Cell Cycle Regulation and Cellular Differentiation in the Developing Ocular Lens

Home , FGF3, FGF4, Lens placode

MIAMI UNIVERSITY The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

Blake Richard Chaffee

Candidate for the Degree:

Doctor of Philosophy

Dr. Michael L. Robinson

Dr. Katia Del Rio-Tsonis

Dr. Paul F. James

Dr. Paul A. Harding

Dr. Eileen K. Bridge, Graduate School Representative ABSTRACT

CELL CYCLE REGULATION AND CELLULAR DIFFERENTIATION IN THE DEVELOPING OCULAR LENS

Blake R. Chaffee

The lens is a simple structure comprised of just two cell types; lens epithelial cells and lens fiber cells. The epithelial cells line, in a single layer, the anterior surface of the lens and remain prolific. The prolific epithelial cells constantly provide the source of new lens fiber cells as the lens epithelial cells differentiate into fiber cells at the lens equator. The fiber cell differentiation process requires an increase in cell cycle inhibitors leading to a permanent withdraw from the cell cycle, the initiation of fiber cell specific genes, and the repression of a subset of epithelial cell specific genes. The most mature fibers in the lens center remove all intracellular organelles in a process that resembles apoptosis. In a single lens section, spatially organized proliferation, cell-cycle withdraw, epithelial cell-to-fiber cell differentiation, and ‘attenuated apoptosis’ occur, making the lens an ideal system to examine the mechanisms controlling gene expression changes during differentiation and the major players involved with cellular proliferation and cellular survival. This dissertation uses the developing ocular lens to elucidate novel mechanisms of 3 proteins proven critical in cell cycle regulation, and cellular survival and differentiation. The third chapter focuses on the essential kinase in mitotic progression, Cyclin dependent kinase 1 (CDK1). Here, we provide further evidence of the requirement of CDK1 in mitotic entry, and more significantly, we reveal a new role for CDK1 in terminal fiber cell differentiation. In the fourth chapter, the Fibroblast Growth Factor Receptor (Fgfr) signaling cascade was explored to further establish the role of the primary downstream arms of Fgfr signaling, PI3K/AKT and MAPK/ERK1/2, in cellular proliferation and differentiation. Lastly, we established the antagonistic relationship of Phosphatase and Tensin Homolog (Pten) and Fgfr signaling on cellular survival in the developing ocular lens. CELL CYCLE REGULATION AND CELLULAR DIFFERENTIATION IN THE DEVELOPING OCULAR LENS

A DISSERTATION

Presented to the Faculty of

Miami University in partial

fulfillment of the requirements

for the degree of

Doctor of Philosophy

Department of Biology in the Program of

Cell, Molecular, and Structural Biology

Blake Richard Chaffee

The Graduate School Miami University Oxford, Ohio

2015

Dissertation Director: Dr. Michael L. Robinson TABLE OF CONTENTS

CHAPTER 1: INTRODUCTION 1.1 Goal/Objective……………………………………………………………………..……...... 1 1.2 Overview of CDK1, FGFRs, and PTEN....……………………………………...…….....…....1 1.2.1 Cyclin-Dependent Kinase 1(CDK1)………....………………………….……....…..1 1.2.2 Fibroblast Growth Factor Receptors (FGFR)………....……..…………….…..….4 1.2.3 Phosphatase and Tensin Homolog (PTEN)………...... ……………………...... ….5 1.2.4 Biological/Clinical Significance…………………...……………………….....…..6 1.3 Overview of Lens Development………………………………...…………………...….….…7 1.3.1 Historical Perspective of Lens Development……………………...... …....………7 1.3.2 From Fertilized Zygote to Ocular Lens………………………….....…….…….…8 1.4 Lens Induction Requires Paracrine signaling from BMPs and FGFs………….…...……10 1.5 Ocular Environment Dictates Lens Epithelial and Fiber Cell Identity: formation of the FGF gradient Hypothesis……...... …………………………………………………………....…12 1.6 Lens Epithelial Cell Proliferation and the Cell Cycle……………………………..….….13 1.6.1 Cell Cycle Progression by the CDKs and Cyclins…...... ………………….....14 1.6.2 Cell Cycle Machinery in the Lens…………………...... ……………………16 1.6.3 Receptor Tyrosine Kinases on Cellular Proliferation in the Lens...... …………17 1.7 Balancing Lens Cell Survival and Death………………………………....……….….….21 1.7.1 FGFRs are Required to Tip the Balance Towards Survival……....……………..21 1.8 Lens Epithelial Cell-to-Fiber Cell Differentiation…………………….……………...... 22 1.8.1 Gene Expression Changes During Differentiation: Growth Factor Control of Gene Regulatory networks…………………………………...... …….23 1.8.2 Mechanism of Nuclear Removal………....……...... ….…………….…..26 1.9 Specific Aims……………………….…………………………...... ……………………..…..28 1.9.1 Nuclear Removal During Terminal Lens Fiber Cell Differentiation Requires CDK1 Activity; Appropriating Mitosis-Related Nuclear Disassembly...... ……...... ….28 1.9.2 FGFR and PTEN signaling interact during lens development to regulate cell Survival...... …………….………………………...... ……..…..

ii 1.10 References ……………………….…………………………………………....……...…….35

CHAPTER 2: MATERIALS AND METHODS 2.1 Mice………………………………………………………………….…………..……..……47 2.1.1 Cdk1 Deletion………………....………………………………………...... …...47 2.1.2 Pten and Fgfr2 Deletions…………....…………………………………...... …..48 2.2 Histology and Immunohistochemistry...... 48 2.3.1Frozen Sections………………………………………………………...... ……...... 48 2.3.2 Paraffin Sections…………….…………………...... ……...…...49 2.3.2a Tissue Processing ………...... …...…49 2.3.2b Immunohistochemistry Protoco...... ……...... ……...….50 2.3.2c Antibodies Used for Immunohistochemistry...... 51 2.4 Immunofluorescence Quantification...... 51 2.5 Whole Mount Epithelial Cell Z-Stacks………………………………..………...…....…52 2.6 Western Blot……………………………………………………………………………....…53 2.6.1 CDK1 Manuscript (Western Blots Completed at Tufts University)………...... …53 2.6.2 PTEN/FGFR2 Manuscript (Western Blots Completed at Miami University)...... 53 2.6.2a Reagents…………….………………………………………………...…..53 2.6.2b Western Blot Protocol………………………………………………....….56 2.7 RT-qPCR……………………………………………………………………………....…….58 2.7.1 Primer List…………………………………………………………………...... …....……59

CHAPTER 3: NUCLEAR REMOVAL DURING TERMINAL LENS FIBER CELL DIFFERENTIATION REQUIRES CDK1 ACTIVITY: APPROPRIATING MITOSIS- RELATED NUCLEAR DISSASSEMBLY 3.1 Summary…………………………………………………………………………………….60 3.2 Introduction……………………………………………………………………………….…61 3.3 Results……………………………………………………………………………………….63 3.3.1 CDK protein expression in epithelial cells and differentiation lens fibers……...63 3.3.2 Removal of CDK1 from the Lens…………………………....………………….63

iii 3.3.3 Loss of CDK1 delays Denucleation of Lens Fiber Cells…………………...... …64 3.3.4 Loss of CDK1 Prevents Entry of DLAD into the Nucleus of Terminally Differentiating Lens Fiber Cells………………………...... ……...... …………65 3.3.5 Relocalization of NuMA during Fiber Cell Differentiation and Maturation…….66 3.3.6 Reduction of Phosphorylated Histone H1 (pH1) in CDK1 Deficient Lenses...... 67 3.3.7 Specificity of Nuclear Retention in CDK1 Deficient Lenses….……………...... 68 3.3.8 Epithelial Cells in CDK1 Deficient Lenses Fail to Undergo Mitosis, and Exhibit DNA endoreduplication………………………………...... …………………68 3.4 Discussion………………………………………………….………………………………...69 3.5 References………………………………………………..…………………………………..95

CHAPTER 4: FGFR AND PTEN SIGNALING INTERACT DURING LENS DEVELOPMENT TO REGULATE CELL SURVIVAL 4.1 Chapter Summary………………………………………………………………….…...…..100 4.2 Introduction…………………………………………………………………………..……..100 4.3 Results…………………………………………………………………………………....…103 4.3.1 Balancing PTEN and FGFR2 in Lens Cell Survival and Proliferation………...... 103 4.3.1a Loss of Pten Restores Lens Size in FGFR-deficient Lenses………….....101 4.3.1b Pten deletion Restores Cell Survival in FGFR-deficient Lenses……..…104 4.3.1c The Effect of Pten Deletion on Lens Cell Proliferation………..………..105 4.3.2 The Impact of FGFR2 and PTEN on Lens Epithelial Cell-to-Fiber Cell Differentiation……………………………………………………………...... ……106 4.3.2a Reduced γ-crystallin expression in Fgfr2Δ/Δ Lenses is partially restored in The Absence of PTEN……………………………………...... ………...…106 4.3.2b Pten Deletion Fails to Rescue Reduced Aquaporin0 Levels in FGFR2- Deficient Lenses……...... ……………………...…………………………..107 4.3.2c Fgfr2Δ/Δ lens fiber cells experience nuclear retention reduced DnaseIIβ expression, neither of which is rescued by simultaneous deletion of Pten…………………...... ………………………………………………107

iv 4.3.2d FGFR2-deficient lenses Exhibit PAX6 retention in posterior lens cells Depleting the lens of PTEN restores the removal of PAX6 from FGFR2 deficient posterior lens cells………………...... ………………………….108 4.3.3 The deletion of Pten normalizes several downstream signaling pathways affected by Fgfr2 loss in the lens…………...... ………………………………….109 4.3.3a Pten Deletion Restores AKT and ERK1/2 Activation in FGFR2-Deficient Lenses………………………………………………...... 109 4.3.3b The Effect of PTEN and FGFR2 on c-Jun N-terminal Protein Kinase (JNK) …………...... …………………………………………..110 4.4 Discussion…………....……………………………………………………………………..110 4.5 References…………....……………………………………………………………………..136

CHAPTER 5: SUMMARY OF CONCLUSIONS AND FUTURE DIRECTIONS

5.1 CDK1 is required for lens fiber cell denucleation…………...... ………………...... 143 5.2 S-Phase entry occurs in the absence of CDK1, yet CDK1 is indispensable for mitosis...... 144 5.3 CDK1 is required for apoptosis?…..……………………...... ………………...145 5.4 Active CDK1 in the lens fiber cell mass opens a new door for future investigations of the role of CDK1 outside of the cell cycle…………………..……...... 145

5.5 FGFR2 is required for lens cell survival and lens differentiation...... ….147 5.6 Pten deletion restores lens cell survival in FGFR2-deficient lenses…………….....….....…148 5.7 Lens development does not require PTEN...... ………………………………………...…...149 5.8 Does lens fiber cell differentiation require Fgfr signaling outside of AKT and ERK?...... 150 5.9 References…………………………………………………………………………....……..152

vi LIST OF TABLES

Table 1.1 CDK and Cycle Expression in the Lens Epithelium and Lens Fiber Cells……...……34 Table 2.1 RT-qPCR Primer List……………………………………………………………..…..59

vi LIST OF FIGURES

Fig. 1.1 PI3K/AKT and MAPK/ERK1/2 are the two main arms of Fgfr signaling…………..…31 Fig. 1.2 Overview of lens development………...……………………...………………….….….32 Fig. 1.3 Spatial Organization of the E18.5 Lens………………………………………….….…..33 Fig. 3.1 CDK Protein Expression in Normal Lens Epithelial Cells and Fiber Cells…….……....73 Fig. 3.2 MLR39 Cre does not Deplete the Lens of CDK1………………………………………75 Fig. 3.3 Little CDK1 Expression Remains in MLR10; Cdk1L/L Lens Cells by E17.5 ...... 77 Fig. 3.4 The Formation of an Organelle Free Zone Requires CDK1……………………………79 Fig. 3.5 CDK Deficiency Decreased the Phosphorylation of Lamin A/C, blocked the entry of DLAD into the Nucleus and Decreased DNA Degradation in Maturing Lens Fiber Cells...... ….81 Fig. 3.6 Lenses Deficient in CDK1 Failed to Phosphorylate NuMA……………………………84 Fig. 3.7 Nuclear Architecture is Disrupted with CDK1 Depletion………………………...….…86 Fig. 3.8 CDK1 Deficiency Decreased the Phosphorylation of Histone H1 in Lens Fiber Cells………………………………………………………………..…...... …..…..87 Fig. 3.9 MLR10; Cdk1L/L Lenses Remove both Mitochondria and Endoplasmic Reticulum Despite Retaining Nuclei……………………………………………………….…...... …….88 Fig. 3.10 CDK1-Deficient Lenses Exhibited Large, Sparse Epithelial Cell Nuclei…….….……90 Fig. 3.11 MLR10; Cdk1L/L Lens Epithelial Cells Continue to Synthesize DNA but Fail to Enter Mitosis…………………………………………………………………….…...... ……92 Fig. 3.12 Cdk1 Deletion does not Result in Epithelial Cell Apoptosis……………………..……94 Fig. 4.1 Le-Cre Efficiently Deletes LoxP-Flanked Pten and Fgfr2………………………….…115 Fig. 4.2 Pten deletion rescues the lens size and elongation defects in Fgfr2Δ/Δ lenses…….....116 Fig. Fig. 4.3 Pten Deletion Restores Lens Cell Survival in FGFR2-Deficient Lenses……..…..118 Fig. 4.4 PtenΔ/Δ,Fgfr2Δ/Δ, and Pten/R2Δ/Δ Lenses Display Early Cell Cycle Withdrawal Defects. None of the knockouts exhibited lens epithelial cell proliferation Alterations later in development………………………………………………...... …….….120 Fig. 4.5 Pten Deletion Restores E12.5 γ-crystallin Expression. Later in Development Fgfr2Δ/Δ, and Pten/R2Δ/Δ lenses display normal transcript levels of γ-crystallin and c-maf…………………………………………………………………………...... …..…...122

vii Fig. 4.6 Fgfr2Δ/Δ lenses exhibit reduced Aquaporin0 and a nuclear retention phenotype that is not rescued by Pten deletion…………………………………………………...... ….....124 Fig. 4.7 E12.5 Fgfr2Δ/Δ lenses maintain PAX6 expression in the posterior fiber cells and Pten deletion restores the normal removal of PAX6……………………………...... …...126 Fig. 4.8 Deleting Pten restore pAKT and pERK1/2 in FGFR2-deficient lenses…………….....128 Fig. 4.9 FGFR2 deficiency leads to increased activation of C-JUN and p53………………...... 129 Fig. 4.10 The expression of many fiber cell differentiation markers were not altered in PtenΔ/Δ , Fgfr2Δ/Δ, or Pten/R2Δ/Δ lenses at E18.5……………………………………...... …..130 Fig. 4.11 PTEN deletion did not inhibit apoptosis at E10.5……………………………………132 Fig. 4.12 Le-Cre Hemizygosity did not alter fiber cell differentiation, but did increase apoptosis………………………………………………………………………...... ……..133 4.13 Le-Cre Hemizygosity did not alter downstream PI3K/AKT or MAPK/ERK1/2…………135

viii ACKNOWLEDGEMENTS

I would like to first thank Dr. Michael Robinson for giving me a chance. In his lab, I transformed from a college athlete with mediocre grades and average academic ambitions to an inspired scientist excited to impact medical genomics as a postdoctoral fellow at the Cleveland Clinic. I would also like to thank our research associate Brad Wanger; my fellow graduate students, Bhavani Madakashira, Thanh Hoang, Evan Horowitz, and Phuong Lam; and the undergraduates, Devin Bruney, Melissa Leonard, Rich Dowd, Sara Perkins and Ben Schwartz. Brad Wagner kept me sane in my years as a graduate student. Brad also helped me/saved me when experiments were going wrong, or even when I needed a reagent or help while I was instructing lab courses. Thanh provided an enormous amount of expertise in the realm of molecular/biochemical techniques. Evan was always a great person to talk with during stressful days, and saved me by covering my lab course while I was interviewing for my postdoc positions. Devin provided help on both the FGFR2/PTEN project and the CDK1 project. Melissa Leonard, Sara Perkins, and Rich Dowd provided me with help on the FGFR2/PTEN project. I would also like to thank my committee members, Dr. Paul James, Dr. Paul Harding, Dr. Eileen Bridge, and Dr. Katia Del Rio-Tsonis. They provided excellent advice, expertise, and support throughout my years as a graduate student. Lastly, I want to thank my friends and family. First, I want to thank my fiancé, Ashley Dean. She supported me in all of my decisions, even the ones that directly affected her. She listened to hours upon hours of “science” talk and science venting. She kept me going when I questioned my career path or myself. I also have the two most supportive parents, Roger and Thelma. They have been my role models, my support system, and the people directing me my entire life.

viii CHAPTER 1: INTRODUCTION

1.1- GOAL/OBJECTIVE

The goal of this dissertation is to advance the understanding of Cyclin Dependent Kinase 1(Cdk1), Phosphatase and Tensin Homolog (Pten), and Fibroblast growth factor receptors (Fgfrs) in cell cycle control, cellular survival, and terminal differentiation. All three of these genes are crucial in the development and homeostatic maintenance of nearly every mammalian tissue. Due to their essential role in cell cycle regulation, cellular growth and survival, these genes have been of particular interest to not only developmental biologists, but also clinical scientists attempting to provide therapeutic treatment to cancer. Inhibiting FGFRs or CDK1 represent potential therapies to halt cancer survival and proliferation, which received attention from clinical investigators in previous reports. Pten mutations often lead to unrestricted cell proliferation and this property makes Pten one of the most frequently mutated genes in cancer. To further elucidate the molecular roles of these 3 genes, we will use the developing murine ocular lens as the model system. This chapter provides an overview of the established biological roles of Cdk1, Fgfrs, and Pten. Secondly, I hope to provide an appreciation for using the ocular lens as a tool to study developmental and molecular biology. To do this, I will highlight the historical importance of the lens and how using the lens has the potential to reveal new molecular mechanisms in a broader biological context-with particular emphasis on CDK1, FGFRs, and PTEN. Lastly, this chapter will clarify the hypotheses addressed in this dissertation and the specific aims carried out to support these hypotheses.

1.2 OVERVIEW OF CDK1, FGFRs, and PTEN

1.2.1 Cyclin-Dependent Kinase 1

1 Cyclin-dependent kinase 1 (CDK1), formerly called Cdc2 or p34cdc2 is a serine-threonine kinase with roughly 200 known substrates. Many of these substrates trigger progression through mitosis (Ubersax et al., 2003). As the level of CDK1 protein expression levels remain fairly constant throughout the cell cycle, the differential activation of CDK1 is regulated by the abundance of a co-activor cyclin, which varies with different stages of the cell cycle. CDK1 activity is low during the initiation of DNA replication (Synthesis phase or S-phase). However, CDK1 activity rises through gap 2 (G2) and is responsible for pushing cells through the G2/M transition. During mitosis, CDK1 with its mitotic coactivator cyclin B, trigger many of the hallmark mitotic events. Of note, CDK1/cyclin B1 promotes spindle assembly by phosphorylating the nuclear mitotic apparatus (NuMA), chromatin condensation by phosphorylating the linker histone H1, and nuclear envelope breakdown (NEBD) by phosphorylating the nuclear lamin proteins (Kotak et al., 2013; reviewed in Nigg, 1993). Following mitosis, CDK1 activity falls, which is important for ending mitosis Despite primarily driving mitotic progression, CDK1 can supplement any other CDK during G1-S-G2-M, and cyclins A, B, E, and D can activate CDK1. In fact, CDK1 can supplement the loss of any other CDKs in the cell cycle, as mice with deletions of Cdk2, Cdk3, Cdk4, and Cdk6 are viable, yet deletion of Cdk1 is embryonic lethal at the blastocyst stage (Kaldis and Aleem, 2005; Malumbres and Barbacid, 2005; Satyanarayana and Kaldis, 2009). The role of CDK1 in non-cycling cells is not clear; yet, CDK1 may be important in apoptotic and differentiation events (Castedo et al., 2002; He et al., 1998). In both apoptosis and mitosis, cell rounding and nuclear envelop disassembly occur, although the process by which these events occur in each process may be fundamentally different. In mitosis, the promotion of NEBD by lamin phosphorylation requires CDK1/CyclinB1, whereas in apoptosis, caspase-6 cleaves lamin (Oberhammer et al., 1994; Ruchaud et al., 2002). There is evidence that, in certain cellular contexts, apoptosis requires CDK1. For example, taxol (a microtublule inhibitor) can enhance cell death in cancer cells, but requires CDK1 activity to do so (Shen et al., 1998). Most likely, CDK1/cyclin B promotes apoptosis by phosphorylating the pro-apoptotic BAD protein, causing the translocation of BAD to mitochondria. Here, BAD antagonizes Bcl-2 like proteins (pro-survival factors) and activates BAX-like proteins leading to mitochondrial membrane permeabilization and cell death (Konishi et al., 2002). On the other hand, CDK1 demonstrated the ability to promote cell survival in other cellular contexts. In several tumor cell lines,

2 inhibiting CDK1 enhanced apoptosis. Treatment of prostate carcinoma, head and neck cancer, non-Hodgkin’s lymphoma, and leukemia with the CDK1 inhibitor, Flavopiridol, enhanced dell death (Kelland, 2000; Zhai et al., 2002). The inhibition of CDK1 likely promotes survival by phosphorylating Survivin, resulting in the dissociation of the Survivin-Caspase 9 complex, resulting in apoptosis (O'Connor et al., 2002). It remains unclear whether CDK1 or the targets of CDK1 participate in cellular differentiation, yet several studies implicate targets of CDK1 as regulators of differentiation. Several studies suggest that the nuclear lamin proteins, which provide the shape and mechanical stability of the nucleus, play a role in differentiation. At the onset of mitosis, lamins are depolymerized by CDK1 phosphorylation. In mice, lamin B is expressed throughout embryogenesis, whereas lamin A and C do not initiate expression until E9, and lamins A and C predominate in the nuclear lamina of differentiated cells (reviewed in Dechat, 2008). Moreover, cells do not express lamin A/C until they lose their pluripotency, thus suggesting a role of lamin A/C in cellular differentiation, although the mechanism of this role remains unclear (reviewed Dechat, 2008). The role of lamin A/C in differentiation may be through epigenetic mechanisms as both lamin A/C and its associated binding proteins are capable of binding DNA (reviewed in Dechat, 2008). To further support the impact of lamin A/C on epigenetic shifts, patients with Huchinson-Gilford Progeria syndrome (HGPS), a well established laminopathy resulting from mutations in the gene encoding lamin A (LMNA), exhibit reduced trimethylation of both Histone H3 lysine 9 (H3K9me3) and Histone H3 lysine 27 (H2K27me3) as well as increased Histone H4 lysine 20 trimethylation (H4K20me3) (Columbaro et al., 2005; Scaffidi and Misteli, 2005; Shumaker et al., 2006). Like Lamin A/C, the CDK1 substrate NuMA may play a role in nuclear structural changes required for cellular differentiation. Several reports revealed a presence of NuMA localized to the nucleus of cells withdrawn from the cell cycle. Changes in NuMA distrubution during epithelial cell differentiation first suggested a role for NuMA in chromatin reorganization (Abad et al., 2007; Lelievre et al., 1998). One such example of the NuMA distribution shift are in human mammary epithelial cells (HMECs). HMECs that had not withdrawn from the cell cycle exhibited a diffuse distribution of NuMA within the nucleus and upon further morphogenesis; NuMA distribution became concentrated into large foci (Lelievre et al., 1998). Further supporting the impact of NuMA on differentiation, using an antibody to target the C terminus of

3 NuMA on mammary acini cells grown in 3D culture generated a shift in expression from the NuMA aggregates to diffuse NuMA expression across the nuclei (Lelievre et al 1998). Accompanying the shift in NuMA expression patterns was a redistribution of acetyl-Histone 4 (acetyl H4) and H4K20 methylation, and an inhibition of acinar differentiation (Abad et al., 2007; Lelievre et al., 1998). One question remaining is whether or not the phosphorylation of NuMA alters the expression patterns seen in differentiation. In thymocytes undergoing apoptosis, NuMA distrubtion changes from a being diffuse across the nuclei to being reduced to a single small intense patch of NuMA expression (Weaver et al., 1996). Accompanying the shift in NuMA expression during apoptosis, NuMA becomes phosphorylated and cleaved prior to the DNA degradation during apoptosis (Weaver et al., 1996). The role of CDK1 outside of the cell cycle remains elusive. It is possible that CDK1 participates in both apoptosis and cellular differentiation, yet the precise mechanism by which CDK1 may induce apoptosis or differentiation is incomplete. Furthermore, the kinase activity of CDK1 may alter the NuMA and Lamin A/C during apoptosis and differentiation.

