) cells, also form during reprogramming of mouse somatic cells by ectopic expression of the transcription factors OCT4, SOX2, KLF4, and MYC (OSKM). The mechanisms leading somatic cells to adopt differing stem cell fates are challenging to resolve given that formation of either stem cell type is slow, stochastic, and rare. For these reasons, fluorescent gene expression reporters have provided an invaluable tool for revealing the path from the somatic state to pluripotency. However, no such reporters have been established for comparable studies of iXEN cell formation. In this study we examined the expression of multiple fluorescent reporters, including Nanog, Oct4, and the endodermal genes Gata4, and Gata6 – alone, and in combination, during reprogramming. We show that only the simultaneous evaluation of Nanog and Gata4 reliably distinguishes iPS and iXEN cell colonies during reprogramming. Section 6.2. Introduction Since the initial report that somatic cells can be reprogrammed to induced pluripotent stem (iPS) cells (Takahashi & Yamanaka, 2006), much interest has focused on how to distinguish iPS cell colonies from undesirable colonies, such as partially reprogrammed cell colonies (Buganim et al., 2012; E. M. Chan et al., 2009; Mikkelsen et al., 2008; Sridharan et al., 2009; Velychko et al., 2019). We and others have reported that cells 164 expressing endodermal genes arise during reprogramming of mouse and human somatic cells (Guan et al., 2022; He et al., 2020; Parenti et al., 2016; Schiebinger et al., 2019; Y. Zhao et al., 2015). Moreover, viral transduction of pluripotency transcription factors Oct4, Sox2, Klf4, and Myc (OSKM) leads to formation of stable induced extraembryonic endoderm (iXEN) stem cell lines in murine and canine fibroblasts (Nishimura et al., 2017; Parenti et al., 2016). Unlike partially reprogrammed cells, iXEN cells are capable of unlimited proliferation and lineage-specific differentiation, and therefore meet both criteria of authentic stem cell lines. However, little is known regarding the mechanisms that guide alternative reprogramming outcomes, nor how to distinguish iPS cells and iXEN cells during the reprogramming process. Fluorescent reporters, such as knock-in reporters of endogenous Oct4 or Nanog expression, are commonly used for quantifying reprogramming efficiencies (Brambrink et al., 2008; Buganim et al., 2012; Dos Santos et al., 2014; Huangfu et al., 2008; Judson et al., 2009; Pour et al., 2015; Y. Shi et al., 2008; Tsubooka et al., 2009; Xiao et al., 2016; X. Y. Zhao et al., 2009). In this application, the number or proportion of fluorescent colonies is often reported. However, the expression of these reporters has not been evaluated in the context of iXEN cells. This seems important especially given that Oct4 is associated with both pluripotent and extraembryonic endodermal lineages during embryogenesis (Frum et al., 2013; Le Bin et al., 2014; Palmieri et al., 1994) and is a component of the transcription factor cocktails that produce iXEN cells (Nishimura et al., 2017; Parenti et al., 2016). Conversely, a fluorescent reporter that specifically labels iXEN cell colonies during reprogramming has not been identified. 165 The identification of fluorescent reporters to reliably distinguish iPS and iXEN cell colonies would enable new approaches to the discovery of the mechanisms underlying the reprogramming process. To identify specific fluorescent reporters requires a more systematic evaluation of the expression and co-expression of pluripotency and endodermal gene expression reporters over the course of somatic cell reprogramming. We therefore evaluated the expression dynamics of fluorescent reporters of the transcription factors important for pluripotency (OCT4 and NANOG) and extraembryonic endoderm (GATA6 and GATA4), in conjunction with colony morphology, during mouse somatic cell reprogramming. We chose to focus on these transcription factors because they are all involved in the earliest stages of segregating pluripotent and endodermal cell fates during development (Artus & Hadjantonakis, 2012; Bassalert et al., 2018; Frum & Ralston, 2015). Section 6.3. Materials and Methods Mouse Lines All animal research was conducted in accordance with the guidelines of the Michigan State University Institutional Animal Care and Use Committee. The following alleles were maintained in a CD-1 background: Gata4H2B-eGFP (Simon et al., 2018), Gata6tm1Hadj (Freyer et al., 2015), Pou5f1tm2Jae (Lengner et al., 2007), and NanogmCherry (reported here). The NanogmCherry mouse was created via a CRISPR/Cas9-mediated knock-in of mCherry immediately downstream of, and in frame with, the Nanog coding region using 166 a published donor plasmid (Addgene 59995) (H. Yang et al., 2013). CAS9 protein and guide RNA (5’CGTAAGTCTCATATTTCACC-3’) (50-200 ng/µl), and circular donor plasmid (5-20 ng/µl) were injected into C57Bl/6 zygotes, which were then transferred to pseudo-pregnant female mice. Offspring were then screened by Nanog locus-specific PCR ('5-TACCACCATGCCAGGCTGAGAATGT-3' and 5'- TCAACAGGGAGAAGTTAGTGGCGCT-3'). Using this approach, a 2,105 bp PCR product can only be generated if homologous recombination has occurred between the donor plasmid's upstream homology arm and the Nanog locus and 9 of 22 offspring were positive using this criterion, for an overall knock-in efficiency of ~40%. The Nanog locus of a single founder mouse was confirmed by sequencing. The NanogmCherry mouse line was maintained in a CD-1 background. Immunofluorescence and Confocal Microscopy Mice carrying NanogmCherry were naturally mated, and then embryos were obtained from pregnant moms around embryonic day (E) E3.75. Embryos were then processed for immunofluorescence as previously described (O’Hagan et al., 2021), using the following reagents: goat-anti-SOX17 (1:2000, R&D, AF1924), goat-anti-SOX2 (1:2000, Neuromics, GT15098), donkey-anti-goat Alexa488 (1:400, Invitrogen, A-11055), and DRAQ5 (1:400, Cell Signaling Technology, 4084). Embryos were imaged using an Olympus FluoView FV1000 Confocal Laser Scanning Microscope system with a 20x UPlanFLN objective (0.5 NA) and 3x digital zoom. For each embryo, z-stacks were collected with 5-μm intervals between optical sections. Optical sections are displayed as an intensity projection over the z-axis. 167 Cell lines and cellular reprogramming R1 ES cell lines (ATCC, SCRC-1011) and XEN cell lines (derived in-house from CD-1 blastocysts) were cultured in the presence of MEFs and Leukemia Inhibitory Factor (LIF) as previously described (Blij et al., 2015; Moauro & Ralston, 2022; Parenti et al., 2016). To derive MEF lines, mice carrying one or more fluorescent reporter alleles were naturally mated, and then embryos were collected from pregnant females at E13.5. MEF lines were established and genotyped from individual E13.5 embryos as previously described (Moauro & Ralston, 2022). Cellular reprogramming was achieved using the modified Moloney Murine Leukemia Virus (MMLV) retrovirus as previously described (Moauro & Ralston, 2022). pMXs plasmids expressing Oct4, Klf4, Sox2 or Myc cDNAs were obtained from Addgene (13366, 13367, 13370, and 13375). Throughout reprogramming, expression of fluorescent markers was detected using a Lumen Prior 200 camera and Leica microscope. Our reprogramming efficiency ranged from 0.44- 0.63% +/- 0.025% (total colonies per MEFs plated). The Nanog-mCherry knock-in ES cell line was generated by electroporation of ES cells with a plasmid carrying Nanog-2A-mCherry (Addgene, p59995) and pSpCas9(BB)-2A- Puro (Addgene, pX459), which was modified to carry the Nanog guide RNA and then cultured in the presence of 1.25 µg/mL puromycin (Gibco A1113803). Fluorescent colonies were isolated, expanded, and then PCR screened as described above. 168 RNA isolation and qPCR RNA was harvested using 1:6 chloroform to Trizol (Invitrogen), and then 1 μg RNA was reverse transcribed to create cDNA using QuantiTect Reverse Transcription Kit (Qiagen), following manufacturer instructions. For qPCR, cDNA was amplified in quadruplicate using transcript-specific primers (Table 6.1) and quantified on Lightcycler 480 (Roche). Primer amplification efficiencies were determined empirically by generating a standard curve using extraembryonic endoderm (XEN) cells and embryonic stem (ES) cell cDNA libraries. Section 6.4. Results The making of a Nanog-2A-mCherry fluorescent reporter mouse line Our goal was to determine whether Nanog, together with other reporters, reliably distinguish iPS and iXEN cells during reprogramming (Fig. 6.1A). Because only GFP- like reporters of many different genes are available, we first created a new Nanog- mCherry mouse line (Supp. Fig. 6.1) that would enable the simultaneous evaluation of multiple fluorescent reporters. To evaluate the specificity of this reporter for the pluripotent lineage, we examined blastocysts carrying Nanog-mCherry at embryonic day E3.75. At this stage, embryos possess pluripotent and non-pluripotent cell types, including the pluripotent epiblast and the primitive endoderm (progenitors of XEN cells) (Chazaud et al., 2006; Kunath et al., 2005). Indeed, we observed expression of NANOG-mCherry only in epiblast cells, and not within the non-pluripotent cell types (Supp. Fig. 6.1C). Moreover, NANOG-mCherry was detected in embryonic stem (ES) 169 cells, but not in differentiated ES cells (Supp. Fig. 6.1D). We therefore conclude that the NanogmCherry allele is a faithful reporter of endogenous Nanog expression. Nanog-mCherry and Oct4-eGFP do not reliably identify iPS cell colonies Next, we evaluated the expression of NANOG-mCherry during reprogramming. We reprogrammed mouse embryonic fibroblasts (MEFs) carrying one allele of Nanog- mCherry by retroviral delivery of OSKM, and then tracked colony morphology and fluorescence. During reprogramming, presumptive iPS cell colonies appear as round, compact colonies with smooth borders (Meissner et al., 2007). Using these criteria, we first determined that around 20% of all observable colonies possessed morphological features of iPS cell colonies staring around day 8, and at three-day intervals until day 20 (Fig. 6.1B). We next quantified the proportion of colonies expressing NANOG-mCherry, with the expectation that if Nanog is a specific reporter of iPS cell colonies, then the proportion of fluorescent colonies should be equivalent to the proportion of morphologically-apparent iPS cell colonies. However, at several points, the proportion of NANOG-mCherry-expressing colonies appeared to be greater than the proportion of colonies that morphologically resembled iPS cell colonies (Fig. 6.1B), suggesting that Nanog is expressed in non-iPS cell colonies during reprogramming. We therefore evaluated the expression of NANOG-mCherry in non-iPS cell colonies, including iXEN colonies. As we previously reported, presumptive iXEN colonies appear flatter and more spread, with less defined borders than iPS cells (Moauro & Ralston, 2022; Parenti et al., 2016). 170 Additionally, we observed a third type of colony that displayed morphological features of both iPS and iXEN cell colonies, which we termed Mixed colonies. Mixed colonies possessed domed, smooth-edged, iPS cell-like clusters, which cascaded into flatter, rough-edged, iXEN cell-like colonies (Fig. 6.1C). Consistent with our prior observations, we observed that NANOG-mCherry expression was not always restricted to presumptive iPS cell colonies, but was also observed in presumptive iXEN and Mixed colonies (Fig. 6.1C-D) during reprogramming. However, NANOG-mCherry was expressed evenly throughout presumptive iPS cell colonies, while NANOG-mCherry was observed in diffuse patches of presumptive iXEN cell and Mixed colonies (Fig. 6.1D). Moreover, expression of NANOG-mCherry was largely restricted to iPS cell-like regions of the Mixed colonies during reprogramming. To evaluate the expression of NANOG-mCherry within stable iPS and iXEN cell lines, we manually picked presumptive iPS and iXEN colonies and derived stable cell lines as previously described (Parenti et al., 2016; Takahashi & Yamanaka, 2006). Ultimately, iPS cell lines expressed NANOG-mCherry, while iXEN cell lines did not (Fig. 6.1E). These stem cell lines also expressed appropriate markers (Fig. 6.1F), authenticating their identities as iPS or iXEN cell lines, and confirming the validity of our morphological criteria for selecting stem cell colony subtypes. Taken together, these observations indicate that while Nanog-mCherry is detected in established iPS cell lines, it is also detected in cells that do not appear to be part of iPS cell colonies during reprogramming. This observation underscores the importance of using both morphology and fluorescence to identify emerging iPS cell colonies. 