UNDERSTANDING THE KEYS TO INSIDE AND OUTSIDE CELL POSITIONING AND GENE EXPRESSION IN PREIMPLANTATION MOUSE EMBRYOS By Tayler Morgan Murphy A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Genetics – Doctor of Philosophy 2023 PUBLIC ABSTRACT Pregnancy has the seemingly impossible task of building a whole baby from a single cell. The embryo goes through innumerable incremental changes on its journey. Some of the most crucial steps of development occur before a person even knows they are pregnant. Defects in these crucial steps can lead to early miscarriage or severe birth defects. We can use mouse embryos as models for human embryo development, which allows us to better understand the genes and proteins that are crucial for normal development at these early stages and eventually possibly prevent early pregnancy loss. This dissertation focuses on the factors and signals that start and maintain development during the first and second cell fate decision when the mouse embryo has just 16 cells. Though the embryo has 16 cells it has only two cell types, inside and outside cells. We wanted to know what happens when a cell cannot decide if it is an inside or outside cell. We found that an on/off switch called HIPPO can help to resolve this cellular confusion. Furthermore, we studied embryos at the second cell fate decision, when the inside cells become two distinct groups. This leaves the embryo with three cell types total. We hoped to discover more about how two proteins that are expressed at this stage, OCT4 and GATA6 impact the development of these three cell types. We turned off OCT4 and GATA6 and evaluated the changes that occurred. We found that in addition to their known roles in promoting inside cell fate, they are actively preventing inside cells from becoming outside cells. Additionally, in the outside cells, OCT4 and GATA6 are required for early maintenance of outside cell factors, and prevention of inside cell factors. ABSTRACT Preimplantation embryo development is highly complex. Cells must manage changes to potency, cell positioning, and lineage specific gene expression. The embryo must first differentiate between inside and outside cells, where inside cells remain pluripotent (able to become any cell within the organism), and the outside cells lose potency to become multipotent stem cells (able to become only a few cell types). The differentiation of outside cells relies partially on the inactivation of the HIPPO signaling pathway that drives expression of the protein CDX2 in outside cells. Inside cells have active HIPPO signaling, maintain pluripotency and express SOX2. However, it is unknown how specific HIPPO pathway members like YAP1/WWTR1 regulate expression of these lineage- specific proteins. The second cell fate decision differentiates the inside cells into pluripotent epiblast and multipotent primitive endoderm. Transcription factor OCT4 has previously been shown to be necessary for development of both cell types. This dual requirement of OCT4 in two distinct lineages raises the question, how can OCT4 drive two cell different cell fates simultaneously. OCT4 has previously been shown to work with cofactors like SOX2, which is specific to pluripotent epiblast, so I hypothesized that OCT4 may also work with the primitive endoderm specific factor GATA6. My transcriptome analysis showed that OCT4 and GATA6 repress trophectoderm gene transcripts in the inner cell mass (ICM), which may help indirectly drive primitive endoderm fate. Additionally, I noted that loss of GATA6 and OCT4 caused reduced expression of trophectoderm specific factor CDX2 in early blastocyst and mid-blastocyst stages, respectively. Gata6 null mid-blastocyst stage embryos also exhibited prolonged OCT4 expression in the trophectoderm, indicating a novel and stage specific role for OCT4 and GATA6 in regulating trophectoderm gene expression. To my Grandma Joyce, who always encouraged my love of science and learning. I miss you. iv ACKNOWLEDGEMENTS I would like to say thank you to everyone that has encouraged me, embraced me, and pushed me through grad school. Family Firstly, to my beautiful wife Hannah. You are an incredible support system. You make me laugh often, even on hard days. Thank you for listening to me constantly talk about OCT4, GATA6, mouse embryos, copulatory plugs, and everything else. I couldn’t have done this without you. Logan, thank you for being the best brother anyone could ask for. You have always pushed me to be my best. Dad, you have always been my biggest supporter. Thank you so much for being endlessly encouraging, loving, and accepting. Mom, thank you for your encouragement and love throughout my whole life. And finally, thank you to my cats; Lupine, Cassia, and Oliveri, for being the best honorary co-writers and emotional support ever. Friends To my grad school cohort, thank you for the emotional support throughout the years, especially all the BMB 801 study sessions. Nicky, Alice, Ana-Maria, and Aiko thank you for the occasional friend dinners, and frequent in the hall convos. To my lab mates, past and present; Dr. Tristan Frum, Robert Fidis, Dan O’Hagan, Dr. Jennifer Watts, Dr. Alex Moauro, Dr. Shannon Walsh, Dr. Michael Halbisen, Robin Kruger, Marcelio Shammami, Farina Aziz, Ian McCrary, and Barb Makela, thank you for being wonderful and extremely supportive people. Thank you for all of your constructive feedback on papers, posters and presentations over the years. You have all played an important role in my development as a scientist. Additionally, to Angie, Willow, Becca, and Rachana, thank you for being the most wonderful friends ever. I am so lucky to have v known you all for so long. You are all like my sisters, thank you for being so understanding with my busy schedule throughout grad school and thank you for still trying to find time to see me. Committee Members To my committee members; Dr. David Arnosti, Dr. Peggy Petroff, and Dr. Julia Ganz. Thank you for all your mentorship and time. Thank you for asking thought provoking questions and teaching me how to problem solve. To Dr. Amy Ralston, thank you for accepting me into your lab, sharing all your knowledge and wisdom. Thank you for always encouraging me. vi TABLE OF CONTENTS CHAPTER 1: Mouse Embryos, Transcription Factors, and Hippo Signaling, Oh My!.......1 CHAPTER 2: Hippo Signaling Resolves Embryonic Cell Fate Conflicts During Establishment of Pluripotency in vivo…………………………………………..……….…...17 CHAPTER 3: GATA6 and OCT4 Direct Proper Gene Expression in Trophectoderm Cells in Morula and Blastocyst Stage Mouse Embryos…………..…………….…………………53 CHAPTER 4: Further Examination of OCT4 and GATA6 in Preimplantation Mouse Embryos…………………………………………………………………..………………….....84 REFERENCES………………………..………………………………………………………..97 vii CHAPTER 1: Mouse Embryos, Transcription Factors, and Hippo Signaling, Oh My! 1.1: An overview of preimplantation mouse embryogenesis Mouse embryos offer a unique window into mammalian embryo and stem cell development. The stages of mouse embryo development, prior to implantation, are very similar to humans but occur over an accelerated period, which has recently been attributed to species-specific differences in protein stability (Molè et al., 2020; Rayon et al., 2020). The short gestational period of mouse embryos is an advantage for researchers. We can study mouse embryo development over their 21-day gestation and use those discoveries as starting points for understanding mechanisms that may be used in human gestation. For the first few cell divisions after fertilization the embryo is made up of identical cells called blastomeres (Kelly et al., 1978; Balakier and Pedersen, 1982; Fujimori et al., 2003). However, controversially, other research has shown bias in lineage contributions in subsets of blastomeres (Piotrowska-Nitsche and Zernicka-Goetz, 2005; Tabanski et al., 2013). At around 3-days post-fertilization blastomeres compact to form a morula (Fig. 1.1). Shortly after compaction the first cell fate decision occurs, separating inside cells from outside cells (Tarkowski and Wroblewska, 1967). Next is the early blastocyst stage, characterized by the formation of the blastocoel (Fig. 1.1). The blastocoel is a fluid-filled space in the embryo that exhibits an ion gradient mediated by the outside cells (Manejwala et al., 1989). After a few more rounds of cell division, when there are around 64-cells the embryo begins its second cell fate decision (Gardner, 1982). This second cell fate decision separates the inside cells into epiblast and primitive endoderm 1 (Gardner, 1982). The blastocyst continues to expand until it hatches from its protective layer, the zona pellucida, using a combination of its expansive force and enzyme production (Perona and Wassarman, 1986; Sawada et al., 1990; Thomas et al., 1997; Leonavicius et al., 2018). At about 4.5-days post-fertilization, the embryo has hatched and has over 100 cells, and the inner cell mass has fully separated into independent sections of epiblast and primitive endoderm. Finally at about 5-days post-fertilization the embryo implants into the uterus. The second cell fate decision is of particular interest because the embryo must maintain a balance of stem cell potency and differentiation. The embryo begins this stage being composed of multipotent outside cells called the trophectoderm (TE), and pluripotent inside cells called the inner cell mass (ICM). Multipotency refers to cells that can only go on to form a very limited set of cells like the trophectoderm and primitive endoderm. Pluripotency refers to cells that can become all the cells within the organism. The second cell fate decision sees part of the ICM lose potency and become multipotent primitive endoderm (PE), while the other part of the ICM retains its pluripotent state, the epiblast (EPI). This causes a brief “salt and pepper” effect in the ICM, with PE and EPI randomly organized (Chazaud et al., 2006). Similarly, to TE, the PE is limited in the types of cells it can become. With two out of its three cell types having lost pluripotency, it is remarkable that the embryo can still maintain pluripotency in the epiblast. Eventually, the blastocyst reaches implantation stage where it grows ever more complex. But all the cells can be traced back to these three cell lineages found in the preimplantation embryo. The trophectoderm will go on to form all the cells that make up the fetal placental contributions (Cross et al., 1994). The primitive endoderm will become 2 extra-embryonic supporting cells, the visceral endoderm and yolk sac endoderm (Nadijcka and Hillman, 1974; Enders et al., 1978). The primitive endoderm has also been shown to make minimal contributions to the gut tube (Kwon et al., 2008; Nowotschin et al., 2019). The epiblast makes up all of the cells in the fetus itself, both somatic and germ (Gardner, 1985; Lawson and Hage, 1994; Saitou et al., 2002). To form a fully functioning organism, an embryo has to be 100% efficient and proficient at stem cell formation, maintenance and eventually differentiation. This unique ability of embryogenesis is something researchers can only dream of achieving, which is why studying it is so vital. We can seek answers to questions of stem cell creation and maintenance from the embryo. We can learn which factors contribute to healthy stem cells and apply that knowledge to further optimize the use of stem cells and reprogramming of non-pluripotent cells to be induced pluripotent stem cells (iPSCs). 1.2: The keys to locking and unlocking cell differentiation in the mouse embryo 1.2.1: Overview of gene expression regulating factors in the very early mouse embryo At fertilization, the maternal and paternal DNA combine to create the new zygotic genome. Therefore, the embryo survives the first cleavage using maternally deposited mRNA and proteins that were present in the egg at fertilization. Once the embryo is comprised of two-cells, zygotic genome activation occurs. When the zygotic genome is active, the maternally deposited mRNA begins to degrade and is largely absent by the four-cell stage (Zhang et al., 2020). The subsequent stages of preimplantation development are reliant on the embryo’s own proper genome initiation and expression of 3 cell type specific factors. Pioneer factors play a key role in initiating the expression of cell type specific factors. The mouse zygotic genome, like almost all eukaryotic genomes, contains histones. Due to the relatively massive amount of DNA in a cell, it needs to be efficiently packaged to fit in the nucleus. DNA is therefore coiled around histone proteins. In the areas where histones are present, the DNA is unable to be transcribed, or be accessed by standard transcription factors. However, pioneer factors are a distinctive type of transcription factor that can open inaccessible chromatin and allow other factors to bind to DNA. Pioneer factors can actively and passively access DNA bound to histones. Some pioneer factors have regions that mimic histone proteins, these domains move the histone out and open the DNA, others act as flags that recruit additional factors which can access the DNA such as SWI/SNF remodelers (Clark et al., 1993; Cirillo et al., 2002). Transcription factors help to regulate gene transcription in the developing embryo. Once the DNA is made accessible, transcription factors can bind to the gene to activate or repress transcription. Transcription factors frequently have affinity to specific binding motifs, a sequence of nucleotides that the transcription factor can recognize and bind. Very generally, transcription factors can be classified as activators or repressors. Both repressing and activating transcription factors have many diverse modes of regulating transcription. Some transcription activators can bind to promoter regions and/ or enhancer regions, then through conformational changes in the DNA, where the enhancer region bends toward the promoter, they enable interactions between co-regulators to form complexes and ultimately allow transcription. Similarly, some transcription 4 repressors can bind to promoters and/ or silencers. Some cis-regulatory elements (CRE) function as both silencers and promoters depending on context. If a repressor binds the CRE it can block binding by activators, leading to transcription being inactive (Chopra et al., 2012; Segert et al., 2021). Silencer regions have binding motifs that are bound by transcriptional repressors, which after binding can work by anti-looping, or preventing the interaction of enhancer and promoter regions and their bound factors (Chopra et al., 2012). Overall, transcriptional regulation via transcription factors is highly complex and dynamic, which allows for the essential finely tuned gene expression present in embryo development. 