1.2.2 Fibroblast Growth Factor Receptors

Fibroblast growth factor receptors (FGFR) are receptor tyrosine kinases (RTKs) encoded by four different genes (Fgfr1-5) in mammals. Full-length FGFRs are comprised of 3 extracellular immunoglobulin (Ig)-like domains, a transmembrane domain, and an intracellular tyrosine kinase domain. In mammals, most of the receptors are activated by one of the 22 known FGF ligands with specificity of the FGFRs dictated mainly by alternative splicing which generates numerous receptor isoforms (Johnson and Williams, 1993; Lee et al., 1989; Ornitz et al., 1996; Zhang et al., 2006). Heparin sulfate proteoglycans (HSPG) facilitate the binding of FGF ligands to their complementary receptors (Mohammadi et al., 2005). Common to many RTKs, activation of FGFRs result in multiple signal transduction cascades including the RAS- RAF-MEK-ERK1/2 (also referred to as MAPK/ERK1/2), PI3K-AKT, PLCγ, SRC, and STAT pathways. Although RTKS and FGFRs activate many downstream cascades, numerous studies suggest the primary importance of PI3K/AKT and MAPK/ERK1/2 in growth factor mediated responses. Activation of ERK1/2 and AKT result in a diverse set of cellular responses, such as, enhanced cellular survival, proliferation, and differentiation. The diversity of response depends

4 on tissue type, ligand/receptor combination, the amount of activation, and whether the activation is transient or sustained (Mansour et al., 1994; Nguyen et al., 1993; Shaul and Seger, 2007; Traverse et al., 1994). Nearly every mammalian tissue requires FGFR signaling during development and homeostatic maintenance during adulthood. Over activation of FGFR signaling plays a pathogenic role in many human cancers, including but not limited to, cancers of the breast, endometrium, liver, lung, prostate, and urinary bladder (Ahmad et al., 2012; Wesche et al., 2011). The specific adaptor proteins used in FGFR signal transduction distinguishes FGFRs from other RTKs. Of particular note, specific to just FGF, TRK, RET, and ALK receptors, FRS2α binds and promotes the activation of PI3K and a sustained activation of ERK1/2 (Hadari et al., 2001). Upon FGFR activation, six tyrosine residues of FRS2α are phosphorylated. Although FRS2α does not have catalytic activity, the phosphorylated tyrosine resides of FRS2α attract SH2-containing proteins, including GRB2 and SHP2. Binding of GRB2 and SHP2 results in downstream activation of PI3K/AKT and the RAS-RAF-MEK-ERK1/2 pathways (Fig 1.1). GRB2/SOS complexes attract RAS. Following RAS activation, RAF is recruited to the plasma membrane, where RAF is phosphorylated, and RAF subsequently phosphorylates MEK (Fig. 1.1). MEK is a dual specific kinase that phosphorylates both the tyrosine and serine/threonine residues of ERK1/2, which results in ERK1/2 activation (Fig. 1.1). Once activated, ERK1/2 transports to the nucleus to activate over 100 distinct cellular targets that mediate gene expression. In regard to the PI3K/AKT pathway, FRS2α and GAB1 activation result in PI3K activation. PI3K phosphorylates PtdIns(4,5)P2 (PIP2), and thereby converting it to the PtIns(3,4,5)P3 (PIP3) (Fig. 1.1). PIP3 recruits AKT and its activator, PDK, to the plasma membrane. Akt is activated by a series of phosphorylations by PDK and full activation requires additional phosphorylation by mTORC2. Activated AKT is known to have over 100 targets, which primarily lead to anti-apoptotic pathways, and cellular proliferation (Carracedo and Pandolfi, 2008).

1.2.3 Phosphatase and Tensin Homolog

Phosphatase and tensin homolog (PTEN) is a dual specific phosphatase capable of dephosphorylating both lipids and proteins. PTEN is a known tumor suppressor, inhibiting

5 tumorigenicity and cell growth by promoting G1 cell cycle arrest, apoptosis, or both (Furnari et al., 1998; Li and Sun, 1998). PTEN mainly resides in the cytoplasm, where it carries out its principle catalytic activity. Cytoplasmic PTEN dephosphorylates PIP3-converting it back to its inactive form PIP2, directly counteracting PI3K, and therefore, inhibiting AKT activation (Fig. 1.1) (Chung and Eng, 2005). The predominate result of AKT inhibition is apoptosis (Chung and Eng, 2005). Although the inhibition of AKT activation represents the most well established role of PTEN, PTEN reduces ERK activation (Chung and Eng, 2005; Weng et al., 2001). PTEN is capable of inhibiting ERK activation both in the cytoplasmic and in the nucleus (Chung and Eng, 2005; Weng et al., 2001). In the cytoplasm, PTEN antagonizes IRS-1 phosphorylation, disrupting the IRS-1/Grb2/SOS complex. Like FGFRs and other RTKs, IRS-1 activation stimulates the activation of ERK1/2, and therefore, the inhibition of IRS-1 phosphorylation by PTEN reduced ERK activation (Weng et al., 2001). Additionally, PTEN resides in the nucleus during the G0-G1 phases of the cell cycle and reduces nuclear activated ERK and cyclin D1 expression resulting in cell cycle arrest (Chung and Eng, 2005). Cytoplasmic PTEN often participates in cell cycle arrest as AKT inhibits the CDK inhibitors, p21CIP1 and p27KIP1; therefore, the antagonism of AKT activation by PTEN can result in elevated CDK inhibition and cell cycle arrest (Chung and Eng, 2005). PTEN represents a potent antagonist of the two primary downstream FGFR induced pathways. Recent studies established the antagonistic relationship of FGFR2 and PTEN in osteoprogenitor proliferation (Guntur et al., 2011), adipocyte differentiation (Scioli et al., 2014) and skin tumorigenesis (Hertzler-Schaefer et al., 2014). In both the Gunter 2011 study on osteoprogenitor proliferation and the Hertzler-Schaefer et al. 2014 study on skin tumorigenesis, deletion of just Fgfr2 rescued the phenotype resulting from PTEN deficiency (Guntur et al., 2011; Hertzler-Schaefer et al., 2014).

1.2.4 Biological/Clinical Significance Cdk1, Fgfrs, and Pten are critical in nearly every mammalian tissue from the early stages of embryogenesis throughout adulthood. Despite their importance, many unanswered questions remain regarding all three of these genes. The role of CDK1 in post-mitotic cells remains elusive. The nuclear lamins represent one of the primary targets of CDK1, and defects in the

6 nuclear lamins represent the driver of several diseases such as, Emery-Dreifuss muscular dystrophy (EDMD), dilated cardiomyopathy (DCM), limb-girdle muscular dystrophy and Hutchinson-Gilford progeria syndrome. Most of these “laminopathies” affect non-proliferating cells (Ho and Lammerding, 2012). Also, the mechanism by which Fgfrs elicit a diversity of cellular responses in different cell types remains incomplete. Similarly, Pten mutations result in a variety of different cellular outcomes. Despite being widely regarded as a tumor suppressor, Pten mutations play a pathological role in a wide array of other diseases, including Cowden Syndrome, Bannayan-Reily-Ruvalcaba Syndrome, Proteus Syndrome, Proteus-like syndrome, and Autism Spectrum Disorder, and yet, not all Pten mutations lead to tumorigenesis (Orloff and Eng, 2008). The murine ocular lens provides for an ideal system to further elucidate the roles of Cdk1, Fgfrs, and Pten. One key feature of using the mouse lens is the existing transgenic tools that make it possible to delete genes at different stages of mouse lens development. Lens epithelial cell explants and culture systems complement these in vivo approaches. Furthermore, as the organization of lens contains compartmentalized regions of cellular proliferation, and terminal differentiation, it provides for a unique system to study the mechanisms involving cellular proliferation, cell cycle withdrawal, and cellular differentiation. As the bulk of the lens is comprised of cells permanently withdrawn from the cell cycle, and these post-mitotic lens cells possess both CDK1 and cyclin B1 protein, the lens represents a potential system to discover and define a post-mitotic role for CDK1. FGFR signaling plays a predominant role in nearly every process of lens development, and in vitro lens systems revealed diverse sets of responses sensitive to differing concentrations of FGF2. Taken together, the lens is a premier system for investigating Fgfr-induced responses in vivo.

1.3 OVERVIEW OF LENS DEVELOPMENT

1.3.1 Historical Perspective of Lens Development

At the turn of the 20th century, tissue induction represented a developmental event of particular interest to biologists. The developing eye remained at the forefront of experiments on tissue induction. The optic vesicle (OV) inducing lens formation quickly became a classic model

7 of tissue induction. In more recent times, the realization of the therapeutic potential of embryonic stem cells (ES cells) and induced pluripotent stem cells (IPS cells) in tissue repair generated an elevated interest in the molecular mechanisms responsible for driving pluripotent cells to become populations of highly specialized cells. Like the study of tissue induction, the study of how paracrine signaling directs cell fate uses the lens as an important model. As recently as 2010, one of the first examples of directing human embryonic stem cells to a highly specialized cells occurred in the lab of Dr. Cvekl, where he directed the differentiation of human embryonic stem cells into lens progenitor cells and lentoid bodies (Yang et al., 2010). Over the century following the initial investigations of lens induction, developmental biologists formed a complete picture outlining the journey of lens formation from the fertilized zygote to the inner cell mass to ectoderm to neural plate border to the preplacodal ectoderm to the lens vesicle to the lens epithelium and fiber cells. To properly place the development of the ocular lens into a broader context of the field of developmental biology and to expand the appreciation of using the ocular lens to study molecular mechanism, this next section is dedicated to outlining lens development from the zygote all the way to the lens epithelial cell and lens fiber cell formation.

1.3.2 From Fertilized Zygote to the Ocular Lens

The fertilized zygote is capable of giving rise to over 200 different cell types. At first, the fertilized zygote undergoes several rounds of cleavage without increasing the overall size of the embryo-defined as the morula stage. Here at this stage, all the cells are still capable of forming all cell types, including extra embryonic tissue; therefore, considered totipotent. Following the morula stage, compaction occurs where the embryo continues to divide and simultaneously forms into a spherical shape. During compaction some cells reside inside of the cell mass- which give rise to the inner cell mass, and other cells on the cell surface-giving rise to the trophoblast. This event marks the first definitive differentiation event in the embryo. Following the compaction of the morula, a blastocyst forms as the mural trophoectoderm secretes an enzyme dissolving the zona pellucida. The blastocyst contains an inner cell mass (ICM)-giving rise to all the different cell types within the body, and the trophoblast- which forms the extra-embryonic tissue. Prior to the time of implantation of the embryo into the uterus, the ICM is divided into either primitive endoderm or primitive epiblast. The primitive epiblast remains pluripotent, and it

8 is this population of cells that give rise to the embryo proper, or ectoderm, mesoderm and endoderm. The lens derives from the region of ectoderm between the neural and presumptive epidermal ectoderm, also known as the neural plate border (NPB). At the neural plate border, both the cells that give rise to the neural crest (NC) and the preplacodal ectoderm (PPE) reside (reviewed in Cvekl and Ashery-Padan, 2014; reviewed in Robinson, 2014). The cells of the NC contain progenitors that will eventually form peripheral neurons, glia, smooth muscle cells, melanocytes, cells of the adrenal medulla, cardiac cells, and the cartilage and bones of the face (reviewed in Robinson 2014). The anterior PPE forms the lens, olfactory and anterior pituitary placodes, whereas the posterior PPE forms the trigeminal, epibranchial, and otic placodes (reviewed in Cvekl 2014; Robinson, 2014). An outgrowth of the neural ectoderm brings the OV in close proximity to the population of surface ectoderm cells that will give rise to the lens, and this region is referred to as the presumptive lens ectoderm (PLE) (Fig. 1.2A-embryonic day 9 (E9) in mice). A fibrillar network starts to form between the OV and the PLE- promoting strong adhesion between the two (Fig. 1.2B- E9.5 in mice). The strong adhesion between the OV and PLE results in a restricted lateral movement of the cells in the PLE. Cells within this restricted region continue to proliferate, thus, causing the area to thicken, forming the lens placode (Fig. 1.2B- E9.5 in mice). Inductive interactions of the OV signal the PLE to invaginate forming a lens pit (Fig. 1.2C- E10.5 in mice). As the lens pit deepens, the association between the two tissues breaks, most likely through apoptosis, forming a hollow sphere of proliferating cells called the lens vesicle (Fig. 1.2D- E11 in mice). Primary fiber cells begin to form at the posterior hemisphere of the lens vesicle. These primary fiber cells withdraw from the cell cycle, initiate fiber cell specific gene production and elongate until they touch the cuboidal epithelial cells at anterior hemisphere of the lens (Fig. 1.2E to F- E12 to E13.5 in mice). The primary fiber cells remain in the lens center throughout the life of an animal. Two different cell types comprise a fully formed lens, lens epithelial cells and lens fiber cells (Fig. 1.3A-black arrows indicating lens epithelium). Lens epithelial cells line, in a single layer, the anterior hemisphere of the lens (Fig. 1.3A). The lens epithelial cells remain capable of proliferation throughout the mammalian lifespan, but normal proliferation eventually localizes to a zone termed the germinative zone, slightly above the lens equator (Fig. 1.3B-white boxes). Approaching the equator of the lens, the epithelial cells begin to differentiate into

9 secondary fiber cells (Fig. 1.3C). The upregulation of cell cycle inhibitors, such as p27KIP1 (Fig. 1.3C), p57KIP2, and pRb promote cell cycle withdrawal, which marks the initiation of fiber cell differentiation. The region of the lens receiving heightened expression of the cell cycle inhibitors defines the transition zone (Fig. 1.3C-white brackets). Differentiating fiber cells then migrate toward the lens equator, elongate, express fiber cell specific genes, such as β (Fig. 1.3D) and γ−crystallin (Fig. 1.3E) and finally, lose all of their organelles (Fig. 1.3F). The organelles, including but not limited to the nuclei, mitochondria, and endoplasmic reticulum, must be removed due to their ability to scatter light. In mice, the degradation of these organelles initiates between E16 and E18, whereby the first fiber cells to denucleate are the primary fiber cells (Kuwabara and Imaizumi, 1974). Retention of any of these light scattering components generates an opaque lens or cataract. The TUNEL assay, a common method to detect apoptotic cells, permits the labeling of nuclear removal in the central lens fiber cell mass (Fig. 1.3F). The overall process of organelle loss is often referred to as attenuated apoptosis (reviewed in Bassnett, 2009; Wride, 2011).

1.4 LENS INDUCTION REQUIRES PARACRINE SIGNALING FROM BMPs AND FGFs

FGF signaling plays a role in directing cell fate decisions as early as the blastocyst stage where FGF signaling in the inner cell mass pushes cells toward a primitive endoderm fate (Yamanaka et al., 2010). The induction of the PPE requires FGF signaling, and a blockage of both WNT and Bone Morphogenic Protein (BMP) signaling (reviewed in Robinson, 2014). Following the initial induction of the PPE, the PPE requires BMP signaling and the expression of the transcription factors, AP2α, FOXI, GATA, and DLX, which together initiate Six1 and Eya1/2 expression (reviewed in Robinson 2014). Six1 and Eya1/2 expression define the PPE (reviewed in Robinson 2014). After PPE specification, individual placodes require different signals to specify their final fate. Lens placode formation requires inductive signals from the OV. The surface ectoderm that does not directly contact the OV receives signals from the periocular mesenchyme (POM) that inhibit lens placode formation (reviewed in Robinson 2014). WNT/β- catenin activity represents the inhibitory signals from the POM and WNT signaling negatively impacts, Pax6 expression. PAX6 is often referred to as the master regulator of eye formation, and

10 upregulation of PAX6 is required during lens induction (reviewed in Cvekl and Ashery-Padan, 2014; reviewed in McAvoy et al., 1999). In addition to a lens inducer by paracrine signaling, the OV prevents WNT signaling from reaching the PLE, therefore, providing a protective barrier to inhibitors of lens formation. Evidence suggests that BMP signaling represents one of the inducers of lens placode formation provided by the OV. BMPs are members of the TGF superfamily, and activate a heteromeric complex of type I and type II receptors. Upon receptor activation, the receptors phosphorylate cytoplasmic Smad (Smad 1/5/8) proteins, which then associate with co-Smad, Smad4. Smad4 localizes to the nucleus to regulate transcription (Heldin et al., 1997). The optic vesicle secretes BMP4, and mice deficient in BMP4 fail to form a lens placode. Interestingly, exogenous BMP4 addition did not restore lens placode formation in the absence of the OV, therefore, the OV must be providing an additional role in lens placode formation (Furuta and Hogan, 1998). In part, the additional role may be to provide a protective barrier to prevent WNT signaling from reaching the PLE. Moreover, BMP7 may play an imperative and non-redundant role with BMP4, as mice deficient in BMP7 exhibit anophthalmia, and a loss of Pax6 expression in the lens placode (Luo et al., 1995; Wawersik et al., 1999). Experiments knocking out the BMP receptors further supports the critical impact of BMP4 and 7 on lens induction. Conditionally knocking out the type I BMP Bmpr1a in the lens placode, prior to placode thickening, resulted in increased apoptosis and deletion of the other type I BMP receptor, ACVR1 at the same stage, resulted in decreased in cellular proliferation (Rajagopal et al., 2009). Combined deletion of both receptors lead to reduced placode thickening, a failure of placode invagination, an inability to express αA-Crystallin and FOXE3, and a reduced expression of SOX2 (Rajagopal et al., 2009). Interestingly, the deletion of both these receptors did not lead to a reduction in PAX6 expression (Rajagopal et al., 2009). In mice, FGF may provide another inductive signal from the OV. Depletion of FGFR1 and FGFR2, at the lens placode stage, resulted in massive apoptosis, yet proliferation in this zone was not altered (Garcia et al., 2011). Depleting the lens placode of the FGFR associated docking protein; FRS2α, also resulted in apoptosis at the lens placode stage. The expression of a dominant-negative FGFR led to a poorly developed lens placode and a delay in subsequent invagination (Faber et al., 2001). Interestingly, the impact on FGF signaling on PAX6 expression remains controversial. Deletion of Fgfr1 and 2 at the lens placode stage resulted in

11 normal PAX6 expression levels at E9.5, yet PAX6 expression declined at E10.5 (Garcia et al., 2011). Deletion of Frs2α in the lens placode did not result in any alteration in Pax6 expression (Madakashira et al., 2012). The expression of the dominant-negative FGFR resulted in a decrease in Pax6, Sox2, and FoxE3-all essential genes in lens induction (Faber et al., 2001). Both the Garcia (2011) and Faber (2001) investigated the interaction of BMP signaling and FGF signaling in lens formation (Faber et al., 2001; Garcia et al., 2011). Faber generated mice with the dominant-negative FGFR that were also heterozygous null for Bmp7. Although heterozygosity for a null Bmp7 mutation does not produce a lens phenotype, the addition of a dominant negative FGFR led to a lens with more severe defects than solely expressing the dominant negative FGFR (Faber et al., 2001). In further support of this interaction, the lab of David Beebe conditionally deleted both Fgfr2 and the BMP receptor Acvr1, which enhanced the cell death observed in lens placodes depleted of just FGFR2 (Garcia et al., 2011). The Faber 2001 manuscript concluded that FGF signaling and BMPs work cooperatively upstream of FoxE3, Pax6, and Sox2 to establish their placodal expression (Faber et al., 2001).

1.5 OCULAR ENVIRONMENT DICTATES LENS EPITHELIAL AND FIBER CELL IDENTITY: FORMATION OF THE FGF GRADIENT HYPOTHESIS

As the previous section indicated, the lens requires paracrine signaling from the OV for lens placode formation. Later in development, the ocular lens still requires paracrine signaling from the surrounding environment to regulate the proliferation and differentiation that continues throughout adulthood. The anterior hemisphere of the lens bathes in the aqueous humor, whereas the posterior hemisphere of the lens is bathed in the vitreous humor. Both the aqueous and vitreous body provide a complement of growth factors directing lens epithelial cell proliferation and differentiation. Coulombre and Coulombre in 1963 were the first to highlight the importance of the aqueous and vitreous humor on lens cell fate as they flipped an embryonic chick lens, so that, the epithelium faced the vitreous side and presumptive retina, and the lens fiber cell mass faced the aqueous humor and presumptive cornea (Coulombre and Coulombre, 1963). Inverting the lens resulted in the lens epithelial cell layer differentiating into lens fiber cells, providing the first evidence that the ocular environment may influence epithelial cell proliferation, cell cycle withdraw and subsequently fiber cell differentiation (Coulombre and Coulombre, 1963).

12 The McAvoy lab extended the work of Coloumbre and Coloumbre by growing rat lens epithelial explants in media supplemented with either aqueous humor, or vitreous (Lovicu et al., 1995). The lenses grown in vitreous supplemented media underwent morphological events characteristic of lens fiber cell differentiation, such as fiber cell elongation, organelle loss, and the formation of ball-socket junctions (Lovicu et al., 1995). One of the main differences between the aqueous humor and vitreous was the concentration of FGF, as the vitreous humor contained much higher levels of FGF ligand (reviewed in McAvoy et al., 1999). McAvoy demonstrated that lens epithelial cell explants supplemented with low concentrations of FGF, proliferated (3ng/mL), whereas increasing FGF2 (40ng/mL) promoted cell cycle withdraw and differentiation (McAvoy and Chamberlain, 1989). These experiments led to the FGF gradient hypothesis. It was hypothesized that low concentration of FGF provided by the aqueous humor stimulates cellular survival and proliferation. Intermediate levels of FGF concentation initiates cellular migration, and high levels of FGF supplied by the vitreous elicits a fiber cell differentiation response (reviewed in McAvoy et al., 1999). The experiments done by Coloumbre and Coloumbre and McAvoy initiated the interest of FGF signaling in the lens, and following these experiments, the lens was utilized as a premier tool to study the mechanistic role of how FGF signaling promotes survival, proliferation, and differentiation. Furthermore, numerous other experiments followed, utilizing the lens to further the understanding of other essential growth factors and signaling pathways and their impact on cellular proliferation, survival, and differentiation.

1.6 LENS EPITHELIAL CELL PROLIFERATION AND THE CELL CYCLE

Every multicellular organism requires the control of cellular proliferation to ensure proper embryogenesis, tissue formation and patterning, postnatal growth and maintenance, and the replacement of damaged cells. Like every other tissue, the development of the ocular lens requires precise control of the cell cycle. Historically, the lens has been used as a crucial tool to study growth factor signaling and the impact on cellular proliferation and cell cycle withdraw. One of the main advantages of using the lens derives from the extensive histological mapping of lens proliferation throughout lens development. The invaginating lens placode to the early lens vesicle stage relies on high levels of proliferation for rapid tissue growth (reviewed in Griep and

13 Zhang, 2004). Moreover, the high levels of proliferation in the PLE bound by fibrillar networks to the OV promote the thickening of the lens placode (reviewed in Griep and Zhang, 2004). Later in development, as posterior cells in the lens vesicle initiate their fiber cell differentiation process, cell cycle withdraw becomes essential for proper fiber cell differentiation (reviewed in Griep and Zhang). As the epithelial layer and fiber cell mass is established, maintaining mechanisms that ensure proper cell cycle withdraw as lens epithelial cells approach the equator is crucial for the maintenance of lens clarity. Cells remaining prolific in the fiber cell mass contribute to lens cataract (Zhang et al., 1997; Zhang et al., 1998). Furthermore, the mature lens displays low levels of proliferation in the central lens epithelium, and contain another population of cells in the epithelium exhibiting ongoing proliferation in the “germinative zone” (Fig. 1.3 B) providing the constant source of new fiber cells as the lens slowly grows throughout the life of the animal. At the lens equator, differentiating epithelial cells permanently withdraw from the cell cycle and express the cell cycle inhibitors, p27KIP1 and p57KIP2, at this zone (Fig. 1.3 C). Taken together, the lens represents an ideal system to observe factors partaking in promoting proliferation and initiating permanent cell cycle withdraw and the requirement of regulating these processes to allow for proper differentiation. This section will outline the cell cycle and regulatory components. Moreover, it will highlight the cell cycle regulatory components within the lens, and the growth factor impact of cell cycle control.