171 Finally, we asked whether the combined use of two pluripotency reporters could more reliably identify iPS cell colonies during reprograming than NanogmCherry alone. We reprogrammed MEFs carrying alleles of both Oct4-eGFP (Lengner et al., 2007), and Nanog-mCherry as before. Overall, we observed concordance between the proportion of morphologically apparent iPS cells and eGFP/mCherry double-positive colonies (Fig. 6.2A). However, closer examination revealed that double-positive cells were present in all three colony types (Fig. 6.2B-C). Interestingly, OCT4-eGFP and NANOG-mCherry appeared to be coexpressed within individual cells of the double-positive colonies (Fig. 6.2B). Our observations indicate that these pluripotency markers label cells that are not part of presumptive iPS cells colonies and/or that colony subtypes are potentially highly heterogeneous. Gata6-H2B-Venus is expressed in multiple colony subtypes during reprogramming Because our previous results suggested that pluripotency reporters alone are not sufficient to distinguish iPS and iXEN cell colonies during reprogramming, we next sought to identify a reliable reporter for iXEN cell fate. In the embryo, GATA6 is considered to be one of the earliest-acting regulators of extraembryonic endoderm development (Artus et al., 2011; Chazaud et al., 2006; Koutsourakis et al., 1999; Morrisey et al., 1996; Schrode et al., 2014). Additionally, GATA6 is sufficient to induce an extraembryonic endoderm phenotype in ES cells (Capo-Chichi et al., 2005; Fujikura et al., 2002; Shimosato et al., 2007; Wamaitha et al., 2015). These observations suggest that GATA6 could be a marker of XEN cell fate during somatic cell 172 reprogramming. A mouse knock-in line carrying Gata6-H2B-Venus has been established, and Venus was observed within extraembryonic endoderm lineages (Freyer et al., 2015). We therefore evaluated the expression of GATA6-H2B-Venus during reprogramming of MEFs carrying Gata6-H2B-Venus. We began by quantifying the proportion of presumptive iXEN colonies present throughout reprogramming, using morphological criteria defined above (Fig. 6.3A). Next, we quantified the proportion of all colonies in which GATA6-H2B-Venus was detected. Starting around day 14, we observed expression of GATA6-H2B-Venus in 10-20% of all colonies (Fig. 6.3A), suggesting that not all presumptive iXEN cell colonies express GATA6-H2B-Venus. We next evaluated the expression of GATA6-H2B-Venus within iPS, Mixed, and iXEN cell colonies. Surprisingly, GATA6-H2B-Venus was expressed diffusely in all three colony types (iPS, iXEN and Mixed), localizing to subsets of cells within each colony subtype (Fig. 6.3B-C). To test whether GATA6-H2B-Venus eventually becomes restricted to iXEN cell lines, presumptive iPS and iXEN cell colonies were picked and passaged to create stable cell lines. After passaging, all cell lines maintained appropriate morphologies (Fig. 6.3D) and expression of key lineage-determining genes (Fig. 6.3E). Notably, GATA6-H2B-Venus was only expressed in some iXEN cell lines, despite qPCR evidence of Gata6 expression (Fig. 6.3D-E). These observations are consistent with the observation that Gata6-H2B-Venus was not detected in all extraembryonic endoderm cells during development (Freyer et al., 2015). For these 173 reasons, we conclude that Gata6-H2B-Venus may not be useful for distinguishing stem cell colony subtypes during reprogramming. Gata4-H2B-eGFP is expressed in iXEN and not iPS cell colonies Like GATA6, the closely-related factor GATA4 is also thought to play an essential and instructional role in extraembryonic endoderm development in vivo and in ES cell lines (Artus et al., 2011; Capo-Chichi et al., 2005; Fujikura et al., 2002; Kuo et al., 1997; Molkentin et al., 1997; Shimosato et al., 2007). The Gata4-H2B-eGFP reporter has been shown to faithfully recapitulate expression of Gata4 during development (Simon et al., 2018). During reprogramming of MEFs carrying the Gata4-H2B-eGFP allele, we observed expression of GATA4-H2B-eGFP in very few colonies (Fig. 6.4A). However, GATA4-H2B-eGFP was only detected in presumptive iXEN and Mixed colonies, and not in iPS cell colonies (Fig. 6.4B-C), indicating that GATA4-H2B-eGFP may be more informative than GATA6-H2B-Venus for identifying presumptive iXEN cells during reprogramming. However, like GATA6-H2B-Venus, GATA4-H2B-eGFP was detected non-uniformly throughout presumptive iXEN and Mixed colonies, raising questions about the fidelity of this marker and the identity of the GATA4-H2B-eGFP-positive cells within these colonies. To investigate this further, presumptive iXEN colonies were picked and passaged to create stable cell lines. Encouragingly, all iXEN cell lines expressed GATA4-H2B-eGFP in conjunction with appropriate morphology and gene expression (Fig. 6.4D-E). Moreover, all iXEN cell lines carrying an allele of Gata4-H2B-eGFP 174 expressed GATA4-H2B-eGFP by passage 5, regardless of whether Gata4 had been expressed initially, unlike iXEN cell lines carrying an allele of Gata6-H2B-Venus (Fig. 6.4F). We conclude that Gata4-H2B-Venus is expressed in iXEN and not iPS cell colonies. We next sought to examine the expression of GATA4-H2B-eGFP and NANOG-mCherry simultaneously. As expected, based on their individual expression patterns, these two reporters were co-expressed in very few colonies during reprogramming (Fig. 6.4G). Strikingly, however, only Mixed colonies co-expressed both GATA4-H2B-eGFP and NANOG-mCherry (Fig. 6.4H). In addition, these two reporters exhibited complementary expression patterns within the Mixed colonies, consistent with cell type-specific expression of these two genes in embryos (Artus et al., 2011; Chazaud et al., 2006; Niakan & Eggan, 2013). We therefore propose that the combined use of GATA4-H2B- eGFP and NANOG-mCherry can be used to resolve specific stem cell colony subtypes during reprogramming. Section 6.5. Discussion Although transcription factor-mediated somatic cell reprogramming has been possible for over a decade, we still have only a rough understanding of the molecular events that each cell undergoes as it acquires iPS or iXEN cell fates. As single cell genomic approaches become more commonplace, we are beginning to understand the stages of the reprogramming process. However, most genomic approaches do not permit the long-term evaluation of cellular changes over time in real time. For these reasons, 175 fluorescent reporters, which permit live imaging, individual