1.2.2: OCT4, potency or not to potency that is the question The transcription factor OCT4 has an extensive history including multiple aliases. In addition to “OCT4” this transcription factor is known as “POU5F1”, “OCT3” or “OCT3/4”. OCT4 is a POU-family member, POU standing for “Pit”, “Oct”, and “Unc”. The POU family is a diverse group of transcription factors that share the POU DNA-binding domain. POU factors have been found in many different animals from zebrafish to humans (Takeda et al., 1994; Cauffman et al., 2005). The conservation of the octamer- binding sequence and POU factors across species means that research on OCT4 may be widely applicable. OCT4 is a well-known transcription factor that plays multiple important roles in preimplantation embryo development. Additionally, OCT4 has been suggested to be a pioneer factor, though this has been disputed (Soufi et al., 2015; King and Klose, 2017; Echigoya et al., 2020). OCT4 was the first Octamer-binding protein to be found in preimplantation mouse embryos (Schӧler et al., 1989). Prior to fertilization, the oocyte is 5 loaded with OCT4 protein and mRNA (Schӧler et al., 1989; Rosner et al., 1990). The presence of OCT4 in the oocyte suggests it plays a role in oocyte formation and/or fertilization however, we and others have shown that maternal OCT4 is not necessary for fertilization nor oocyte maturation (Frum et al., 2013; Wu et al., 2013; Le Bin et al., 2014). Oct4 is not immediately activated in the embryo during zygotic genome activation at the 2-cell stage (Palmieri et al., 1994). Starting just before the 8-cell stage Oct4 starts to be transcribed in all cells (Palmieri et al., 1994). Therefore, during the first cell fate decision, when inside and outside cells differentiate from each other, OCT4 is expressed in all cells no matter their position (Dietrich and Hiiragi, 2007). By the second cell fate decision, Oct4 is repressed in the trophectoderm but remains active in the ICM (Palmieri et al., 1994, Strumpf et al., 2005; Niwa et al., 2005; Ralston and Rossant, 2008; Schrode et al., 2013). The epiblast has been shown to produce fibroblast growth factor 4 (FGF4) which is necessary for PE differentiation (Fig. 1.1) (Niswander and Martin, 1992; Rappolee et al., 1994; Chazaud et al., 2003). Accordingly, the cell surface receptor for FGF4, FGFR2, is expressed on PE cells and is required for proper specification (Kurimoto et al. 2006; Arman et al., 1998). Fgf4 has been shown to contain binding motifs for OCT4 and SOX2 (Curatola and Basilico, 1990; Ma et al., 1992; Yuan et al., 1995). Embryos lacking OCT4 have decreased Fgf4 expression (Nichols et al. 1998). OCT4 is frequently considered solely a pluripotency factor and is even being used as a marker of pluripotency in cellular reprogramming. In the famous mouse fibroblast reprogramming paper, OCT4 was one of 4 factors needed to create induced pluripotent stem cells (iPSCs) from mouse embryonic fibroblast cells (Takahashi and Yamanaka, 2006). This paper specifically showed that without OCT4, iPSCs would not form 6 (Takahashi and Yamanaka, 2006). In pluripotent embryonic stem cells, that are derived from the epiblast, it has been shown that OCT4 is required for SOX2 and NANOG expression (Tomioka et al., 2002; Loh et al., 2006; Kuroda et al., 2005). SOX2 is one of the first pluripotency markers, as it is expressed strictly in the inside cells at the first cell fate decision (Guo et al., 2010; Wicklow et al., 2014). NANOG is strictly relegated to the pluripotent epiblast after the second cell fate decision (Fig. 1.1) (Chambers et al., 2003). Sox2 null ES cells lose their pluripotency, and can be rescued by expressing OCT4, which indicates that SOX2 keeps the amount of OCT4 in balance to maintain pluripotency (Masui et al., 2007). Additionally, ES cells are unable to be derived from Oct4 null embryos (Nichols et al., 1998). Taken together all this evidence argues that OCT4 is a pluripotency factor. However, more recent evidence suggests that OCT4 is not required for the initiation of pluripotency nor is it only driving pluripotent cell fate. Maternal and zygotic Oct4 null embryos can successfully develop to the blastocyst stage, including proper initiation of both SOX2 and NANOG (Frum et al., 2013). This indicates that OCT4, though maternally loaded in the oocyte, does not seem to be necessary for oocyte maturation, fertilization, or success of the embryo prior to zygotic genome activation. Contradicting the previous induced pluripotent stem cell research, it has been shown that leaving out OCT4 from the reprogramming scheme allows higher success in producing living offspring from the iPSCs (Velychko et al., 2019). This increased success without OCT4 suggests that induction of pluripotency in a reprogramming context is sensitive to concentration of OCT4 present, and that excess OCT4 is detrimental to regaining pluripotency. Similarly, sensitivity to excess OCT4 has also been seen in ES cells 7 expressing varying levels of OCT4 via its up and down regulation (Niwa et al., 2000). Overall, it seems that OCT4 is not needed to initiate pluripotency but, some expression is needed to maintain it. Furthermore, OCT4 has been shown to drive non-pluripotent primitive endoderm cell fate in ex vivo embryo outgrowths, as indicated by the absence of all ICM cells in Oct4 null outgrowths (Nichols et al., 1998). This requirement for OCT4 in non-pluripotent PE development was further confirmed in vivo using Oct4 null embryos (Frum et al., 2013; Le Bin et al., 2014). An early hypothesis for how OCT4 worked was that relative levels of Oct4 expression drove cell fate; high OCT4 led to PE, medium OCT4 led to epiblast and low led to TE, though this was seen in vitro in ES cells not embryos (Niwa et al., 2000). However, mRNA levels of Oct4 have been shown to be very similar in PE and EPI cells (Chazaud et al., 2006; Guo et al., 2010). Another early hypothesis was that OCT4 drove PE fate non-cell-autonomously upstream of FGF4. If this were the case, it would indicate that OCT4 may not actively drive PE fate but passively by producing FGF4 in pluripotent cells. This type of passive cell fate specification would not necessarily exclude the possibility of OCT4 being a pluripotency factor. Congruently with the outgrowth experiments done by Nichols et al., Oct4 null embryos lack the ability to express PE specific factors GATA6, SOX17, and GATA4 (Frum et al., 2013). Attempts to rescue PE gene expression with exogenous FGF4 or complementation with wild type ES cells were not successful (Frum et al., 2013). The inability to rescue PE gene expression suggests that OCT4 has a role downstream of FGF4 in the response of PE cells to FGF4 signaling. This is further expanded to show that late loss of OCT4 and addition of exogenous FGF4 leads to rescue of SOX17 expression and suggests that OCT4 is 8 needed for priming PE cells to respond to FGF4 but is expendable after that priming (Le Bin et al., 2014). The ability of OCT4 to co-bind with SOX factors is well established, particularly SOX2 in the epiblast, and it has been suggested that OCT4 can switch to bind with SOX17 to direct primitive endoderm fate (Aksoy et al., 2013). The “Yamanaka factors” used in reprogramming; OCT4, SOX2, KLF4, and c-MYC have also been shown to produce not only iPSCs but also induced extraembryonic endoderm (iXEN) stem cells (Parenti et al., 2016). The removal of OCT4 from this reprogramming scheme has also been shown to promote less off target gene expression in resulting iPSCs (Velychko et al., 2019). Altogether this contradicts the notion that OCT4 can or should purely be used as a marker for pluripotency. In addition to the hypotheses stated above, OCT4 was thought to repress TE cell fate in turn allowing for PE development (Nichols et al., 1998). Loss of OCT4 in embryos has been shown to cause ectopic expression of TE specific markers CDX2 and GATA3 (Ralston et al., 2010). Isolated ICMs of Oct4 null have also been shown to express TROMA-1, another TE specific marker (Nichols et al., 1998). Because the ICM contains two distinct cell types it was important to determine if, in Oct4 null embryos, ectopic TE genes are being ectopically expressed in only PE cells. However, both non-NANOG- expressing (PE cells) and NANOG-expressing (EPI cells) gain CDX2 expression and not all PE gained ectopic CDX2 (Frum et al., 2013). The lack of ectopic CDX2 expression in all PE cells indicates that OCT4 is not promoting PE fate by repressing TE gene expression. 9 1.2.3: GATA6 a key component of primitive endoderm cell fate GATA6 is part of the GATA family of proteins, which are zinc-finger transcription factors. One of the best-known roles of GATA6 is later in embryo development where it helps drive liver, gut, and other organ development (Laverriere et al., 1994; Morrisey et al., 1996; Morrisey et al., 1998; Zhao et al., 2005). Like OCT4, GATA factors have been proposed to be pioneer factors. There are multiple GATA factors that play diverse roles in preimplantation embryo development. GATA3 is a trophectoderm specific GATA factor, while GATA6 and GATA4 are relegated to the primitive endoderm. In the preimplantation embryo, GATA6 is expressed from the eight-cell stage and can be found in the trophectoderm and the inner cell mass but is then restricted to the PE (Plusa et al., 2008; Koutsourakis et al., 1999). Interestingly, it has been seen that even after the restriction of GATA6 to the PE, a seemingly significant step toward differentiation, these cells can contribute to any lineage until OCT4 is down regulated (Grabarek et al., 2012). This suggests that GATA6-expressing PE cells before E4.5 are still relatively plastic (Grabarek et al., 2012). GATA6 is repressed in non-PE track ICM cells directly by NANOG, and in PE-fated cells the expression of SOX17 and GATA4, but not GATA6, requires NANOG mediated FGF4 signaling (Fig. 1.1) (Frankenberg et al., 2011; Kang et al., 2013). In fact, GATA6 is necessary for PE-track ICM cells to respond to FGF4 signaling, suggesting that GATA6 works early in the pathway between FGFR2 and PE gene expression (Schrode et al., 2014). Embryos that have lost GATA6, in some ways, phenocopy embryos that lack OCT4. Primitive endoderm factors SOX17 and GATA4 are lost in Gata6 null embryos (Schrode et al., 2014). Like Oct4 null embryos, Gata6 null embryos survive to 10 implantation, however Gata6 null embryos do not die until between E5.5 and E7.0 (Morrisey et al., 1998; Koutsourakis et al., 1999; Schrode et al., 2014). As stated in the previous section, OCT4 is necessary for both EPI and PE development, and is hypothesized to work with cell type specific co-factors. Because Gata6 null and Oct4 null embryos both lose SOX17 and GATA4 expression, GATA6 may act as a cofactor of OCT4. 1.3: HIPPO signaling, cell polarization and cell localization in mouse embryogenesis Transcription factors are important for development but, how do cells know what transcription factors to turn on to become specific cell types in the context of a rapidly growing and changing embryo? Signaling pathways help to instruct what cells to select specific differentiation pathways based on their position. The first cell fate decision that needs to be settled in mouse embryos is outside versus inside cells, and the HIPPO pathway plays a pivotal role. The HIPPO signaling pathway, originally discovered in Drosophila, is conserved across mammalian embryo development (Justice et al., 1995; Xu et al., 1995). Signaling pathways are characterized by protein-ligands that act as signals, and protein-receptors that act as receivers. In general, signals coming from outside the cell bind to the transmembrane receptor. The binding of the ligand causes a conformational change in the receptor, which will trigger downstream enzymatic effects from the cytoplasmic side of the receptor. Cell polarity and position are essential for HIPPO signaling and the cell fates of the mouse embryo. Originally the inside and outside cells fates were proposed to be based on position alone, so that cells that happened to be on the outside became 11 trophectoderm and cells that were on the inside became ICM (Tarkowski & Wroblewska 1967). A second model proposed that cell fate was based on symmetrical or asymmetrical cell divisions (Johnson et al., 1981). In this model, symmetrical division indicates a cell divides in such a way that both daughter cells maintain the presence of a polar apical domain; an asymmetrical division indicates one where one of the daughter cells retains that apical domain and the other does not (Johnson et al., 1981). Cells with the apical domain would become TE and those without would become ICM (Johnson et al., 1981). A third model considers both position and polarity (Yamanaka et al., 2006). The Par-aPKC system has been shown to control cell polarity (Hirate et al., 2015). Cell polarity is a key to HIPPO signaling and HIPPO signaling drives expression of call fate factors (Fig. 1.2) (Hirate et al., 2013; Frum et al., 2018; Nishioka et al., 2009). The HIPPO signaling pathway in mouse development has many components. MST1 and MST2 are the mouse orthologs of HIPPO (Harvey et al. 2003), however because Mst1 null; Mst2 null embryos survive until around E9.0 these proteins are probably not involved in the pre-implantation HIPPO pathway (Oh et al., 2009). Instead, the pre- implantation HIPPO pathway is likely responding to physical signals such as, force, tension, and polarity. In outside cells, which are polarized, HIPPO signaling is inactive (Fig. 1.2). The protein AMOT is sequestered to the polar apical domain, which prevents it from binding to adherens junctions and being phosphorylated by LATS, leading to nuclear YAP localization in outside cells (Fig. 1.2) (Hirate et al., 2013). LATS acts as a kinase in inside cells where it phosphorylates YAP, therefore preventing nuclear localization. When YAP is phosphorylated, it cannot interact with its DNA-binding partner TEAD4, preventing TE gene expression, instead allowing SOX2 expression (Fig. 1.2) 12 (Nishioka et al., 2009; Frum et al., 2018). In outside cells, nuclear localization of YAP allows it to interaction with TEAD4 on the Cdx2 promoter, enabling expression of Cdx2, a TE specific gene (Fig. 1.2) (Nishioka et al., 2009). Overall, HIPPO signaling activity is dependent on cell polarity, allowing ICM gene expression when active and allowing Cdx2 expression when inactive. 1.4: Dissertation objectives Altogether, the previous research on OCT4 in preimplantation embryo development indicates that OCT4’s role is complex. OCT4 has cell-autonomous roles in the development of both EPI (pluripotent) and PE (non-pluripotent) cells. The overall goal of my dissertation is to understand how OCT4 can simultaneously drive distinct cell fates. To do this I will explore a possible co-factor relationship between OCT4 and PE specific protein GATA6. Both OCT4 and GATA6 are known transcription factors that when lost seemingly lead to ectopic expression of TE specific protein CDX2 (Ralston et al., 2010; Schrode et al., 2014). Therefore, I hypothesize that OCT4 and GATA6 may allow PE gene expression by repressing TE specific factors. I also will explore the possibility of GATA6 and OCT4 being necessary for HIPPO signaling activation in ICM cells, as the absence of HIPPO signaling normally leads to CDX2 expression, like seen in TE cells. Through these studies I hope to better understand the complexity of OCT4; how it drives PE fate in inside cells, if it is working in conjunction with GATA6, and do these factors play a role in HIPPO pathway activation in the ICM. 13 FIGURES Figure 1.1: Illustrated Mouse Embryo Development Timeline, Highlighting Stages of Interest. Mouse embryonic development from just after fertilization to blastocyst stage, and then midgestation to illustrate where these cells contribute to. Dark blue indicates trophectoderm cells which give rise to the placenta after implantation. Green indicates pluripotent inner cell mass prior to differentiation into pluripotent epiblast (EPI) (yellow) and multipotent primitive endoderm (PE) (red). EPI eventually gives rise to all fetal cell types as seen in midgestation image(yellow). PE gives rise to supporting cells called the yolk sac seen in midgestation image (red). 14 Figure 1.2: Illustrated overview of Key HIPPO Signaling Components in a 16-cell Mouse Morula. Trophectoderm (TE) cells (blue), have a polar apical domain (purple) which sequesters AMOT from interacting with LATS, therefore it is unable to phosphorylate YAP. Unphosphorylated YAP is able to enter the nucleus of the outside TE binding with its co- factor TEAD4 and promoting Cdx2 expression. Inner cell mass (ICM) cells (green) do not have a polar apical domain, because they are surrounded by neighboring cells. (See next page) 15 Figure 1.2 (cont’d): Being unpolarized allows AMOT to bind to the adherens junctions (grey boxes), and be phosphorylated by LATS. LATS also phosphorylates YAP, preventing it from entering the nucleus. Without YAP nuclear localization, TEAD4 cannot promote Cdx2, which allows Sox2 expression. 