1.6.1 Cell Cycle Progression by the CDKs and cyclins

Progression through the stages of the cell cycle requires the action of cyclin-dependent kinases (CDKs) and their complementary activators, the cyclins. CDKs are serine-threonine kinases that are always present in actively cycling cells, but their cyclin co-activator, fluctuate with the phase of the cell cycle (Griep and Zhang, 2004). Upon CDK activation by their complementary cyclin, CDKs phosphorylate specific targets; driving progression through the cell cycle. The mammalian cell cycle is divided into four different phases, gap 1 (G1), synthesis (S), gap 2 (G2), and mitosis (M). The two points of most stringently regulated cell cycle transitions occur at the end of the two gap phases, immediately prior to the S and M phases. In G1, before entering S-phase, the cell enters a “restriction point.” At this moment, the cell makes a commitment to proliferate or exit from the cell cycle (reviewed in Griep and Zhang,

14 2004). If the cell passes the restriction point, thus entering S phase, DNA replication occurs. The replication of the DNA generates the DNA content required to provide for 2 resulting daughter cells. The complex of CDK4 and cyclin D are critical for the transition from G1 to S-phase. cyclinD expression is sensitive to mitogenenic activity, representing one mechanism by which growth factor signaling controls cell cycle progression. More specifically, the catalytic activity of activated ERK1/2 enhances cyclinD expression, whereby, inhibition of ERK1/2 leads to the down-regulation of cyclinD expression. Upon activation of CDK4/cyclinD, CDK4 phosphorylates the gene product of retinoblastoma (Rb), pRb. The phosphorylation of pRb promotes disassociation of pRb from the E2F transcription factor, which promotes transcriptional activation of E2F target genes (reviewed in Griep and Zhang, 2004; Dyson, 1998; Mulligan and Jacks, 1998; Nevins, 1998). E2F target genes drive the expression of cyclin E, which in turn, activates CDK2. Upon activation, CDK2 also phosphorylates pRb resulting in a positive feedback enhancement of cyclinE expression. Other cell cycle regulators, such as, dihyrofolate reductase, Cdk1, B-myb, c-myc, and N-myc rely on E2F driven transcription (reviewed in Griep and Zhang, 2004; Dyson, 1998; Mulligan and Jacks, 1998; Nevins, 1998). In addition to targeting pRb, CDK2/cyclinE activity drives centrosome duplication and histone transcription (Lacy and Whyte, 1997; Ma et al., 2000; Matsumoto et al., 1999; Zhao et al., 2000). The complex of CDK2/CyclinE drives the cell into S-phase (reviewed in Dulic et al., 1992; Griep and Zhang, 2004; Koff et al., 1992; Ohtsubo et al., 1995). Cyclin B accumulates in the cytoplasm from the S to G2 phase. The activation of CDK1 by cyclin B marks the onset of mitosis. Upon activation, CDK1/cyclinB transports to the nucleus and immediately initiates cell rounding (Gavet and Pines, 2010). Subsequently, CDK1 phosphorylation of Eg5 kinesin increases the microtubule binding activity of Eg5 and facilitates centrosome separation (Blangy et al., 1995; Sawin and Mitchison, 1995). The phosphorylation of NuMA by CDK1/cyclin B coordinates the spindle assembly and organization during both prophase and metaphase (Kotak et al., 2013). Moreover, CDK1 phosphorylation of Histone H1 promotes chromosome condensation (reviewed in Nigg, 1993). Major nuclear architecture shifts occur during mitosis, which too, are mediated in a CDK1/cyclin B dependent manner. The nuclear importation of CDK1/cyclinB1 promotes lamin phosphorylation which leads to the disassembly of the nuclear envelope (reviewed in Nigg, 1993). Lower levels of CDK1 activity can stimulate the chromosome structure shifts, cell rounding, and centrosome movement, but

15 higher levels of CDK1 activity are required for nuclear envelope breakdown (NEBD) which is stimulated by CDK1 phosphorlation of nuclear lamins (reviewed in Nigg, 1993). The highest level of CDK1 activity is required for the phosphorylation of the anaphase promoting complex/cyclosome (APC), which leads to the subsequent inactivation of CDK1 by the degradation of cyclin B (Kraft et al; 2003; Rudner and Murray 2000). Entry into anaphase requires the inactivation of CDK1. Upon inactivation of CDK1, the phosphatase, PPP2CA dephosphorylates NuMA, which generates cortically localized NuMA, and thus, permits spindle elongation during anaphase (Kotak et al., 2013). Throughout the remainder of mitosis, CDK activity remains low. Stringent regulation of cell cycle entry prevents continuous proliferation. As there exists many molecules responsible for halting the cell cycle, it is presumably a difficult process to halt, as disruptions of nearly any cell cycle inhibitor contribute to tumorigenesis. The master brake of the cell cycle is pRB and following mitosis, pRB becomes dephosphorylated and forms a transcriptionally repressive complex with E2Fs (Lundberg and Weinberg, 1999). The p21CIP1 family and p16INK4 family represent two additional classes of CDK inhibitors. The p21CIP1 family includes p21CIP1, p27KIP1, and p57KIP2 and these inhibit all CDKs involved with the G1-S transition (reviewed in Griep and Zhang 2004). The p16INK4 family includes p15, p16, p18, and p19 and directly binds and inhibits CDK4 and CDK6 (reviewed in Robinson 2004; Harper, 1996).

1.6.2 Cell-Cycle Machinery in the lens

It is of no surprise that the main CDKs and cyclins are present in the lens, as the lens epithelium continually has a region of cells actively cycling. Cyclins A , B, D, and E, as well as CDK1, 2, and 4 are found in the lens epithelium (Table 1) (Fromm and Overbeek, 1996; Gao et al., 1995; Gao and Zelenka, 1997; Griep and Zhang, 2004; Hyde and Griep, 2002). Moreover E2F 1, 2, 3, 4, and 5 are expressed in the lens epithelium (Rampalli et al., 1998). Counteracting the cell cycle machinery in the lens epithelium, cell cycle inhibitors are also prevalent. The cell cycle inhibitors present in the lens include the CDK inhibitors, p57KIP2, p27KIP1 (Zhang et al., 1998). Additionally, the E2F inhibitors, pRB, p107, and p130 are displayed in the lens epithelial cell layer (Griep and Zhang, 2004). As the lens epithelial cells approaching the equator of the

16 lens require permanent withdrawal from the cell cycle, these equatorial cells upregulate the expression of pRb, p57KIP2 and p27KIP1 (Lovicu and McAvoy, 1999; Zhang et al., 1998). Furthermore, disruption of p27KIP1 and p57KIP2, or loss of pRb leads to cells remaining prolific in the fiber cell compartment of the lens (reviewed in Griep and Zhang, 2004; Zhang et al., 1998). Despite being permanently withdrawn from the cell cycle, the lens fiber cells continue to contain active cyclin B and CDK1 (Table 1) (He et al., 1998). It is surprising that cyclin B expression is maintained in the fiber cell mass, as transcription of each cyclin relies on the particular phase of the cell cycle, so in non-mitotic cells, cyclin B is rarely found. Additionally, the ubiquitin proteasome pathway normally targets cyclin B for degradation prior to anaphase. Interestingly, the kinase activity of CDK1 in lens fiber cells peaks between embryonic day E18-E19 in mice- representing the stage where nuclear removal initiates (He et al., 1998).

1.6.3- Receptor Tyrosine Kinase Activity Drives Cellular Proliferation in the Lens

Entry and exit from the cell cycle frequently rely on growth factor signaling. Gradients of FGF ligand provided by the vitreous and aqueous humor dictate both cellular proliferation and cell cycle withdrawal (reviewed in McAvoy et al., 1999), yet other growth factors receptors, in synergism with FGF/FGFR signaling, provide cues regulating cellular proliferation and withdrawal. Acting downstream of FGF/FGFR signaling, the PI3K/AKT and MAPK/ERK1/2 signaling cascades, primarily regulate cellular proliferation in the context of the lens. Other RTKs expressed in the lens are capable of stimulating both the MAPK/ERK1/2 and PI3K/AKT. The RTKs expressed in the lens include members of the insulin and insulin-like growth factor receptors (IGF), platelet-derived growth factor (PDGF) receptors, epidermal growth factor (EGF) receptors, and hepatocyte growth factor (HGF) receptors (reviewed in Lovicu et al., 2014; Robinson, 2006). Although IGF signaling is most commonly associated with cellular metabolism, IGF signaling participates in cell cycle regulation in the lens epithelium. Not only is the aqueous concentration of IGF twice of the vitreous, IGFRs are prominent in the mitotically active compartments of the lens epithelium (Alemany et al., 1990; Arnold et al., 1993). Similar to adding FGF to lens epithelial cell cultures (LECs), IGF addition to in vitro lens systems demonstrated the ability to enhance proliferation of lens epithelial cells (Liu et al., 1996).

17 Moreover, overexpressing IGF-1 in mouse lenses caused an expansion of both the germinative zone and transition zone (Shirke et al., 2001). One prominent hypothesis is that IGF/IGFR signaling provides the spatial cue to specify the boundaries of the germinative zone and regulates proliferation in this zone (Griep and Zhang, 2004). The ciliary body and iris strongly express PDGF (reviewed in Lovicu et al., 2014). As both the iris and ciliary body are two structures in close proximity to the germinative zone, PDGF supplied by these two structures may participate in regulating proliferation in the germinative zone of the lens epithelium (reviewed in Lovicu et al., 2014). Furthermore, the PDGF receptor PDGFα primarily localizes to the germinative zone in the postnatal lenses (Reneker and Overbeek, 1996). In lens epithelial cell explants, PDGF addition, like IGF and FGF, promoted lens epithelial proliferation (Iyengar et al., 2009). Supplementing the explant studies, transgenic PDGF overexpression in mouse lenses increased cyclin A, cyclin D2, and S- phase entry in the lens epithelium (Reneker and Overbeek, 1996). Lastly, PDGF stimulation may be more potent than FGF signaling on proliferation. An antibody antagonizing PDGF-D reduced lens epithelial cell proliferation to a greater degree than antagonizing FGF-2 in rat eye anterior segment organ cultures (Ray et al., 2005). The aqueous humor surrounding the lens also contains EGFs, and the lens epithelium expresses EGFR receptors. Like the other RTKs mentioned, EGFs are implicated in promoting lens epithelial cell proliferation and addition of EGF to lens explants and whole lens cultures increases epithelial cell proliferation (Arora et al., 1996; Haque et al., 1999; Reddan and Wilson- Dziedzic, 1983). The RTKs expressed in the lens may act in “synergism” with each other to promote cellular proliferation. This synergism can be attributed, at least in part, to their different ways they active the same downstream signals. Both PI3K/AKT and MAPK/ERK1/2 elicit an incredibly diverse response dependent upon the activation level and surrounding cellular context, yet both are capable of regulating cellular proliferation. Activated MAPK/ERK1/2 stimulates cellular proliferation by several distinct mechanisms. One-way ERK1/2 activation increases cellular proliferation is by increasing ribosomal biogeneis by activating ribosomal S6-kinase-1 (RSK1). Activation of the ribosomal biogenesis permits the protein synthesis required for cellular growth and proliferation (Felton-Edkins et al., 2003; Stefanovsky et al., 2001). Activated ERK1/2 also moves into the nucleus where it phosphorylates numerous transcription factors

18 leading to the transcription of genes involved in cellular proliferation (Weng et al., 2001a). Amongst the most direct link to cell cycle control by MAPK signaling is through the transcriptional activation of the essential CDK1 coactivator, cyclin D1. RAS activation promotes Sp1-mediated cyclin D1 expression (Klein and Assoian, 2008). RAC activation, upstream of NF- κB, strongly induces cyclin D1 (Klein and Assoian, 2008). ERK1/2 may also directly stimulate cyclin D1 expression as mouse embryonic fibroblasts (MEFs) treated with the ERK1/2 inhibitor, U0126 at mid G1, exhibited fewer cyclin D1 transcripts (Villanueva et al., 2007). In the context of the lens, the impact of ERK1/2 on cellular proliferation has received frequent attention in both in vitro and in vivo systems. The increased proliferation resulting from FGF-2 addition in rat lens epithelial cell cultures (rLEC) relies on ERK activity. U0126 treatement of FGF-2 treated rLEC abolished the FGF-induced proliferation. Furthermore, in vivo depletion of ERK2 in mouse lenses disrupted lens epithelial cell proliferation in the germinative zone, and these lenses displayed significantly reduced amounts of cyclin D1 protein (Upadhya et al., 2013). AKT activates both glycogen synthesis kinase 3 (GSK3) and mammalian target of rapamycin (mTOR), which promote cellular proliferation. More directly to cell cycle control AKT, like ERK, regulates cyclin D1 levels. Despite the precise mechanism remaining vague, inhibiting AKT activation with LY2994002 severely inhibits cyclin D1 expression (Muise- Helmericks et al., 1998). Additionally, AKT activation can result in reduced levels of the CDK inhibitors, p27KIP1 and p57KIP2 (Testa and Bellacosa, 2001). Again, similar to ERK activation, the FGF-2 induced proliferation in the lens epithelial cell culture relies on AKT activation, although AKT activation on proliferation may be secondary to the impact of AKT on ERK (Iyengar et al., 2006). LY294002 treatment to FGF-2 treated epithelial cell cultures resulted in a reduction of proliferation and reduced ERK activation (Iyengar et al., 2006). As the impact of AKT on cellular proliferation shares many similarities to ERK activity, it seems reasonable to question whether the impact of AKT on proliferation is due to increasing ERK activity. As noted, many RTKs can enhance cellular proliferation in the lens epithelium. Most likely, the combined effort of all the RTKs, acting in synergism with each other, provide for the appropriate regulatory cues for cellular proliferation. Many of the lens culture studies demonstrated enhanced proliferation when adding multiple RTKs to the culture system. For

19 example, the addition of IGF and FGF to lens epithelial explants enhanced the lens epithelial cell proliferation (Liu et al., 1996). Furthermore, blocking different RTKs in lens epithelial cell cultures grown in an aqueous humor culture provided insight on the impact of each RTK on cellular proliferation. Blocking FGF did not inhibit lens epithelial cell proliferation, but blocking FGF in the presence of blocking either IGF, PDGF or EGF blocked the proliferative response (Iyengar et al., 2006; Iyengar et al., 2009; Iyengar et al., 2007). Regarding ERK1/2 activation, blocking just PDGF resulted in an immediate reduction of ERK (within 20 minutes), yet ERK1/2 activity was not altered in progressing hours. Furthermore, FGF addition leads to 6 hours of ERK activation whereas additions of IGF, PDGF or EGF lead to only 1 hour of ERK1/2 activation (Iyengar et al., 2009). Similarly, blocking FGF resulted in ERK1/2 being activated for only 1 hour (Iyengar et al., 2009). Taken together, it is possible that the initial ERK activation requires stimulation by PDGF, IGF or EGF, yet this response is only transient, and in order to display a sustained ERK1/2 response to promote proliferation, FGFR activation is required. In vivo functional analysis of FGFRs in the lens makes it difficult to definitively state that FGFR signaling provides an indispensable proliferative cue in the lens epithelium. Conditional deletion of Fgfr2 at the lens placode stage results in an impairment of cellular survival and differentiation, yet does not result in profound proliferative defects (Garcia et al., 2005). Combined deletion of all three Fgfrs at the lens vesicle stage, again leads to cell death and fiber cell differentiation defects, but does not lead to reduced proliferation at E11.5 or E12.5 (Zhao et al., 2008). By E14.5, these Fgfr triple knockout lenses experienced reduced BrdU incorporation (Zhao et al., 2008). Similarly, deletion of the FGFR docking protein Frs2α at the lens placode stage did not alter lens epithelial cell proliferation until E15.5 (Madakashira et al., 2012). Taken together, this could support the idea that FGFR provides for the sustained ERK1/2 activation necessary for continued proliferation at later stages of lens development, whereas PDGF, IGF or EGF provide for immediate ERK1/2 activation. On the other hand, the lack of proliferation at E14.5 may be secondary to the apoptosis occurring in the FGFR1-3 or Frs2α deficient lenses (Madakashira et al., 2012; Zhao et al., 2008). To provide further support of the in vivo necessity of FGFRs in lens epithelial cell proliferation, overexpression of a dominant-negative form of Fgfr1, which was expected to eliminate all FGFR signaling, led to a reduction of epithelial cell proliferation at E12.5 (Faber et al., 2001). With the conflicting results of inactivating aspects of FGFR signaling on proliferation, it is difficult to determine if FGFR signaling has an

20 indispensable role in lens epithelial cell proliferation, or if other RTKs can compensate for FGFR loss.

1.7 BALANCING LENS CELL SURVIVAL AND DEATH

The development of the ocular lens requires both survival and apoptotic cues to direct the morphological changes occurring throughout development. The detection of apoptotic cells occurs at E9 in mice and apoptosis is prominent in the anterior cells of lens pit at E10.5 (Aso et al., 1998). Moreover, apoptosis is required for proper separation of the lens vesicle from the overlying surface ectoderm, and a failure of the lens vesicle separation leads to Peter’s Anomoly (Ozeki et al., 2001). Later in development, lens cells favor survival, and very few apoptotic cells are present in a normal lens from E14.5 to birth. Despite the lows levels of apoptosis present in the lens at later stages of development, the lens still contains active apoptotic pathways, which partake in the fiber cell differentiation process. Although the apoptotic machinery initiates during fiber cell differentiation, lens fiber cells do not completely go through apoptosis. As apoptosis is required for lens vesicle separation, and apoptotic machinery is present during fiber cell differentiation, the lens requires a fine balancing act between survival factors and apoptotic factors. If the balance is tipped too far towards apoptosis severe lens defects occur, such as anopthalmia, microphthalmia, and cataract formation, thus making it a unique tool for understanding mechanisms of survival factors in the lens.

1.7.1 FGFRs are required to tip the balance towards survival

Depletion of FGFR2, FGFR1 and FGFR2, or FRS2α at the lens placode stage results in apoptosis and anophthalmia or microphthalmia (Garcia et al., 2011; Garcia et al., 2005; Madakashira et al., 2012). Depletion of all three FGFRs at the lens vesicle stage also leads to massive apoptosis and small lenses (Zhao et al., 2008). As a result of FGFR deficiency, activated ERK1/2 and AKT are significantly reduced, which is the likely cause of the enhanced apoptosis (Garcia et al., 2011; Madakashira et al., 2012). The lab of Lixing Reneker supported the requirement of ERK1/2 in lens cell survival. Dr. Reneker showed that in vivo depletion of ERK2 at the lens vesicle stage increased apoptosis (Upadhya et al., 2013). The apoptosis resulting from

21 ERK2 depletion was likely due to decreased Survivin protein levels. Survivin inhibits apoptosis by blocking caspase 3, 7, and 9 activity (Upadhya et al., 2013). Although direct in vivo functional characterization of AKT in the lens does not exist, AKT has a well-established survival role in nearly every tissue. One way AKT prevents apoptosis is through phosphorylating MDM2. Once phosphorylated, MDM2 transports to the nucleus and mediates the degradation of the proapoptotic factor p53 and, therefore, prevents Pten transcription, as p53 is a transcription factor known to drive the transcription of Pten (Endersby and Baker, 2008; Mayo et al., 2002; Mayo and Donner, 2002). Interestingly, amongst the most prominent ways AKT protects against apoptosis is by inhibiting the most well characterized inhibitor of AKT, PTEN. The functional role of PTEN in lens development has yet to be explored, although mutations in PTEN lead to lens defects. A common pathology of patients with mutations in Pten, or PTEN harmartoma tumor syndrome (PHTS) is lens cataract. Furthermore, deletion of Pten at the lens vesicle stage lead to lens swelling, opacities and eventual organ rupture (Sellitto et al., 2013). The phenotype resulting from Pten deletion at the lens vesicle was a result of high levels of AKT activity that inhibited Na+/K+-ATPase, thus disrupting the Na+ driven microcirculatory system in the lens that is responsible for carrying nutrients to the fiber cells and disposing waste (Sellitto et al., 2013). Despite this characterization of the impact of PTEN on lens physiology, it has not yet been established the impact PTEN has on lens cell survival throughout the development of mouse lenses from the lens placode to birth.

1.8 LENS EPITHELIAL CELL-TO-FIBER CELL DIFFERENTIATION

Drastic changes occur to lens epithelial cells during the epithelial-to-fiber cell differentiation. The lens fiber cell differentiation process initiates after the completion of the final cell division at the germinative zone where they subsequently enter the transition zone of the lens. These differentiating epithelial cells are either displaced, or start to migrate towards the lens equator, although because certain levels of FGF stimulation initiate migration in rat lens epithelial cell cultures, these precursors are likely migrating towards the posterior (Chamberlain and McAvoy, 1989). As these cells migrate, they initiate the expression of cell cycle inhibitors

22 KIP2 KIP1 p57 , p27 , and pRb (Lovicu and McAvoy, 1999; Rampalli et al., 1998; Zhang et al., 1998). Although fiber cell elongation, which represents the morphological hallmark of fiber cell differentiation, does not occur in the transitional zone, elongation requires cell cycle withdraw. Disrupting both p57KIP2 and p27KIP1, or pRB lead to fiber cell elongation defects (Hyde and Griep, 2002; Zhang et al., 1998). Given this, it is apparent that cell cycle withdrawal and lens fiber cell differentiation are intrinsically linked. Following the onset of cell cycle inhibitor expression and, as the differentiating lens epithelial cells approach the lens equator, the cells begin to elongate. At the equator, differentiating fiber cells significantly increase their transcriptional activity, as these differentiating cells not only require the production of fiber cell specific proteins, but these differentiating fibers must prepare for a massive increase in cell volume during elongation (Bassnett and Beebe, 2004). Most of the proteins produced during fiber cell differentiation are structural, including both crystallin proteins, which ensure lens clarity, and intermediate filament proteins and membrane proteins that ensure cellular contact at the lens epithelium and lens capsule (Bassnett and Beebe, 2004). Following elongation, the fiber cells move interiorly. Here, they restructure their lateral membrane complexes, creating ball socket junctions between fiber cells and lose their contact between the capsule and epithelium (Bassnett and Beebe, 2004). Aquaporin0, the most prominent membrane protein in lens fiber cells, is required for the formation of the ball socket junctions (Bassnett and Beebe, 2004). The last stage of lens fiber differentiation is organelle loss to form a zone in the center of the lens free of all organelles, including but not limited to, the nuclei, mitochondria, and endoplasmic reticulum (referred to as the organelle free zone, OFZ). Formation of the OFZ completes the maturation of the lens fiber cells. It is the point that they are considered mature fibers (reviewed in Bassnett, 2009; Wride, 2011). The organelles refract light, so lens clarity requires organelle loss.

1.8.1 Gene Expression Changes During Differentiation; Growth Factor control of gene regulatory networks.

The control of gene expression lies at the heart of developmental biology. The lens provides a unique system to determine the molecular mechanism of chromatin environment

23 changes mediated by growth factor signaling. As FGF-2 addition to lens epithelial cell cultures resulted in a fiber cell differentiation response, most focus on fiber cell gene activation centered on how the downstream factors of FGF signaling promote a chromatin environment favorable for the transcription of fiber cell genes. Inhibiting either AKT or ERK1/2 block the differentiation response induced by FGF-2, and because of this, many of the proposed mechanisms of FGF induced gene regulatory networks involve the action of PI3K/AKT, and MAPK/ERK1/2 (Lovicu and McAvoy, 2001; Wang et al., 2009). Mouse in vivo studies support the impact of FGF signaling on lens epithelial to fiber cell differentiation. FGFR activation in the lens epithelium by transgenic overexpression of Fgf1, Fgf3, Fgf4, Fgf7, Fgf8, Fgf9 or ectopically overactivating Frs2α leads to fiber cell differentiation in the lens epithelium (Lovicu and Overbeek, 1998; Madakashira et al., 2012; Robinson et al., 1998; Robinson et al., 1995). Supplementing the in vivo evidence that over stimulated FGF signaling in the lens initiates lens fiber cell differentiation, disrupting FGFR signaling in mouse lenses resulted in lens fiber cell differentiation defects. Transgenic expression of a self-dimerizing, secreted version of the extracellular domain of FGFR1 and 3 in the lens were created to bind and sequester FGF ligands to reduce FGFR signaling (Govindarajan and Overbeek, 2001). Although the secreted FGFR1 did not produce an inhibition of fiber cell differentiation, the secreted FGFR3 resulted in postnatal fiber cell differentiation defects (Govindarajan and Overbeek, 2001). These defects included a posterior displacement of the transition zone of the lens, and what appeared to be lens epithelial cells nearly enclosing both the anterior and posterior sides of the lens (Govindarajan and Overbeek, 2001). Depletion of just FGFR2 at the lens placode stage leads to a delay in differentiation marked by an early fiber cell elongation defects, and posterior cells remaining prolific at E12.5 (Garcia et al., 2005). Deletion of all three FGFRs in the lens (FGFR1-3) leads to reduced expression of Prox1 and c-maf, which are transcription factors important for the expression of the cell cycle inhibitors p57KIP2 and p27KIP1, and all lens crystallin proteins (reviewed in Cvekl and Ashery-Padan, 2014). Furthermore, the FGFR triple knockout lenses failed to express p57KIP2 and p27KIP1 γ and β-crystallin (Zhao et al., 2008). As FGFR signaling requires heparin sulfate proteoglycans (HSPG) for effective FGFR activation, mutants null for both HPSG genes, Ndst1 and Ndst2 lead to lower levels of γ and β-crystallin protein expression (Qu et al., 2011).