cell tracking, and readout of gene expression, are an appealing complement to genomic studies of somatic cell reprogramming. We have identified a combination of two mouse gene expression fluorescent reporter lines that can reliably distinguish colonies of presumptive iPS and iXEN cells. One limitation of this approach is the need to derive MEF lines from mice carrying multiple distinct reporter alleles. However, the reliable identification of early iXEN and iPS cell lines will lead to discovery of more facile approaches for cell isolation, such as unique cell surface markers that label live iXEN and iPS cells early in their formation. Nevertheless, the ability to select cells on the basis of the expression of potent developmental regulators of pluripotent and extraembryonic endoderm cell fate provides additional advantages. First, we note that, while Gata4 was detected within presumptive iXEN cell colonies, it did not appear to be expressed in all cells of those colonies. The reasons for the limited expression of Gata4 within iXEN cell colonies is not yet clear. One possibility is that Gata4 expression is dynamically regulated in nascent iXEN cells, giving the illusion, in snapshots, that Gata4 is only expressed in a subset of iXEN cells. Similarly, expression of Gata4 may be progressive, consistent with the gradual adoption of iXEN cell fate. Another possibility is that the Gata4 reporter labels a unique subtype of iXEN cells. These possibilities could be investigated using live imaging and fluorescent cell sorting in future studies. 176 Second, we were surprised that the Nanog reporter was detected within non-iPS cell colony subtypes, including iXEN and Mixed colonies. While this pattern could raise some concern for the fidelity of Nanog as a marker for the emergence of pluripotency, we do not yet understand the dynamics of Nanog expression outside of iPS cell colonies. One possibility is that Nanog is expressed very transiently in most cells that undergo reprogramming – regardless of their ultimate fates. Another possibility is that bona fide, Nanog-expressing iPS cells arise within Mixed and iXEN cell colonies. These possibilities could again be addressed using live imaging and fluorescent cell sorting in future studies, which could reveal whether Nanog expression is stable or transient and whether Nanog-expressing cells give rise to iPS, iXEN, Mixed, or failed cell lines. Lastly, we observed a significant number of Mixed colonies in all of our reprogramming experiments. These colonies could be considered to have failed or stalled during reprogramming. However, it is intriguing that Mixed colonies possess characteristics of both iPS and iXEN colonies, including morphology and fluorescent marker expression. Therefore, is tempting to speculate that Mixed colonies also give rise to bona fide iPS and iXEN cells. However, the quality and properties of stem cell lines that can be derived specifically from the Mixed colonies awaits further investigation. Similarly, these reporters can be used to determine when and how iXEN and iPS cell fates first diverge from each other in future studies because they will enable identification of differences in transcriptional signature and developmental potential in single-cell studies. Ultimately, the use of fluorescent gene expression reporters will likely enable us to address these and other exciting questions about how and why reprogramming works. 177 Section 6.6. Acknowledgements We would like to thank Anna-Katerina Hadjantonakis and her lab for providing Gata4H2B-eGFP and Gata6tm1Hadj/J mice and the Michigan State University Transgenic Core for assistance creating the Nanog-mCherry mouse line. 178 Gene Target Forward Primer Oct4 GTTGGAGAAGGTGGAACCAA CCAAGGTGATCCTCTTCTGC TABLES Reverse Primer Nanog Sox2 Gata6 Gata4 Sox7 Sox17 ActinB ATGCCTGCAGTTTTTCATCC GAGGCAGGTCTTCAGAGGAA GCGGAGTGGAAACTTTTGTCC CGGGAAGCGTGTACTTATCCTT ATGCTTGCGGGCTCTATATG GGTTTTCGTTTCCTGGTTTG CTGGAAGACACCCCAATCTC ACAGCGTGGTGGTGGTAGT GGCCAAGGATGAGAGGAAAC TCTGCCTCATCCACATAGGG CTTTATGGTGTGGGCCAAAG GCTTCTCTGCCAAGGTCAAC CTGAACCCTAAGGCCAACC CCAGAGGCATACAGGGACAG Table 6.1. Quantitative Polymerase Chain Reaction Primers for Detecting Endogenous Transcripts for Chapter 6. Gene Target NanogmCherry mCherry Pou5f1eGFP Pou5f1 wild-type allele Gata6H2B-Venus Gata6 wild-type allele Gata4H2B-eGFP Gata4 wild-type allele Forward Primer CCACTAGGGAAAGCCAT GCGCATTT AGGACGGCGAGTTCATC TAC CCAAAAGACGGCAATAT GGT TGCCAGACAATGGCTAT GAG CCAGGGAGCTCTGAGA AAAAG CCAGGGAGCTCTGAGA AAAAG GTTTCTGCTTTGATGCT GGA GTTTCTGCTTTGATGCT GGA Reverse Primer GGAAGAAGGAAGGAAC CTGGCTTTGC TGGTGTAGTCCTCGTTG TGG CAAGGCAAGGGAGGTA GACA CAAGGCAAGGGAGGTA GACA CCTTAGTCACCGCCTTC TTG GTCAGTGAAGAGCAACA GGT TGCTCAGGTAGTGGTTG T CGGAGTGGGCACGTAG AC Table 6.2. Genotyping Primers for Chapter 6. 179 FIGURES Figure 6.1. NANOG-mCherry expression is detected in colonies bearing non-iPS cell colony morphology during reprogramming. 180 Figure 6.1 (cont’d). A) Workflow for all experiments described in this study, including the derivation of MEF lines carrying one or more fluorescent reporter, evaluation of colonies during the 21-day reprogramming process, and isolation of colonies at the end of reprogramming to derive stable stem cell lines. B) The proportion of iPS cell colonies is not equivalent to the proportion of mCherry-positive colonies during days 11–17 of reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). C) Representative images of NANOG-mCherry expression among colony subtypes during reprogramming (scale bar = 200 µm). D) Expression of NANOG-mCherry is observed in all three colony subtypes throughout reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). E) Expression of NANOG-mCherry expression is observed in established iPS, and not iXEN, cell lines (scale bar = 200 µm). F) Heat map summary of qPCR analysis of established iXEN and iPS cell lines (n = 3 biological replicates each), relative to positive control cell lines: XEN or ES cell lines (n = 3 replicate wells per cell line), confirms expected expression patterns of key lineage markers. Scale bar = 200 µm. ES, embryonic stem; iPS, induced pluripotent stem; iXEN, induced extraembryonic endoderm; MEF, mouse embryonic fibroblast; qPCR, quantitative polymerase chain reaction; XEN, extraembryonic endoderm. 