16 CHAPTER 2: HIPPO Signaling Resolves Embryonic Cell Fate Conflicts During Establishment of Pluripotency in vivo Tristan Frum1, Tayler Murphy2,3, Amy Ralston1-3 1) Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824 2) Genetics and Genomic Sciences Graduate Program, Michigan State University, East Lansing, MI, 48824 3) Reproductive and Developmental Biology Training Program, East Lansing, MI, 48824 Published as Tristan Frum, Tayler M. Murphy, Amy Ralston (2018) HIPPO signaling resolves embryonic cell fate conflicts during establishment of pluripotency in vivo, in eLife. Tristan Frum performed experiments in figures 2.1-2.6, performed data and statistical analysis in figures 2.2-2.6, wrote and edited the manuscript. Tayler Murphy performed data and statistical analysis in figure 2.1, reviewed and edited the manuscript. Amy Ralston wrote, reviewed, and edited the manuscript. The manuscript was changed to follow the formatting structure of this dissertation. Supplemental figures can be found in the online version at https://doi.org/10.7554/eLife.42298. 17 2.1: Abstract During mammalian development, the challenge for the embryo is to override intrinsic cellular plasticity to drive cells to distinct fates. Here, we unveil novel roles for the HIPPO signaling pathway segregates pluripotent and extraembryonic fates by controlling cell positioning as well as expression of Sox2, the first marker of pluripotency in the mouse early embryo. We show that maternal and zygotic YAP1 and WWTR1 repress Sox2 while promoting expression of the trophectoderm gene Cdx2 in parallel. Yet, Sox2 is more sensitive than Cdx2 to Yap1/Wwtr1 dosage, leading cells to a state of conflicted cell fate when YAP1/WWTR1 activity is moderate. Remarkably, HIPPO signaling activity resolves conflicted cell fate by repositioning cells to the interior of the embryo, independent of its role in regulating Sox2 expression. Rather, HIPPO antagonizes apical localization of Par complex components PARD6B and aPKC. Thus, negative feedback between HIPPO and Par complex components ensure robust lineage segregation. 2.2: Introduction During embryogenesis cells gradually differentiate, adopting distinct gene expression profiles and fates. In mammals, the first cellular differentiation is the segregation of trophectoderm and inner cell mass. The trophectoderm, which comprises the polarized outer surface of the blastocyst, will mainly produce cells of the placenta, while the inner cell mass will produce pluripotent cells, which are progenitors of both fetus and embryonic stem cells. Understanding how pluripotent inner cell mass cells are segregated from non-pluripotent cells therefore reveals how pluripotency is induced in a naturally occurring setting. Progenitors of inner cell mass are first morphologically apparent at the 16-cell 18 stage as unpolarized cells residing inside the morula (reviewed in Frum and Ralston, 2018). However, at this stage, pluripotency genes such as Oct4 and Nanog, do not specifically label inside cells (Dietrich and Hiiragi, 2007; Niwa et al., 2005; Palmieri et al., 1994; Strumpf et al., 2005). Thus, the first cell fate decision has been studied mainly from the perspective of trophectoderm specification because the transcription factor CDX2, which is essential for trophectoderm development (Strumpf et al., 2005), is expressed specifically in outer cells of the 16-cell embryo (Ralston and Rossant, 2008), and has provided a way to distinguish future trophectoderm cells from non- trophectoderm cells. Knowledge of CDX2 as a marker of trophectoderm cell fate enabled the discovery of mechanisms that sense cellular differences in polarity and position in the embryo, and then respond by regulating expression of Cdx2 (Nishioka et al., 2009). However, the exclusive study of Cdx2 regulation does not provide direct knowledge of how pluripotency is established because the absence of Cdx2 expression does not necessarily indicate acquisition of pluripotency. As such, our understanding of the first cell fate decision in the early mouse embryo is incomplete. In contrast to other markers of pluripotency, Sox2 is expressed specifically in inside cells at the 16-cell stage, and is therefore the first marker of pluripotency in the embryo (Guo et al., 2010; Wicklow et al., 2014). The discovery of how Sox2 expression is regulated in the embryo will therefore provide unique insight into how pluripotency is first established in vivo. Genes promoting expression of Sox2 in the embryo have been described (Cui et al., 2016; Wallingford et al., 2017). However, it is currently unclear how expression of Sox2 becomes restricted to inside cells. Interestingly, Sox2 is restricted to inside cells by a Cdx2-independent mechanism (Wicklow et al., 2014), which differs from 19 Oct4 and Nanog, which are restricted to the inner cell mass by CDX2 (Niwa et al., 2005; Strumpf et al., 2005). Thus, Sox2 and Cdx2 are regulated in parallel, leading to complementary inside/outside expression patterns. However, it is not known whether Sox2 is regulated by the same pathway that regulates Cdx2 or whether a distinct pathway could be in use. The expression of Cdx2 is regulated by members of the HIPPO signaling pathway. In particular, the HIPPO pathway kinases LATS1/2 become active in unpolarized cells located deep inside the embryo, where they antagonize activity of the YAP1/WWTR1/TEAD4 transcriptional complex that is thought to promote expression of Cdx2 (Anani et al., 2014; Cockburn et al., 2013; Hirate et al., 2013; Kono et al., 2014; Korotkevich et al., 2017; Leung and Zernicka-Goetz, 2013; Lorthongpanich et al., 2013; Mihajlović and Bruce, 2016; Nishioka et al., 2009, 2008; Posfai et al., 2017; Rayon et al., 2014; Watanabe et al., 2017; Yagi et al., 2007; Zhu et al., 2017). In this way, the initially ubiquitous expression of Cdx2 becomes restricted to outer trophectoderm cells. However, the specific requirements for Yap1 and Wwtr1 in the regulation of Cdx2 has been inferred from overexpression of wild type and dominant-negative variants, neither of which provide the standard of gene expression analysis that null alleles can provide. Nonetheless, the roles of Yap1 and Wwtr1 in regulating expression of Sox2 have not been investigated. Here, we evaluate the roles of maternal and zygotic YAP1/WWTR1 in regulating expression of Sox2 and cell fate during blastocyst formation. 2.3: Results 2.3.1: Patterning of Sox2 is ROCK-dependent To identify the mechanisms regulating Sox2 expression during blastocyst 20 formation, we focused on how Sox2 expression is normally repressed in the trophectoderm to achieve inside cell-specific expression. We previously showed that SOX2 is specific to inside cells in the absence of the trophectoderm factor CDX2 (Wicklow et al., 2014), suggesting that mechanisms that repress Sox2 in the trophectoderm act upstream of Cdx2. Rho-associated, coiled-coil containing protein kinases (ROCK1 and 2) are thought to act upstream of Cdx2 because embryos developing in the presence of a ROCK-inhibitor (Y-27632, ROCKi) exhibit reduced Cdx2 expression (Kono et al., 2014). Additionally, quantitative RT-PCR showed that Sox2 mRNA levels are elevated in ROCKi-treated embryos (Kono et al., 2014), suggesting that ROCK1/2 activity leads to transcriptional repression of Sox2. However, the role of ROCK1/2 in regulating the spatial expression of Sox2 has not been investigated. To evaluate the roles of ROCK1/2 in patterning Sox2 expression, we collected 8- cell stage embryos prior to embryo compaction (E2.5), and then cultured these either in control medium or in the presence of ROCKi for 24 hours (Fig. 2.1A). Embryos cultured in control medium exhibited normal cell polarity, evidenced by the apical localization of PARD6B and basolateral localization of E-cadherin (CDH1) in outside cells (Fig. 2.1B, C) as expected (Vestweber et al., 1987; Vinot et al., 2005). Additionally, SOX2 was detected only in inside cells in control embryos (Fig. 2.1C, D). By contrast, embryos cultured in ROCK inhibitor exhibited defects in cell polarity (Fig. 2.1B’, C’), consistent with prior studies (Kono et al., 2014). Interestingly, in ROCK inhibitor-treated embryos, we observed ectopic SOX2 expression in cells located on the outer surface of the embryo (Fig. 2.1C’, D), indicating that ROCK1/2 participates in the pathway responsible for repressing expression of Sox2 in the trophectoderm. 21 To scrutinize the identity of outside positioned SOX2-positive cells in ROCK- inhibited embryos, we co-stained an additional cohort of control and ROCKi-treated embryos with CDX2 and SOX2 and compared the overlap of lineage marker expression. In control embryos, CDX2 was detected only in outside cells (Fig. S1A) as expected at this stage (Ralston and Rossant, 2008; Strumpf et al., 2005). In ROCKi-treated embryos, CDX2 expression levels were reduced (Fig. S1A’) as was the proportion of outside cells in which CDX2 was detected (Fig. S1B), as previously reported (Kono et al., 2014). However, among outside cells, a substantial proportion coexpressed CDX2 and SOX2 in ROCK-inhibited embryos compared with controls (Fig. 2.1E and S1A), suggesting that ROCK inhibition leads to an increase in outside cells of mixed lineage. Since SOX2 expression does not regulate expression of CDX2 (Wicklow et al., 2014), these observations suggest that ROCK1/2 activity regulate these genes through parallel mechanisms. We next sought to identify mediators that act downstream of ROCK1/2 to repress expression of Sox2 in the trophectoderm. 2.3.2: YAP1 is sufficient to repress expression of SOX2 in the inner cell mass Several direct and indirect targets of ROCK1/2 kinases in the early embryo have been described (Alarcon and Marikawa, 2018; Shi et al., 2017). Among these is YAP1, a transcriptional partner of TEAD4 (Nishioka et al., 2009), since ROCK activity is required for the nuclear localization of YAP1 (Kono et al., 2014). Notably, Tead4 is required to repress expression of Sox2 in the trophectoderm (Wicklow et al., 2014), consistent with the possibility that YAP1 partners with TEAD4 to repress Sox2 expression in the trophectoderm. To test this hypothesis, we overexpressed a constitutively active variant of YAP1 (YAP1CA). Substitution of alanine at serine 112 leads YAP1 to be constitutively 22 nuclear and constitutively active (YAP1CA hereafter) (Dong et al., 2007; Nishioka et al., 2009; Zhao et al., 2007). We injected mRNAs encoding YAP1CA and GFP into one of two blastomeres at the 2-cell stage, and then cultured these to the blastocyst stage (Fig. 2.1F). This mosaic approach to overexpression permitted comparison of YAP1CA- overexpressing with non-injected cells, which served as internal negative controls. We first examined localization of YAP1 in these embryos at the morula stage, with the expectation that YAP1 would be detected in nuclei of both inside and outside cells in YAP1CA-overexpressing cells (Nishioka et al., 2009). As expected, YAP1 was observed in nuclei of all YAP1CA-overexpressing cells (Fig. S1B, C). We next evaluated the consequences of ectopic nuclear YAP1 on expression of SOX2 in inside cells. We observed a conspicuous decrease in the proportion of YAP1CA-overexpressing inside cells lacking detectable SOX2 (Fig. 2.1G, H). Therefore, nuclear YAP1 is sufficient to repress Sox2 expression in the inner cell mass, indicative of a likely role for YAP1 in repressing expression of Sox2 in the trophectoderm downstream of ROCK1/2. 2.3.3: LATS kinase is sufficient to induce inside cell positioning To functionally test of the role of YAP1 in repressing expression of Sox2, we injected one of two blastomeres with mRNA encoding LATS2 kinase, which inactivates YAP1 and, presumably, the related protein WWTR1 by phosphorylation, causing their cytoplasmic retention (Nishioka et al., 2008). We then examined expression of SOX2 after culturing embryos to the blastocyst stage (Fig. 2.2A), predicting that LATS2 kinase would induce the ectopic expression of Sox2 in outside cells. Surprisingly, we observed that almost all Lats2-overexpressing cells ended up within the inner cell mass by the blastocyst stage (Fig. 2.2B, C), in contrast to cells injected with GFP mRNA only, which 23 contributed to both inner cell mass and trophectoderm. Notably, SOX2 was detected in all Lats2-overexpressing cells observed within the inner cell mass (Fig. 2.2D), suggesting that Lats2-overexpressing cells were not only localized to the inner cell mass but also position-appropriate regulation of Sox2. The strikingly increased prevalence of Lats2-overexpressing cells in the inner cell mass was also associated with a stark decrease in the number of Lats2-overexpressing cells detected within the trophectoderm and a decrease in the number of outside cells compared to embryos injected with GFP mRNA alone (Fig. 2.2C, E), suggesting that Lats2-overexpressing outside cells either internalize or undergo cell death. Furthermore, we observed cellular fragments within the trophectoderm of Lats2-overexpressing embryos (Fig. 2.2B, yellow arrowheads), as well as increased TUNEL staining in Lats2- overexpressing embryos compared to embryos injected with GFP mRNA only (Fig. S2A- B, D), consistent with increased death of Lats2-overexpressing cells. In addition to detecting SOX2 in all Lats2-overexpressing cells located inside the embryo, SOX2 was also detected in rare Lats2-overexpressing cells that remained on the embryo surface (Fig. 2.2D). Therefore, LATS2 is sufficient to induce expression of SOX2 in cells regardless of their position within the embryo. Importantly, the kinase-dead variant of LATS2 (Nishioka et al., 2009), did not alter cell positioning, survival, or SOX2 expression (Fig. S3A, B), consistent with a previous report (Posfai et al., 2017). Thus, overexpressed LATS2 influences cell position and gene expression by modulating the activity of YAP1 and possibly WWTR1. We predicted that, if Lats2 overexpression drove cells to adopt inner cell mass fate by influencing YAP1 and WWTR1 activity, then co- overexpression of Yap1CA would enable Lats2-overexpressing cells to contribute to 24 trophectoderm. Consistent with this prediction, co-overexpression of Lats2 and Yap1CA led to a significant decrease in the proportion of Lats2-overexpressing cells contributing to the inside cell position, and a concomitant increase in the proportion of Lats2- overexpressing cells remaining in the outside position (Fig. S3C-F). Moreover, co- overexpression of Lats2 and Yap1CA reduced the number of TUNEL positive nuclei, consistent with Yap1CA rescuing survival of outside positioned Lats2-overexpressing cells (Fig. S2C-D). Collectively, these observations strongly suggest that LATS2 promotes inside cell positioning by regulating the activities of YAP1 and, likely, the related protein WWTR1. To pinpoint when Lats2-overexpressing cells come to occupy the inside of the embryo, we performed a time course, examining the position of injected and non-injected cells from the 16-cell to the blastocyst stage (~80 cells). Surprisingly, between the 16 and 32-cell stages, the proportion of injected and non-injected cells in the total, outside, and inside cell populations were comparable whether embryos had been injected with Lats2 and GFP or GFP mRNA alone (Fig. 2.2F-H). In embryos injected with GFP mRNA alone, the proportion of injected and non-injected cells making up the total, outside, and inside cell populations remained constant throughout the time course. In contrast, starting around the 32-cell stage, the average proportion of Lats2-overexpressing cells making up the inside population began to increase dramatically. This increase was associated with a decrease in the proportion Lats2-overexpressing cells making up the outside population, consistent with internalization of Lats2-overexpressing cells after the 32-cell stage (Fig. 2.2G). After the 32-cell stage, Lats2-injected cells became underrepresented as a proportion of the total cell population (Fig. 2,2H), lending further 25 support to the idea that Lats2-overexpressing cells that fail to internalize undergo cell death. Interestingly, the inside-skewed contribution of Lats2-overexpressing cells did not influence the ability of non-injected cells to contribute to the ICM (Fig. 2.2I), arguing that Lats2-overexpression drives inside positioning cell-autonomously. We therefore conclude that Lats2 overexpression acts on cell position and survival around the time of blastocyst formation. 