24 More directly influencing fiber cell differentiation, the transcription factors c-Maf, Hsf4, Pax6, Pitx3, Prox1, Sox1 and Sox2 control the expression of two or more crystallin genes, fiber cell intermediate filament proteins, and cell cycle inhibitors (reviewed in Cvekl and Ashery- Padan, 2014; reviewed in Cvekl and Duncan, 2007). c-Maf null lenses did not express αA, αB, γ or β-crystallin and experience fiber cell elongation defects (Kawauchi et al., 1999). Mice null for Prox1 exhibit down regulated p57KIP2 and p27KIP1 and have fiber cell elongation defects (Wigle et al., 1999). Hsf4 knockout mice display reduced γ-crystallin, Bfsp1 and 2, DNaseIIβ and exhibit nuclear retention (Fujimoto et al., 2004; He et al., 2010; Shi et al., 2009). Deletion of Pax6 at the lens vesicle stage resulted in cell cycle withdraw failure in the lens fiber cell mass without a reduction in p57KIP2 and p27KIP1 (Shaham et al., 2009). Fgfr signaling, either directly or indirectly, regulates the expression of c-maf, Pax6, Prox1, and Hsf4, but the mechanism by which Fgf signaling regulates these transcription factors remains elusive and contradictory. Using small molecules inhibitors of FGFR activity in mouse explant cultures diminished the expression of Pax6 (Faber et al., 2001). In the same study, transgenic expression of a dominant- negative FGFR in the presumptive lens ectoderm reduced Pax6 expression in the lens placode (Faber et al., 2001). Contradictory to this, the FGFR triple knockout at the lens vesicle stage experienced retained PAX6 expression in the posterior cells of the lens (Zhao et al., 2008). Similarly, Ndst mutants expressed high levels PAX6 in posterior lens cells (Qu et al., 2011). It remains to be determined the way in which FGF signaling impacts PAX6 in the lens, but their relationship appears complex. PAX6 may be FGF-responsive in an ERK1/2 or AKT-dependent manner. Inhibition of FGF signaling by retinoic acid in explanted caudal regions decompacts the chromatin near the PAX6 locus; therefore, FGF signaling compacts the PAX6 locus inhibiting its expression (Patel et al., 2013). On a protein level of control, PAX6 is a phospho-protein with 4 highly conserved phosphorylation sites, which can be phosphorylated by ERK1/2-which promotes the transactivation of PAX6 (Mikkola et al., 1999). As PAX6 is essential for early lens development, and can act as both an inhibitor and activator of genes associated with lens differentiation, it will would be interesting to see what the requirement of PAX6 phosphorylation has on lens development. Furthermore, as both FGFR signaling and PAX6 regulate lens, retina, pancreas, and brain development it is crucial to further understand, in an in vivo context, the relationship PAX6 has with FGFR signaling on both its activation through phosphorylation and the transcriptional activation of the Pax6 gene. The regulation of c-maf by FGF signaling is not

25 as contradictory. FGF signaling is known to control c-Maf expression by its promoter (reviewed in Cvekl and Ashery-Padan, 2014). Although knocking out just Fgfr2 at the lens placode stage did not alter c-maf expression, knocking out all three Fgfrs in the lens vesicle significantly reduced c-maf expression (Zhao et al., 2008). In the Fgfr1-3 triple knockout lenses at the lens vesicle stage, Prox1 did not experience the normal upregulation at the equator (Zhao et al., 2008). miRNAs likely participate in fiber cell differentiation and are responsive to FGF signaling. Conditional knockout of Dicer1, using either the Le-Cre or the MLR10 Cre transgene (initiating Cre expression at the lens vesicle stage) initiated the interest in miRNAs in lens development (Li and Piatigorsky, 2009; Wolf et al., 2013). Dicer is responsible for cleaving double-stranded RNA or pre-microRNA producing microRNAs. MLR10 Cre –mediated deletion of Dicer1 led to fiber cell elongation defects, fiber cell organization defects, and nuclear retention (Wolf et al., 2013). The lab of Dr. Cvekl recently published late in 2013 that upon FGF2 addition to rat LECs, there are 131 FGF responsive miRNAs-targeting over 3000 transcripts (Wolf et al. 2013). FGF signaling is a key participant in the many transcriptional changes that occur during fiber cell differentiation, although the manner by which FGF signaling results in the activation or repression of certain genes remains incomplete. The dependency of FGF on differentiation may be downstream of ERK1/2 and AKT or can be through a separate pathway. The endogenous role of FGF signaling in vivo, may be primarily to maintain lens survival, and the variable readouts of lens fiber cell differentiation resulting from FGFR deficiencies may be a secondary effect of apoptosis. FGF signaling could potentially alter gene expression changes away from ERK1/2 or AKT, and might be due to the shift of the miRNAs in the lens. Future studies are required to depict more precise mechanisms on how FGFR signaling initiates fiber cell differentiation.

1.8.2 Mechanism of Nuclear Removal.

Of particular interest and focus during the fiber cell differentiation process has been the unique aspect of organelle and nuclear removal of central lens fiber cells. The initial interest in the mechanism of nuclear removal initiated over a century ago, when Rabl discovered that central lens fiber cells remove their nucleus (Rabl, 1899). Failure to remove the light scattering

26 nuclei in the lens center results in lens cataract (reviewed Wride, 2011). Cataracts caused by a number of different mutations commonly result in fiber cell nuclear retention. These include lenses lacking Hsf4b (Fujimoto, 2004), Fgfr2 (Garcia et al., 2005), Frs2α (Madakashira et al., 2012), DNaseIIβ (De Maria and Bassnett, 2007; Nakahara et al., 2007; Nishimoto et al., 2003), Cdk1 (Chaffee et al., 2014) and transgenic mice expressing K6W-ubiquitin (Caceres et al., 2010). All of these proteins may directly contribute to the mechanism of fiber cell denucleation. The lysosomal enzyme, DNaseIIβ (DLAD), is the essential nuclease for the removal of nuclear content in lens fiber cells. Mice lacking DLAD exhibit cataracts resulting from a failure to remove lens fiber cell nuclei (De Maria and Bassnett, 2007; Nakahara et al., 2007; Nishimoto et al., 2003). Both PAX6 and HSF4b binding sites are present on both the DLAD promoter, as well as a 3’ conserved region (He et al., 2010). The manuscript by He et al. 2010 postulated that a both PAX6 and HSF4b bind to the DLAD promoter (He et al., 2010). Moreover, PAX6 and HSF4 demonstrated the ability to recruit the chromatin-remodeling enzyme, BRG1, to initiate DLAD transcription (He et al., 2010; Tu et al., 2006). Both PAX6 and HSF4b transcriptional activation are controlled by MAPK activation, through ERK1/2 (Hu et al., 2013; Mikkola et al., 1999). It is tantalizing to speculate that ERK1/2 activation is required to bring HSF4b, Pax6, and BRG1 to the DLAD promoter to generate a chromatin environment favorable for DLAD transcription. The mechanism by which lysosomal DLAD gains access to fiber cell nuclear DNA remained elusive prior to the start of this dissertation. Preceding large-scale destruction of fiber cell nuclear DNA, the nuclear lamina appears to dissolve (Bassnett, 1997). Although the mechanism by which fiber cells break down their nuclear lamina remained unknown, mitotic cells experience similar disassembly of their lamina (nuclear envelope) prior to karyokinesis. The phosphorylation of nuclear lamin proteins by CDK1 (in conjunction with cyclin A or B) mediates the disassembly of the nuclear envelope in mitotic cells (reviewed in Nigg, 1993). Typically, post-mitotic cells fail to maintain expression of CDK1 or cyclins A or B (reviewed in King et al., 1994; Tommasi and Pfeifer, 1995). However, post-mitotic lens fiber cells unusually express both CDK1 and cyclin B (He et al., 1998), but the nuclear envelop in immature fiber cells remains intact. Lens cell cycle withdrawal at the transition zone requires the expression of CDK1 inhibitors, p57KIP2 and p27KIP1, (Lovicu and McAvoy, 1999; Zhang et al., 1997; Zhang et al.,

27 1998). The presence of these CDK inhibitors in post-mitotic lens epithelial cells, and immature fiber cells, may prevent CDK1 activity which would otherwise result in nuclear envelop breakdown. Several recent reports link the ubiquitin proteasome pathway (UPP) to fiber cell denucleation. The lab of Allen Taylor discovered that poly-ubiquitinated conjugates increase in the equatorial epithelial cell nuclei, and eventually localize to differentiating fiber cell nuclei (Shang et al., 1999). Zebrafish containing a mutation in the 26S proteasome gene, Psmd6, experience abnormal retention of lens fiber cell nuclei as well as a number of cell cycle alterations in the lens epithelium (Imai et al., 2010). Caceres provided evidence that the sixth lysine (K6) of the ubiquitin tail targets p27KIP1 for destruction in the lens (Caceres et al., 2010). In this study, they drove the expression of a ubiquitin K6W mutation in the lens and observed the accumulation of p27KIP1 and the lack of nuclear removal from the center of lens (Caceres et al., 2010). Organelle loss in mature fibers mimics apoptotic processes, particularly in regards to the morphological nuclear changes. Of note, the nuclear lamin disintegrates promoting nuclear envelope breakdown (reviewed in Bassnett, 2009). Closely following the initiation of nuclear breakdown, mitochondria and the endoplasmic reticulum disintegrate, which occurs more rapidly than nuclear removal.

1.9 SPECIFIC AIMS

1.9.1 Nuclear Removal During Terminal Lens Fiber Cell Differentiation Requires CDK1 Activity; Appropriating Mitosis-Related Nuclear Disassembly.

Chapter 3 focuses on the role of CDK1 in the embryonic mouse lens. Despite being a protein almost exclusively associated with actively cycling cells, CDK1 and its co-activator, cyclin B, maintain their expression in the terminally differentiated lens fiber cells (He et al., 1998). As it is unusual for CDK1/Cyclins to retain their expression in terminally differentiated cells, we questioned whether CDK1 has a role in the differentiation process. Within the cell cycle, CDK1 phosphorylates many targets essential for chromosome condensation, spindle apparatus assembly, cytoskeletal rearrangements involved in cytokinesis, and nuclear lamins promoting nuclear envelope breakdown (reviewed in Nigg, 1993). All of these processes prove crucial for mitosis, yet the role of Cdk1 in post-mitotic cells remained elusive. Interestingly, as

28 mitotic cells require the disassembly of the nuclear lamins preceding karyokinesis, terminally differentiated fiber cells require nuclear lamina disassembly prior to nuclear removal (Bassnett, 1997). Here, we hypothesized that CDK1 phosphorylates nuclear lamin proteins, as it does in mitosis, to promote nuclear lamina disassembly and nuclear removal. Further, the disassembled

Through the use of MLR10 Cre, we specifically deleted Cdk1 in the lens to characterize the role of CDK1 in denucleating lens fiber cells.

Aim 1: To characterize the role of CDK1 in post-mitotic lens fiber cells. a) To determine the expression pattern of CDK1 in lens fiber cells. b) To determine if CDK1 activity is required for lens fiber cell denucleation. c) To determine if CDK1 activity is required for removal of mitochondria and endoplasmic reticulum from the lens fiber cells.

Aim 2: To determine if lens epithelial cells require CDK1 activity for DNA synthesis and cell cycle progression.

1.9.2 FGFR and PTEN signaling interact during lens development to regulate cell survival

Despite the ability of FGF to induce fiber cell differentiation in lens epithelial cultures and in vivo studies, removal of FGFRs in vivo results in BOTH apoptosis and lens differentiation defects (Garcia et al., 2011; Garcia et al., 2005; Madakashira et al., 2012; Zhao et al., 2008). Due to the reliance on FGFR signaling for lens cell survival, it has been challenging to decouple the apoptotic phenotype from the differentiation phenotype in lenses with compromised FGFR signaling. The endogenous, mechanistic role of FGFR signaling in the lens may in fact be to primarily promote cell survival with the differentiation defects resulting as a secondary response to apoptosis. PTEN is a dual specific phosphatase capable of dephosphorylating both lipids and proteins. The phosphatase capability of PTEN antagonizes cytoplasmic AKT activation, and sends the cell towards apoptosis (Chung and Eng, 2005). Due to the antagonistic relationship between PTEN and FGFR2 in several tissue types, and the apoptotic influence of PTEN, we predicted that depleting the lens of PTEN could restore the cell death associated with FGFR2 loss in the lens. Furthermore, if PTEN restores survival in FGFR2-deficient lenses, it would

29 provide for a system to elucidate the impact of FGFR signaling on differentiation away from its role as a survival factor.

Aim 1: To determine if Pten deletion restores lens size and survival in the presence and absence of Fgfr2 deletion.

Aim 2: To determine if FGFR2 and PTEN regulate lens epithelial cell-to-fiber cell differentiation:

Aim 3: To determine if FGFR2 and PTEN deletions disrupt downstream PI3K/AKT and MAPK/ERK1/2 pathways.

30 Figure 1.1 PI3K/AKT and MAPK/ERK1/2 are the two main arms of Fgfr signaling The ligand-mediated dimerization of FGFRs induced tyrosine phosphorylation and the assembly of signaling complexes leading to the activation of several common pathways including RAS-RAF-MEK-ERK1/2 (also referred to as MAPK/ERK1/2) and PI3K-AKT pathways. Upon ligand binding, FRS2α is phosphorylated and through Grb2/Sos adaptor complexes attracts Ras. Following Ras activation, Raf is recruited to the plasma membrane, where Raf is phosphorylated, and subsequently phosphorylates MEK. MEK is a dual specific kinase that phosphorylates both the tyrosine and serine/threonine residues of ERK1/2. PI3K is dependent upon Frs2α phosphorylation, which recruits proteins with SH2 binding domains, such as Grb2. Grb2 recruits PI3K, and the major role of the following PI3K activation is to add a phosphate group to PtdIns(4,5)P2 (PIP2)and thereby converting it to the PtIns(3,4,5)P3 (PIP3). PIP3 then activates PDK1 activates phosphoinositide dependent kinase (PDK1) and in return, recruits Akt to the plasma membrane where Akt is activated by a series of phosphorylations PTEN dephosphorylates PIP3-converting it back to its inactive form PIP2, therefore inhibiting AKT activation.

31 Figure 1.2 Overview of Lens Development. Lens development initiates at E9.5 when the surface ectoderm overlaying the neural ectoderm (A) starts to thicken to form a lens placode (B). The placode invaginates to form a lens pit (C), which eventually pinches off from the rest of the surface ectoderm forming a lens vesicle (D). Cells at the posterior half of the lens vesicle become primary lens fiber cells, and start to elongate (E) eventually contacting the cells at the anterior half, which have become the lens epithelium (F). Mature lenses are comprised of epithelial cells that line the anterior of the lens, and proliferate in a zone termed the germinative zone (G). Epithelial cells approaching the lens equator withdraw from the cell cycle (transition zone), begin elongate, and express fiber cell specific genes, generating secondary fiber cells (G). Fiber cells approaching the lens center must remove their nucleus, and all other light refracting. Figure A organelles (G). Figure adapted from Lovicu and McAvoy 2005

32 Figure 1.3 The lens is a simple organ, comprised of 2 cell types, the lens epithelium (A-black arrows) and lens fiber cells (A). The lens epithelial cells line the anterior surface of the lens (A); whereas the bulk of the lens is comprised of lens fiber cells (B). The epithelial cells continually proliferate-staining positive for the S-phase marker, BrdU (B-green) with eventual localization to a zone anterior to the equator of the lens called the germinative zone (B-white box). At the equator of the lens, p27KIP1 levels rise; whereby immunological detection can display the characteristic rise of p27KIP1 marking the withdraw of cells from the cycle cycle (C-white brackets). Following cell cycle withdraw, fiber cell specific genes initiate expression. Antibodies raised against β (D) and γ-crystallin (E) are commonly used as readouts of fiber cell differentiation. Lastly, the fiber cells undergo an “attenuated apoptosis” and lose their organelles and nuclei to form a zone free of light scattering components (F-dashed line). TUNEL analysis, detecting exposed 3’OH groups, is a useful tool in visualizing the nuclear removal of fiber cells (F-inset).

33 Table 1.1 CDK and Cyclin Expression in the Lens Epithelium and Lens Fiber Cells.

(Table adapted from Gao et al 1995; He et al. 1998)

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46 CHAPTER 2

MATERIALS AND METHODS

2.1 MICE

2.1.1 Cdk1 deletion

Mice were used in accordance with the ARVO statement for the Use of Animals in Ophthalmic and Visual Research. Transgenic mice expressing Cre in the lens fiber cells were generated to mediate the deletion of loxP flanked genes. The MLR10 and MLR39 mice were used to mediate delete loxP flanked Cdk1 in either the lens epithelium and lens fiber cells (MLR10) or solely the lens fiber cells (MLR39). The MLR10 mice drive Cre expression using the -282/+43 region of the αA-crystallin promoter with a 32 base pair Pax6 consensus binding sequence in a 5’ to 3’ orientation at -86 (Zhao et al., 2004). The MLR10 mice drive Cre initiating at E10.5 in both the lens epithelial cells and lens fiber cells. The MLR39 mice drive Cre expression with the -282/+43 region of the αA-crystallin promoter. MLR39 initiates Cre expression at E12.5 in the lens fiber cells (Zhao et al., 2004). LoxP flanked Cdk1 (Cdk1L/L) mice were generated in the lab of Mitch Eddy. Using a conditional targeting vector assembled on a pBluescript II KS(+) backbone (Stratagene). The targeted region of Cdk1 contained exon 3 originally amplified with Cdk1 primers containing engineered restriction sites (forward, CGG GGT ACC TAG ATA GCT AGG GAA TCC GCA CCT GA; reverse, GCG TCC GGA GGC AGC TAC CAG AGG TGC TAA GTA AG) with flanking LoxP sites. The Cdk1 5’ arm contained exon 2, while the 3’ arm contained exons 4 and 5. The neomycin gene flanked by FRT sites was inserted into intron 2 as a selectable marker. Transfection of linearized pBluescript, screening of targeted TC-1 embryonic stem (ES) cells, and injection of blastocysts to produce chimeric males were performed as described previously

47 (Dix et al., 1996). Agouti male chimera offspring were mated with C57BL/6NCrl females and then backcrossed onto the 129S genetic background. Experimental mice were maintained on a mixed genetic background segregating for alleles originating in FVB/N, 129S and C57BL/6NCrl strains.

2.1.2 Pten and Fgfr2 deletions

Dr. Ruth Ashery-Padan at Tel Aviv University kindly provided the transgenic mice expressing Cre recombinase in the ocular surface ectoderm at E 9.5 (LeCre) (Ashery Padan et al. 2000). LeCre utilizes the PAX6 P0 promoter to drive Cre expression in all surface ectoderm derived ocular structures, including the lens, corneal epithelium, lacrimal gland, periocular epidermis, and the pancreas (Ashery-Padan et al. 2000). LeCre mice were used to conditionally delete Fgfr2, Pten or the combination of the two. Mice engineered with loxP sites flanking exons 4 and 5 of Pten were obtained from Dr. Gustavo Leone and Dr. Michael C. Ostrowski, Molecular Biology and Cancer Genetics Program, The Ohio State University, Columbus, OH and were previously described (Trimboli et al. 2009). Mice engineered with loxP sites flanking exon 8, 9, and 10 were obtained from Dr. David M. Ornitz from the Department of Molecular Biology and Pharmacology, Washington University Medical, and previously described (Yu et al. 2003).

2.3 HISTOLOGY AND IMMUNOHISTOCHEMISTRY

Gestational age of experimental embryos was determined by vaginal plug detection, set at embryonic day 0.5 (E0.5). One hour prior to embryo collection pregnant dams were administered (0.1 mg/g body weight) 5-bromo-2'-deoxyuridine (BrdU) dissolved in phosphate buffered saline (PBS) at a concentration of 100 mg/ml.

2.3.1 Frozen Sections

Lenses were fixed in 4% neutral buffered paraformaldehyde for 90 minutes at 4°C, embedded in OCT, frozen and sectioned at 10 µm thickness. Cryosections were permeablized in 0.05% Triton X-100/PBS for 2 minutes, blocked in 5% donkey serum and 5% BSA in PBS for 30 minutes at

48 room temperature before being incubated with DLAD, phosphorylated lamin A/C, or PDI antibodies. The primary antibody for DLAD was generated as previously described (Nakahara et al., 2007) and used at a dilution of 1: 500 overnight at 4oC. The secondary antibody used for visualization of DLAD was 1: 250 diluted conjugated goat-anti hamster antibodies (Jackson ImmunoResearch Labs, 127-035-160). Detection of DLAD was via DAB peroxidase substrate kit (Vector labs, SK4100). Images were collected using an Olympus light microscope BX51. The primary antibody for phosphorylated lamic AC (pLaminA/C) was obtained from Abcam (ab58528). The primary antibody raised against PDI was obtained from Millipore (16-189). The primary antibody of DLAD was used at a 1:50 dilution. The primary antibodies for PDI and pLaminA/C were used at a 1:100 dilution. The secondary antibody used to detect the pLaminA/C and PDI were attached to fluorescent probes (Alexafluor 488 anti-rabbit).

2.3.2 Paraffin Sections

For paraffin sections, embryos were collected and fixed in 10% neutral buffered formalin (NBF). The embryos were collected at E10.5, E12.5, E15.5, E17.5, E18.5 and P0. The embryos obtained at E10.5 and E12.5 were fixed for 4 hours. The embryos collected at E15.5-P0 were fixed for 12- 16 hours. The fixation process took place at room temperature. Two washes of 1X Phosphate Buffered Saline followed the fixation.

2.3.2a Tissue Processing

For paraffin infiltration, the tissue was placed in a cassette and subjected to the below washes for 15 minutes (E10.5-E12.5), 30 minutes (E15.5-E17.5), or 1 hour (E18.5-P0).

1. 70% EtOH 2. 80% EtOH 3. 95% EtOH 4. 3 x 100% 5. 3 x Xylenes 6. 2 x Paraffin

Following these sequential washes and paraffin infiltration, the tissue was embedded in paraffin blocks. The tissue was section at 5 µm thickness for further analysis by immunohistochemistry or hematoxylin and eosin staining

2.3.2b Immunohistochemistry Protocol

1. DePariffinization 2 X 5 minutes in xylene

2. Dehydrate 2 X 5 minutes 100% EtOH 1 minute 95% EtOH 1 minute 70% EtOH

3. Rehydrate

1 minute 50% EtOH Pour out half of the 50% and replace with ddH20. 1 minute per wash. Continue until the slides are in 100% ddH20. 2, 1 minute washes in solely ddH20

4. Antigen Retrieval

.01 M Sodium Citrate Buffer put slides in slide holders, and in the rice cooker. Get the water to boiling, and once its on boiling I usually just put the cooker on warm and it’ll stay just barely below boiling (so the slides don’t tip over but remains around 100 degrees C)

30 minutes in the rice cooker. 10 minutes out of rice cooker -cool to room temp.

5. Nuclear Permeability

Wash in either 1% PBS + Saponin or 2N HCl in PBS for 5-10 minutes.

3 X 5 minutes in PBS

6. Blocking

Use a hydrophobic pen and surround the tissue with the hydrophobic ink.

50 Put 50-100 uL of the 10% Normal Horse Serum (NHS) or Normal Goat Serum (NGS) in 1% PBST on the tissue-enclosed by the hydrophobic ink.

Place slides in an environmental chamber container in 37 degree C incubator for 1 hr.

7. Primary Antibody Incubation

Replace the blocking solution with the primary antibody solution (10% NHS in 1%PBST and anywhere from a 1:50-1:250 antibody dilution dependent upon the antibody). Incubate the tissue with the primary antibody for8-12 hours (overnight) at 4 degrees C or 4-6 hours at room temperature.

8. Secondary Antibody Incubation

Wash off the primary antibody (3 x5 minutes in PBS)

While your washing you should make up 10% NHS 1%PBST and 1:100 dilution of secondary body which should be the anti species (whichever species you used putting on your primary)

e*Note - th secondary antibody should always be handled in minimal light.

Put secondary on for 2 hrs at room temp.

Wash-off the secondary antibody (3 x 5 minutes-limited lighting)

Place a drop of vectashield+dapi on tissue, and cover slides with a coverslip.