181 Figure 6.2. NANOG-mCherry and OCT4-eGFP are coexpressed in multiple colony subtypes during reprogramming. A) The proportion of iPS cell colonies is roughly equivalent to the proportion of NANOG-mCherry and OCT4-eGFP double-positive colonies during reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). B) Expression of NANOG-mCherry and OCT4-eGFP is observed in all three colony subtypes during reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). C) Representative images of NANOG-mCherry and OCT4-eGFP expression during reprogramming, scale bar = 200 µm. 182 Figure 6.3. GATA6-H2B-Venus expression is detected in colonies bearing non- iXEN cell colony morphology during reprogramming. A) The proportion of GATA6- H2B-Venus-positive colonies is low relative to the proportion of colonies exhibiting iXEN cell morphology during reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). B) Expression of GATA6-H2B-Venus is observed in all three colony subtypes during reprogramming, scale bar = 200 µm. C) Expression of GATA6-H2B- Venus is observed in all three colony subtypes during reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). D) Expression of GATA6-H2B-Venus is restricted to established iXEN, and not iPS, cell lines, but is not expressed in all iXEN lines. 183 Figure 6.3 (cont’d). E) Heat map summary of qPCR analysis of established iXEN and iPS cell lines (n = 3 biological replicates each), relative to positive control cell lines: XEN or ES cell lines (n = 3 replicate wells per cell line), confirms expected expression patterns of key lineage markers. 184 Figure 6.4. GATA4-H2B-eGFP, in combination with NANOG-mCherry, distinguishes iPS and iXEN cell colonies during reprogramming. 185 Figure 6.4 (cont’d). A) The proportion of GATA4-H2B-eGFP-positive colonies is low relative to the proportion of iXEN cell colonies observed during reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). B) Expression of GATA4B-H2B-eGFP is observed in iXEN and Mixed, but not iPS, cell colonies during reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). C) Expression of GATA4B-H2B- eGFP is observed in iXEN and Mixed, but not iPS, cell colonies during reprogramming, scale bar = 200 µm. D) Expression of GATA4B-H2B-eGFP is observed in established iXEN, and not iPS, cell lines, scale bar = 200 µm. E) Heat map summary of qPCR analysis of established iXEN and iPS cell lines (n = 3 biological replicates each), relative to positive control cell lines: XEN or ES cell lines (n = 3 replicate wells per cell line), confirms expected expression patterns of key lineage markers. F) All established iXEN cell lines carrying the Gata4-H2B-eGFP allele expressed eGFP, whereas only half of the iXEN cell lines carrying Gata6-H2B-Venus expressed Venus, error bar = standard error (n = 6 and 8 iXEN cell lines, respectively). G) Percentage of all colonies in which expression of both NANOG-mCherry and GATA4-H2B-eGFP was observed during reprogramming, error bar = standard error (n = 3 MEF lines reprogrammed). H) Expression of both NANOG-mCherry and GATA4-H2B-eGFP is observed only in Mixed colonies, scale bar = 200 µm. 186 Supplemental Figure 6.1. Genetic engineering and characterization of Nanog- mCherry knock-in in early embryos and ES cell lines. 187 Supplemental Figure 6.1 (cont’d). A) Targeting scheme to generate mice expressing a NANOG-mCherry fusion protein. Schematic shows mCherry inserted in frame near the stop codon (magenta TGA) of the Nanog locus. The protospacer-adjacent motif (PAM) sequence (green GGT) and the sgRNA target sequence (purple) are also highlighted. Arrows represent the genotyping primer locations. B) Genotypes of ear punches of wild-type and Nanog-mCherry/+ distinguished by PCR using locus-specific primers (as shown in panel A and Table 6.2). C) Confocal cross-sections of Nanog-mCherry and wild type E3.75 blastocysts immunofluorescently stained with antibodies specific for pluripotent epiblast (SOX2) or primitive endoderm (SOX17) cells, demonstrate that SOX2 and NANOG are coexpressed (yellow arrows), while SOX17 and NANOG are complementary in expression. Scale bar = 25 µm. D) Expression of Nanog-mCherry in undifferentiated mouse ES cells and in ES cells following withdrawal of MEFs and LIF. Scale bar = 100 µm. 188 CHAPTER 7. IDENTIFICATION OF OCT4 BINDING SITES IN ICM CELLS BY uliCUT&RUN Robin E. Kruger1,2, Sarah Hainer3, and Amy Ralston2,4 1) Cell and Molecular Biology Ph.D. Program, Michigan State University, East Lansing, MI, 48824, USA 2) Reproductive and Developmental Sciences Training Program, Michigan State University, East Lansing, MI, 48824, USA 3) Department of Biological Sciences, University of Pittsburgh, Pittsburgh, PA, 15260, USA 4) Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA This study was supported by the National Institutes of Health R35 GM131759 and T32 HD087166. 189 Section 7.1. Abstract The transcription factor OCT4 is necessary for the specification of both epiblast (EPI) cells and primitive endoderm (PrE) cells in the inner cell mass of the mouse blastocyst. However, it is unknown how the same transcription factor promotes the emergence of a pluripotent cell type and a non-pluripotent cell type in the same tissue. I hypothesize that OCT4 binds different cis-regulatory elements in these two different cell types. To investigate OCT4 binding sites in EPI and PrE cells, I performed ultra-low-input CUT&RUN for OCT4 in blastocyst-stage embryos enriched for either EPI or PrE cells. I was able to identify unique peaks of OCT4 binding locations in EPI-enriched and PrE- enriched blastocysts, demonstrating the effectiveness of this technique to identify transcription factor binding sites in samples with limited genetic material. Section 7.2. Introduction The transcription factor OCT4 is generally considered to be a marker of pluripotency (Frum et al., 2013; Le Bin et al., 2014; Palmieri et al., 1994). OCT4 is expressed in early mouse embryos beginning at the 8-cell stage, and becomes restricted to the pluripotent inner cell mass after blastocyst formation (Palmieri et al., 1994). Oct4-knockout embryos die at ~E4.5 with severe inner cell mass (ICM) defects (Nichols et al., 1998). It is also considered one of the key factors that maintain pluripotency in embryonic stem cells, forming a network with NANOG and SOX2 (M. Li & Belmonte, 2017; Masui et al., 2007; Mitsui et al., 2003). When ESCs are differentiated and lose their pluripotency, OCT4 is rapidly downregulated. Altogether, these observations demonstrate that this transcription factor plays a key role in maintaining pluripotent cells. 