2.3.4: LATS2 induces positional changes independent of Sox2 Our observation that Lats2-overexpression induces both the expression of SOX2 and cell repositioning to inner cell mass prompted us to investigate whether SOX2 itself drives cell repositioning downstream of Lats2. In support of this hypothesis, SOX2 activity has been proposed to bias inner cell mass fate (Goolam et al., 2016; White et al., 2016). We therefore investigated whether Sox2 is required for the inner cell mass- inducing activity of LATS2 by overexpressing Lats2 in embryos lacking maternal and zygotic Sox2 (Fig. 2.3A), as previously described (Wicklow et al., 2014). However, we observed that Lats2-overexpressing cells were equally likely to occupy inside position in the presence and absence of Sox2 (Fig. 2.3B, C). Furthermore, Lats2-overexpressing cells were equally unlikely to occupy outside position in the presence and absence of Sox2 (Fig. 2.3D). Therefore, although Lats2 overexpression is sufficient to induce expression of Sox2, LATS2 acts on cell positioning/survival independently of Sox2. 2.3.5: LATS2 antagonizes formation of the apical domain Trophectoderm cell fate has been proposed to be determined by apically localized membrane components that maintain the position of future trophectoderm cells on the embryo surface (Anani et al., 2014; Korotkevich et al., 2017; Maître et al., 2016, 2015; 26 Samarage et al., 2015; Zenker et al., 2018). For example, the apical membrane components aPKC and PARD6B are required for maintaining outside cell position and trophectoderm fate (Alarcon, 2010; Dard et al., 2009; Hirate et al., 2015; Plusa et al., 2005). Because Lats2 overexpression led cells to adopt an inside position, this raised the testable possibility that LATS2 antagonizes localization of aPKC and PARD6B. Since Lats2 overexpression leads to cell positioning starting around the 32-cell stage, we examined the localization of aPKCz and PARD6B in embryos just prior to the 32-cell stage. At this stage, apical membrane components PARD6B and aPKCz were detected at the apical membrane of non-injected outside cells and outside cells injected with GFP only (Fig. 2.4A-D). By contrast, most Lats2-overexpressing outside cells lacked detectable aPKCz and PARD6B (Fig. 2.4A-D). Therefore, LATS2 is sufficient to antagonize localization of key apical domain proteins in outside cells, providing a compelling mechanism for the observed repositioning of Lats2-overexpressing outside cells. We also examined other markers of apicobasal polarization in Lats2- overexpressing outside cells prior to the 32-cell stage. Curiously, other markers of apicobasal polarization were properly localized in all cells examined. For example, CDH1 was restricted to the basolateral membrane (Fig. 2.4E), while filamentous Actin and phospho-ERM were restricted to the apical domain in outside cells of both Lats2- overexpressing and non-injected outside cells (Fig. 2.4F, G). Thus, we propose that Lats2-overexpressing outside cells initially possess hallmarks of apicobasal polarization, but aPKC and PARD6B fail to properly localize, leading to their eventual depolarization and internalization. 27 2.3.6: YAP1 and WWTR1 restrict Sox2 expression to the inner cell mass Our overexpression data suggested that the activities of YAP1 and WWTR1 are important for regulating cell fate and gene expression. Next, we aimed to test the requirement for Yap1 and Wwtr1 in embryogenesis. Yap1 null embryos survive until E9.0 (Morin-Kensicki et al., 2006), suggesting that oocyte-expressed (maternal) Yap1 (Yu et al., 2016), or the Yap1 paralogue Wwtr1 (Varelas et al., 2010) are important for preimplantation development. However, embryos lacking maternal and zygotic Wwtr1 and Yap1 have not been analyzed. To generate embryos lacking maternal and zygotic Wwtr1 and Yap1, we deleted Wwtr1 and Yap1 from the female germ line using mice carrying conditional alleles of Wwtr1 and Yap1 (Xin et al., 2013, 2011) and the female germ line-specific Zp3Cre (de Vries et al., 2000). We then crossed these females to males heterozygous for deleted alleles of Wwtr1 and Yap1 (see Methods). From these crosses, we obtained embryos lacking maternally provided Wwtr1 and Yap1 and either heterozygous or null for Wwtr1 and/or Yap1 (Table S1). At E3.25 (≤32 cells), SOX2 and CDX2 are normally mutually exclusive (Fig. 2.5A). However, with decreasing number of wild type zygotic alleles of Wwtr1 and Yap1, we observed worsening phenotypes (Fig. 2.5B-F). In the complete absence of Wwtr1 and Yap1, we observed a severe loss of CDX2 and expansion of SOX2 in outside cells (Fig. 2.5D-F), phenocopying Lats2 overexpression. However, in embryos of intermediate genotypes, we observed expanded SOX2 and persistent, yet lower, expression levels of CDX2 (Fig. 2.5C, E-F). Thus, regulation of Sox2 expression is more sensitive to Wwtr1 and Yap1 dosage than is Cdx2. Moreover, these observations indicate that intermediate doses of Wwtr1 and Yap1 produce outside cells expressing 28 markers of mixed cell lineage at E3.25. 2.3.7: YAP1 and WWWTR1 maintain outside cell positioning Based on our observations of Lats2-overexpressing embryos, we anticipated that defects in cell positioning in embryos lacking maternal and zygotic Wwtr1 and Yap1 could arise after the 32-cell stage. We therefore examined embryos lacking Wwtr1 and Yap1 at E3.75, when embryos possess more than 32-cells. Indeed, we observed skewed lineage contributions, correlating with the dosage of Wwtr1 and Yap1 (Fig. 2.6A- D). Embryos with one or fewer wild type alleles of Wwtr1 or Yap1 exhibited an increase in the number of inside cells and a reduction in the number of outside cells (Fig. 2.6A-B), consistent with altered cell positioning. Although the average total number of cells was also reduced in these embryos (Fig. 2.6C), the reduction in total cell number did not alone account for the loss of cells on the outside of the embryo (Table S2). This observation suggested that, similar to Lats2-overexpressing cells, cells with reduced Wwtr1 and Yap1 exhibit an increased frequency of outside cell death, in addition to increased outside cell internalization. Consistent with this, embryos with one or fewer wild type alleles of Wwtr1 or Yap1 exhibited an increase in the ratio of inside to outside cells (Fig. 2.6D) and an increase in cells undergoing apoptosis by TUNEL assay (Fig. 2.6G and S6A, B). Critically, the fewer outside cells apparent in embryos lacking Wwtr1 and Yap1, which appeared stretched over the mass of inside cells, exhibited ectopic expression of SOX2 (Fig. 2.6E-F). Therefore, Wwtr1/Yap1 repress inner cell mass fate, downstream of LATS kinases. Intriguingly, our data also indicate that WWTR1 is a more potent repressor of Sox2 at E3.75 than YAP1 since embryos with a single wild type allele of 29 Wwtr1 had significantly fewer cells expressing ectopic SOX2 then embryos with a single wild type allele of Yap1 (Fig. S4). Since loss of Wwtr1 and Yap1 phenocopied Lats2 overexpression in terms of Sox2 expression, cell death, and cell repositioning, we next evaluated apical domain and cell polarization in outside cells of embryos lacking Wwtr1 and Yap1 at E3.75. We observed greatly reduced aPKC at the apical membrane of outside cells in embryos with one or fewer doses of Wwtr1 or Yap1 (Fig. 2.6H and S6C). In addition, we evaluated the localization of the tight junction protein ZO-1, which suggested failure in tight junction formation in embryos with 1 or fewer doses of Wwtr1 and Yap1 (Fig. 2.6I and S6D). Notably, however, other markers of apicobasal polarity, such as CDH1 and pERM were correctly localized in outside cells of mutant embryos at this stage (Fig. 2.6J and S6E). Our observations indicate that WWTR1 and YAP1 play a crucial role in the formation of the apical domain and maintaining the positioning and survival of outside cells while repressing expression of Sox2. 2.4: Discussion During preimplantation development, lineage-specific transcription factors are commonly expressed in ‘noisy’ domains before refining to a lineage-appropriate pattern (Simon et al., 2018). For example, Oct4 and Nanog are expressed in both inner cell mass and trophectoderm until after blastocyst formation (Dietrich and Hiiragi, 2007; Strumpf et al., 2005). Similarly, CDX2 is detected in inner cell mass, as well as trophectoderm, until blastocyst stages (McDole and Zheng, 2012; Ralston and Rossant, 2008; Strumpf et al., 2005). In striking contrast to these genes, SOX2 is never detected in outside cells (Wicklow et al., 2014), indicating that robust mechanisms must exist to 30 minimize noise and prevent its aberrant expression in trophectoderm. Here, we identify YAP1/WWTR1 as key components that repress Sox2 expression in outside cells of the embryo. Notably, manipulations known to antagonize YAP1/WWTR1 activity, including chemical inhibition of ROCK and overexpression of LATS2 lead to ectopic expression of SOX2 in outside cells, reinforcing the notion that YAP1/WWTR1 activity are crucial for repression of Sox2 in outside cells. Additionally, we find that Sox2 expression is more sensitive than is Cdx2 to YAP1/WWTR1 activity, since intermediate doses of active YAP1/WWTR1 yields cells that coexpress both SOX2 and CDX2 (Fig. 2.7A). This observation is consistent with the fact that CDX2 is initially detected in inside cells of the embryo during blastocyst formation (Dietrich and Hiiragi, 2007; McDole and Zheng, 2012; Ralston and Rossant, 2008), where SOX2 is also expressed (Wicklow et al., 2014). Thus, inside cells could initially possess intermediate doses of active YAP1/WWTR1 at this early stage. By contrast, outside cells would have greatly reduced YAP1/WWTR1 activity, owing to elevated LATS activity. In this way, the HIPPO pathway ensures robust developmental transitions, by rapidly nudging SOX2-expressing cells into their correct and final positions inside the embryo (Fig. 2.7B). Consistent with our proposed model, the timing of HIPPO-induced cell internalization coincides with loss of cell fate plasticity around the 32-cell stage (Posfai et al., 2017). This timing also coincides with the formation of mature tight-junctions among outside cells (Sheth et al., 1997), which reinforce and intensify differences in HIPPO signaling activity between inside and outside compartments of the embryo (Hirate and Sasaki, 2014; Leung and Zernicka-Goetz, 2013). Our observations indicate that HIPPO 31 signaling can, in turn, interfere with trophectoderm epithelialization. Therefore, we propose that HIPPO engages in a negative feedback loop with cell polarity components (Fig. 2.7B). We propose two mechanisms by which HIPPO signaling eliminates cells from the trophectoderm, both of which are downstream of YAP1/WWTR1 (Fig. 2.7C). First, a small proportion of conflicted cells undergo cell death. This is in line with the observed increase in the level of apoptosis detected after the 32-cell stage (Copp, 1978). We showed that cell lethality due to elevated HIPPO can be rescued by increasing levels of nuclear YAP1, suggesting that YAP1 activity normally provides a pro-survival signal to trophectoderm cells, consistent with the proposed role of YAP1 in promoting proliferation in non-eutherian mammals (Frankenberg, 2018). Moreover, deletion of Sox2 did not rescue survival of outside cells in which HIPPO signaling was artificially elevated, arguing that HIPPO resolves cell fate conflicts independently of lineage-specific genes. The second way that conflicted cells are eliminated from the trophectoderm is that cells with elevated HIPPO signaling drive their own internalization. This is consistent with the observation that cells in which Tead4 has been knocked down become internalized (Mihajlović et al., 2015). However, in contrast to Tead4 loss of function, which preserves the polarization of outside cells (Mihajlović et al., 2015; Nishioka et al., 2008), we observed that Yap1/Wwtr1 loss of function leads loss of apical PARD6D/aPKC. These observations suggest that YAP1/WWTR1 could partner with proteins other than TEAD4 to regulate apical domain formation. Consistent with this proposal, TEAD1 has been proposed to play an essential role in the early embryo (Sasaki, 2017). Nevertheless, since PARD6B/aPKC are essential for outside cell positioning (Dard et al., 2009; Hirate 32 et al., 2015; Plusa et al., 2005), the loss of the apical domain could affect cell positioning in several ways. For instance, loss of PARD6B/aPKC would eventually lead to cell depolarization (Alarcon, 2010), which could influence any of the processes normally governing the formation of inside cells, such as oriented cleavage, cell contractility, or apical constriction (Korotkevich et al., 2017; Maître et al., 2016; Samarage et al., 2015). Identifying the downstream mechanisms by which HIPPO drives cells to inner cell mass will be a stimulating topic of future study. Our studies also revealed that SOX2 does not play a role in cell positioning. This observation sheds light on a recent study, which showed that SOX2 dwells longer in select nuclei of four-cell stage embryos that are destined to contribute to the inner cell mass (White et al., 2016). We propose that SOX2 is associated with future pluripotent state but does not alone contribute to all aspects of pluripotency, such as inside positioning. It is therefore still unclear why it is important to establish the inside cell- specific SOX2 expression during embryogenesis. Identification pathways that function downstream of YAP1/WWTR1 and in parallel to SOX2 to promote formation of pluripotent cells will provide meaningful insights into the natural origins of mammalian pluripotent stem cell progenitors. 2.5: Methods 2.5.1: 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. Wild type embryos were derived from CD-1 mice (Charles River). The following alleles or transgenes were used in this study, and maintained in a CD-1 background: Sox2tm1.1Lan 33 (Smith et al., 2009), Yaptm1.1Eno (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 mice carrying floxed alleles and mice carrying ubiquitously expressed Cre, 129-Alpltm(cre)Nagy (Lomelí et al., 2000). 2.5.2: Embryo collection and culture Mice were maintained on a 12-hour light/dark cycle. Embryos were collected by flushing the oviduct or uterus with M2 medium (Millipore). For embryo culture, KSOM medium (Millipore) was equilibrated overnight prior to embryo collection. Y-27632 (Millipore) was included in embryo culture medium at a concentration of 80 µM with 0.4% DMSO, or 0.4% DMSO as control, where indicated. Embryos were cultured at 37ºC in a 5% CO2 incubator under light mineral oil. 2.5.3: Embryo microinjection LATS2 and YAPS112A mRNA was synthesized from cDNAs cloned into the pcDNA3.1-poly(A)83 plasmid (Yamagata et al., 2005) using the mMESSAGE mMACHINE T7 transcription kit (Invitrogen). EGFP or nls-GFP mRNA were synthesized from EGFP cloned into the pCS2 plasmid or the nls-GFP plasmid (Ariotti et al., 2015) using the mMESSAGE mMACHINE SP6 transcription kit (Invitrogen). mRNAs were cleaned and concentrated prior to injection using the MEGAclear Transcription Clean-Up Kit (Invitrogen). Lats2, Lats2KD and YAPCA mRNAs were injected into one blastomere of two-cell stage embryos at a concentration of 500 ng/µl, mixed with 350 ng/µl EGFP or nls-GFP mRNA diluted in 10 mM Tris-HCl (pH 7.4), 0.1 mM EDTA. 2.5.4: Immunofluorescence and Confocal Microscopy Embryos were fixed with 4% formaldehyde (Polysciences) for 10 minutes, 34 permeabilize with 0.5% Triton X-100 (Sigma Aldrich) for 30 minutes, and then blocked with blocking solution (10% Fetal Bovine Serum (Hyclone), 0.2% Triton X-100) for 1 hour at room temperature, or overnight at 4ºC. Primary Antibodies used were: mouse anti- CDX2 (Biogenex, CDX2-88), goat anti-SOX2 (Neuromics, GT15098), rabbit anti- PARD6B (Santa Cruz, sc-67393), rabbit anti-PARD6B (Novus Biologicals, NBP1-87337), mouse anti-PKC (Santa Cruz Biotechnology, sc-17781), rat anti-CDH1 (Sigma Aldrich, U3254), mouse anti-YAP (Santa Cruz Biotechnology, sc101199), rabbit anti phospho- YAP (Cell Signaling Technologies, 4911), chicken anti-GFP (Aves, GFP-1020). Stains used were: Phallodin-633 (Invitrogen), DRAQ5 (Cell Signaling Technologies) and DAPI (Sigma Aldrich). Secondary antibodies conjugated to DyLight 488, Cy3 or Alexa Flour 647 fluorophores were obtained from Jackson ImmunoResearch. Embryos were imaged using an Olympus FluoView FV1000 Confocal Laser Scanning Microscope system with 20x UPlanFLN objective (0.5 NA) and 5x digital zoom. For each embryo, z-stacks were collected, with 5 µm intervals between optical sections. All embryos were imaged prior to knowledge of their genotypes. 