2.3.2c Antibodies used for Immunohistochemistry

Primary antibodies for phosphorylated NuMA (at Threonine 2055) were previously described (Kotak et al., 2013). Primary antibodies for Aquaporin0, BrdU, p57KIP2, and CDK1 (ab15077, ab6326, ab4058, and ab7953, respectively) were obtained from Abcam. The primary antibody for p27KIP1 (BD610241) was obtained from BD Biosciences. Primary antibodies for Tom20 (sc- 11415) and total NuMA (sc-48773) were obtained from Santa Cruz Biotechnology, while the antibody for phosphorylated histone H3 (Ser10) was obtained from Millipore (16-189). The primary antibodies for β and γ-crystallin were kind gifts from Samuel Zigler at Johns Hopkins University School of Medicine. The primary antibodies for phosphorylated AKT, and

51 phosphorylated ERK1/2 were purchased from Cell Signaling Technolgies (9101, 9271 respectively). All primary and secondary antibodies were used at a 1:100 dilution, with the exception of total NuMA and phosphorylated histone H3 which were used at a 1:50 dilution and β and γ-crystallin which were used at 1:250 dilution. Primary antibodies were detected using secondary antibodies attached to fluorescent probes (Alexafluor 488 goat anti-rabbit IgG, Alexafluor 546 goat anti-rat IgG, FITC for donkey anti-rabbit IgG, 711-095-152 and Cy3 for donkey anti-mouse IgG). Cells undergoing DNA degradation/apoptosis was detected using the In Situ Cell Death Detection Kit (TMR Red, Roche AppliedScience, 2156792). Sections were counterstained with DAPI (Vector Laboratories, H-1200). Photomicrographs were captured on a Zeiss 710 Laser Scanning Confocal System at the Center for Advanced Microscopy and Imaging at Miami University. Standard hematoxylin and eosin stained sections were used to analyze the structure of the lens, and images were captured using a Nikon TI-80 microscope.

2.4 Immunofluorescence Quantification

Quantifying indirect immunoflourescent labeling on tissue sections was previously described (Garcia et al., 2011; Plageman et al., 2011; Madakashira et al., 2012). All immunoflourescent assays were photographed with identical exposure times. IMAGEJ software was used to measure the signal intensity of the pixels (RGB) and given areas. IMAGEJ software was used to select the plot to be measured. The BrdU, and TUNEL index represented the ratio of the lens cell nuclei positive for the mentioned markers over the total DAPI stained nuclei in the ocular lens.

2.5 Whole Mount Epithelial Cell Z-Stacks

Lenses from MLR10; Cdk1L/L and Cdk1L/L mice at embryonic day 17.5 were immediately fixed in 10% NBF for one hour. After fixation, the lenses were washed in PBS, and stained with DAPI. The lenses were then placed in between two coverslips with a drop of PBS and a series of images were collected at varying depths using the Zeiss 710 Laser Scanning Confocal System, and finally reconstructed into a three-dimensional image. Cross sectional area of each individual nucleus was determined using IMAGEPRO software at the Center for Advanced Microscopy and Imaging at Miami University.

2.6 WESTERN BLOT

2.6.1 CDK1 manuscript- Western blots were completed at Tufts University

Cdk1L/L, MLR39; Cdk1L/L, and MLR10; Cdk1L/L lenses were taken at birth. The epithelial cell layer and fiber cell mass were physically separated in Cdk1L/L, MLR39; Cdk1L/L lenses and homogenized in RIPA buffer (50mM Tris-HCl pH 8.0, 150 mM NaCl, 1% NP40, 0.5% Sodium deoxycholate, 0.1% SDS) with phosphatase and protease inhibitors (Pierce, Rockford, IL, Cat No: 78440). The protein concentration was determined by BCA assay (Pierce, Rockford, IL, Cat No: 23227). Protein lysates were separated on a 10% SDS-polyacrylamide gels and transferred to PVDF membranes (Millipore, Billerica, MA, Cat No: IPVH10100), blocked with 5% non-fat dry milk for one hour at room temperature and incubated overnight at 4°C with antibodies to CDK1 (1:2000, Abcam, ab7953), or pNuMA (1:1000). After incubation with HRP- conjugated secondary antibody (1:1000) for 2 hours (Cell Signaling Technology, 7074), the proteins were analyzed on X-ray films following the addition of the chemiluminescent substrate Lumiglo (Cell Signaling Technology, 7003).

2.6.2 PTEN/FGFR2 manuscript-western blots were completed at Miami University

Lenses were dissected from embryos and homogenized in RIPA buffer (50mM Tris HCl pH 8, 150nM NaCl, 1% NP40, 0.5% sodium deoxycholate) with protease inhibitor (Sigma). Proteins were analyzed on 12% Tris-glycine gels and transferred to PVDF membrane. Membranes were incubated with PBST with 3% BSA, probed with primary antibodies overnight at 40C and then incubated with secondary antibody for 2 hours at room temperature. The expression of GAPDH was used as the loading control. Western blot images were acquired; and ImageJ quantified the intensity of immunoreactivity. 2.6.2a Reagents

Acrylamide-Bis Stock (30-.8%) (we have commercial from BIORAD Cat#161-0156) 100mL Acylamide 30.0g Bis 0.8g

53 75% Sucrose

75g Sucrose, Bring to 100mL final Volume

5% Ammonium Persulfate (APS)

0.5g Ammonium persulfate, bring to 10mL final volume in water

4X Upper Gel Buffer 100mL 200mL 400mL Tris Base 6.06g 12.12g 24.24g SDS .4g .8g 1.6g *Bring to 80% FV with water, adjust pH to 6.8 at 20-21 C with HCL and add water to final volume (FV)

4X Upper Gel Buffer (100mL)-no need to manually calculate pH. 100mL 200mL 400mL Tris Base 0.3g 0.6g 1.2g Tris-HCl 7.5g 15g 30g SDS 0.4 0.8g 1.6

8X Lower Gel Buffer-Adjust pH to 8.8 at 20-21 C with HCL bring to FV 100mL 400mL Tris Base 36.34 145.36g SDS 0.8g 3.2g

8X Lower Gel Buffer (pH 8.8), 100mL –no need to manually pH 100mL 400mL Tris Base 29.7g 118.8g Tris-HCl 10.98g 43.92 SDS .8g 3.2g *bring to 100mL final volume with water…pH will be 8.8

7.5% SDS Page # of Gels 2 3 4 Total Volume (mL) 10.00 15.00 18.00 8x lower buffer 1.25 1.88 2.25 (mL)

30% 1.88 2.81 3.38 Acrylamide/Bis Solution 29:1 (mL) 75% Sucrose (mL) 1.33 2.00 2.40 Water (mL) 5.54 8.31 9.98 5% Ammonium 50.00 75.00 90.00 Persulfate (µL) TEMED (µL) 6 8 10.8

10%SDS PAGE # of Gels 2 3 4 Total Volume (mL) 10.00 15.00 18.00 8x lower buffer 1.25 1.88 2.25 (mL) 30% 2.50 3.75 4.50 Acrylamide/Bis Solution 29:1 (mL) 75% Sucrose (mL) 1.33 2.00 2.40 Water (mL) 4.92 7.38 8.85 5% Ammonium 50.00 75.00 90.00 Persulfate (µL) TEMED (µL) 6 8 10.8

12.5% SDS PAGE # of Gels 2 3 4 Total Volume (mL) 10.00 15.00 18.00 8x lower buffer 1.25 1.88 2.25 (mL) 30% 3.13 4.69 5.63 Acrylamide/Bis Solution 29:1 (mL) 75% Sucrose (mL) 1.33 2.00 2.40 Water (mL) 4.29 6.44 7.73 5% Ammonium 50.00 75.00 90.00 Persulfate (µL) TEMED (µL) 6 8 10.8

15% SDS PAGE # of Gels 2 3 4 Total Volume (mL) 10.00 15.00 18.00

8x lower buffer 1.25 1.88 2.25 (mL) 30% 5.00 7.5 9 Acrylamide/Bis Solution 29:1 (mL) 75% Sucrose (mL) 1.33 2.00 2.40 Water (mL) 2.40 3.6 4.3 5% Ammonium 50.00 75.00 90.00 Persulfate (µL) TEMED (µL) 6 8 10.8

4.5% Stacking Gel # of Gels 3 (fast) 3 6 Total Volume (mL) 5.00 5 10 4x Upper Buffer 1.25 1.25 2.50 (mL) Acrylimide .56 .56 1.13 Water (mL) 3.19 3.19 6.38 5% Ammonium 45 30 60 Persulfate (µL) TEMED (µL) 22 15 30

10X Electrode Buffer Stock for SDS-Page (AKA Running Buffer)

1 Liter 4 Liter Tris Base 30.3g 121.2g Glycine 144.1g 576.4g SDS 10.0g 40.0g

10X Transfer Buffer for Western Blots (1L)

Tris Base Aka Tris 30.385g Glycine 144.134g Water to 100mL FV

* Note: 1x transfer buffer contains 20% methanol. *Additional Note: if you just make the 10X transfer buffer stock. Then you can add SDS (to make Running Buffer) or 20% methanol (to complete the transfer buffer)

56 2.6.2b Western Protocol

1. Protein Isolation.

a) Pool 6 lenses in 200 µL of RIPA buffer. RIPA buffer ordered from Santa Cruz (sc-24948) b) Provide constant agitation for 30 minutes at 4 degrees C (sonicate if needed) c) Centrifuge samples (16,000xg) for 20 min at 4 degrees C d) Remove centrifuge tube and place it on ice. e) Transfer Supernatant to a fresh tube and discard the pellet.

2. Protein Concentration bicinchoninic acid assay (BCA)

3. Acrylamide Gel a) Make 9mL of SDS Page Gel per gel (see reagent list-this is an oxygen free reaction) i. Stacking gel is placed on top (roughly 2 mL) ii. Wait for an Hour for the gel to solidify

4. Protein Preparation a) Use a 1:1 dilution of Laemmli sample buffer to sample. Optimal amount of lens protein is roughly 15 µg of protein i. Add β-mercaptoethanol to Laemmli Buffer before use b) Boil each sample at 95 degrees C for 5 min c) Centrifuge at 16,000 x g for 1 min.

5. Load samples- a. Make Running Buffer with 10x transfer buffer and SDS (100mL of 10x transfer buffer, 5mL of SDS) b. Fill the middle with the Running buffer (to top)-make sure no leaks. c. Ladder in lane 1-try to keep ladder at same volume as rest of samples i. So the ladder we have- use 6 µL of of Ladder, with Laemmli and Ringers at 1:1. Essentially act like the ladder is your protein and treat everything the same. ii. Use long pipets-try not to generate any “bubbles” while pipetting iii. Run for 20 mins at 90V. 1-1.5 hrs at 150V

6. Transfer protein to membrane a. Use a PVDF or Nitrocellulose membrane b. Activate membrane in methanol. Make a cut on a corner in order to keep tract of membrane orientation. c. Cut filter paper to go over membrane. d. Place stacking equipment and filter paper in transfer buffer (700mL of Water, 100mL of 10x transfer buffer, 200mL methanol).

57 e. Place a magnetic spinner on bottom of the transfer apparatus, and an ice pack in the back. f. Cut out the wells of SDS page gel. g. Put together the stacking “sandwich”-placing the black side of the “sandwich” on the bottom, followed by stacking foam, filter paper, SDS page gel, membrane, filter paper, stacking foam, and finallywhite part of stacking equipment. h. Put in Cold room on magnetic spinner, set at 280V (1.5 hrs) or 20V (Overnight)

7. Block

a. Remove membrane from the stacking equipment with forceps b. (optional) use Ponceau to make sure protein transferred evenly (5-10min) c. Use either 5% nonfat dry milk or 3% BSA (phosphoproteins)

8. Antibody Incubation

Primary Antibody- incubate primary antibody overnight at 4 degrees C)-usually around a 1:2000 dilution (antibody:5%BSA in PBST or 5% nonfat milk in PBST)

Wash in PBST

Secondary Antibody- Incubate Secondary antibody at room temp (5% BSA in PBST or 5% nonfat milk in PBST)

Wash in PBST

9. Chemiluminescent

a) Obtain Chemiluminescent Substrate (BioRad) b) Obtain AutoRadiography Film (if box is open make sure ALL lights are off) c) Go to Room 79 in the basement d) Turn on machine (needs 10 minutes to power up) e) Make 1 mL of chemilumiescent (BioRad substrate by mixing the substrate at 1:1 conc. f) Tape down membrane g) Put substrate on membrane h) Make sure lights are off, and take out a sheet of autoradiography film. Place Autorad film on membrane for anywhere between 5 seconds and 5 minutes. Take autorad film off membrane with forceps (quickly) and put in machine.

2.7 QUANTITATIVE RT-qPCR

The expression levels of selected genes were analyzed by RT-qPCR. Total RNA was extracted

58 from the lenses and used to synthesize cDNA by reverse transcription using random primers and the superscript II reverse transcriptase (Invitrogen), according to the manufacturer’s instructions. qPCR assays were performed on the cDNA using Gotaq Green Master Mix (Promega) following the manufacturer’s instruction and read using CFX connect (BioRad). Intron-spanning primers were designed to specifically quantify targeted mRNA transcripts (Table 1). Each biological sample was analyzed in triplicate by qPCR. The expression of GAPDH was used as an endogenous control. The cycling conditions consisted of 1 cycle at 95° C for 100s for denaturation, followed by 40 three-step cycles for amplification (each cycle consisted of 95° C incubation for 20s, an appropriate annealing temperature for 10s, and product elongation at 70° C incubation for 20s). The melting curve cycle was generated after PCR amplification. The reaction specificity was monitored by determination of the product melting temperature, and by checking for the presence of a single DNA band on agarose gels from the RT-qPCR products.

PRIMER LIST

59 CHAPTER 3

NUCLEAR REMOVAL DURING TERMINAL LENS FIBER CELL DIFFERENTIATION REQUIRES CDK1 ACTIVITY: APPROPRIATING MITOSIS-RELATED NUCLEAR DISASSEMBLY Published by: Chaffee, B.R., Shang, F., Chang, M.L., Clement, T.M., Eddy, E.M., Wagner, B.D., Nakahara, M., Nagata, S., Robinson, M.L., Taylor, A., 2014. Development 141, 3388-3398.

3.1 SUMMARY

Lens epithelial cells and early lens fiber cells contain the typical complement of intracellular organelles. However, as lens fiber cells mature they must destroy their organelles, including nuclei, in a process that has remained enigmatic for over a century, but which is crucial for the formation of the organelle-free zone in the center of the lens that assures clarity and functionto transmit light. Nuclear degradation in lens fiber cells requires the nuclease DNase IIβ (DLAD) but the mechanism by which DLAD gains access to nuclear DNA remains unknown. In eukaryotic cells, cyclin-dependent kinase 1 (CDK1), in combination with either activator cyclins A or B, stimulates mitotic entry, in part, by phosphorylating the nuclear lamin proteins leading to the disassembly of the nuclear lamina and subsequent nuclear envelope breakdown. Although most post-mitotic cells lack CDK1 and cyclins, lens fiber cells maintain these proteins. Here,we show that loss ofCDK1 from the lens inhibited the phosphorylation of nuclear lamins A and C, prevented the entry of DLAD into the nucleus, and resulted in abnormal retention of nuclei. In the presence of CDK1, a single focus of the phosphonuclear mitotic apparatus is observed, but it is not focused in CDK1-deficient lenses. CDK1 deficiency inhibited mitosis, but did not prevent DNA replication, resulting in an overall reduction of lens epithelial cells, with the remaining cells possessing an abnormally large nucleus. These observations suggest thatCDK1-dependent phosphorylations required for the initiation of nuclear membrane disassembly during mitosis are adapted for removal of nuclei during fiber cell differentiation.

60 3.2 INTRODUCTION

The ocular lens and cornea are the only clear tissues in the body. Opacification of the normally clear lens(called cataract), afflicts 80% of the elderly population and remains the most common cause of blindness worldwide, afflicting over 19,000,000 people (WHO, 1998). As such, cataracts contribute significantly to the $139 billion spent annually in the USA alone due to compromised vision (Preventblindness.org, 2013). Mammalian lenses develop from a surface ectoderm-derived vesicle. The anterior cells of the vesicle differentiate into lens epithelial cells while the cells comprising the posterior half of the vesicle differentiate into primary fiber cells. A single layer of epithelial cells lines the anterior hemisphere of the lens. Although, initially, all lens epithelial cells proliferate, only a band of epithelial cells slightly anterior to the lens equator undergo cell division in the mature lens. Posterior to this zone, the epithelial cells begin to differentiate to form secondary fiber cells. These fibers elongate and eventually form the bulk of lens tissue. The lens continues to grow throughout life such that the original fibers, or oldest cells, occupy the lens center with progressively younger fiber cells found closer to the lens surface. Proper differentiation of fiber cells also involves formation of an organelle free zone (OFZ) comprised of fiber cells from which light scattering intracellular organelles, including the cell nucleus, are removed (reviewed in Wride, 2011). Failure to form the OFZ results in a cataractous lens. Although denucleation was observed more than a century ago, the molecular mechanisms leading to lens fiber cell denucleation remain poorly understood (Rabl, 1899). The lysosomal DNAse, DNAse IIβ (DLAD), is essential for breaking down lens fiber cell DNA and establishing an OFZ and clear lens (Nishimoto et al., 2003; De Maria and Bassnett, 2007; Nakahara et al., 2007). Lysosomal/cytoplasmic DLAD gains access to and destroys chromatin DNA upon disassembly of the nuclear membrane (Bassnett and Mataic, 1997). This is reminiscent of mitosis. In proliferating cells, mitosis requires cyclin dependent kinase 1 (CDK1) in conjunction with cyclins A or B to phosphorylate nuclear membrane lamins to destabilize the nuclear envelope (reviewed in Nigg, 1993). Activated CDK1 also aids in regulating mitotic chromatin (reviewed in Nigg, 1993; Kotak et al.; Orthwein et al., 2014; Zheng et al., 2014). In contrast, post-mitotic cells rarely exhibit CDK1 and cyclin A/B expression (King et al., 1994;

61 Tommasi and Pfeifer, 1995). However, post-mitotic lens fiber cells contain both CDK1 and cyclin B protein (He et al., 1998), suggesting that these proteins might initiate nuclear envelope disassembly to provide access for DLAD during terminal differentiation. CDK inhibitors, p57KIP2 and p27KIP1 also regulate CDK1 and the G1/S transition- regulating kinase, CDK4 (Sherr and Roberts, 1999). Increased synthesis of p57KIP2 and p27 KIP1 characterizes the withdrawal from cell cycle and initiation of lens fiber cell differentiation (Zhang et al., 1997; Zhang et al., 1998; Lovicu and McAvoy, 1999; Nagahama et al., 2001; Reza et al., 2007). As differentiation progresses, fiber cells continue to elaborate crystallins, the major fiber cell gene products. Several lines of evidence suggest that a ubiquitin proteasome system participates in nuclear breakdown. First, poly-ubiquitinated conjugates increase in equatorial epithelial cell nuclei just prior to fiber cell differentiation, and localize to differentiating fiber cell nuclei (Shang et al., 1999). Second, zebrafish containing a mutation in the 26S proteasome gene Psmd6 experience abnormal retention of fiber cell nuclei as well as a number of cell cycle alterations in the lens epithelium (Imai et al., 2010). Proteosomal degradation of cyclins A and B during mitosis inactivates CDK1 facilitating reformation of the nuclear membrane in daughter cells subsequent to karyokinesis. Cyclins A and B re-accumulate during the G2 phase of the cell cycle to activate CDK1 in preparation for the next mitosis. However, transgenic mice expressing a mutated ubiquitin (K6W-Ubiquitin) in the lens fiber cells accumulated p27KIP1, decreased phosphorylation of nuclear lamins A and C, retained nuclei within the usual OFZ, delayed synthesis of crystallins, and were cataractous (Caceres et al., 2010). Together, these data suggested that CDK1 is prominent in directing lens fiber cell denucleation. Here, we tested the hypothesis that, as in mitotic cells, the disassembly of the nuclear envelope in terminally differentiating fiber cells requires CDK1. We suggest that in contrast with cycling cells, where CDK activators and inhibitors are cyclically regulated, in lens fibers there is a unidirectional pathway in which high levels of p57KIP2 and p27KIP1 keep CDK1 inactive in immature fiber cells. However, prior to the formation of the OFZ, diminishing levels of these CDK inhibitors lead to CDK1 activation, lamin phosphorylation, disassembly of the nuclear membrane, entry of DLAD, reorganization of chromatin and destruction of the nucleus. Deletion of Cdk1 from the lens lineage facilitated the testing of this hypothesis.

62 3.3 RESULTS

3.3.1 CDK1 protein expression in epithelial cells and differentiating lens fibers.

Although previous studies documented the presence of CDK1 protein in lens fiber cells (He et al., 1998), the subcellular localization of CDK1 remained unknown. As expected, the lens epithelium expressed abundant CDK1 and much of the enzyme appeared to be localized to nuclei in epithelial and outer cortical fiber cells (Figure 3.1, zones 1, 2). In secondary lens fiber cells, the overall level of CDK1 expression declined as development advanced (compare right to left side, lower panels). While CDK1 was obvious in both the cytoplasm and nuclei of elongating cells (Fig. 3.1, zone 2), it remained most concentrated in the nuclei of the deeper fiber cells (Fig. 3.1, zones 3 and 4).

3.3.2 Removal of CDK1 from the lens.

Transgenic mice homozygous for the loxP-flanked (floxed) allele of Cdk1 and hemizygous for the MLR10 Cre transgene (MLR10; Cdk1L/L) were created to remove CDK1 from the lens. Cre expression in MLR10 transgenic mice initiates at E10.5 and this transgene can effectively delete loxP-flanked alleles in both lens epithelial cells and lens fiber cells (Zhao et al., 2004). In the MLR10; Cdk1L/L lenses, the overall expression of CDK1 protein became mosaic by E15.5 (Fig. 3.2, D-F) with few CDK1 positive epithelial cells remaining by E17.5 (Fig. 3.3 B, D, white arrows). By comparison with expression in lens, MLR10; Cdk1L/L retinas displayed no alterations in CDK1 expression relative to control littermates (Fig 3.2 F, Fig. 3.3 E), indicating that the Cre transgene properly targeted the lens without affecting other tissues within the optic cup. CDK1 protein persisted in postnatal epithelial cells and fiber cells from both control lenses. Western blots corroborated the diminution of CDK1 at E18.5 in MLR10; Cdk1L/L lenses (Fig. 3.3 F). The remaining protein indicates that some epithelial cells escape Cre-mediated deletion or, alternatively, represents persistent CDK1 protein produced from transcripts that existed prior to the deletion of Cdk1.

63 To isolate Cdk1 deletion to lens fiber cells, MLR39 transgenic mice were bred to Cdk1L/L animals to generate MLR39; Cdk1L/L mice. Cre expression in MLR39 mice initiates at embryonic day 12.5 (E12.5), and within the lens, remains exclusively in the fiber cell compartment (Zhao et al., 2004). Immunohistochemical and western blot comparisons between the lenses from MLR39; Cdk1L/L mice and those of Cre negative control littermates (Cdk1L/L) failed to reveal significant diminution of CDK1 in fiber cells from the P0 MLR39; Cdk1L/L mice (Fig. 3.2, compare B to A, C, G). Also, MLR39; Cdk1L/L lenses remained clear and appeared histologically identical to control lenses (data not shown). The persistence of CDK1 protein in the lens fibers indicated that the MLR39 transgene failed to delete the Cdk1 gene early enough to significantly reduce CDK1 protein in the fiber cell compartment. Therefore, all subsequent analyses employed MLR10; Cdk1L/L mice.

3.3.3 Loss of CDK1 delays denucleation of lens fiber cells.

The gross morphology of MLR10; Cdk1L/L lenses appeared similar to control lenses at E12.5, prior to fiber cell denucleation, (Fig. 3.4 A, B). However, by E15.5, the nuclei in the differentiating secondary fibers at the bow region of MLR10; Cdk1L/L lenses appeared 152% larger by cross sectional area, relative to those of control littermates (Fig. 3.4, compare the size of nuclei in the dashed ovals in D to C). Mouse fiber cells normally begin losing their nuclei at approximately E16-E18 (Kuwabara and Imaizumi, 1974). Consistent with this observation, lenses with intact Cdk1 excluded nuclei from central fibers from E17.5 onward (Fig. 3.4 E, dashed oval, E’ zone 6). In contrast, E17.5 MLR10; Cdk1L/L lenses retained nuclei in the center of the lens resulting in a failure to form an OFZ (Fig. 3.4 F, F’ see persistent nuclei in zone 6). Bassnett and others documented descriptive criteria for nuclei during the lens fiber cell differentiation process. We used these criteria to distinguish the nuclei in E17.5 MLR10; Cdk1L/L and control lenses (Bassnett, 2009). The youngest secondary fiber cells, near the lens equator in zone 1, possessed densely packed elongated oval nuclei in control lenses (Fig. 3.4 E, E’ zone 1). In zone 1 of the MLR10; Cdk1L/L lens, the fiber cell nuclei were also oval, but fewer in number and their average cross sectional area was 172% larger than those of control lenses (Fig. 3.4 compare zone 1 in F’

64 to E’). From E17.5 on, distinctly fewer cells appeared to undergo secondary fiber cell differentiation in the MLR10; Cdk1L/L lenses. In zone 2, the nuclei in the fibers of control lenses were more sparse and less elongated than in zone 1 (Fig. 3.4 E’ compares zone 1 and 2). CDK1-deficient lenses also contained relatively fewer nuclei in zone 2 but they appeared considerably larger than the comparable region in the control lens (Fig. 3.4, F’, E’ compare zones 2) and virtually unchanged in size from zone 1 (Fig. 3.4, F’ compare zones 1 and 2). In zone 3, the control fiber nuclei assumed a smaller, more rounded shape (compared to zones 1 and 2) and also stained more darkly with hematoxylin (suggestive of nuclear condensation). The CDK1-deficient fibers of zone 3 contained a mixture of large, elongated, more rounded nuclei appearing similar, in size and shape, to the zone 3 nuclei of control fiber cells (Fig. 3.4 F’, E’ compare zones 3). Sparsely packed spherical nuclei, similar in size to those in zone 3, persisted in the control lenses in zones 4 and 5 (Fig. 3.4 E’ zone 4- 5). The MLR10; Cdk1L/L fiber nuclei appeared fewer in number with more variable size in zones 4 and 5 (compared to the control) and this pattern persisted into zone 6 (Fig. 3.4 F’). In contrast, zone 6 of the control lens was devoid of nuclei (Fig. 3.4 E’, zone 6).