190 However, more recent studies have shown that OCT4 does not solely promote pluripotency in preimplantation mouse embryos. Rather, OCT4 is essential for the specification of a non-pluripotent cell type, primitive endoderm (PrE). PrE cells are specified in the second cell fate decision, when the ICM differentiates into pluripotent epiblast (EPI), which forms the fetus, and multipotent PrE, which contributes mostly to the extraembryonic yolk sac (Cockburn & Rossant, 2010, see Chapter 1). Interestingly, knockouts of Oct4 do not express PrE markers such as Sox17 (Frum et al., 2013; Le Bin et al., 2014), and reprogrammed XEN cells, the in vitro stem cell model of PrE, also express OCT4 (Moauro et al., 2022). Therefore, it appears that OCT4 promotes both EPI and PrE cell fate in the ICM. How a single transcription factor can promote two different cell fates within the same tissue remains an open question. Genomic analyses are capable of providing extremely rich information about transcription and protein-DNA interactions down to the single cell level (Wen & Tang, 2022). Knowing where a transcription factor or chromatin modifier is bound on the DNA – be that on a promotor, enhancer, or silencer sequence – can help us build gene regulatory networks to determine how trans-acting factors ultimately affect the actions of a cell. Chromatin immunoprecipitation followed by sequencing (ChIP-seq) is a commonly used technique to map transcription factor binding sites. However, this technique has some shortcomings (Dahl & Gilfillan, 2018). ChIP-seq and its variations generally require protein crosslinking, which may introduce artifacts in the data (Baranello et al., 2016). More importantly, ChIP-seq involves shearing the entire genomic chromatin and the precipitating out the bound chromatin, which leads to loss of 191 material and a high background. For this reason, ChIP-seq techniques are not effective or reproducible with small (<10,000 cells) or highly heterogeneous samples (Hainer et al., 2019; Jakobsen et al., 2015). As preimplantation mouse embryos are typically fewer than 100 cells, the inefficiency of ChIP-seq based techniques has thus far prevented effective mapping of transcription factor binding sites in these rare samples. As an alternative to ChIP-seq, Cleavage Under Targets and Release Using Nuclease (CUT&RUN) uses a different approach that is far more compatible with small sample sizes (Skene & Henikoff, 2017). In CUT&RUN, an antibody against the protein-of- interest first detects the protein bound to the DNA. A fusion protein A-micrococcal nuclease (MNase) binds to the antibodies and the nuclease cleaves the chromatin precisely on either side of the bound protein. Once released from the protein complex, these short DNA fragments go into solution while the bulk chromatin remains insoluble, allowing it to be easily removed. This drastically improves the signal-to-noise ratio, decreasing the requirement for high reads and large samples to overcome the background. An adaptation to this method, called ultra-low-input (uli) CUT&RUN, has been shown to be effective in profiling transcription factor binding sites specifically in preimplantation mouse embryos, even at the single-cell level (Hainer et al., 2019; Hainer & Fazzio, 2019; Patty & Hainer, 2021). Here, I used uliCUT&RUN to generate a dataset of OCT4 binding sites in mid-blastocyst stage mouse embryos. To compare OCT4 binding sites between EPI and PrE cells, I pooled embryos enriched for either EPI or PrE cells in the ICM. These two pools of 192 embryos produced different lists of OCT4-bound genes, providing evidence for the hypothesis that OCT4 regulates different targets in EPI and PrE cells. Section 7.3. Materials and Methods Mouse Strains and Genotyping All animal research was conducted in accordance with the guidelines of the Michigan State University Institutional Animal Care and Use Committee under approved protocol 202300108. Wild type embryos were obtained from CD-1 mice (Charles River). Mice were maintained on a 12 hour light/dark cycle. Embryo Collection and Culture Preimplantation (E2.5 or E3.75) embryos were collected by flushing the oviduct or uterus with M2 medium (Sigma M7167). For embryo culture, KSOM medium (Millipore MR-121-D) was equilibrated overnight prior to embryo collection. E2.75 embryos were collected and cultured at 37°C in a 5% CO2 incubator under light mineral oil (Millipore ES-005-C). To enrich for PrE cells, 1 µg/mL recombinant FGF4 in PBS with 0.1% BSA (R&D 235-F4) + 1 µg/mL heparin (Sigma H3149) was added to the culture media. To enrich for EPI cells, 1 µM PD173074 in DMSO (Selleckchem S1264) + 5 µM PD0325901 in DMSO (Stemgent 04-0006) was added to the culture media. 0.2% DMSO (New England BioLabs B0515A) was added to the culture media in control samples. 193 Immunofluorescence and Confocal Microscopy Preimplantation embryos were fixed with 4% formaldehyde (Polysciences 04018) for 10 minutes, permeabilized with 0.5% Triton X-100 (Sigma Aldrich X100) for 30 minutes, and then blocked with blocking solution (10% Fetal Bovine Serum (HyClone SH30396.02), 0.1% Triton X-100) overnight at 4°C. Embryos were incubated with primary antibody overnight at 4°C. The next day, embryos were washed in blocking solution for 30 minutes, incubated in secondary antibody diluted in blocking solution for 1 hour, washed in blocking solution for 30 minutes, then stained with nuclear stain diluted in block for 10 minutes or overnight. Antibodies used are listed in Table 1. Embryos were imaged using an Olympus FluoView FV1000 Confocal Laser Scanning Microscope system with 60X PlanApoN oil (NA 1.42) objective. For each embryo, z- stacks were collected, with 5 µm intervals between optical sections. Embryo Analysis For each embryo, z-stacks were analyzed using Fiji (ImageJ), which enabled the labeling, based on DNA stain, of all individual cell nuclei. Statistical analysis was performed using GraphPad Prism (v. 9.5.1). Figure images were assembled using Adobe Illustrator. Embryo Preparation for uliCUT&RUN Wild-type mouse embryos were collected at E3.75 or collected at E2.75 and cultured 36 hours in the treatment conditions described above. The zona pellucida was removed by Acid Tyrode’s treatment. Embryos were transferred into Nuclear Extraction buffer (20 194 mM HEPES-KOH, pH 7.9; 10 mM KCl; 0.5 mM