2.5.5: Embryo Analysis For each embryo, z-stacks were analyzed using Photoshop or Fiji, which enabled the virtual 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., SOX2 or CDX2 positive or negative), protein localization (e.g., aPKC or CDH1 apical, basal, absent, or unlocalized), and cell position, where cells making contact with the external environment were considered ‘outside’ and 35 cells surrounded by other cells were considered ‘inside’ cells. 2.5.6: TUNEL Assay Embryos were fixed, permeabilized, and blocked as described for immunofluorescence. Zonae pellucida were removed using Tyrode’s Acid treatment prior to performing the TUNEL assay (In Situ Cell Death Detection Kit, Fluorescein, Millipore- Sigma). Embryos were incubated in 200 µl of a 1:10 dilution of enzyme in label solution for 2 hours at 37 ºC. Embryos were then washed twice with blocking solution for 10 minutes each, and then mounted in a 1 to 400 dilution of DRAQ5 in blocking solution to stain DNA. 2.5.7: Embryo Genotyping To determine embryo genotypes, embryos were collected after imaging and genomic DNA extracted using the Extract-N-Amp kit (Sigma) in a final volume of 10 µl. Genomic extracts (1-2 µl) were then subjected to PCR using allele-specific primers (Table S3). 2.6 Acknowledgements We are grateful to Dr. Hiroshi Sasaki for providing expression constructs, to Dr. Randy L. Johnson for providing mice carrying conditional alleles of Yap1 and Wwtr1, and to Dr. Jason Knott for embryo microinjection training. We also thank Dr. Ripla Arora, Dr. Julia Ganz, and members of the Ralston Lab for comments. This work was supported by NIH R01 GM104009 and the Eunice Kennedy Shriver National Institute of Child Health & Human Development of the National Institutes of Health under Award Number T32HD087166. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. We thank 36 anonymous reviewers for insightful questions and suggestions. 37 FIGURES Figure 2.1: ROCK1/2 and nuclear YAP1 repress expression of SOX2. A. Experimental design: embryos were collected at E2.5 and treated with ROCK inhibitor Y-27632 (ROCKi) or DMSO (control) for 24 hours. B-B’. Confocal images of apical (PARD6B) and basolateral (CDH1) membrane components in control and ROCKi-treated embryos. As expected, PARD6B and CDH1 are mislocalized to the entire cell membrane of all cells in ROCKi-treated embryos, demonstrating effective ROCK inhibition (n = number of embryos examined). (See next page) 38 Figure 2.1 (cont’d): C-C’. In control embryos, SOX2 is detected only in inside cells, while in ROCKi-treated embryos, SOX2 is detected in inside and outside cells (arrowheads, outside cells; n = embryos). D. Quantification of ectopic SOX2 detected in outside cells of control and ROCKi-treated embryos (p, student’s t-test, n = embryos). E. SOX2 and CDX2 staining in outside cells of control and ROCKi-treated embryos. ROCK-inhibitor treatment leads to outside cells with mixed lineage marker expression (CDX2+/SOX2+). F. Experimental design: embryos were collected at E1.5 and one of two blastomeres injected with mRNAs encoding YAPCA and GFP. Embryos were cultured for 72 hours, fixed, and then analyzed by immunofluorescence and confocal microscopy. G. SOX2 is detected non-injected inside cells. SOX2 is not detected in YAPCA- overexpressing inside cells (arrowheads), n = embryos. H. Across multiple embryos, all non-injected inside cells express SOX2, whereas the vast majority of YAPCA-injected inside cells fail to express SOX2. 39 Figure 2.2: LATS2 kinase is sufficient to direct cells to inner cell mass fate. A. Embryos were collected at E1.5 and one of two blastomeres was injected with mRNAs encoding Lats2 and GFP or GFP alone. Embryos were cultured for 72 hours, fixed, and then analyzed by immunofluorescence and confocal microscopy. (See next page) 40 Figure 2.2 (cont’d): B. Cells injected with GFP (dotted line) contributed to trophectoderm and inner cell mass, while cells injected with Lats2 and GFP (dotted line) contributed almost exclusively to the inner cell mass, leaving only cellular fragments in the trophectoderm (arrows), suggestive of cell death (n = embryos). C. Proportion of inside, outside, and total cell populations across multiple embryos, which were comprised of non-injected cells, or cells injected with either GFP or GFP/Lats2 mRNAs. Cells injected with GFP/Lats2 were overrepresented within the inside cell population and underrepresented in the outside and total cell populations, relative to cells injected with GFP alone (P, chi-squared test). D. Percentage of SOX2-positive cells within non-injected and GFP-injected or Lats2/GFP-injected populations observed inside and outside of the embryo. SOX2 was detected in all of the Lats2/GFP-injected inside cells, and in half of the rare, Lats2/GFP- injected outside cells (same number of embryos as in panel C) (p, student’s t-test). E. Average number of outside and total cells per embryo. The average number of outside cells is reduced in embryos injected with Lats2/GFP, relative to GFP-injected (p, student’s t-test). F. Proportion of GFP and Lats2/GFP-injected cells, relative to total cell number, over the course of development to the ~80-cell blastocyst (Solid lines = average of indicated data point and four previous data points). G. Data as shown in panel H, shown relative to outside cell number. H. Data as shown in panel H, shown relative to inside cell number. (See next page) 41 Figure 2.2 (cont’d): I. Contribution of injected and non-injected cells to the inside cell population, following injection with GFP or Lats2/GFP. Injection with Lats2/GFP increases the overall number of inside cells compared to injection with GFP only through increasing the number of injected cells contributing to the inside cell population, without affecting the number of non-injected cells contributing to the inside cell population (p, student’s t-test). 42 Figure 2.3: LATS2 directs inner cell mass fate independently of Sox2. A. Lats2 and GFP or GFP alone were overexpressed in embryos lacking maternal or maternal and zygotic Sox2. B. Lats2/GFP-overexpressing cells (dotted line) contribute almost exclusively to the inner cell mass in the presence or absence of Sox2 (n = embryos). C. Proportion of non-injected cells and cells injected with Lats2/GFP mRNAs contributing to inner cell mass in the indicated genetic backgrounds. No significant differences were observed based on embryo genotype, indicating that Sox2 is dispensable for inside positioning by Lats2-overexpression (P, chi-squared test; n = embryos). D. Proportion of non-injected cells and cells injected with the indicated mRNAs contributing to trophectoderm in the indicated genetic backgrounds. No significant differences were observed based on embryo genotype (P, chi-squared test; n = embryos). 43 Figure 2.4: LATS2 antagonizes formation of the apical domain. A. In embryos at 16-32 cell stages, PARD6B is detectable in GFP-overexpressing and in non-injected cells, but not in Lats2-overexpressing cells (arrowheads, n = embryos). B. At 16-32 cell stages, aPKCz is detectable in GFP-overexpressing and in non-injected cells, but not in Lats2-overexpressing cells (arrowheads, n = embryos). C. Quantification of embryos shown in panel A (p, student’s t-test). D. Quantification of embryos shown in panel B (p, student’s t-test). E. At 16-32 cell stages, CDH1 is localized to the basolateral membrane in both Lats2- overexpressing and non-injected cells (n = embryos). F. At 16-32 cell stages, Phalloidin staining demonstrates that filamentous Actin is apically enriched in Lats2-overexpressing and non-injected cells (n = embryos). (See next page) 44 Figure 2.4 (cont’d): G. At 16-32 cell stages, pERM is localized to the apical membrane in both Lats2- overexpressing and non-injected cells (n = embryos). 45 Figure 2.5: Wwtr1 and Yap1 are required to repress SOX2 expression in outside cells. A. CDX2 and SOX2 in wild type embryos at E3.25 (16-32 cell stages). CDX2 staining is more intense in outside cells than inside cells and SOX2 staining is specific to inside cells (n = embryos). B. Embryos lacking maternal Wwtr1 and Yap1 with and heterozygous for Wwtr1 and Yap1 (which we consider to have 2 doses of WWTR1/YAP1) exhibit normal CDX2 and SOX2 expression (n = embryos). (See next page) 46 Figure 2.5 (cont’d): C. Embryos lacking maternal Wwtr1 and Yap1 and heterozygous for either Wwtr1 or Yap1 (1 dose of WWTR1/YAP1) exhibit a high degree of ectopic SOX2 in outside cells (arrowheads), but continue to express CDX2, although the levels appear reduced (n = embryos). D. Embryos lacking maternal and zygotic Wwtr1 and Yap1 (0 doses of WWTR1/YAP1) have the most severe phenotype, with a high degree of ectopic SOX2 in outside cells (arrowheads) and little or no detectable CDX2 (n = embryos). E. Quantification of the percentage of outside cells in which ectopic SOX2 is detected in the presence of decreasing dose of Wwtr1 and Yap1 (t = student’s t-test, n = embryos). F. Quantification of the percentage of outside cells in which CDX2 is detected in the presence of decreasing dose of Wwtr1 and Yap1 (t = student’s t-test, n = embryos). 47 Figure 2.6: Positioning and epithelialization defects in embryos with Wwtr1 and Yap1 null alleles. A. Quantification of the average number of inside cells per embryo with decreasing dose of Wwtr1 and Yap1. The number of inside cells increases as the dose of wild type Wwtr1 and Yap1 alleles is reduced (p, student’s t-test, n = embryos). (See next page) 48 Figure 2.6 (cont’d): B. Quantification of the average number of outside cells per embryo with decreasing dose of Wwtr1 and Yap1. The number of outside cells decreases as the dose of wild type Wwtr1 and Yap1 alleles is reduced (p, student’s t-test, n = embryos). C. Quantification of the average number of total cells per embryos with decreasing dose of wild type zygotic Wwtr1 and Yap1. The number of total cells decreases as the dose of wild type Wwtr1 and Yap1 is reduced (p, student’s t-test, n = embryos). D. Quantification of the average ratio of inside to outside cells per embryo with decreasing dose of Wwtr1 and Yap1. The ratio of inside to outside cells increases as the dose of wild type Wwtr1 and Yap1 is reduced (p, student’s t-test, n = embryos). E. Wild type embryos at E3.75 exhibit inner cell mass-specific expression of SOX2 (n = embryos). E’. E3.75 embryos lacking maternal Wwtr1 and Yap1 and heterozygous for zygotic Wwtr1 and Yap1 cavitate and repress Sox2 in outside cells, leading to inner cell mass- specific expression of SOX2 similar to wild type embryos (n = embryos). E’’. Embryos lacking maternal Wwtr1 and Yap1 but with only one wild type allele of Wwtr1 or Yap1 fail to cavitate and repress Sox2 in outside cells, leading to ectopic SOX2 in outside cells (arrowheads, n = embryos). E’’’. Embryos lacking maternal and zygotic Wwtr1 and Yap1 fail to cavitate and repress Sox2 in outside cells, leading to ectopic SOX2 in outside cells (arrowheads, n = embryos). (See next page) 49 Figure 2.6 (cont’d): F. Quantification of ectopic SOX2 detected in embryos such as those shown in panels E- E’’’. The percentage of outside cells with ectopic SOX2 increases as the dose of wild type Wwtr1 and Yap1 alleles is reduced (p, student’s t-test, n = embryos). G. TUNEL analysis of embryos lacking maternal Wwtr1 and Yap1 heterozygous for zygotic Wwtr1 and Yap1 or lacking maternal and zygotic Wwtr1 and Yap1. Extensive TUNEL staining is observed in embryos lacking maternal and zygotic Wwtr1 and Yap1 indicative of cell death. Max projections of all confocal sections from a single embryo are shown (n = embryos). H. aPKCz staining in embryos lacking maternal Wwtr1 and Yap1, either heterozygous for zygotic Wwtr1 and Yap1 or with no zygotic Wwtr1 and Yap1. aPKC is not localized to the apical membrane of embryos with no zygotic Wwtr1 and Yap1 (n = embryos). I. ZO-1 staining in embryos lacking maternal Wwtr1 and Yap1, either heterozygous for zygotic Wwtr1 and Yap1 or with no zygotic Wwtr1 and Yap1. ZO-1 is disorganized in embryos with no zygotic Wwtr1 and Yap1, suggesting that formation of a mature epithelium depends on Wwtr1 and Yap1 (n = embryos). J. pERM and CDH1 staining in embryos lacking maternal Wwtr1 and Yap1, either heterozygous for zygotic Wwtr1 and Yap1 or with no zygotic Wwtr1 and Yap1. pERM is localized to apical membranes and CDH1 to basolateral membranes regardless of the dose of wild type Wwtr1 and Yap1 alleles (n = embryos). 50 Figure 2.7: Resolution of cell fate conflicts in the preimplantation mouse embryo. A. The expression of Sox2 and Cdx2 is differentially sensitive to YAP1/WWTR1 activity, leading to co-expression of both lineage markers in cells when YAP1/WWTR1 activity levels are intermediate. (See next page) 51 Figure 2.7 (cont’d): B. During division from the 16- to the 32-cell stage, cells that inherit the apical membrane repress HIPPO signaling and maintain an outside position. However, cells that inherit a smaller portion of the apical membrane would initially elevate their HIPPO signaling. We propose that elevated HIPPO then feeds back onto polarity by further antagonizing PAR- aPKC complex formation, leading to a snowball effect on repression of Sox2 expression, and thus ensuring that SOX2 is never detected in outside cells because these cells are rapidly internalized or apoptosed. C. A closeup of the boxed region in panel B. In most outside cells, low LATS2 activity enables high levels of YAP1/WWTR1 activity, which repress Sox2 and apoptosis and promote Cdx2 expression and apical localization of aPKC and PARD6B, which in turn repress the HIPPO pathway. In rare outside cells, LATS2 activity becomes elevated, leading to lower activity of YAP1/WWTR1, which then leads these cells to become internalized or to undergo apoptosis. 52 CHAPTER 3: Gata6 and Oct4 Direct Proper Gene Expression in Trophectoderm Cells in Morula and Blastocyst Stage Mouse Embryos Tayler Murphy1, Tristan Frum2,3, Stephanie Hickey3, Michael A. Halbisen3, and Amy Ralston1,3 1) Genetics and Genome Sciences Graduate Program, Michigan State University, East Lansing, MI, 48824 2) Department of Internal Medicine, University of Michigan Medical School, Ann Arbor, MI, 48109 3) Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824 Tayler Murphy wrote and edited the chapter, performed all experiments except RNA- sequencing and created figures, performed data and statistical analysis. Tristan Frum performed RNA-sequencing and reviewed the chapter. Stephanie Hickey compiled gene sets for gene set enrichment analysis from Nowotschin et al. 2019. Michael Halbisen performed bioinformatics processing and analysis of raw RNA-seq alignments. Amy Ralston helped to write the abstract and results section and edited the chapter. This chapter is in preparation for submission to Developmental Biology. 