3.3.4 Loss of CDK1 prevents entry of DLAD into the nucleus of terminally differentiating lens fiber cells.

The entry of DLAD into the nuclear compartment requires the disassembly of the nuclear membrane which normally occurs as fiber cells approach the OFZ. By analogy to mitotic events, we postulated that the nuclear membrane might remain intact in fiber cells lacking CDK1, thus preventing the entry of DLAD. CDK activity is controlled by cyclin dependent kinase inhibitors. As lens epithelial cells begin to differentiate, they express p27KIP1 and p57KIP2 , and both of these CDK inhibitors cooperate in the initiation of cell cycle withdrawal during embryonic fiber cell differentiation (Zhang et al., 1998; Lovicu and McAvoy, 1999; Nagahama et al., 2001; Kase et al., 2005). In keeping with these reports, both control and MLR10; Cdk1L/L lens epithelial cells express p57KIP2 prior to reaching the lens equator, however, expression in both genotypes abruptly ends as the cells begin to detach from the lens capsule (Fig. 3.5 compare bracketed region in A and D). In comparison, the expression of p27KIP1 extends further into the fiber cell

65 mass than p57KIP2 in control lenses (Fig. 3.5 B, bracketed region), and becomes undetectable just prior to nuclear degradation in the OFZ. Additional coordinated events that occur prior to the formation of the OFZ in control lenses include the nuclear concentration of CDK1 (Fig. 3.1 zones 3, 4), the adoption of a spherical nuclear morphology (Fig. 3.4 E’ zones 4 and 5), the disappearance of p27KIP1 (Fig. 3.5 B), and the phosphorylation of nuclear lamins A and C (Fig. 3.5 C). In CDK1-deficient lenses, p57KIP2 and p27KIP1 are also observed in the outer regions but p27KIP1 expression is below detection limits in the inner fiber cell mass (Fig. 3.5 compare E to B). Also, CDK1-deficient nuclei fail to exhibit phosphorylation of Lamin A/C (Fig. 3.5 compare F to C). Concurrent with the onset of phosphorylation of lamin A/C in control lenses, DLAD moves from the periphery of the nucleus to within the nucleus (Fig. 3.5 H). As indicated in the top inset panel, there is little DLAD evident in early nucleated fiber cells. In contrast, as indicated in the bottom inset panel, uniform DLAD staining is observed throughout the nuclei just prior to the formation of the OFZ. The entrance of DLAD in control lenses anticipates the DNA degradation as shown by abundant TUNEL positive foci in the developing OFZ (Fig. 3.5 I, yellow staining, arrows). Although DLAD generates 3’-phosphoryl/5’-hydroxyl ends following endonucleic cleavage of DNA (Shiokawa and Tanuma, 1999), endogenous phosphatases rapidly

- convert the 3’-PO4 ends to 3’-OH ends that can be labeled by the TUNEL assay (De Maria and Bassnett, 2007). In contrast, in MLR10; Cdk1L/L lenses DLAD accumulated around the periphery of central fiber cell nuclei, rather than entering the nucleus as seen in the control lenses (Fig. 3.5, compare the lower inset of K with the lower inset of H), and fewer fiber cell nuclei demonstrate degradation of DNA (Fig. 3.5 compare yellow nuclei in L to I).

3.3.5 Relocalization of NuMA during fiber cell differentaition and maturation.

The nuclear mitotic apparatus protein (NuMA) is mechanistically involved in chromosome segregation that preceeds nucelar disassembly and mitosis (Gribbon et al., 2002; Abad et al., 2007; Kotak and Gonczy, 2014). During the metaphase-anaphase transition CDK1- induced phosphorylation on threonine 2055 (T2055) results in movement of NuMA from the cell membrane to the spindle poles, resulting in chromosome segregation (Kotak et al., 2013).

66 Western analysis indicated the presence of pNuMA in control lenses (Fig. 3.6 C). Immunohistochemical analyses confirmed the presence of pNuMA in control, CdkL/L, lenses (Fig. 3.6 A1-3). In these lenses pNuMA is observed distributed or as multiple foci throughout the entire nuclei of epithelial cells and early differentiating fiber cells (Fig. 3.6 A1). Consistent with a role for pNuMA in organizing chromatin, fewer prominent pNuMA puncta are observed in more differentiated fibers (Fig. 3.6 A2) of the control lens and this is echoed by the punctate pattern of chromatin staining (Fig. 3.4 E1-3, Fig. 3.6 A1-3, Fig. 3.7 B, specifically #2-4). Strikingly, however, rather than coalescing to a few or two prominent foci as they do in mitosis, in Cdk1L/L control mouse lenses they appear to coalesce to a single large focus in the denucleating cells (Fig. 3.6 A3). In comparison, pNuMA was present at considerably lower levels in western blots in MLR10; Cdk1L/L lenses and it was below the limit of immunofluorescent detection in the CDK1-deficient lens fiber cells (Fig. 3.6 B1-B3). Some pNuMA was observed in a few epithelial cells in these lenses (Fig. 3.6 B). Consistent with an absence of pNuMA foci in the MLR10; Cdk1L/L lens, the nuclei remain larger and the chromatin remains heterogeneously spread or less focused throughout the nucleus (Fig 3.7 compare C to A, D to B).

3.3.6 Reduction of phosphorylated histone H1 (pH1) in CDK1 deficient lenses

The linker histone H1 becomes phosphorylated by CDK1 to increase the attraction between histone H1 and the chromosome prior to mitosis. The phosphorylation of H1 has been used as a readout of CDK1 enzymatic activity (He et al. 1998). In control lenses, high levels of pH1 were detected in nuclei of cells at at the germinative zone of the lens epithelium, and in differentiating fiber cell nuclei (Fig. 3.8 A, B-white circle). CDK1 deficient lenses displayed pH1 in nuclei of the lens epithelium, yet was absent from the differentiating fiber cell nuclei (Fig. 3.8 C, D-white circle). The alteration in chromatin organization in CDK1-deficient lenses (Fig. 3.7 compare C to A D to B) may be either to loss of the phosphorylation of NuMA or Histone H1, or both.

67 3.3.7 Specificity of nuclear retention in CDK1 deficient lenses.

Despite retaining fiber cell nuclei, CDK1-deficient lenses removed mitochondria from central fiber cells as indicated by diminished Tom20 (an outer mitochondrial membrane protein) immunoreactivity in fiber cells (Fig. 3.9, asterisks in A, D). Some mitochondria remained in outer fibers (white arrows) but disappeared in the central fiber cells in both genotypes (Fig. 3.9, white arrows in A, D). Immunostaining for the endoplasmic reticulum marker, protein disulfide isomerase, PDI, also appeared similar between the MLR10;Cdk1L/L and the control mice (Fig. 3.9 C, F), and no endoplasmic reticulum is indicated in mature central fiber cells (Fig. 3.9 inside dotted border C, F). This contrasts with the prolonged expression of PDI in lenses from dominant negative Ncoa6 transgenic mice that also exhibit retention of nuclei in lens fiber cells (Wang et al., 2010). Thus, while mitochondria (Fig. 3.9 A, D) and endoplasmic reticulum (Fig. 3.9 C, F) were destroyed in both genotypes, only MLR10; Cdk1L/L lenses retained nuclei in central fiber cells (Fig. 3.9 E, yellow arrowheads), consistent with the hypothesis that CDK1 deficiency specifically inhibits denucleation rather than causing a generalized inhibition of organelle destruction.

3.3.8 Epithelial Cells in Cdk1 deficient lenses fail to undergo mitosis, and exhibit DNA endoreduplication.

Although the number and density of lens epithelial cell nuclei were similar in the control and MLR10; Cdk1L/L lenses at E12.5 (Fig. 3.10 E), by E15.5 there were fewer total epithelial cell nuclei, and fewer epithelial cell nuclei per unit area, in MLR10; Cdk1L/L lenses than in control lenses (Fig. 3.10 compare B to A, D to C, E). There was, however, no increase in epithelial cell apoptosis in CDK1 deficient lenses at E15.5 or E17.5 (Fig. 3.5 I, L; Fig. 3.12B-D). Therefore, decreased cell survival fails to explain the loss of epithelial cell population density in the MLR10; Cdk1L/L lens. Since CDK1 plays an essential role in cell cycle regulation, assays for BrdU incorporation (an S-phase indicator) and phosphorylated histone H3 (pH3) (a marker of late G2 phase, immediately prior to mitosis) were used to assess the cell cycle in MLR10; Cdk1L/L lenses. MLR10; Cdk1L/L lenses exhibited an S-phase index (proportion of BrdU positive nuclei) similar

68 to that of control lenses at both E12.5 and E15.5 (Fig. 3.11 compare B to A, solid bars in I). However, by E17.5 epithelial cells in the MLR10; Cdk1L/L lens displayed a significantly higher S- phase index, despite the decreased cell density of the epithelial cell layer. Likewise, starting at E15.5 and continuing to at least E17.5, a significantly higher percentage of lens epithelial cells in MLR10; Cdk1L/L mice exhibit pHH3 immunoreactivity than in age-matched control mice (Fig. 3.11 compare F to E, H to G, shaded bars in I). Since MLR10; Cdk1L/L lens epithelial cells exhibited both BrdU incorporation and pH3 expression, the cell cycle appeared to be active in these epithelial cells. However, the widespread expression of pH3 in MLR10; Cdk1L/L lens epithelial cells suggested that although the mutant lens nuclei prepare to leave G2 they may not actually enter mitosis. Consistent with this hypothesis we observe a 265% increase in nuclear cross sectional area in comparison to control lens epithelial cell nuclei (Fig. 3.11, compare K to J, L). The reduced number of lens epithelial cells coupled with the increased nuclear size suggests that CDK1 deficient lens epithelial cells bypass mitosis and simply undergo endoreduplication of their DNA during the cell cycle. This also explains the reduced number of, and larger, nuclei in the differentiating secondary fiber cells of MLR10; Cdk1L/L lenses.

3.4 DISCUSSION

In mammals, destruction or extrusion of nuclei occurs as a normal event during differentiation only in erythrocytes, keratinocytes and lens fiber cells. Of these, only lens fiber cells destroy nuclei within the cell. Despite many investigations over the last century, the molecular mechanism(s) by which fiber cell denucleation occurs has remained a mystery (Vrensen et al., 1991; He et al., 1998; Vrensen et al., 2004; Ivanov et al., 2005; Xie et al., 2007; Rivera et al., 2009; He et al., 2010; Wang et al., 2010; Gupta et al., 2011; Ma et al., 2011; Jarrin et al., 2012; Rodrigues et al., 2013). Preliminary work suggested that the lens “appropriates” normal mitotic mechanisms in order to accomplish denucleation; specifically that nuclear membrane disassembly occurs after phosphorylation of nuclear lamins and that stabilization of the CDK1 inhibitor p27KIP1 delays denucleation (Caceres et al., 2010). Furthermore, the persistence of CDK1 and its activator, cyclin B, and entry of DLAD, while p27KIP1 levels decline (He et al., 1998) (Fig. 3.5), as well as observations of delayed denucleation in DLAD -/- mice (Nishimoto et al., 2003) suggested that CDK1 directs fiber cell denucleation. In proof of

69 principle experiments, here we show that the phosphorylation of lamin, entry of DLAD in to the nuclear compartment, and denucleation per se requires CDK1, thus, elucidating upstream events leading toward lens fiber denucleation. Focalization of pNuMA appears to be a consequence of CDK1 activity. Since germline deletion of Cdk1 results in pre-implantation lethality (Santamaria et al., 2007), the MLR10 CRE transgene was exploited to remove Cdk1 specifically from the lens (Fig. 3.2, 3.3). Whereas normal fiber cells exhibited phosphorylation of the known CDK1 substrate, lamin A/C, in the region just prior to fiber cell denucleation, it remained unphosphorylated in CDK1 deficient fiber cells. Furthermore, in the absence of CDK1, DLAD remained outside the fiber cell nuclei (Fig. 3.5 K, lower inset) and these cells failed to denucleate (Fig. 3.4 D, F, F’). Taken together these findings indicate that fiber denucleation requires CDK1 activity. The specificity of the requirement of CDK1 for removal of nuclei per se is implied by observations that breakdown of the mitochondria and endoplasmic reticulum occurred on schedule in MLR10; Cdk1L/L lens fibers (Fig. 3.9). NuMA is a CDK1-dependent regulator of mitosis. Albeit elucidation of regulation of its function remains in progress, it is clear that during mitosis CDK1-dependent phosphorylation causes NuMA to concentrate at spindle poles and induce a redistribution of dynein that results in chromosome segragation and eventally division of the nucleus (Lelievre et al., 1998; Abad et al., 2007; Kotak et al., 2013; Zheng et al., 2014). Data in Fig. 3.6 indicate that NuMA is also be involved in the chromatin organization that preceeds lens fiber denucleation. (Fig. 3.6 A1-A3, Fig. 3.7). Interstingly, whereas in mitotic cells two foci are formed, in denucleating lens cells, only a single large pNuMA focus is observed in these WT mice. In contrast, NuMA T2055 remained largely unphophorylated in CDK1-deficient fiber cells, and chromatin was less consistently organized (Fig. 3.6 B1-B3, Fig. 3.7). Together the data suggest that lens fiber cells appropriate from normal mitosis critical functions for CDK1-driven phosphorylation of lamin and NuMA in directing lens cell denucleation and development. Whereas in mitosis CDK1 drives the coordinated dissassembly of the nuclear membrane and organization of chromatin to allow for formation of daughter cells, in lens fibers chromatin and nuclei are destroyed. In addition to directing phosphorylation and localization of NuMA CDK1 seems to regulate the levels of the native protein and this may be related to the levels of the pNuMA. The relationship between CDK1 and NuMA is consistent with a feedback mechanism. Elucidation of additional

70 steps in regulation of these events during lens development will be the topic of future investigations. Interestingly, MLR10; Cdk1L/L lenses contained fewer nuclei in both the epithelium (Fig. 3.10) and fiber cells (Fig. 3.4) than controls, despite exhibiting retention of fiber cell nuclei in the putative OFZ. The reduction in total fiber cells and nuclei in the MLR10; Cdk1L/L lens appears to result from a reduction in epithelial cells required to fuel continued secondary fiber cell differentiation (Fig. 3.4). Furthermore, the loss of lens epithelial cells in CDK1 deficient lenses occurred without detectable increases in apoptosis, suggesting that CDK1 is dispensable for lens cell survival (Fig. 3.5, Fig. 3.12). Also, in MLR10; Cdk1L/L lenses, there was no extensive migration of the lens posterior epithelium as observed in lens fibers in which DNA damage repair is compromised (Wang et al., 2010). To the extent to which apoptosis is a consequence of DNA damage, these data suggest CDK1-driven entry of DLAD into the nucleus, and initial disassembly of DNA, operate upstream of the requirements for retention of DNA integrity. Another remarkable feature of MLR10; Cdk1L/L lenses is the disparate size of primary versus secondary fiber cell nuclei (Fig. 3.4 D, F vs. C, E). Nuclei within inner fiber cells of both genotypes (including nuclei of what should be the OFZ in MLR10; Cdk1L/L) are of similar dimensions. (Fig. 3.4 compare zones 4-6 in F’ with zones 4-6 in E’). However, nuclei in outer fibers of CDK1-deficient lenses, though fewer in number, appear distinctly larger (Fig. 3.4 compare zones 1,2 in F’ with zones 1,2 in E’, Fig. 3.5 compare the p57KIP2-positive nuclei in D with those in A). We posit that this difference in the size of primary versus secondary fiber cell nuclei in MLR10; Cdk1L/L lenses results from a difference in the number of genomic duplications experienced by the two different populations of precursor cells, as well as poorer organization due to limited NuMA, as noted above. Since the deletion of the floxed Cdk1 allele commences at E10.5, the older, primary fiber cells would largely have been in the process of withdrawing from the cell cycle before the knock down of CDK1 was taking effect. In contrast, the future secondary fiber cells would still be epithelial cells at E10.5 and would likely go through one or more rounds of DNA synthesis before withdrawing from the cell cycle. The large secondary fiber cell nuclei in MLR10; Cdk1L/L lenses precisely match the phenotype expected if lens epithelial cells underwent endoreduplication of DNA without mitosis in the absence of CDK1 prior to differentiation. This is supported by the higher proliferation index of CDK1- deficient lens epithelial cells (Fig. 3.11). Furthermore, the higher proportion of MLR10; Cdk1L/L lens

71 epithelial cells in S-phase or G2 phase and that are enlarged (Figs. 3.11) is consistent with previous studies documenting the requirement of CDK1 for nuclear disassembly in mitosis and meiosis during development (Adhikari et al., 2012). Likewise, Cdk1 null pre-implantation mouse embryos that reach the blastocyst stage and mouse embryonic fibroblasts conditionally deleted for Cdk1 exhibit a reduced number of- but abnormally large- nuclei (Santamaria et al., 2007; Diril et al., 2012). In conclusion, this discovery of a requirement for CDK1 activity for a terminal differentiation pathway, including removal of nuclei and establishing an OFZ, expands the known functions for this protein beyond those for mitosis and meiosis. In the lens, CDK1 deficiency fails to induce apoptosis or prevent the onset of secondary fiber cell differentiation. The fundamental process of nuclear disassembly apparently requires lamin phosphorylation by CDK1 and includes NuMA-related chromatin organization. The finding that these processes can occur independently from cell division implies that CDK1 may play important roles in other aspects of nuclear function. There are several disease related laminopathies including Emery- Dreifuss muscular dystrophy (EDMD), dilated cardiomyopathy (DCM), limb-girdle muscular dystrophy, and Hutchinson-Gilford progeria syndrome, most of which profoundly affect non- proliferating cells (reviewed in Ho and Lammerding, 2012). This work suggests that CDK1, and perhaps other regulators of nuclear structure during mitosis, may play an unappreciated role in terminally differentiated cells.

72 CONTROL (Cdk1L/L) 1 CDK1 CDK1/DAPI epi 4 1 1 2

epi epi 3

fiber fiber

fiber 50µm

BOW CENTER

2 3 4 CDK1

2 3 4

CDK1/ DAPI 50µm

73 Figure. 3.1 CDK1 protein expression in normal lens epithelial cells and fiber cells. Anti CDK1 antibodies detected CDK1 protein in control (Cre negative) mice where the floxed Cdk1 allele (Cdk1L/L) remained intact. Zones of the E17.5 lens were subdivided into central epithelium (1), and fiber cells (2-4) with higher numbers representing progressively older fiber cells. These zones (depicted in the upper left panel) are shown in higher magnification with (CDK1/DAPI) and without (CDK1) nuclear counterstaining with DAPI. Note, that while CDK1 protein appeared in both the cytoplasm and nucleus of epithelial cells and fiber cells of the bow region (regions 1 and 2), CDK1 protein exclusively localized to the nucleus of deeper (older) fiber cells (arrowheads in region 1, regions 3-4). epi and fiber represent the lens epithelial and fiber cell compartments, respectively.

74 CDK1 Immunofluorescence Relative CDK1 Cdk1L/L MLR39; Cdk1L/L Fluorescence Intensity

A B 120 MLR39;Cdk1L/L le le 100

80 P0 P0 60 OFZ OFZ 40 100µm 20 0 Lens Retina

Cdk1L/L Cdk1L/L MLR10; Cdk1L/L 140 MLR10;Cdk1L/L F 120 D E 100

E15.5 60

E15.5 le le 40 re 20 re 200µm 0 Lens Retina

G Fibers Epithelium

MLR39 MLR39 Cdk1L/L Cdk1L/L Cdk1L/L Cdk1L/L

Cdk1 34kDA

GAPDH 37kDA

Fig. 3.2 MLR39 Cre does not deplete the lens of CDK1 Cdk1L/L and MLR39; Cdk1L/L lenses were compared at P0 (birth) for CDK1 expression (A-C). CDK1 protein is evident in the lens epithelium and nuclei of fiber cells in Cdk1L/L lenses (A). The MLR39 transgene did not reduce immunologically detectable CDK1 expression in the fiber cells of MLR39; Cdk1L/L mice (B, C). Western blot analysis supported the immunofluorescent data, as MLR39; Cdk1L/L lenses retained CDK1 protein in the fiber cell mass (G). Cdk1L/L and MLR10; Cdk1L/L lenses were compared at E15.5 for the expression of CDK1 (D-F). At E15.5 CDK1 was detected throughout the entire epithelium of Cdk1L/L lenses and in early differentiating fiber cell nuclei (D); whereas MLR10; Cdk1L/L lenses showed a mosaic pattern of CDK1 expression in the epithelium and almost no detectable CDK1 in the fiber cells (E). The fluorescent intensity for immunological detection of CDK1 was reduced 50% in MLR10; Cdk1L/L lenses relative to the lenses of the of Cre negative controls lenses (F). However, the relative fluorescent intensity of CDK1 detection in the retina did not differ significantly between either the MLR39; Cdk1L/L, and Cdk1L/L mice (C) or the MLR10 Cdk1L/L and Cdk1L/L mice (F). Relative fluorescent intensity of CDK1 in MLR39; Cdk1L/L, and MLR10; Cdk1L/L lens and retina were measured by ImageJ software and normalized to the control lens and retina (C, F). Scale bars represent 100µm in A and B; 200µm in C-F.

76 Cdk1L/L MLR10; Cdk1L/L

A epi B epi le

OFZ CDK1

epi D C epi le le

OFZ CDK1/ DAPI

200µm

E F L/L MLR10; Cdk L/L

INTENSITY Cdk1

CDK1

Relative amount 1.0 0.29 FLUORESCENT

GAPDH CDK1

77 Figure 3.3 Little CDK1 expression remains in MLR10; Cdk1L/L lens cells by E17.5. Cdk1L/L (A, C) and MLR10; Cdk1L/L (B, D) lenses were compared at E17.5 for the expression of CDK1 with (C, D) and without (A, B) nuclear counterstaining with DAPI. Abundant CDK1 was detected throughout the entire epithelium (epi) and in early differentiating fiber cell nuclei of Cdk1L/L lenses (A,C). In contrast CDK1 was absent from most of the MLR10; Cdk1L/L lens epithelium and only a few CDK1 positive nuclei are observed in early differentiating fiber cells (B, D, arrows). Relative CDK1 levels were estimated via immunofluorescent intensity measurements (E). CDK1 protein levels were similar in the retinas of Cdk1L/L and MLR10; Cdk1L/L mice. le and OFZ represent the lens and organelle free zone, respectively. Western blotting of total lens protein from E18.5 lenses revealed a marked reduction in CDK1 in MLR10; Cdk1L/L lenses with GAPDH as a loading control (F).

L/L L/L Cdk1 MLR10; Cdk1

A B E12.5

100µm

E15.5

E17.5

E’

L/L

Cdk1

F’

L/L MLR10; Cdk1

79 Figure 3.4 The formation of an organelle free zone requires CDK1. Hematoxylin and Eosin staining of the lenses at E12.5 (A, B), E15.5 (C, D), and E17.5 (E, F) with boxed zones designated 1-6 indicating comparable areas within the fiber cell mass shown at 10 fold higher magnification (E’, F’). At E12.5 Cdk1L/L control (A) and MLR10; Cdk1L/L (B) lenses appeared morphologically indistinguishable. At E15.5 the MLR10; Cdk1L/L lens fibers in the bow region contained larger nuclei than comparable fiber cells in the control bow region (compare nuclei in the dashed ovals of D and C). The cortical secondary fiber cell nuclei of MLR10; Cdk1L/L lenses were fewer in number and larger than those of the control lens at E17.5 (F’ and E’, compare zones 1 and 2). Although the nuclear size of MLR10; Cdk1L/L fiber cells normalized, the nuclear density remained lower than the control lens in zones 3-5 (F’, E’). By E17.5 an organelle free zone (OFZ), devoid of nuclei, had formed in the control lenses (dashed oval in E and zone 6 in E’) but nuclei persisted in the MLR10; Cdk1L/L lenses (zone 6, F’). Scale bars represent 100 µm in A, B, C, D; 200 µm in E, F and 20 µm in E’ and F’.