Spermidine; 0.1% Triton X-100; 20% glycerol; 1X HALT protease inhibitors (ThermoFisher 78439)) and incubated on ice for 10 minutes. Samples were then centrifuged at 600xg for 10 minutes at 4°C to pellet the nuclei. Once completed, the supernatant was removed. After nuclear extraction, samples were either flash-frozen in liquid nitrogen and stored at -80°C, or the nuclei were fixed for 10 minutes in formaldehyde solution (1.5 mL 37% formaldehyde, 0.1mL 5M NaCl, 0.01mL 0.5M EDTA, 0.25mL 1M HEPES 7.6, and up to 5mL water), then quenched in 0.25M glycine for 5 minutes. Crosslinked nuclei were stored in quenched fix at 4°C for up to 2 weeks. uliCUT&RUN on Embryo Nuclei Prepared samples were shipped on ice to Dr. Sarah Hainer, who performed uliCUT&RUN for CTCF and OCT4 as described in Hainer & Fazzio, 2019. Briefly, each sample of frozen or fixed nuclei was affixed to concanavalin A beads. The samples were then treated for 2 hours with a primary antibody against either CTCF or OCT4. One or more negative control samples were not treated with primary antibody (no Ab control) and washed. All samples were then treated with pA-MNase fusion protein and washed. The MNase activity was then activated by treatment with calcium chloride for 30 minutes. The DNA digestion was stopped by treatment with Stop buffer (see Hainer & Fazzio, 2019), including a known concentration of S. cerevisiae spike-in DNA as a positive control. The samples were incubated at 37 °C for 10 minutes to release the fragments of DNA into solution. After incubation, the samples were centrifuged at 16,000xg for 2 minutes, then placed on a magnetic rack to separate the intact 195 chromatin. The supernatant containing the enriched, fragmented samples was transferred to new microcentrifuge tubes. Then, in fixed samples, the crosslinking was reversed by treatment with 10% SDS and Proteinase K. Input chromatin was then fragmented by sonification and purified by DNA spin column. Sample library construction and multiplexing was performed using DNA Library Prep Kit for Illumina (ChIP-seq, CUT&RUN) with Multiplex Oligos for Illumina (ChIP-seq, CUT&RUN), according to the manufacturer’s protocol for CUT&RUN DNA. Before multiplexing, proper library construction was confirmed using 5 µL of each sample for analysis by agarose gel electrophoresis. Bioinformatic Analyses Bioinformatic analyses were performed as described in Hainer & Fazzio, 2019. Briefly, sample sequences were aligned to the mouse or S. cerevisiae genomes using bowtie2. Low-quality reads (MAPQ<10) and PCR duplicates were removed. The remaining aligned sequences were filtered for size between 1-120 bp. To assess peak enrichment over known genomic loci, deepTools was used to compare aligned sequences to known binding sites in embryonic stem cells for either CTCF (X. Chen et al., 2008) or OCT4 (Marson et al., 2008). To identify all binding sites of interest, peaks located within 2kb of transcriptional start sites were called using the algorithm Sparse Enrichment Analysis for CUT&RUN (SEACR, Meers et al., 2019). 196 Section 7.4. Results and Conclusions Our goal for these experiments was to map OCT4 binding sites in EPI and PrE cells in preimplantation mouse embryos. However, to optimize the protocol for isolating nuclei and performing uliCUT&RUN on embryos, we first tried mapping the binding sites for CTCF, using an antibody already validated for uliCUT&RUN (Hainer et al., 2019). To determine the optimal number of embryos per sample, we performed uliCUT&RUN on samples of E3.75 wild-type embryos from single embryos, pools of five embryos, and pools of ten embryos. The resulting peaks were tested for enrichment over known CTCF and OCT4 binding sites in embryonic stem cells (X. Chen et al., 2008; Marson et al., 2008). We were successful in using uliCUT&RUN to detect CTCF enrichment over known binding sites in 100% of samples from 5- or 10-embryo pools, but only 50% of samples from single embryos (Fig. 7.1A). This suggested that pools of 5 embryos would maximize success in later experiments. However, we were unable to map OCT4 binding sites in any samples using frozen nuclei, regardless of embryo number (Fig. 7.1B). We hypothesized that the failure to detect OCT4 binding sites may be due to thawing of flash-frozen nuclei during shipping; therefore, we repeated the experiment to map OCT4 binding sites in wild-type E3.75 embryos using fixed nuclei instead of frozen. We also tested the efficiency of two different anti-OCT4 antibodies. We were able to detect enrichment of OCT4 peaks over ESC OCT4-binding sites in all fixed samples, regardless of the antibody used (Fig. 7.2). Similar to our results from the first experiment, larger embryo pools resulted in the detection of more OCT4 binding sites, likely due to increased read depth (Fig. 7.2). This helped us determine that OCT4 197 binding sites could be mapped in preimplantation mouse embryos, and that crosslinking the nuclei would give us the best outcome for later experiments. Finally, we were able to examine OCT4 binding sites in EPI- and PrE-enriched embryos. Treatment with exogenous FGF4 promotes differentiation of the ICM to a PrE fate in mouse embryos, while inhibiting FGF signaling in mouse embryos promotes EPI cell fate (Yamanaka et al., 2010). I used this protocol to generate pools of embryos which were enriched for either EPI or PrE cells (see Methods). I performed immunofluorescence for markers of EPI and PrE on a separate group of treated embryos to confirm the effectiveness of the treatments (Fig. 7.3A). The Hainer lab performed uliCUT&RUN in triplicate on fixed nuclei from pools of 5 embryos from each treatment group. Both the FGF4-treated samples and FGF inhibitor-treated samples showed excellent peak enrichment over known OCT4 binding sites compared to no- antibody controls (Fig. 7.3B). In analysis of called peaks (see Methods), FGF4-treated embryos showed 170 total OCT4-binding peaks, while FGF-inhibited embryos showed 11,994 (Fig. 7.4B). Unfortunately, the no-treatment control samples had much sparser read depth and did not show high peak enrichment compared to no-antibody controls (Fig. 7.3B). This prevented more in-depth analysis of the called peaks, since there was no positive control to validate that the sites identified in the treated embryos were also bound by OCT4 in wild-type embryos. We were able to ascertain that all samples (even the poor quality