53 3.1: Abstract During mammalian development, the first cell fate decision leads trophectoderm cells (placenta lineage) to surround an inner cell mass (ICM – epiblast and extraembryonic endoderm lineages). Lineage-specific changes in gene expression are fundamental to this process and to healthy pregnancy. Trophectoderm versus ICM- specific gene expression differences are known to be induced by differential activity of the HIPPO signaling pathway, via a transcriptional complex that includes YAP1. In trophectoderm cells, the nuclear YAP1 complex drives expression of trophectoderm genes such as Cdx2, while the YAP1 complex is predominantly cytoplasmic within ICM cells. Additionally, we and others previously reported aberrant expression of CDX2 within the ICM of embryos lacking either OCT4 or GATA6 – two transcription factors that are essential for early embryo development. These observations raise the possibility that OCT4 and GATA6, which are coexpressed in ICM cells, could act cooperatively to repress trophectoderm gene expression in the ICM. Consistent with this hypothesis, our whole-transcriptome analysis of ICM cells lacking either factor reveal remarkable similarities. To understand the mechanism by which OCT4 and GATA6 regulate trophectoderm gene expression in the ICM, we evaluated YAP1 subcellular localization ICM cells in the absence of either factor. We observed that nuclear localization of YAP1 and CDX2 expression are detected in even control ICM cells of early and mid- blastocysts. Curiously, we found that expression of CDX2 and YAP1 are repressed in TE cells when GATA6 is lost and CDX2 is repressed when OCT4 is lost. In Gata6 null TE cells we also observed prolonged expression of OCT4 when compared to control. These observations suggest that GATA6 and OCT4 may be important factors for regulating the 54 expression of YAP1 in trophectoderm cells. Altogether, our studies deepen our understanding of the roles of OCT4 and GATA6 in ICM mouse embryonic development and reveal previously unknown roles in TE development. 3.2: Introduction Embryogenesis is one of the most important and complex biological phenomena. The ability to create a successful embryo is literally a matter of life and death. From zygote to fetus the embryo undergoes many stages, many changes, and many opportunities for failure or success. Understanding what makes a successful embryo can influence stem cell studies, human fertility treatment outcomes, and basic understanding of genetics. Mouse embryos undergo the same cell fate decisions as human embryos but at a much faster rate, making them a good starting point in studying embryogenesis. The first cell fate decision causes embryos to differentiate, inner cell mass (ICM), and outside, trophectoderm (TE) cells. The next cell fate decision further differentiates the ICM into primitive endoderm (PE) and epiblast (EPI). This cell fate decision occurs just before implantation in the uterus. Problems with cell fate decisions at this stage can impact implantation success. Therefore, it is vital that we understand the factors, conditions and mechanisms required for initiation and maintenance of embryonic cell fates. Transcription factors in the preimplantation mouse embryo are key to cell fate decisions as they help maintain the delicate balance of what is being expressed. At the blastocyst stage, CDX2 has been implicated in proper differentiation of TE from ICM (Strumpf et al. 2005). Loss of CDX2 causes the failure to restrict OCT4 and NANOG expression to the ICM (Strumpf et al. 2005). OCT4 has been shown to be necessary for 55 proper development of the ICM, both primitive endoderm and epiblast cells (Nichols et al. 1998, Frum et al. 2013, Le Bin et al. 2014). The loss of OCT4 leads to ectopic expression of CDX2 in some but not all cells within the ICM (Frum et al. 2013). Additionally, previous studies have shown that loss of GATA6 also leads to ectopic expression of CDX2 in some cells of the ICM, at the same stage as the loss of OCT4 phenotype (Schrode et al. 2014). However, the mechanism(s) linking OCT4 and GATA6 to TE gene expression are unknown. Mouse embryo development relies on signaling pathways that can affect the actions of transcription factors. Proper differentiation and localization of inside and outside cells in mouse embryos requires HIPPO pathway signaling (Nishioka et al. 2009, Frum et al. 2018). When HIPPO signaling is on, the LATS kinase phosphorylates YAP, causing it to remain in the cytoplasm and suppresses TEAD4, leading to inside cell gene expression (Nishioka et al. 2009). Outside cells have inactive HIPPO signaling, nuclear YAP and active TEAD4, which actively drives CDX2 expression (Nishioka et al. 2009). HIPPO signaling in inside cells has also been shown to regulate pluripotency marker SOX2 expression and resolve cell fate conflicts via changes to cell localization (Frum et al. 2018). However, it is not fully understood if there are differences in the HIPPO signaling pathway between the inside epiblast and primitive endoderm cell subtypes. Our previous work has shown that the FGF pathway is responsible for driving proper inside cell fate; epiblast cells produce FGF4, which is secreted and bound by FGFR2 on the membrane of primitive endoderm cells (Frum et al., 2013). This causes a signaling cascade, eventually leading to upregulation of PE specific factors, SOX17, GATA6, GATA4 and the downregulation of CDX2 (Frum et al., 2013). 56 Studies of OCT4 and GATA6 frequently focus only on ICM phenotypes, however, it is well known that GATA6 has sustained expression in the TE at mid-late blastocyst stage (Schrode et al., 2014; Meng et al., 2018; Cai et al., 2008), and OCT4 is not completely repressed in the TE until after early blastocyst stage (Palmieri et al., 1994). We previously showed loss of OCT4 to be detrimental to the TE, but this was attributed to overall embryo death and not a specific TE phenotype (Frum et al., 2013). We have also noted that previous studies from us and others have shown intriguing changes to the TE in Gata6 null and Oct4 null embryos, though not commented on in those papers. Loss of OCT4 seems to cause increased and/or prolonged expression of GATA6 in TE at E3.75 and E4.0 compared to wild type (Frum et al., 2013). Based on visual interrogation of immunofluorescence intensity, loss of OCT4 may also be causing decreased levels of CDX2 in TE cells (Frum et al., 2013). Similarly, loss of GATA6 may have caused decreased levels of CDX2 in E3.75 embryos, based on our visual interrogation of immunofluorescence images in Bessonnard et al., 2014. In this study we aim to explore the possible role of GATA6 and OCT4 in TE development, as well as further understand their role in repressing TE gene expression in the ICM. 3.3: Results 3.3.1: Whole Transcriptome Analysis of ICMs from Gata6 Null Embryos and Oct4 Null Embryos Show Increase in Trophectoderm Specific Gene Expression To investigate the role of GATA6 and OCT4 in the development of primitive endoderm we performed whole transcriptome analysis of GATA6 knock-out (KO) and OCT4 KO ICM cells. We performed statistical analyses to find genes that were differentially expressed in our Gata6 null or Oct4 null embryos (Fig. 3.1A,F). Gata6 null 57 ICMs had >300 differentially expressed genes (Fig. 3.1B). We noticed multiple trophectoderm specific genes (Mt1, Tagln2, Krt18, and Id2) were significantly and consistently upregulated in Gata6 null embryo ICMs. Prior study of Gata6 null embryos using immunofluorescence revealed ectopic expression of the TE protein CDX2 in a rare subset of ICM cells (Schrode et al., 2014). However, according to our transcriptional analysis Cdx2 was not significantly upregulated in Gata6 null ICMs. We repeated this analysis with RNA-seq data from Oct4 null ICMs. We observed that Oct4 null ICMs exhibited substantially fewer differentially expressed genes than the Gata6 null ICMs (60 genes compared to >300) (Fig. 3.1G). Consistent with our prior report (Frum et al., 2013), individual trophectoderm genes were not significantly upregulated in ICM at this stage. These observations suggested that GATA6 and OCT4 play different roles in ICM development. To further characterize the gene expression differences among these mutants, we performed gene set enrichment analysis (GSEA) on our transcriptome data. Our GSEA focused on the three early lineages: TE, PE, and EPI. We created EPI, PE, and TE gene sets using published single cell RNA-seq data from blastocysts (Nowotschin et. al., 2019). As expected, in the Gata6 null ICM transcriptome, the PE gene set was negatively enriched (Fig. 3.1C). In addition, the EPI gene set was not strongly enriched for in the Gata6 null ICM transcriptome (Fig 3.1D). However, the TE gene set was enriched in our Gata6 null ICM transcriptome (Fig. 3.1E). This observation is consistent with our differential gene expression analysis described above. In Oct4 null ICMs, the GSEA showed that PE genes were negatively enriched (Figure 1H), as expected (Frum et al., 2013; Le Bin et al., 2014). Some EPI genes were 58 slightly enriched, and others were slightly negatively enriched (Fig. 3.1I). Curiously, TE genes were enriched in Oct4 null ICMs (Fig. 3.1J). This was somewhat surprising since we had not detected any individual TE gene that was significantly different in our RNA- seq data. However, the advantage of performing GSEA is that it reveals cohorts of genes that, while not individually significantly different, do differ in expression as a set. Moreover, this observation reveals that Gata6 and Oct4 null do indeed phenocopy in terms of ectopic TE gene expression. These observations raise the possibility that GATA6 and OCT4 work together to repress expression of TE genes in the ICM. 3.3.2: Morula and Blastocyst Stage Embryos have Rare Instances of CDX2 Expression and Nuclear YAP Localization Our analysis failed to show increased Cdx2 expression in Gata6 null ICMs, which was contrary to the immunofluorescence analysis performed by Schrode et al. To begin to reconcile this apparent discrepancy, we repeated the immunofluorescence analysis of Gata6 null embryos. We observed a slightly increased ratio of inside to outside cells in embryos with ≤ 64 cells which is resolved after the 64-cell stage (Fig. 3.2A). This increased proportion of inside cells lead us to consider HIPPO signaling involvement, because we have seen an increased proportion of inside cells related to HIPPO pathway mutations (Frum et al., 2018). Therefore, in addition to examining CDX2 in Gata6 null embryos we also examined localization of YAP. Typically, YAP exhibits nuclear localization in the TE and cytoplasmic localization in ICM cells. Like Schrode et al., we observed rare instances of CDX2 expression in ICM cells of Gata6 null ≤ 64 cell embryos; however, this was also seen in our controls (Fig. 3.2B). We also noted rare nuclear localization of YAP (nYAP) in both control and Gata6 null ICMs (Fig. 3.2C). 59 However, only some of those cells expressing CDX2 co-expressed nYAP (Fig. 3.2D), indicating that the rare CDX2 expression seen in the ICM are not triggered by instances of nYAP localization. Additionally, Schrode et al. indicated that the ectopic CDX2 phenotype was present from the 64 cell to the 128 cell stage. To further investigate this phenotype, we examined CDX2 in Gata6 null embryos at this stage. Our analysis showed that, in Gata6 null embryos with > 64 cells there was expression of CDX2 in some ICM cells; however, this was also seen in our control embryos (Fig. 3.3A,B). Additionally, we investigated the localization of YAP in the CDX2+ ICM cells. Similarly to the ≤ 64 cell embryos, we observed rare expression of nYAP in both control and Gata6 null embryos with no difference between number of cells expressing nYAP (Fig. 3.3D,E). There was a similar amount of CDX2 and nYAP co-expression in control and Gata6 null >64 cell embryos. (Fig. 3.3G). Therefore, we surmise that the rare CDX2 expressing cell in the ICM may be a normal feature of morula and early blastocyst embryos, which is consistent with our RNA-seq analysis, as it did not show an increase in Cdx2 expression between control and Gata6 null ICMs at E3.75. 3.3.3: GATA6 Promotes Expression of Trophectoderm Genes and Repression of ICM Genes in the TE of Early Blastocysts Interestingly, we observed consistent defects in trophectoderm cell gene expression in Gata6 null early blastocyst. Firstly, the number of cells expressing CDX2 was lower in the trophectoderm of Gata6 null embryos with ≤64 cells than in control embryos at the same stage (Fig. 3.2E, F). Additionally, the levels of CDX2 expression were lower in Gata6 null embryos, based on fluorescence intensity of litter mate control 60 (Fig. 3.2F,G). Trophectoderm marker KRT8 expression was also lower in Gata6 null embryos compared to litter mate controls (Fig. 3.2H,I). These findings reveal a compelling role of GATA6 in promoting trophectoderm gene expression within the trophectoderm in the early mouse blastocyst. Subsequently, we decided to further explore this CDX2 phenotype in Gata6 null embryos at a later stage. It is well established that Gata6 null embryos successfully implant in the uterus but ultimately die between E6.5 and E7.5 (Morrisey et al., 1998). Is the lack of CDX2 inconsequential to successful implantation, or does CDX2 expression recover prior to that stage via a non-GATA6 mediated mechanism? In Gata6 null embryos with > 64 cells (65-109 cells) CDX2 expression in trophectoderm was comparable to control embryos (Fig. 3.3A,C). Additionally, expression of nYAP in the trophectoderm of Gata6 null > 64 cell embryos was the same as in controls (Fig. 3.3D,F). The amount of trophectoderm cells co-expression nYAP and CDX2 in these embryos were also similar to control embryos (Fig. 3.3H). This suggests that the CDX2 phenotype we noted in earlier stage Gata6 null embryos is transient in nature. Additionally, the slight increase in inside:outside cell ratio in Gata6 null ≤ 64 cell embryos is resolved after the 64 cell stage (Fig. 3.2A), which suggests a possible compensating mechanism that promotes CDX2 in trophectoderm, in the absence of GATA6. Having parallel mechanisms to ensure proper trophectoderm development at this stage would provide robustness since it is just prior to implantation for which the TE is a crucial component. Additionally, we analyzed the expression of CDX2 and NANOG expression in Gata6 null embryos. We observed increased NANOG expression in Gata6 null embryos, both in number of cells expression NANOG and intensity of NANOG fluorescence, as 61 early as the 8-cell stage (Fig. 3.4A-C). The increased number of NANOG expressing cells in Gata6 null continued to 32-63 cell embryos. However, only in outside cells, the number of NANOG expressing cells in the inner cell mass of Gata6 null embryos were similar to controls (Fig. 3.4 D,E). A previous study found ectopic NANOG expression in the trophectoderm at early and mid-blastocyst stage Gata6 null embryos (Schrode et al., 2014). This previous study also concluded that the loss of CDX2 does not seem to be caused by gain of NANOG in Gata6 null embryos, as not all NANOG expressing trophectoderm cells lost CDX2 expression (Schrode et al., 2014). Our results taken with the previous study indicating that loss of CDX2 in outside cells does not correlate to acquisition of NANOG expression (Schrode et al., 2014), suggests that GATA6 regulates CDX2 expression in the trophectoderm independent of its repression of NANOG. Given the NANOG phenotype we decided to investigate another ICM marker, OCT4. OCT4 expression is normally restricted to ICM cells at around the blastocyst stage. We decided to evaluate expression of OCT4 at both early and mid-blastocyst stage control and Gata6 null embryos. Firstly, the expression of OCT4 in the ICM of our ≤ 60 cell Gata6 null embryos was normal compared to control (Fig. 3.2K). Expression of OCT4 in outside cells was also normal in ≤ 60 cell Gata6 null embryos (Fig. 3.2J). However, we observed prolonged OCT4 expression in the TE of > 60 cell Gata6 null embryos compared to controls, which had already downregulated OCT4 in the TE (Fig. 3.2J). This suggests that GATA6 is necessary for repression of OCT4 in the trophectoderm. 