80 p57KIP2/DAPI p27KIP1/DAPI pLaminAC/DAPI A B C L/L Cdk1

E1 7.5 D E F L/L MLR10; Cdk1 100µm 100µm 100µm

DAPI DLAD Inset TUNEL/DAPI G H I L/L Cdk1

E17.5 J K L/L MLR10; Cdk1 200µm

200 µm 20 µm

81 Figure 3.5 CDK1 deficiency decreased the phosphorylation of lamin A/C, blocked the entry of DLAD into the nucleus and decreased DNA degradation in maturing lens fiber cells. Primary antibodies to p57KIP2 (A, D), p27KIP1 (B, E), pLamin A/C (C, F) and DLAD (H, K) were used on E17.5 Cdk1L/L (A, B, C, G, H) and MLR10; Cdk1L/L (D, E, F, J, K) lens sections to detect appropriate antigens. TUNEL analysis on E17.5 Cdk1L/L (I) and MLR10; Cdk1L/L (L) lens sections revealed DNA degradation. p57KIP2 expression (green stained nuclei) initiated in transitional zone epithelial cells in both control (Cdk1L/L) and Cdk1 deficient (MLR10; Cdk1L/L ) lenses but declined quickly in as fiber cells elongated (bracketed region in A, D). In contrast, p27KIP1 persisted in the nuclei of fiber cells deep into the cortex of the control lens (bracketed region in B) but remained more cortical in the MLR10; Cdk1L/L lenses (E). Lamin A/C phosphorylation (green foci in C, dashed circle) initiated near the center of the lens where the organelle free zone is forming. In the same region where pLamin A/C is detected in the control lenses, both DLAD positive nuclei (lower right inset, H), and TUNEL positive foci (I, yellow staining, arrows) are found. MLR10; Cdk1L/L lenses do not contain pLamin A/C (F), display reduced TUNEL positive fiber cell foci (L, yellow staining, arrow) and exhibit DLAD accumulation around rather than within late fiber cell nuclei (lower right inset, K). Upper right insets in H and K are high magnifications of cortical fiber cells were DLAD expression is comparatively weak whereas lower right insets are high magnifications of more mature fiber cells where DLAD expression is clearly evident. Nuclei are counterstained with DAPI which is blue in A-G, J but pseudocolored red in I and L to enhance the contrast for the TUNEL assay. Scale bars represent 100µm in A-F; 200µm in G, H, I, J, K,L. In the insert of H and K the scale bar represents 20µm.

83 Cdk1L/L MLR10; Cdk1L/L

A B C MLR10;

L/L Cdk1L/L Cdk1

DAPI 2 3 pNuMA 1 1 2 3 1.0 0.48 Relative amount

pNuMA / GAPDH

100µm 100µm

pNuMA/DAPI

A1 A2 A3 L/L Cdk1

20µm

B1 B2 B3 L/L MLR10; Cdk1

20µm

BOW CENTER

84 Figure 3.6 Lenses deficient in CDK1 failed to phosphorylate NuMA. The monoclonal anti- phosphorylated threonine 2055 of NuMA (pNuMA) antibody detected the presence of pNuMA in Cdk1L/L lenses (A, C), and MLR10; Cdk1L/L lenses at E16.5 (B, C). Immunoflourescent (A, B) and western blot analysis (C) revealed reduced pNuMA in MLR10; Cdk1L/L lenses (B, C) relative to Cdk1L/L lenses (A, C). Three regions of Cdk1L/L (A,white boxes 1-3) and MLR10; Cdk1L/L (B, white boxes 1-3) lenses were selected for magnification (A1-A3, B1-B3 respectively). At the bow region of Cdk1L/L lenses, pNuMA diffusely spread across the entire nucleus (A1). As fiber cells mature toward the center of Cdk1L/L lenses, pNuMA localization appeared more punctate, finally converging on a single focus in the most mature fiber cell nuclei (A2-A3). In contrast, MLR10; Cdk1L/L lenses exhibit low levels of pNuMA post-mitotically, as both peripheral (B1) and central fibers (B3) lack pNuMA staining. Scale bars represent 100µm in A and B; 50µm in A1-A3; B1-B3.

85 Bow Center A B 3

L/L 2

Cdk1 1 4

E17.5 C D L/L

5 7 8 6 MLR10; Cdk1 50µm

Fig. 3.7 Nuclear architecture is disrupted with CDK1 depletion DAPI staining was implemented for nuclear structure analysis of CdkL/L (A, B) and MLR10; Cdk1L/L (C, D) E17.5 lenses. Consistent nuclear structure changes occur moving inwardly from the lens bow (A) towards the lens center (B) in Cdk1L/L lenses. At the lens bow, control lens nuclei are oval in shape and exhibit intense DAPI foci (A, nucleus 1 and 2). Moving towards the lens center, nuclei become spherical (B, nucleus 3), and contain an intense, even DAPI stain throughout the nucleus (B nucleus 4). MLR10; Cdk1L/L lenses do not exhibit consistent nuclear changes. Some nuclei near the bow region exhibit an intense DAPI stain throughout the nucleus, as if it were about to denucleate (C, nucleus 5), whereas some nuclei near the lens center are enlarged and oval shaped (D, nucleus 7 and 8). Scale bar represents 50µm A-D.

86 Fig. 3.8 CDK1 deficiency decreased the phosphorylation of Histone H1 in the lens fiber cells. A primary antibody to pH1 was used on E17.5 Cdk1L/L (A, B) and MLR10; Cdk1L/L (C, D) lens sections to determine the impact of CDK1 on phosphorylated Histone H1. Control lenses displayed intense pH1 staining in epithelial cell nuclei, presumably in proliferative cells of the germinative zone (A, B-green). Additionally, control lenses contained enhanced pH1 staining in the bow region of the lens (A, B- green). MLR10; Cdk1L/L lenses contained intense pH1 staining the lens epithelial cell layer, but the pH1 staining was absent in the fiber cell mass (compare white circle in D to white circle in B). Scale bar represents 100 µm A-D.

87 TOM20 TOM20/DAPI PDI A B C epi cf L/L

Cdk1 O F * Z

E17.5 D E F c L/L epi f

Mitochondrial Free

MLR10; Cdk1 * 100µm 200µm

Figure 3.9 MLR10; Cdk1L/L lenses remove both mitochondria and endoplasmic reticulum despite retaining nuclei. Mitochondria were detected by Tom20 (A, B, D, E) and endoplasmic reticulum by PDI (C, F) immunofluorescence (green staining) in E17.5 lenses. Cdk1L/L (A, B, C) and MLR10; Cdk1L/L (D, E, F) lens fiber cells lose their mitochondria (A, B, D, E) and endoplasmic reticulum (C, F) prior to reaching the center of the lens. Tom20 staining drops precipitously subsequent to the most peripheral fiber cells (asterisks in A, D) but remains as punctate foci until the deep fiber cells of the central zone (arrows in A, D). Cdk1L/L lenses form an organelle free zone (OFZ) lacking both mitochondria and nuclei (B). The MLR10; Cdk1L/L central lens fibers lack mitochondria, but retain nuclei (E, yellow arrowheads). Likewise, both control (C) and MLR10; Cdk1L/L (F) lenses remove PDI-staining endoplasmic reticulum from mature nuclear fiber cells (lack of green staining within the dotted line border in C, F). Nuclei were counterstained with DAPI (B, E). Scale bar represents 100µm A, B, D, E or 200µm C, F. epi and cf denote the lens epithelium and central fiber cells, respectively.

88 Cdk1L/L MLR10; Cdk1L/L

A B

epi epi E15.5

fiber fiber

C D

epi epi E17.5

fiber fiber 20µm

E p-value < 0.0001 * L/L * Cdk1 * NUMBER/SECTION

CELL

MLR10; Cdk1L/L EPITHELIAL

90 Figure 3.10 CDK1-deficient lenses exhibited large, sparse epithelial cell nuclei. Cdk1L/L (A, C) and MLR10; Cdk1L/L (B, D) lenses at E15.5 (A, B) or E17.5 (C, D) were stained with hematoxylin and eosin. At E15.5 the MLR10; Cdk1L/L lens epithelium contained fewer nuclei that appeared larger (B) than those in the Cdk1L/L control epithelium (A). This relative epithelial cell reduction continued in MLR10; Cdk1L/L lenses at E17.5 (Compare D to C). Since the MLR10; Cdk1L/Land Cdk1L/L lenses were similar in size, the number of lens epithelial cell nuclei per section was used as an indicator of lens epithelial cell population size. At embryonic day 12.5 (E12.5) lens epithelial sections contained comparable numbers of nuclei in both control and CDK1-deficient lenses, but by E15.5 the number of lens epithelial cell nuclei present in the MLR10; Cdk1L/L lenses actually declined, while the epithelial cell number increased in the control lenses (E). The lens epithelial nuclei per section continued to diverge between MLR10; Cdk1L/L and control lens through P0. Scale bars represent 20 µm A-D. epi and fiber represent the lens epithelium and fiber cell compartment, respectively. Arrows highlight selected epithelial nuclei to emphasize the difference in nuclear size and population density between the MLR10; Cdk1L/L and control lens epithelium.

91 Cdk1L/L Cdk1L/L; MLR10 Cdk1L/L Cdk1L/L; MLR10

epi A B E epi F epi epi

le E15.5 le le le re

re DAPI

epi epi C D G epi H epi pH3/ DAPI BrdU/

le le

E17.5 re le le re

100µm 200µm

P-value < 0.001 I Cdk1L/L * MLR10; Cdk1L/L

P-value < 0.001 P-value < 0.01 * * Proliferation Index Proliferation

BrdU pH3 BrdU pH3 BrdU pH3 DAPI J

L/L p-value < 0.0001

Cdk1 *

K MLR10 ; ; L/L

Cdk1 20µm

92 Figure 3.11 MLR10; Cdk1L/L lens epithelial cells continue to synthesize DNA but fail to enter mitosis. BrdU incorporation and phosphorylated histone H3 (pHH3) immunohistochemistry (green nuclei) were used to determine the proportion of cells in S-phase (A-D) and late G2 phase (E-H), respectively. Nuclei were counterstained with DAPI. The proliferation index (S-phase fraction) did not differ significantly between MLR10; Cdk1L/L and Cdk1L/L lenses at E12.5 or at E15.5 (compare B to A, solid bars in I) but significantly increased in MLR10; Cdk1L/L lenses by E17.5 (compare D to C, solid bars in I). Note: although there were fewer overall BrdU positive nuclei in the MLR10; Cdk1L/L lenses, the proportion of total nuclei that were BrdU positive was relatively increased at E17.5. The proportion of pHH3 positive cells levels were significantly higher in Cdk1-deficient lenses beginning at E15.5 (compare E to F, shaded bars in I) and most remaining epithelial cells in MLR10; Cdk1L/L lenses stained positive for pHH3 by E17.5 (compare H to G, shaded bars in I). Whole lenses from Cdk1L/L (J) and MLR10; Cdk1L/L mice (K) were stained with DAPI and the intact epithelium was visualized by confocal microscopy to visualize the size and density of epithelial nuclei. MLR10; Cdk1L/L lenses exhibited a significant increase in nuclear size (K, L), and an increased DAPI staining foci in each cell. Scale bars represent 100µm in A-D; 200µm in E-H; 20 µm J and K. le, re and epi represent lens, retina and lens epithelium, respectively.

93 TUNEL/DAPI

L/L Cdk1 MLR10 Cdk1L/L A * B

epi epi E15.5 fiber fiber

C D epi

epi

E17.5 fiber fiber

100µm

Fig. 3.12 Cdk1 deletion does not result in epithelial cell apoptosis. TUNEL analysis was implemented on E15.5 (A, B) and E17.5 (C, D) Cdk1L/L (A, C) and MLR10; Cdk1L/L (B, D) lenses to determine if increased apoptosis led to the reduction in epithelial cell number observed in MLR10; Cdk1L/L lenses. At E15.5 neither Cdk1L/L (A) or MLR10; Cdk1L/L (B) exhibited many TUNEL positive nuclei (green) in the epithelial cell layer, despite E15.5 MLR10; Cdk1L/L lenses already exhibiting noticeably fewer nuclei in the epithelial cell layer as indicated by the nuclear stain DAPI (red). Although the lens center of Cdk1L/L displayed numerous TUNEL positive foci by E17.5, the epithelial cell layer of both E17.5 Cdk1L/L (C) and MLR10; Cdk1L/L (D) lacked any detectable TUNEL positive signal. Sections were counterstained with DAPI (red) in A-D. Scale bar represents 100 µm in A-D.

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99 CHAPTER 4

FGFR and PTEN signaling interact during lens development to regulate cell survival

4.1 SUMMARY: Lens epithelial cells express many receptor tyrosine kinases (RTKs) that stimulate PI3K-AKT and RAS-RAF-MEK-ERK intracellular signaling pathways. These pathways ultimately activate the phosphorylation of key cellular transcription factors and other proteins that control proliferation, survival, metabolism and differentiation in virtually all cells. Among RTKs in the lens, only stimulation of Fibroblast growth factor receptors (Fgfrs) elicits a lens epithelial cell to fiber cell differentiation response. Moreover, despite expressing 3 Fgfrs in the lens, depleting the lens of just FGFR2 leads to both a survival and lens fiber cell differentiation phenotype. Phosphatase and tensin homolog (Pten), commonly known for its role as a tumor suppressor, inhibits ERK and AKT activation and primarily initiates apoptotic pathways, and cell cycle arrest. Here, we show that when we deplete the lens of FGFR2 and PTEN, the cell death phenotype normally associated with FGFR2 loss disappeared. Additionally, PTEN abrogation in the presence and absence of FGFR loss increased AKT and ERK activation above the levels of controls, yet does not stimulate ectopic differentiation in the Pten isolated deletion or restore Aquaporin0 and DnaseIIβ when deleted in the presence of FGFR2 loss. In conclusion, lens cell survival absolutely requires AKT and ERK activation, however lens fiber cell differentiation requires signaling specifically through FGFR2.

4.2 INTRODUCTION

The relative developmental simplicity of the ocular lens makes it an important model to study developmental mechanisms controlling cellular growth, survival, differentiation, and

100 proliferation (Wormstone and Wride, 2011). Invaginations of surface ectoderm overlying the optic vesicles create bilateral lens vesicles during early mammalian development (reviewed in Robinson, 2014). Cells in the posterior hemisphere of the lens vesicle withdraw from the cell cycle, elongate, and turn on fiber cell specific genes as they differentiate into the primary fiber cells reviewed (reviewed in Bassnett and Beebe, 2004) . The lens vesicle cells in the anterior hemisphere differentiate into the lens epithelium. Only lens epithelial cells can proliferate, and as the lens matures, cell proliferation ceases in all areas except the germinative zone, a narrow band of epithelial cells slightly anterior to the lens equator (Harding et al., 1971; McAvoy, 1978). The proliferation within the germinative zone displaces epithelial cells toward the lens equator where they differentiate into secondary fiber cells. Proliferation in the germinative zone and secondary fiber cell differentiation provide a constant source of new lens fibers throughout the mammalian lifespan. Among the numerous receptor tyrosine kinases (RTKs) expressed in the developing lens, fibroblast growth factor receptors (FGFRs) play a unique and indispensable role in lens development (Garcia et al., 2011; Garcia et al., 2005; Madakashira et al., 2012; Robinson, 2006; Zhao et al., 2008). As with most RTKs, ligand (FGF) binding by FGFRs leads to downstream activation of intracellular phosphorylation cascades culminating in the activation by phosphorylation of ERK1/2 and AKT kinases. Activation of these kinases leads to many of the cellular responses associated with growth factor stimulation (Lemmon and Schlessinger, 2010). In lens explants, or cultured lens epithelial cells, AKT and ERK1/2 phosphorylation results in enhanced cell survival, growth, proliferation, and differentiation (Chandrasekher and Sailaja, 2004a, b; Iyengar et al., 2006; Le and Musil, 2001a, b; Lovicu and McAvoy, 2001; Wang et al., 2009; Weber and Menko, 2006). However, eliminating or inhibiting FGFR signaling in vivo leads to dramatically decreased lens cell survival and inhibition of differentiation without significantly altering cell proliferation (Chow et al., 1995 et al., 1995; Garcia et al., 2011; Garcia et al., 2005; Madakashira et al., 2012; Robinson et al., 1995; Stolen and Griep, 2000; Zhao et al., 2008). The mouse lens specifically expresses three FGFR genes, Fgfr1, Fgfr2 and Fgfr3 (Hoang et al., 2014). Lenses lacking Fgfr2 prior to the lens vesicle stage undergo degeneration marked by both apoptosis and differentiation defects, while simultaneously removing Fgfr1 exacerbates this phenotype (Garcia et al., 2005; Zhao et al., 2008). Conditional deletion of Fgfr1, Fgfr2 and

101 Fgfr3 receptors in the lens, subsequent to the lens placode stage, causes massive apoptosis and arrest of fiber cell differentiation (Zhao et al., 2008). Conversely, lenses that overexpress FGFs in vivo undergo ectopic fiber cell differentiation in the lens epithelium (Lovicu and Overbeek, 1998; Robinson et al., 1998; Robinson et al., 1995). FGFR activation requires heparan sulfate in a ternary complex with FGF. The loss of heparan sulfate synthesizing enzymes Ndst1 and Ndst2 also causes lens cell apoptosis, reduced proliferation and defective fiber cell differentiation (Qu et al., 2011). However, expression of a constitutively active Ras allele in these lenses increased ERK1/2 phosphorylation and reversed the Ndst1/Ndst2 deficient phenotypes. Since lens cells rely on FGFR signaling for survival, decoupling the apoptotic phenotype from the differentiation phenotype in lenses with compromised FGFR signaling remains a challenge. During normal development, FGFR signaling in the lens may primarily promote cell survival with defective differentiation in FGFR-deficient lenses resulting as a secondary response to apoptosis. AKT enhances cell survival by a variety of mechanisms, including inhibiting FOXO transcription factors and destabilizing the pro-apoptotic BAD/Bcl-XL complex (reviewed in Zhang et al., 2011). FGFR stimulation activates phosphoinositide 3-kinase (PI3K) which converts the cell membrane lipid PtdIns (4,5)P2, hereafter referred to as PIP2, into PtdIns(3,4,5)P3, hereafter referred to as PIP3. PIP3 then recruits AKT to the cell membrane where phosphorylation by mTORC2 and PDK1 activates AKT (Sarbassov et al., 2005). The tumor suppressor protein, Phosphatase and tensin homolog (PTEN), counteracts PI3K by dephosphorylating PIP3 back to PIP2, leading to reduced AKT activation. In addition to inhibiting AKT activation, PTEN acts as a tumor suppressor by inhibiting cell proliferation and promoting apoptotic pathways (Chung and Eng, 2005; Franke et al., 2003; Weng et al., 2001a). Given the antagonism between PI3K and PTEN, we hypothesized that PTEN may act as an important negative regulator of FGFR activity during lens development. In particular, PTEN activity may drive lens cells toward apoptosis by exacerbating decreased PIP3 levels in FGFR- deficient lens cells, which indirectly prevents the activation of AKT. Studies in both osteoprogenitor cells and keratinocytes reveal the importance of balancing FGFR and PTEN signaling. Deletion of Fgfr2 rescues overproliferation in osteoprogenitors caused by the loss of Pten (Guntur et al., 2011). Likewise, skin tumorgenesis resulting from Pten deletion requires Fgfr2 (Hertzler-Schaefer et al., 2014). To specifically determine whether increased AKT activation might rescue cell survival in FGFR-deficient lens cells, Cre recombinase facilitated

102 the lens-specific removal of both Pten and Fgfr2 during early lens development. We reasoned that the restoration of survival in FGFR-deficient lens cells would also reveal survival- independent aspects of FGFR-mediated fiber cell differentiation. Given the central importance of FGFR signaling in the development of many different tissues and organs (reviewed in Carter et al., 2015; Teven et al., 2014), it comes as no surprise that aberrant FGFR signaling causes numerous developmental disorders and drives the pathogenesis of many human cancers (reviewed in Ahmad et al., 2012; Katoh and Nakagama, 2014; Wesche et al., 2011). Often, the same mutations that give rise to developmental disorders in the germline lead to specific cancers in somatic tissues, suggesting that FGFRs regulate different processes in a context dependent manner. Likewise, PTEN mutations drive the genesis and malignancy of several human tumors (reviewed in Mester and Eng, 2013). Revealing how FGFR and PTEN signaling interact in the context of lens development may facilitate the discovery of new targets for therapeutic intervention to treat diseases or conditions caused by FGFR and/or PTEN dysfunction.

4.3 RESULTS: 4.3.1 Balancing PTEN and FGFR2 in lens cell survival and proliferation. 4.3.1 a Loss of Pten restores lens size in FGFR2-deficient lenses. To generate the required mice, the Le-Cre transgene mediated the deletion of homozygous loxP- flanked (floxed) alleles of Fgfr2 (Fgfr2Δ/Δ), Pten (PtenΔ/Δ), or both Fgfr2 and Pten (Pten/R2Δ/Δ) in mouse lens precursor cells at embryonic day 9.5 (E9.5) of development. Control mice consisted of animals lacking the Le-Cre transgene while homozygous for floxed alleles of Pten, Fgfr2, or both Pten and Fgfr2. Since the Le-Cre transgene possesses the ability to independently affect eye development on some genetic backgrounds (Dora et al., 2014), our experiments controlled for any phenotypes resulting solely from the Le-Cre transgene with hemizygous Le-Cre mice containing wild-type alleles of Fgfr2 and Pten. Quantitative reverse transcriptase PCR (RT- qPCR) confirmed the deletion of the floxed alleles (Fig 4.1 A, B). Both Fgfr2Δ/Δ and Pten/R2Δ/Δ lenses contained very few Fgfr2 transcripts at E15.5, and Pten deletion alone failed to alter the abundance of Fgfr2 transcripts compared to that in the control lenses (Fig. 4.1 A). Likewise, Pten/R2Δ/Δ and PtenΔ/Δ lenses exhibited efficient deletion of the Pten transcript, but lenses

103 lacking Fgfr2 alone displayed an increased number of Pten transcripts compared to that in Cre negative lenses (Fig. 4.1 B). Although lenses lacking Fgfr2 failed to achieve normal size or morphology, lenses lacking Pten resembled control lenses. The gross morphology of PtenΔ/Δ lenses was virtually indistinguishable from that of Le-Cre negative control lenses at E12.5 (compare Fig. 4.2 B to A), E15.5 (compare Fig. 4.2 F to E), and E18.5 (compare Fig. 4.2 J to I). Moreover, PtenΔ/Δ lenses failed to display a significant difference in lens section planar area at any stage examanined (Fig. 4.2 M). In contrast, lenses depleted of FGFR2 displayed a drastic size reduction at E12.5 (Fig. 4.2 compare C to A). In addition to decreased size, the presumptive fiber cells in the posterior of E12.5 Fgfr2Δ/Δ lenses exhibited drastically reduced elongation toward the anterior epithelium (Fig. 4.2 compare C’ to A’ white arrows). Interestingly, Fgfr2-deficient lenses possessed a consistent tilt towards the nasal side of the head (Fig 4.2 C’ black arrow). By E15.5, the Fgfr2Δ/Δ lenses remained significantly smaller than control lenses and contained vacuolated central fiber cells (Fig. 4.2 compare G to E, G’ to E’, M). By E18.5, Fgfr2Δ/Δ lenses remained abnormally small compared to control, PtenΔ/Δ or Pten/R2Δ/Δ lenses (Fig. 4.2 compare K to I, J and L, M). Although isolated Pten deletion failed to alter lens morphology or size, deletion of both Pten and Fgfr2 significantly rescued the reduced size of Fgfr2Δ/Δ lenses (Fig. 4.2 compare D, H, L to C, G, K; M). Moreover, by E12.5 primary fiber cell elongation in Pten/R2Δ/Δ lenses advanced well beyond that seen in lenses missing only Fgfr2 (Fig. 4.2 compare D’ to C’). At E15.5, Pten/R2Δ/Δ lenses were virtually indistinguishable from control lenses, in both size and morphology (Fig. 4.2 compare H and H’ to E and E’). Like control lenses, E15.5 Pten/R2Δ/Δ lenses contained a single layer of epithelial cells lining the anterior surface with differentiating fiber cells at the equator, or bow region, of the lens (Fig. 4.2 Compare H’ to E’). At E18.5, lenses lacking both PTEN and FGFR2 reached a size only slightly smaller than control lenses (Fig. 4.2 compare L to I, M), but significantly larger than Fgfr2Δ/Δ lenses. Although the deletion of Pten permitted FGFR2-deficient lenses to reach nearly normal size, normal eyelid closure failed to occur in either Fgfr2Δ/Δ or Pten/R2Δ/Δ mice (Fig. 4.2 compare dashed box of L and K to I).