control samples) showed peak profiling over the Pou5f1 promoter, the gene which encodes OCT4 (Fig. 7.4A). Since OCT4 is known to bind its own promoter in mouse embryonic stem cells (Chew et al., 2005; Loh et al., 2006), this provides some 198 quality control evidence that the uliCUT&RUN detected true OCT4-binding sites in the treated samples. The generation of this dataset shows that this technique can be used efficiently to detect transcription factor binding sites in preimplantation embryos. It also suggests that this dataset could be a useful starting point in helping to determine how OCT4 may regulate both EPI and PrE cell fate within the ICM. Future studies will want to optimize the protocol further to get a reliable high read depth to be able to test useful hypotheses from these data. 199 TABLES Antibody/Stain Source Identifiers Dilution Fluorophore Goat-anti-SOX17 R&D Systems AF1924 1:2000 Rabbit-anti-NANOG Reprocell RCAB002P-F 1:400 Donkey-anti-goat Invitrogen A-11055 1:400 Alexa488 IgG Donkey-anti-rabbit Jackson Immuno 711-165-152 1:400 Cy3 IgG Research DRAQ5 Cell Signaling 4084 1:400 Table 7.1. Antibody Table for Chapter 7. 200 FIGURES Figure 7.1. Pools of five embryos maximize successful uliCUT&RUN experiments. 201 Figure 7.1 (cont’d). A) uliCUT&RUN for CTCF was performed on nuclei extracted from wild-type E3.75 embryos and flash-frozen in liquid nitrogen. The heat maps show CTCF binding between 2kb upstream and 2kb downstream of CTCF binding sites mapped in embryonic stem cells (X. Chen et al., 2008). Graphs show average signal across the same region. A darker line in the center of the heat map shows peak enrichment over background at the known CTCF binding site. Heat maps display data from a representative single experiment performed in 2-4 replicates (No-antibody control (N=2); Single embryos (N=4); Pools of 5 embryos (N=2); Pools of 10 embryos (N=2)). No- antibody control represents background detected without enrichment from a primary antibody. 50% of single-embryo and 100% of pooled embryo experiments show enrichment over no-antibody control. B) uliCUT&RUN for OCT4 was performed on nuclei extracted from wild-type E3.75 embryos and flash-frozen in liquid nitrogen. Heat maps display data from representative single experiments of each type of sample (No- antibody control (N=1); Single embryos (N=2); Pools of 5 embryos (N=1); Pools of 10 embryos (N=1)). No experimental samples show enrichment over no-antibody control. 202 Figure 7.2. Crosslinking nuclei improves successful mapping of OCT4 binding sites. uliCUT&RUN for OCT4 was performed on nuclei extracted from wild-type E3.75 embryos and crosslinked. Heat maps represent single samples of each experiment (N=1 for all). All experimental samples show enrichment over no-antibody control. Inv=anti-OCT4 antibody from Invitrogen, Diag=anti-OCT4 antibody from Diagenode. 203 Figure 7.3. Detection of OCT4 binding sites in EPI-enriched and PrE-enriched blastocysts. 204 Figure 7.3 (cont’d). A) Wild-type embryos were collected at E2.75 and cultured for 36 hours in either unsupplemented media, media containing 1 μg/mL FGF4 + 1 μg/mL heparin, or media containing 1 μM FGFR inhibitor + 5 μM MEK inhibitor (2i, Yamanaka et al., 2010). Changes in ICM composition were confirmed by immunofluorescence for EPI marker NANOG and PrE marker SOX17. B) uliCUT&RUN for OCT4 was performed on nuclei extracted from pools of 5 embryos cultured according to the conditions in A and crosslinked. Heat maps present data from representative single experiments from each sample (No-antibody control (N=1); Control media embryo pools (N=3); FGF4-treated embryo pools (N=3); FGF inhibitor-treated embryo pools (N=3)). FGF4-treated and FGF inhibitor-treated samples show enrichment over no-antibody control. 205 Figure 7.4. Analysis of OCT4-bound peaks from EPI-enriched and PrE-enriched blastocysts. A) Profiles of peaks for each sample from Figure 7.3 over the Pou5f1 promoter region from the UCSC Genome Browser. All samples show expected enrichment on the Pou5f1 promoter. B) Number of OCT4-bound peaks in FGF4-treated and FGF inhibitor treated embryos, sorted by genome location type. C) Percentage of total peaks from B by genome location type. 206 CONCLUSION This work presents several advancements in the fields of embryology and stem cell biology (Fig. 8.1). Firstly, I identify novel regulatory mechanisms of embryonic development by intercellular molecular signaling. Conflicting reports in the literature have made the role of BMP signaling in mouse embryonic development unclear. My work shows that BMP signaling is not required for preimplantation development in mouse embryos. However, my studies uncovered a previously unknown role for SMAD4-mediated signaling immediately after implantation, in which SMAD4 attenuates FGF signaling to promote proper epiblast scaling and morphogenesis. This role for SMAD4 does not depend on BMP signaling activity; therefore, identification of the exact mechanism by which SMAD4 interacts with FGF signaling and how unknown factors downstream of these pathways regulate epiblast morphogenesis will be of particular interest in future studies. I also demonstrate that Hippo effectors YAP1 and WWTR1 are essential for maintaining blastomere polarization during preimplantation development, but that these factors are dispensable for initiating blastomere polarization. Future studies are needed to determine the mechanisms which regulate the timing of Hippo signaling dependence in blastomere polarization. Finally, I present three novel techniques for studying early embryonic development. The GOGREEN system provides an efficient technique for generating fluorescently-tagged proteins in early embryos. A combination of fluorescent reporters more accurately distinguished iPSC and iXEN colonies during cellular reprogramming. uliCUT&RUN can be used to detect transcription factor binding sites in preimplantation mouse embryos. Altogether, my work 207 addresses many previously outstanding questions in the field of mammalian embryology and provides new technical tools for future studies in this fascinating field of research. Figure 8.1. Summary of Findings. My work presents several advancements in the study of early embryonic development. I newly identify several molecular regulatory mechanisms in murine development at several early embryonic stages. 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