62 3.3.4: OCT4 Promotes CDX2 Expression and Nuclear YAP Localization in Trophectoderm Cells Therefore, we decided to investigate whether OCT4 plays a role in TE gene expression prior to it being repressed. First, we examined TE markers in Oct4 null >64 cell embryos by immunofluorescence. We detected expression of CDX2 in ICM cells of Oct4 null embryos, similarly to our observation in Gata6 null embryos, it was not more than seen in controls (Fig. 3.5B). Additionally, nuclear YAP expression was seen occasionally in Oct4 null >64 cell embryos but not significantly over control levels (Fig. 3.5C). Co-expression of nYAP and CDX2 was also seen occasionally in the ICM of Oct4 null embryos; however, not significantly different than in controls (Fig. 3.5D). Interestingly, the number of TE cells expressing CDX2 was lower in Oct4 null >64 cell embryos than in control embryos (Fig. 3.5F), which has not been previously reported. However, the number of cells expressing nYAP and co-expressing CDX2 and nYAP in the TE of in Oct4 null >64 cell embryos was similar to control (Fig. 3.5F,G). These observations suggest a previously unrecognized role for OCT4 in promoting gene expression in the TE. This provides evidence that OCT4 and GATA6 are working in conjunction to promote CDX2 expression in the TE. 3.4: Discussion Our observations implicate GATA6 and OCT4 in guiding proper trophectoderm gene expression in early mouse blastocysts. Previous studies and our transcriptome analyses of Gata6 null and Oct4 null embryo ICMs suggest that these factors may be doing the opposite within ICM cells, by repressing TE gene expression. The possibility that OCT4 and GATA6 may regulate expression of CDX2 and repression of ICM genes 63 in the TE is interesting. Further work needs to be done to better understand possible mechanisms of how these factors are impacting TE gene expression in the TE. It has previously been shown that the ICM signals to the TE via BMP signaling (Graham et al., 2014; Frias-Aldeguer et al., 2019). It has also been shown that GATA6 regulates Bmp4/7 by binding enhancer regions (Whissell et al., 2014). Moreover, BMP4 leads to nuclear localization of TEAD4, which can bind to YAP that is also localized in the nucleus (Home et al., 2012). One possible non-cell-autonomous mechanism we propose is that loss of GATA6 leads to loss of BMP4/7 ligand production in the ICM, preventing TEAD4 and YAP from localizing in the nucleus to allow for CDX2 expression (Fig. 3.6A). Another significant signaling pathway working in the pre-implantation mouse embryo is FGF4 signaling. A previous study, using exogenous FGF4 in cultured Gata6 null embryos seems to indicate that GATA6 is not promoting CDX2 expression in the TE through FGF4 signaling if our interpretation of their data is correct (Bessonard et al., 2014). This study does not explicitly note a reduction in CDX2 expression in Gata6 null embryos, however CDX2 appears fainter in E3.75 Gata6 null embryos compared to their controls, and this seems to continue in their embryos cultured in FGF4 (Bessonard et al., 2014). However, OCT4 may non-cell-autonomously promote CDX2 expression through FGF4 signaling (Fig. 3.6B). It has previously been shown that TE cells express relatively high FGFR2, the receptor that binds FGF4 (Molotkov et al., 2017). FGF4 is produced in and secreted from epiblast cells in the ICM, and Oct4 null embryos have been shown to have reduced FGF4, so it is possible that OCT4 may be involved in FGF signaling between the ICM and TE (Nichols et al., 1998). This proposed mechanism is not mutually exclusive with our proposed GATA6/BMP4 pathway. 64 Additionally, it must be considered that GATA6 and/or OCT4 are acting cell- autonomously in the TE. It has previously been shown that GATA6 is expressed in the TE (Molotkov et al., 2017), and OCT4 is initially expressed in all cells before being relegated to the ICM. This leads to the possibility that GATA6 and/or OCT4 may be working cell-autonomously as pioneer factors/ transcription factors in the early TE to allow for expression of CDX2 (Fig. 3.6C,D). Overall, our findings suggest there are still important things to be learned about how and what drives ICM and TE fate. 3.5: Materials and Methods 3.5.1: Animal Usage and Genotyping All animals were used and kept in accordance with Michigan State University Institutional Animal Care and Use Committee. These experiments were completed using embryos from matings of Gata6-/+ (Gata6tm2.2Sad/+) mice (Sodhi et al. 2006) and matings of Oct4-/+ (created using Oct4tm1Scho x 129-Alpltm1(cre)Nagy) mice (Frum et al., 2013; Kehler et al., 2004; Lomeli et al., 2000). The Gata6- allele and Oct4- allele were maintained in a CD-1 background. Mice were genotyped using genomic DNA extracted from mouse ear punches, using the Extract-N-Amp tissue PCR kit (Sigma). Wild type and Gata6 null PCR were performed in a BioRad S1000 Thermocycler using the primers in the table below (Battle Lab). Wild type and Oct4 null PCR was performed using the primers in the table below (Kehler et al., 2004). 65 Gata6 wild type primers Oct4 null primers Gata6 null primers F: GCTCCACCCTACTATGACCAATTCC R: CCCGGTTTAAAAATCTGCTTGAGTC F: GTGGTTGTAAGGCGGTTTGT R: ACGCGAGCTCCAGAAAAAGT F: AACTGGTTTGTGAGGTGTCCG R: GTATCCACTCGCACCTTGTTC F: TTGTTACTGAAGAGGTTGGGTGTGACTGG R: GGGGACTCCTGCTACAACAATCGCTAAG Oct4 wild type primers Table 3.1: Genotyping Primers for Gata6 Null, Oct4 Null, and Control Alleles. ~400bp 159bp 245bp 415bp 3.5.2: Embryo Collection Practiced Gata6tm2.2Sad/+ males were mated to Gata6tm2.2Sad/+ females and practiced Oct4-/+ males were mated to Oct4-/+ females. Females were checked for copulatory plugs and day of copulatory plug was counted as E0.5. Embryos were harvested on E3.75 via dissection of the uterus and flushing each horn with M2 medium (Millipore), warmed to 37◦ C. Embryos were rinsed in fresh M2 medium and then immediately fixed at room temp using 4% formaldehyde in PBS for 10 minutes. Embryos are then permeabilized in 0.5% Triton-X in PBS at room temp for 30 minutes, and blocked with 10% Fetal Bovine Serum and 1% Triton-X in PBS overnight at 4◦ C. 3.5.3: Immunofluorescence and Confocal Imaging After fixing, permeabilizing, and blocking; embryos were incubated in primary antibodies, as indicated on figures; mouse anti-OCT4 (1:10) (Santa Cruz, 5279), rat anti- KRT8 (1:10) (Developmental Studies Hybridoma Bank, TROMA-I-c), mouse anti-YAP (1:200) (Santa Cruz Biotechnology, sc101199), rabbit anti-CDX2 (1:200) (Abcam, ab76541), or rabbit anti-NANOG (1:400) (Reprocell, RCAB002P-F) in blocking solution overnight at 4◦ C. Embryos were washed in blocking solution for 30 minutes at room temperature, then moved to corresponding secondary in blocking, donkey anti-mouse Cy3 (1:400) (Jackson Immuno Research, 715-165-150), donkey anti-rabbit Alexa 488 66 (1:400) (Jackson Immuno Research, 711-485-152), or donkey anti-rat Cy3 (1:400) (Jackson Immuno Research, 712-165-150), for 1 hour at room temp in the dark. They were then moved to DRAQ5 stain (1:400) (Cell Signaling, 4084S) for 30 minutes at room temp in the dark. After staining, embryos were imaged using an Olympus Fluoview FV1000 Confocal microscope. Optical sections were taken over the whole embryo in 5um increments. 3.5.4: Embryo Genotyping DNA was extracted from embryos using the Extract-N-Amp tissue PCR kit to a volume of 10ul, then using the same thermocycler, oligos and settings as described above for the PCR. 3.5.5: Embryo Analysis Embryos were analyzed using the software FIJI (Schindelin et al. 2012) and Adobe CC Photoshop 2019. FIJI is used to make montages of each plane of the embryo and each channel plus a merge of all the channels. These montage images were then opened as PNGs in Photoshop and virtually labeled with the brush tool. Once labeled all nuclei were categorized based on the expression pattern and recorded in an Excel spreadsheet. In this study, control embryos include wild type and the respective heterozygotes. Statistical analysis and visualization of expression data was done using Graphpad Prism for Windows version 9.4.1 or 9.5.0. 3.5.6: Immunosurgery and Library Prep To isolate inner cell mass cells from Gata6 null, Oct4 null and wild type embryos for RNA-sequencing, immunosurgery was performed (Solter and Knowles; 1975). Embryos were collected using the method above but at E3.75 and stopping at the M2 67 medium rinse. Embryos were stored in the pre-warmed M2 medium while preparing Tyrode’s Acid and KSOM plate. 3.5.7: RNA Isolation from embryos and qPCR Isolation of RNA and PCR based amplification was performed according to (Tang et al., 2010). Genotyping of embryos used in RNA-sequencing was performed by qPCR on PCR2 product (step 40 of Tang et al., 2010) using primers spanning an intron with one primer located in the floxed exon for each conditional. Actin was used as the housekeeping gene. qPCR Primer Sets used are in the table below. GATA6GT Ex 2 F:TCCACAGCTTACAGGGCC Exon R:CCGTCTCGTCTCCACAGTG 2-3 Pou5f1 exon 1 F:GTTGGAGAAGGTGGAACCAA Exon R:CCAAGGTGATCCTCTTCTGC 1-2 Actb B F:CTGAACCCTAAGGCCAACC Exon R:CCAGAGGCATACAGGGACAG 3-4 Table 3.2: Primers for qPCR Genotyping of the Embryos Used in RNA-sequencing. 3.5.8: RNA-Sequencing and post sequencing processing Sequencing was performed by the RTSF core at Michigan State University on an Illumina HiSeq 4000 to produce single-end reads, using HiSeq 3000/4000 SBS Kit (50 cycles) (FC-410-1001). Post sequencing processing began with Trimmomatic/0.32 (Bolger et al. 2014) and AdapterRemoval v1 (Lindgreen, S. 2012) to remove unwanted adapter sequences. Next, FastQC/0.11.3 (https://www.bioinformatics.babraham.ac.uk/projects/fastqc/) was 68 used for assessing post-trimming quality control. Resultant reads were mapped using TopHat v2.0.12 with the mm9 UCSC genome (https://genome.ucsc.edu/), then HTSeq/0.6.12 (Putri et al. 2021) was used to count reads for each annotated transcript. 3.5.8: Bioinformatic analysis All bioinformatic analyses were performed in R/3.3.1 (R Core Team 2016). 2. EdgeR 3.24.3 (Robinson et al. 2010 , McCarthy et al. 2012, Chen et al. 2016) was used for ordination and MDS plots, as well as differential gene expression analyses. 3. Pairwise Spearman correlations (Spearman 1904, Glasser and Winter 1961) were calculated for each sample, and the heatmap.2 function of gplots 3.0.1 (Warnes et al. 2016) were used to generate heat maps. Volcano plots were generated using the ggplot2 3.3.0 package (Wickam 2016). Results Quality Control Analysis Each sample was sequenced to a depth of ~50 million reads (n = 33.5-74.5), and sample quality was verified with FASTQC. 3.5.9: GSEA Analysis Single-cell RNA-seq data generated from mouse E3.5 blastocysts by Nowotschin et. al. were used to identify primitive endoderm (PrE), epiblast (EPI) and trophectoderm (TE) enriched genes. Specifically, we followed the Satija Lab’s Introduction to SCTransform, v2 regularization vignette, and used scTransfrom v0.3.5 from Seurat v4.3.0.9001 to normalize counts for each single-cell RNA-seq library (Lib1-3 and Lib1-4) and the FindNeighbors function to identify each cell’s 20 nearest neighbors using the first 30 principal components of the gene expression. Data from the two libraries were subsequently integrated using 3000 features to find integration anchors. 69 We used the FindConservedMarkers function with the Wilcoxon rank-sum test to identify genes enriched in each cell type versus all other cells in each library. Only fold changes >.25 from genes expressed in >2 cells and >0.1% of the cells in either of the two populations were reported. The library specific p-values were combined using Tippett’s method and corrected for multiple comparisons using the Benjamini–Hochberg method. We found cell-type enriched genes by comparing the following clusters: 1) Each cluster as defined by Nowotschin plus TE cells: Nowotschin et al. identified five clusters of cells at E3.5, corresponding to nascent PrE (E3.5:0), more advanced PrE (E3.5:4), uncommitted ICM cells (E3.5:2), ICM cells that had started to specify (E3.5:1) and EPI cells (E3.5:3). While the authors identified TE, these cells were not included in the original analysis. We identified cluster enriched genes for all five E3.5 clusters plus the TE cluster by comparing the gene expression in each cluster versus the cells from all other clusters. 2) General cell type clusters: Here, we collapsed the nascent PrE and more advanced PrE clusters into one general PrE cluster and uncommitted ICM cells and ICM cells that had started to specify into one general ICM cluster. We identified cluster enriched genes for general PrE, general ICM, EPI, and TE clusters by comparing the gene expression in each cluster versus the cells from all other clusters. 3) No ICM: This analysis was performed as in analysis #2 without including the general ICM cluster. 4) EPI and PrE only: This analysis was performed as in analysis #2 without including the general ICM cluster or the TE cluster. 70 For comparisons 3 and 4, the data was renormalized and integrated as above before enriched genes were identified. Once genes enriched in each cell type were established (Nowotschin et al. 2019). Gene Set Enrichment Analysis was conducted on the differentially expressed genes found in the RNA-seq of Gata6 null and wild type embryo ICMs. 71 FIGURES Figure 3.1: GATA6 and OCT4 Repress Trophectoderm Gene Expression Within the Inner Cell Mass (ICM) of Mouse Blastocysts. A. Heatmap showing significantly differentially expressed genes in Gata6 null ICMs, FDR <0.10 n= 301 genes. B. Volcano plot of differentially expressed genes in Gata6 null ICMs. C. Gene set enrichment plot of primitive endoderm genes in Gata6 null ICMs. D. Gene set enrichment plot of epiblast genes in Gata6 null ICMs. E. Gene set enrichment plot of trophectoderm genes in Gata6 null ICMs. F. Heatmap showing significantly differentially expressed genes in Oct4 null ICMs, FDR <0.10 n= 63 genes. G. Volcano plot of differentially expressed genes in Oct4 null ICMs. (See next page) 72 Figure 3.1 (cont’d): H. Gene set enrichment plot of primitive endoderm genes in Oct4 null ICMs. I. Gene set enrichment plot of epiblast genes in Oct4 null ICMs. J. Gene set enrichment plot of trophectoderm genes in Oct4 null ICMs. 73 Figure 3.2: GATA6 is Necessary for Proper Expression of CDX2 and KRT8 Prior to the 64-Cell Stage. (See next page) 74 Figure 3.2 (cont’d): A. Plot showing inside:outside cell ratio of control, ≤ 64 n=39; > 64 n= 14, and Gata6 null, ≤ 64 n=19; > 64 n=7, blastocysts at ≤ 64 and > 64 cells. B. Plot of CDX2+ inside cells in control, n= 33, and Gata6 null, n=15, ≤ 64 cell embryos. C. Plot of nYAP+ inside cells in control, n= 36, and Gata6 null, n=7, E3.75 embryos. D. Plot of Double+ inside cells in control, n= 36, and Gata6 null, n=7, E3.75 embryos. E. Plot of the percent of outside cells expressing CDX2 in control, n=33, Gata6 null, n=15, ≤ 64 cell embryos F. Single plane from confocal Z-Stack of control and Gata6 null blastocysts (≤ 64 cells), immunofluorescently stained for CDX2 and DNA (DRAQ5). G. Surface plots showing immunofluorescence intensity of CDX2 in control and Gata6 null blastocysts (≤ 64 cells). H. Single plane from confocal Z-Stack of control and Gata6 null blastocysts (≤ 64 cells), immunofluorescently stained for KRT8 and DNA (DRAQ5). I. Surface plots showing immunofluorescence intensity of KRT8 in control and Gata6 null blastocysts (≤ 64 cells). J. Plot showing percent of outside cells that are OCT4+ in control, n=4, and Gata6 null, n=3, blastocysts (≤ 60 cells). K. Plot showing percent of inside cells that are OCT4+ in control, n=4, and Gata6 null, n=3, blastocysts (≤ 60 cells). L. Single plane from confocal Z-Stack of control and Gata6 null blastocysts (≤ 60 cells), immunofluorescently stained for OCT4 and DNA (DRAQ5). 75 Figure 3.3: GATA6 is Not Necessary for Proper Expression of CDX2 and nYAP after 64 cells stage but is Required for Repression of OCT4 in Trophectoderm Cells after 60 cell stage. (See next page) 76 Figure 3.3 (cont’d): A. Single plane from confocal Z-Stack of control, n=10, and Gata6 null, n=5, blastocysts (> 64 cells), immunofluorescently stained for CDX2 and DNA (DRAQ5). B. Plot showing percent of inside cells expressing CDX2 in control, n=10, and Gata6 null, n=5, blastocysts (> 64 cells). C. Plot showing percent of outside cells expressing CDX2 in control, n=10, and Gata6 null, n=5, blastocysts (> 64 cells). D. Single plane from confocal Z-Stack of control, n=10, and Gata6 null, n=5, blastocysts (> 64 cells), immunofluorescently stained for CDX2 and DNA (DRAQ5). E. Plot showing percent of inside cells expressing nYAP in control, n=8, and Gata6 null, n=4, blastocysts (> 64 cells). F. Plot showing percent of outside cells expressing CDX2 in control, n=8, and Gata6 null, n=4, blastocysts (> 64 cells). G. Plot showing percent of inside cells expressing CDX2 and nYAP in control, n=8, and Gata6 null, n=4, blastocysts (> 64 cells). H. Plot showing percent of outside cells expressing CDX2 and nYAP in control, n=8, and Gata6 null, n=4, blastocysts (> 64 cells). I. Single plane from confocal Z-Stack of control and Gata6 null blastocysts (> 60 cells), immunofluorescently stained for OCT4 and DNA (DRAQ5). Orange arrows indicate TE cells that are OCT4 positive. J. Plot showing percent of inside cells expressing OCT4 in control, n=6, and Gata6 null, n=3, blastocysts (> 64 cells). (See next page) 77 Figure 3.3 (cont’d): K. Plot showing percent of outside cells expressing CDX2 and nYAP in control, n=8, and Gata6 null, n=4, blastocysts (> 64 cells). 78 Figure 3.4: GATA6 is Necessary for Preventing Premature NANOG Expression in 8-Cell Embryos and for Down Regulating NANOG in Outside Cells in Embryos with Between 32 and 63 Cells. A. Single plane from confocal Z-Stack of control, n=8 and Gata6 null, n=8, 6-8 cell embryos. Immunofluorescently stained for NANOG and DNA (DRAQ5). B. Surface plots showing immunofluorescence intensity of NANOG in C. Plot showing the percentage of cells expressing NANOG in control, n=8, and Gata6 null, n=8, 6-8 cell embryos. D. Plot showing the percentage of inside cells expressing NANOG in control, n=6, and Gata6 null, n=5, 32-63 cell blastocysts. E. Plot showing the percentage of outside cells expressing NANOG in control, n=6, and Gata6 null, n=5, 32-63 cell blastocysts. 79 Figure 3.5: OCT4 is Necessary for Maintaining Expression of CDX2 in the Trophectoderm in >64 Cell Blastocysts. A. Single plane from confocal Z-Stack of control, n=6, and Oct4 null, n=5, blastocysts >64-cells. Immunofluorescently stained for CDX2, YAP and DNA. B. Plot showing the percentage of inside cells expressing CDX2 in control, n=6, and Oct4 null, n=5, >64 cell embryos. C. Plot showing the percentage of inside cells expressing nYAP in control, n=6, and Oct4 null, n=5, >64 cell embryos. D. Plot showing the percentage of inside cells expressing CDX2 and nYAP in control, n=6, and Oct4 null, n=5, >64 cell embryos. E. Plot showing the percentage of outside cells expressing CDX2 in control, n=6, and Oct4 null, n=5, >64 cell embryos. (see next page) 80 Figure 3.5 (cont’d): F. Plot showing the percentage of outside cells expressing nYAP in control, n=6, and Oct4 null, n=5, >64 cell embryos. G. Plot showing the percentage of outside cells expressing CDX2 and nYAP in control, n=6, and Oct4 null, n=5, >64 cell embryos. 81 Figure 3.6: Possible Models for GATA6 and OCT4 Promotion of CDX2 Expression in Trophectoderm Cells. A. Possible non-cell-autonomous model of GATA6 promoting CDX2 expression via ICM to TE BMP4/7 signaling. (see next page) 82 Figure 3.6 (cont’d): B. Second possible non-cell-autonomous model of OCT4 promoting CDX2 expression via ICM to TE FGF4 signaling. C. Possible cell-autonomous model of GATA6 promoting CDX2 expression in the early blastocyst TE by acting as pioneer or transcription factor. D. Possible cell-autonomous model of OCT4 promoting CDX2 expression in the mid- blastocyst TE by acting as pioneer or transcription factor. 83 CHAPTER 4: Further Examination of OCT4 and GATA6 in Preimplantation Mouse Tayler M. Murphy Embryos 84 4.1 Introduction The second cell fate decision in preimplantation mouse embryos separates the inner cell mass into primitive endoderm and epiblast. Both epiblast and primitive endoderm cells require OCT4 for proper development (Nichols et al. 1998, Frum et al. 2013, Le Bin et al. 2014). OCT4 being necessary for two distinct cell fates, led me to ask how OCT4 is driving two cell-fates at the same stage. I hypothesized that OCT4 is working with a PE-specific cofactor GATA6. In chapter 3, I noted that OCT4 and GATA6 both individually were needed for repression of TE specific gene transcription in the ICM of E3.75 embryos. Additionally, I observed that loss of OCT4 or GATA6 caused reduction in CDX2 in mid-blastocyst and early-blastocyst stage embryos, respectively. The phenotypes I noted in embryos without OCT4 or GATA6 caused me to consider what would happen if both were lost, and conversely what would happen if either factor were overexpressed. I hypothesized that in Oct4 null; Gata6 null embryos I would observe a more severe phenotype than in the single nulls, such as a more pronounced loss of CDX2 in blastocysts, or worse, early embryonic lethality. Moreover, in OCT4 overexpressing embryos I hypothesized that SOX17 may be ectopically expressed in trophectoderm cells. The trophectoderm continues to express GATA6 at the early blastocyst stage (Koutsourakis et al., 1999; Plusa et al., 2008), therefore if OCT4 and GATA6 were sufficient to drive primitive endoderm development I would expect to see PE gene expression in TE cells forced to express OCT4 ectopically. 4.2 Results 4.2.1 Gata6 null; Oct4 null embryos may have very early lethality I hoped to further investigate the mechanisms by which OCT4 and GATA6 85 mediate trophectoderm gene repression in ICM cells, and ICM gene repression and TE gene expression in the TE. I hypothesized that if OCT4 and GATA6 were working together to drive these expression patterns loss of both would cause more severe phenotypes. However, I was unable to obtain any Oct4 null; Gata6 null embryos from seven litters; two at E2.5, two at E3.0, two at E3.5 and one at E3.75 (Fig. 4.1A). This indicates that loss of both OCT4 and GATA6 may lead to lethality prior to E2.5. Oct4 null embryo lethality has been shown to occur around implantation (Nichols et al., 1998). Gata6 null embryos die between E6.5 and E7.5 (Morrisey et al., 1998). Therefore, loss of OCT4 and GATA6 together seems to have a compounding effect leading to loss prior to either of the single null embryos. 4.2.2 Overexpressing Oct4 does not cause ectopic expression of primitive endoderm specific factor SOX17 Additionally, I attempted to determine the role that OCT4 has in primitive endoderm gene expression by overexpressing OCT4 in blastocysts using a doxycycline inducible promoter. Firstly, I examined expression of OCT4 in control and putative Oct4 overexpressing embryos (Fig. 4.2A,F). Across the whole embryo in both ≤ 50 cell and > 50 cell embryos, OCT4 expression increased (Fig. 4.2A,F). It is normal for most ICM cells to express OCT4 as seen in our control embryos and also seen in our Oct4 overexpressers (Fig. 4.2B,G). Next, to confirm overexpression of OCT4 I evaluated OCT4 expression in trophectoderm cells using immunofluorescence. Compared to expression of OCT4 in control embryo TE; Oct4 overexpressing embryos showed a large increase in expression of OCT4 in the TE (Fig. 4.2C,H). Though there were still some cells not expressing OCT4 in the ICM and TE of my Oct4 overexpressing embryos, the 86 increase seen in the TE is significant and shows that the doxycycline inducible promoter is turning OCT4 on in most TE cells. Furthermore, I examined expression of SOX17 in both ≤ 50 cell and > 50 cell control and Oct4 overexpressing embryos. SOX17 is a primitive endoderm specific factor, not normally found in the trophectoderm or epiblast cells within the ICM (Niakan et al., 2010). There was not an increase in cells expressing SOX17 in the ICM compared to control (Fig. 4.2D,I). Also, I did not see significant ectopic expression of SOX17 in the trophectoderm of ≤ 50 cell nor > 50 cell embryos (Fig. 4.2E,J). This indicated that this level of OCT4 expression is not sufficient to promote SOX17 expression in the epiblast nor the trophectoderm. 4.3 Discussion Overall, this study shows that loss of Oct4 and Gata6 is lethal prior to E2.5 in mouse embryos and that OCT4 expression in the trophectoderm is not sufficient to induce SOX17 expression. Given that Oct4 null and Gata6 null embryos can be successfully recovered past E2.5; this supports a compound effect of losing both proteins at the same time (Nichols et al., 1998; Morrisey et al., 1998). Further research needs to be done to determine when exactly lethality occurs in Oct4 null; Gata6 null embryos and what is leading to the compounded severity of the phenotype. I speculate that OCT4 and GATA6 may work together in parallel to promote proper gene expression in the ICM and TE, where the absence of one may compensate for the other, but when both are lost at the same time there is early lethality. A previous study in ES cells hypothesized that the level of OCT4 expression led to different cell fates; low expression leading to trophectoderm fate, moderate expression maintains epiblast fate, and high expression leads to primitive endoderm fate (Niwa et 87 al., 2000). This study adds to the evidence that this hypothesis is not relevant in vivo; as I did not see ectopic expression of SOX17. Further studies should be done to confirm this result, possibly looking at other PE specific factors like GATA4. 4.4 Methods 4.4.1: Animal Usage and Genotyping All animals were used and kept in accordance with Michigan State University Institutional Animal Care and Use Committee. These experiments were completed using embryos from matings of Gata6-/+ (Gata6tm2.2Sad/+) mice (Sodhi et al. 2006). As well as embryos from matings of Oct4-/+ (created using Oct4tm1Scho x 129-Alpltm1(cre)Nagy) mice (Frum et al., 2013; Kehler et al., 2004; Lomeli et al., 2000). The Gata6- allele and Oct4- allele were maintained in a CD-1 background. Mice were genotyped using genomic DNA extracted from mouse ear punches, using the Extract-N-Amp tissue PCR kit (Sigma). Wild type and Gata6 null PCR were performed in a BioRad S1000 Thermocycler using the primers in the table below (Battle Lab). Wild type and Oct4 null PCR was performed using the primers in the table below (Kehler et al., 2004). Overexpression experiments were completed using embryos from matings of rtTA;tet-on-Oct4 (derived from B6;129- Gt(ROSA)26Sortm1(rtTA*M2)Jae Col1a1tm2(tetO-Pou5f1)Jae/J from Jackson Labs, bred to CD-1) (Hochedlinger et al., 2005). 88 Oct4 wild type primers rtTA primers Gata6 wild type primers Oct4 null primers Gata6 null primers F: GCTCCACCCTACTATGACCAATTCC R: CCCGGTTTAAAAATCTGCTTGAGTC F: GTGGTTGTAAGGCGGTTTGT R: ACGCGAGCTCCAGAAAAAGT F: AACTGGTTTGTGAGGTGTCCG R: GTATCCACTCGCACCTTGTTC F: TTGTTACTGAAGAGGTTGGGTGTGACTGG R: GGGGACTCCTGCTACAACAATCGCTAAG F: AAAGTCGCTCTGAGTTGTTAT R: GCGAAGAGTTTGTCCTCAACC F: AAAGTCGCTCTGAGTTGTTAT R: GGAGCGGGAGAAATGGATATG F: GCAGAAGCGCGGCCGTCTGG R: CCCTCCATGTGTGACCAAGG F: GCACAGCATTGCGGACATGC R: CCCTCCATGTGTGACCAAGG Tet-on-Oct4 primers Col1a1 primers Rosa26 primers ~400bp 159bp 245bp 415bp 340bp 650bp ~500bp 300bp Table 4.1: Primers for Genotyping PCR. 4.4.2: Embryo Collection Practiced Gata6 null het or Oct4 null het males were mated to Oct4 null het or Gata6 null het females, respectively. Females were checked for copulatory plugs and day of copulatory plug was counted as E0.5. Embryos were harvested at the indicated stage via dissection of the uterus and flushing each horn with M2 medium (Millipore), warmed to 37◦ C. Embryos were rinsed in fresh M2 medium and then immediately fixed at room temp using 4% formaldehyde in PBS for 10 minutes. Embryos are then permeabilized in 0.5% Triton-X in PBS at room temp for 30 minutes, and blocked with 10% Fetal Bovine Serum and 1% Triton-X in PBS overnight at 4◦ C. Practiced rtTA;tet-on-Oct4 males were mated to rtTA;tet-on-Oct4 females. Females were checked for copulatory plugs and day of copulatory plug was counted as E0.5. Embryos were harvested at E2.5 via dissection of the oviducts and flushing them with M2 medium (Millipore), warmed to 37◦ C. Embryos were rinsed in fresh M2 medium and then moved to pre-equilibrated culture plates with EmbryoMax® KSOM Mouse 89 Embryo Media (Millipore), and 40ug/ml of doxycycline, covered with embryo culture grade light mineral oil. The embryos were incubated at 37◦ C with 5% CO2 for 36 hours. After culture, embryos were fixed as indicated previously. 4.4.3: Immunofluorescence and Confocal Imaging After fixing, permeabilizing, and blocking; embryos were incubated in one of the primary antibodies listed; mouse anti-OCT4 (1:10) (Santa Cruz, 5279), goat anti-SOX17 (1:200) (R&D Systems, AF1924) in blocking solution overnight at 4◦ C. Embryos were washed in blocking solution for 30 minutes at room temperature, then moved to secondary in blocking, donkey anti-mouse Cy3 (1:400) (Jackson Immuno Research, 715-165-150) and donkey anti-goat Alexa 488 (1:400) (Invitrogen, A-11055) for 1 hour at room temp in the dark. They were then moved to DRAQ5 stain (1:400) (Cell Signaling, 4084S) for 30 minutes at room temp in the dark. After staining, embryos were imaged using an Olympus Fluoview FV1000 Confocal microscope. Optical sections were taken over the whole embryo in 5um increments. 4.4.4: Embryo Genotyping DNA was extracted from embryos using the Extract-N-Amp tissue PCR kit to a volume of 20ul. Then using the same thermocycler, oligos as described above for the PCR. 4.4.5: Embryo Analysis Embryos were analyzed using the software FIJI (Schindelin et al. 2012) and Adobe CC Photoshop 2019. FIJI is used to make montages of each plane of the embryo and each channel plus a merge of all the channels. These montage images were then opened as PNGs in Photoshop and virtually labeled with the brush tool. Once labeled all 90 nuclei were categorized based on the expression pattern and recorded in an Excel spreadsheet. In this study, control embryos include wild type and the respective heterozygotes. Statistical analysis and visualization of expression data was done using Graphpad Prism for Windows version 9.4.1 or 9.5.0. 91 FIGURES Figure 4.1: Gata6 Null; Oct4 Null Embryos Seem to have Early Lethality. A. Plot of embryos by genotype harvested from 7 litters of Oct4 null het; Gata6 null het x Oct4 null het; Gata6 null het matings at various time points between E2.5 and E3.75 (see results). n=70 embryos total. 92 Figure 4.2: Overexpression of Oct4 is not Sufficient to Promote Ectopic Expression of SOX17 in Mouse Trophectoderm Cells. A. Plot showing the percentage of cells expressing OCT4 in the whole embryo, in embryos with ≤ 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on-Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=8. Oct4 overexpressing, n=3. (see next page) 93 Figure 4.2 (cont’d): B. Plot showing the percentage of cells expressing OCT4 in the ICM, in embryos with ≤ 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on-Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=8. Oct4 overexpressing, n=3. C. Plot showing the percentage of cells expressing OCT4 in the trophectoderm, in embryos with ≤ 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on- Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=8. Oct4 overexpressing, n=3. D. Plot showing the percentage of cells expressing SOX17 in the ICM, in embryos with ≤ 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on-Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=8. Oct4 overexpressing, n=3. E. Plot showing the percentage of cells expressing SOX17 in the trophectoderm, in embryos with ≤ 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on- Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=8. Oct4 overexpressing, n=3. (See next page) 94 Figure 4.2 (cont’d): F. Plot showing the percentage of cells expressing OCT4 in the whole embryo, in embryos with > 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on- Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=19. Oct4 overexpressing, n=9. G. Plot showing the percentage of cells expressing OCT4 in the ICM, in embryos with > 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on-Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=19. Oct4 overexpressing, n=9. H. Plot showing the percentage of cells expressing OCT4 in the trophectoderm, in embryos with > 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on- Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=19. Oct4 overexpressing, n=9. I. Plot showing the percentage of cells expressing SOX17 in the ICM, in embryos with > 50 cells. Control includes wild type, rtTA/rtTA, and rtTA/+ embryos cultured in 40ug/ml dox for 36hrs. Oct4 overexpressing includes rtTA/(rtTA or +);tet-on-Oct4/(tet-on-Oct4 or +) cultured in 40ug/ml dox for 36hrs. Control, n=19. Oct4 overexpressing, n=9. (See next page) 95 Figure 4.2 (cont’d): J. Plot showing the percentage of cells expressing SOX17 in the trophectoderm, in embryos with > 50 cells. 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