4.3.1b Pten deletion restores cell survival in FGFR2-deficient lenses 104 The loss of Fgfr2 leads to smaller lenses primarily because of decreased lens cell survival (Garcia et al. 2005). TUNEL analysis made it possible to determine if Pten deletion rescues cell survival in lens cells lacking Fgfr2. As expected, lenses lacking FGFR2 exhibited a significant increase in TUNEL positive nuclei at E12.5 compared to control lenses (Fig. 4.3 compare C to A, I). At this stage, Fgfr2Δ/Δ lenses contained apoptotic cells in both the anterior and posterior sides of the lens (Fig. 4.3 C). At E15.5, Fgfr2Δ/Δ lenses contained fewer TUNEL positive cells than E12.5 Fgfr2Δ/Δ lenses (Fig. 4.3 compare G to C), but still contained a higher apoptotic index than control lenses (Fig. 4.3 compare G to E). At this stage, the Fgfr2Δ/Δ apoptotic nuclei mainly localized to the lens epithelium (Fig. 4.3 G-white arrows), with the fiber cells remaining TUNEL negative. Although deleting Pten in isolation actually resulted in a slight increase in apoptosis as compared to control lenses at E12.5 (Fig. 4.3 compare B to A, I), the simultaneous deletion of both Pten and Fgfr2 significantly increased cell survival relative to Fgfr2Δ/Δ lenses both at E12.5 (Fig. 4.3 compare D to C, I,) and at E15.5 (Fig. 4.3 compare H to G, J). TUNEL positive nuclei nearly disappeared in the control and PtenΔ/Δ lenses by E15.5 (Fig. 4.3 F, J).

4.3.1c The effect of Pten deletion on lens cell proliferation. To determine if an increase in cellular proliferation contributed to the restoration of lens size in Pten/R2Δ/Δ lenses, we labeled embryos with BrdU to determine the S-phase fraction in control, PtenΔ/Δ, Fgfr2Δ/Δ, and Pten/R2Δ/Δ lens cell nuclei. Neither PtenΔ/Δ nor Pten/R2Δ/Δ lenses displayed a significant difference in BrdU incorporation, relative to the control lenses, at E12.5 (Fig. 4.4 compare B to A, D to A; M) or E15.5 (Fig. 4.4 compare F to E, H to E; N). Despite the unchanged proliferation index in PtenΔ/Δ and Pten/R2Δ/Δ lenses, several nuclei in the posterior (fiber cell compartment) incorporated BrdU in the fiber cell mass in both genotypes (Fig. 4.4 B’ and D’-white arrows). At E12.5, the Fgfr2Δ/Δ lenses exhibited increased BrdU incorporation (Fig. 4.4 compare C to A, M), with an overall morphology suggesting that the primary fiber cells had not yet withdrawn from the cell cycle (Fig. 4.4 C’-white arrows). Because fiber cell elongation was impaired in FGFR2-deficient lenses, it was difficult to discern epithelial cells from fiber cells. For this reason, the proliferation index for all E12.5 lenses was calculated for the whole lens, rather than for just lens epithelial cells (Fig. 4.4 C’; M). By E15.5, BrdU positive cells were no longer detected in the fiber cell mass of Fgfr2Δ/Δ lenses, and therefore, at this stage

105 only the epithelial cell proliferation index was calculated (Fig. 4.4 G). By E15.5, Fgfr2Δ/Δ lenses did not display a change in epithelial cell proliferation compared to the Cre negative controls (Fig. 4.4 compare G to E, M). Taken together, the evidence suggests that Pten deletion restores FGFR2-deficient lens size primarily through increased cell survival rather than increased proliferation. Additionally, lens epithelial cell proliferation was not dependent on the presence of either PTEN or FGFR2 at E15.5 and E18.5.

4.3.2 The impact of FGFR2 and PTEN on lens epithelial cell-to-fiber cell differentiation Four hallmarks of fiber cell differentiation are 1) cell cycle withdraw 2) fiber cell elongation, 3) the onset of fiber cell structural protein expression and 4) organelle loss. At E12.5, Fgfr2Δ/Δ lenses exhibited defects in both fiber cell elongation (Fig. 4.2 C’) and cell cycle withdrawal (Fig. 4.4 C, C’- white arrows). Although PTEN deletion largely restored fiber cell elongation in the FGFR2-deficient lenses at E12.5 (Fig. 4.2 compare C’ to D’), several posterior fiber cells failed to exit the cell cycle at this stage (Fig. 4.4 D-white arrows). Given the partial restoration of fiber cell differentiation in the FGFR2-deficient lenses by simultaneous deletion of Pten, we sought to determine more broadly which FGFR2-deficient lens phenotypes depended on the presence of PTEN.

4.3.2a Reduced γ-crystallin expression in Fgfr2Δ/Δ lenses is partially restored in the absence of PTEN. Another hallmark of lens epithelial-to-fiber cell differentiation is the production of fiber cell-specific structural proteins. Crystallin proteins are the most abundant structural components of the mammalian lens fiber cells mass, and permit lens transparency (reviewed in Harding and Dilley, 1976). Most β- and γ-crystallin proteins appear exclusively in lens fiber cells, although β- crystallin expression precedes the onset of γ-crystallin expression (reviewed in McAvoy et al., 1999). Both β- and γ-crystallin depend on FGFR signaling (Madakashira et al., 2012; Zhao et al., 2008). Moreover, ectopic activation of Frs2α in lens epithelial cells induces high levels of pAKT and pERK1/2 accompanied by the onset of β-crystallin expression (Madakashira et al., 2012). FGFR2-deficient lenses display dramatically reduced γ-crystallin (Fig. 4.5 compare C to A), and modestly reduced of β-crystallin (Fig. 4.5 compare K to I) expression at E12.5. As lens

106 development progresses to E15.5 and E18.5, β- and γ-crystallin expression increases in Fgfr2Δ/Δ lenses (Fig. 4.5 G, H; Supp. Fig. 1 G, K). Furthermore, by E15.5 γ-crystallin transcript levels in Fgfr2Δ/Δ lenses is indistinguishable from control lenses (Fig. 4.5 Q). Additionally, the expression of c-maf, which encodes a transcription factor known to regulate crystallin expression in the lens (Kim et al., 1999; Yang et al., 2006 et al. 2006) remains unaltered in Fgfr2 deleted lenses at E15.5 (Fig. 4.5 R). Although Pten deletion alone failed to reduce β- or γ-crystallin expression, (Fig. 4.5 compare B, F, to A, E; Q) Pten deletion increased γ-crystallin expression in FGFR2-deficient lenses at E12.5 (Fig. 4.5 compare D to C). As in E15.5 Fgfr2Δ/Δ lenses, E15.5 Pten/R2Δ/Δ lenses did not display a significant difference in γ-crystallin or c-maf transcripts (Fig. 4.5 Q, R).

4.3.2b Pten deletion fails to rescue reduced Aquaporin0 levels in FGFR2-deficient lenses. Aquaporin0, the most abundant lens fiber cell membrane protein, promotes fiber cell adhesion which minimizes extracellular space (Engel et al., 2008; Kumari et al., 2013). Deletion of Frs2α in the lens placode resulted in a reduced Aquaporin0 expression (Madakashira et al., 2012). Likewise, Fgfr2Δ/Δ lenses experienced a significant reduction in Aquaporin0 protein expression, demonstrated by immunofluorescence and western blot analysis (Fig. 4.6 compare C to A, J, K). Since Aquaporin0 localizes to the fiber cell membranes, immunofluorescent analysis revealed that FGFR2-deficient fiber cells appear structurally different than Cre negative controls lenses. Control and PtenΔ/Δ lenses contain Aquaporin0 expression in columns extending the length of the lens fiber cell mass representing individual lens fiber cells (Fig. 4.6 A and B). In contrast, Fgfr2Δ/Δ lenses exhibited circular Aquaporin0 stained structures in the center of the lens. (compare Fig. 4.6 arrows in C to A). Depleting Pten failed to restore either the normal amount of Aquaporin0 protein or the normal structure of the central fiber cells. Like Fgfr2Δ/Δ lens fiber cells, Pten/R2Δ/Δ lenses contained intense expression of Aquaporin0 in a circular pattern (Fig 4.6. D), and did not restore Aquaporin0 expression to control levels (Fig. 4.6 J, K).

4.3.2c Fgfr2Δ/Δ lens fiber cells experience nuclear retention reduced

DnaseIIβ expression, neither of which is rescued by simultaneous deletion of Pten.

107 In between E16-E18, central lens fiber cells initiate the removal of their nuclei, a process that requires the nuclease DnaseIIβ (Bassnett, 1997, 2009; Chaffee et al., 2014; Nakahara et al., 2007). Both Fgfr2Δ/Δ and Pten/R2Δ/Δ lenses retain their nuclei in central fiber cells (Fig. 4.6 G, and H), representing another differentiation defect present in FGFR2-deficient lenses that simultaneous deletion of Pten fails to restore. TUNEL analysis reveals DNA breakdown that takes place normally as a consequence of fiber cell denucleation. Both control and PtenΔ/Δ lenses contained numerous TUNEL positive foci in the central fiber cell mass, the site of active denucleation (Fig. 4.6 F and E-white arrows). In contrast, neither Fgfr2Δ/Δ nor Pten/R2Δ/Δ lenses exhibited abundant TUNEL positive staining in this region (Fig. 4.6 G, H), although a few Pten/R2Δ/Δ central fiber cells contained TUNEL positive foci (Fig. 4.6 H-white arrow). To determine if the inability to degrade the nuclear content in both Fgfr2Δ/Δ and Pten/R2Δ/Δ lenses resulted from a reduction in the expression of the essential fiber cell nuclease, DnaseIIβ , qRT-PCR was performed standardizing the DnaseIIβ transcript abundance to that of GAPDH. Both Fgfr2Δ/Δ and Pten/R2Δ/Δ lenses exhibited a significant reduction in DnaseIIβ transcript (Fig. 4.6 I). Deleting Pten in isolation did not significantly alter DnaseIIβ (Fig. 4.6 I).

4.3.2d FGFR2-deficient lenses exhibit PAX6 retention in posterior lens cells. Depleting the lens of PTEN restores the removal of PAX6 from FGFR2-deficient posterior lens cells. As epithelial cells differentiate into fiber cells, not only do fiber cell specific proteins, such as Aquaporin0, γ-crystallin, and DNaseIIβ become expressed, certain proteins driving the production of epithelial cell specific genes need to be removed. PAX6 is a key regulator of eye development, where it is necessary for eye formation, and misexpression of PAX6 can initiate ectopic eye development (reviewed in Shaham et al., 2012). Although PAX6 is required to drive the expression of lens epithelial genes, PAX6 must be removed from the promoter of several fiber cell specific genes, such as γ-crystallin, in order for their transcriptional initiation (reviewed in Cvekl and Duncan, 2007). As PAX6 maintains its expression in posterior lens cells in Fgfr1-3 conditional knockouts at the lens vesicle stage, and this retention of PAX6 accompanied the differentiation defects observed in the FGFR1-3 triple knockout, we predicted that FGFR2 deficiency might lead to

108 PAX6 retention in the fiber cell mass contributing to the differentiation defects observed in Fgfr2Δ/Δ lenses. A specific antibody against PAX6 revealed that PAX6 remains in the posterior cells of Fgfr2Δ/Δ lenses at E12.5, whereas control lenses display low amounts of nuclear PAX6 in differentiating fiber cells (Fig. 4.7 compare C to A, C’ to A’-white brackets). At E15.5 PAX6 was largely removed from the fiber cell mass of Fgfr2Δ/Δ lenses, but remained in “islands” of posterior nuclei removed from the rest of fiber cell nuclei (Fig. 4.7 G-inset, white arrows). Interestingly, these “islands” of removed nuclei express high levels of not only PAX6, but p27KIP1 and PROX1. These PAX6 “islands” in Fgfr2Δ/Δ lenses occur in close proximity to folds of the neural retina caused by the abnormally small lens (Fig. 4.7 G-inset, white arrows; Fig. 4.10 O-inset, white arrows; Fig 4.2- dotted circle). Deleting Pten restored the ability of posterior FGFR2-deficient lens cells to remove PAX6 at E12.5 (Fig. 4.7 compare D to C, D’ to C’). Moreover, isolated Pten deletion, had no influence on the pattern of PAX6 expression in comparison to the control in E12.5 lenses (Fig. 4.7 compare B to A) or E15.5 (Fig. 4.7 compare F to E).

4.3.3 The deletion of Pten normalizes several downstream signaling pathways affected by Fgfr2 loss in the lens.

4.3.3a Pten deletion restores AKT and ERK1/2 activation in Fgfr2-deficient lenses. FGFR signaling and PTEN work antagonistically in terms of their impact on both PI3- K/AKT and MAPK/ERK1/2 (Di Cristofano and Pandolfi, 2000) (Weng et al., 2001a; Weng et al., 2001b). The activation of AKT, by phosphorylation, promotes cell survival. In contrast, PTEN initiates apoptotic pathways, in part, by inhibiting AKT activation. The ERK1/2 sensitive cellular responses include, but are not limited to, the promotion of cellular proliferation, migration, and differentiation. At E15.5, PtenΔ/Δ lenses displayed over a 5-fold increase in p-AKT (Fig. 4.8 A, B). Surprisingly, the total level of AKT protein was reduced upon Pten deletion (Fig. 4.8 A, C). PtenΔ/Δ lenses also experienced a significant increase in p-ERK1/2 (Fig. 4.8 A, D), although this increase observed in p-ERK1/2 was more modest than that observed in p-AKT (Fig. 4.8 A, compare differences between PtenΔ/Δ lenses to control lenses in B and D). Pten deletion did not alter the total amount of ERK1/2 protein (Fig. 4.8 E).

109 Fgfr2Δ/Δ lenses exhibited a 29.8% reduction in p-AKT and a 24% reduction in p-ERK1/2 signal in comparison to control lenses (Fig. 4.8 A, B, D). Depleting the lens of PTEN, in addition to the Fgfr2 deletion increased the levels of both p-AKT and p-ERK1/2 beyond the levels of control lenses. The Pten/R2Δ/Δ lenses experienced nearly a 4-fold increase in p-AKT expression when compared to control lenses. Additionally, Pten/R2Δ/Δlenses experienced the same amount of p-ERK1/2 increase as deleting solely Pten (Fig. 4.8 D). The total level of ERK1/2 protein remained unchanged in Pten/R2Δ/Δlenses, but like the single deletion of Pten, knocking out Pten and Fgfr2 resulted in low levels of total AKT protein (Fig. 4.8 A, C, E).

4.3.3b The effect of PTEN and FGFR2 on c-Jun N-terminal protein kinase (JNK) c-Jun represents another critical survival factor as it is capable of activating AKT by increasing PDK1 transcription and suppressing both p53 and Pten (reviewed in Dhanasekaran and Reddy, 2008; Kolomeichuk et al., 2008). Activated c-Jun is also required to mediate the apoptotic response through the activation of the transcription factor, AP-1, which is correlated to the transcription of apoptotic factors such as, Bak, Tnf-α, and Fas-2 (reviewed in Dhanasekaran and Reddy, 2008). In addition to apoptosis, c-Jun activation can initiate cell proliferation and differentiation and can act downstream of FGFR signaling (reviewed in Dhanasekaran and Reddy, 2008). To elucidate the impact of deleting Pten, Fgfr2, and Pten and Fgfr2 on activated p53 and c-Jun western blot analysis was implemented comparing the 3 LeCre mediated conditional knockouts to Cre negative control lenses using antibodies raised against phoshprylated p53 (p-p53) and cJun (p-cJun). FGFR2-deficient lenses up regulate the phosphorylation of both p53 (Fig. 4.9 A, B), and c-JUN (Fig. 4.9 A, C). The deletion of Pten alone did not affect the phosphorylation of either p53 or c-JUN, but Pten deletion restored the normal level of p-c-JUN and p-p53 in FGFR2-deficient lenses (Fig 4.9).

4.4 DISCUSSION: A proper balance between PTEN and FGFR signaling is crucial in both the development and homeostatic maintenance of organs and tissues. Developmentally, the outcome of FGFR signaling is diverse, but includes enhanced proliferation, survival, and at times (as in the case of the lens) promotes cell cycle withdraw and differentiation. Aberrant FGFR signaling can promote tumorigenesis by driving proliferation, enhancing survival, and promoting angiogenesis

110 (reviewed in Turner and Grose 2010). PTEN inhibits two of the major pathways stimulated by FGFR signaling (MAPK/ERK1/2 and PI3K/AKT), and plays an important role in inhibiting proliferation and stimulating apoptosis. In addition to its role as a tumor suppressor, PTEN participates in early developmental processes and morphogenesis. Mice null for Pten experience embryonic lethality, and tissue specific deletions in Pten provide evidence for its role in appropriate apoptosis during embryonic development (Di Cristofano et al., 1998; Li et al., 2002; Tiozzo et al., 2009). As previously reported, the impact of depleting the lens of just FGFR2 with Le-Cre, resulted in both apoptosis and fiber cell differentiation defects (Garcia et al., 2005). To further elucidate the role of FGFR2 in differentiation, we combined the Fgfr2 deletion with additionally deleting Pten, anticipating a restoration of survival, but not differentiation. Additionally, we depleted the lens of just PTEN to determine if depleting the lens of PTEN alone altered fiber cell differentiation-and to characterize the impact of PTEN deficiency on the developing ocular lens. As ectopic over activation of FRS2α activation followed by differentiation in central lens epithelial cells (Madakashira et al., 2012), we questioned whether enhanced AKT and ERK1/2 activation, through PTEN loss, could mimic the effect of over activation of FRS2α. Furthermore, despite the recorded physiological role of PTEN in maintaining the Na+/ATPase activity, the developmental role of PTEN has not been established (Sellitto et al., 2013). By deleting Pten and Fgfr2 at the lens placode stage, we successfully decoupled the apoptotic phenotype with the differentiation phenotype associated with FGFR2-deficient lenses. The Pten deletion likely prevents apoptotic pathways, in part, by increasing p-AKT 5-fold as the activation of AKT, by phosphorylation, promotes cell survival pathways. In addition to promoting cell survival, AKT inhibits apoptotic pathways. One way AKT prevents apoptosis is through phosphorylating MDM2. Once phosphorylated, MDM2 transports to the nucleus and mediates the degradation of the proapoptotic factor p53 and, therefore prevents Pten transcription, as p53 is a transcription factor known to drive the transcription of Pten (Mayo et al., 2002; Mayo and Donner, 2002). Along with a heightened apoptotic index, lenses deficient of FGFR2 experienced low AKT and higher levels of p53 activation (Fig. 4.9 A, B) and Pten transcript (Fig. 4.1 B). Conversely, when PTEN was depleted in the presence of the Fgfr2 deletion, AKT activation increased beyond the level of control lenses, and p-p53 dropped. Taken

111 together, these data suggests that FGFR2 promotes cell survival in the lens, in part, through activating AKT, which prevents against p-p53 activation and Pten transcription. The activation of c-Jun by JNK is tightly correlated with cellular survival, although, the activation of c-Jun has been shown to both enhance and inhibit cell survival dependent on the tissue type and apoptotic stimuli (Hettinger et al., 2007; Morishima et al., 2001; Resnick and Fennell, 2004). For example, in the case of neuronal cells, mutating Serine 63 and 73 to Alanine of c-Jun, preventing c-Jun phosphorylation, resulted in the neuronal cells becoming resistant to kainate-induced apoptosis (Dhanasekaran and Reddy, 2008). Contrarily, in nutrient deprived fibroblasts, apoptosis was significantly higher in the absence of c-Jun (Hettinger et al., 2007). Additionally, JNK signaling contains contradicting roles on Pten and p53 (Dhanasekaran and Reddy, 2008; Hettinger et al., 2007). In many cancer cell lines, and fibroblast cells, c-Jun suppresses Pten transcription by enhancing AKT/MDM2 activation and stimulating p53 degradation (Eferl et al., 2003; Hettinger et al., 2007). On the other hand, the induction of PTEN required the presence of JNK signaling in intestinal epithelial cells (Dhanasekaran and Reddy, 2008). Here, we show that the FGFR2 deficiency results in high levels of p-cJun. Furthermore, deleting Pten restores c-Jun activation back to control levels (Fig. 4.9 A, C). This is suggestive that, in the context of the lens with impaired FGFR signaling, p-cJun stimulates apoptotic pathways. Surprisingly, the upregulation of p-cJun with FGFR2 deficiency requires intact Pten, despite prior reports suggesting that it is Pten that is downstream of p-cJun. Although the apoptosis resulting from FGFR2 deletion requires PTEN, endogenous apoptosis during lens development does not require PTEN. Lenses deficient of PTEN still experience TUNEL positive cells (Fig. 4.3 I) despite the over 5-fold increase in AKT. During normal lens morphogensis, apoptosis occurs as the lens vesicle is forming from the lens pit, whereby, the cells that will eventually close off the lens vesicle and separate from the rest of the surface ectoderm undergo higher amounts of cell death (Fig. 4.11 E-white arrows). Not only do PtenΔ/Δ PtenΔ/Δ exhibit reduced apoptosis in between the lens pit and lens vesicle stage (Fig. 4.3 I; Fig. 4.11). This is in contrast to the role of PTEN in involuting mammary glands and developing lung epithelial cells as PTEN-deficiency enhances cell survival promoting enlarged tissue (Li et al., 2002; Tiozzo et al., 2009).

112 AKT, ERK1/2, and c-Jun activation have received extensive attention with regard to cellular proliferation. High levels of these correlate with increased cellular proliferation, and are often correlated as a driver of tumorigenesis. Although Fgfr2Δ/Δ lenses experience an early cell cycle withdraw phenotype, none of our conditional knockout lines experienced alterations in epithelial cell proliferation by E15.5 or E18.5. This is surprising as both, PtenΔ/Δ and Pten/R2Δ/Δ Fgfr2Δ/Δ lenses display significantly heightened p-c-Jun. Regarding lens fiber cell differentiation, the restoration of a robust, AKT and ERK1/2 activation (beyond levels of control lenses) does not completely restore fiber cell differentiation in the absence of FGFR2. Moreover, high levels of pAKT and pERK1/2 in the lens epithelium of PtenΔ/Δ lens epithelium, whereas ectopic overactivation of FGFR signaling promotes fiber cell differentiation in central lens epithelial cells (Lovicu and Overbeek, 1998; Madakashira et al., 2012; Robinson et al., 1998; Robinson et al., 1995). Taken together, it appears that signaling through FGFRs, specifically, is required for fiber cell differentiation beyond just high levels of AKT and ERK1/2 activation. As the PI3K/AKT and MAPK/ERK1/2 pathway are two of the main pathways associated with FGFR stimulation, particularly in lens-based studies, the question still remains, what else is downstream of FGFR signaling that is essential for fiber cell differentiation? The lab of Dr. Cvekl recently published late in 2013 that upon FGF2 addition to rat LECs, there are 131 FGF responsive miRNAs-targeting over 3000 transcripts (Wolf et al., 2013). Conditional knockout of Dicer1, which is required for miRNA production, using both Le-Cre and the MLR10 Cre transgene (initiating Cre expression at the lens vesicle stage) initiated the interest in miRNAs in lens development (Li and Piatigorsky, 2009; Wolf et al., 2013). MLR10 Cre–mediated deletion of Dicer1 led to fiber cell elongation defects, fiber cell organization defects, and nuclear retention (Wolf et al., 2013). Potentially, the components of fiber cell differentiation that are not restored by Pten deletion in FGFR2-deficient lenses are due to the dependency of differentiation on drastic changes of miRNA stimulated by FGFR signaling. In particular, the FGF-dependent miRNAs may be responsible for Aquaporin0 or expression. To date, only miR- 204 has been functionally characterized in the lens (Conte et al., 2010). The function of miRNAs in lens and their endogenous control through FGFR signaling remains a relatively unexplored topic,

113 yet could explain how FGFR signaling dictates lens fiber cell differentiation outside of AKT and ERK. Recently, a manuscript was published raising the concern on the use of the LeCre transgene, as phenotypes have been reported on homo and hemizygous LeCre lines without containing LoxP flanked alleles (Dora et al. 2013). Although we have seen, and reported independent LeCre phenotypes (Robinson, 2005), we have not observed an independent LeCre affect using mice mostly bred on an FVB background. Moreover, our experiments implemented 3 different knockouts lines of mice mediated by LeCre. Two of our lines (Fgfr2Δ/Δ, Pten/R2Δ/Δ) are hemizygous for the LeCre transgene, and carry loxP-flanked Fgfr2. In these two FGFR2- deficient, LeCre positive lines of mice, there are low levels of Aquaporin0 protein and DnaseIIb transcript, and nuclear retention. When comparing the two of the lines alone, it would be reasonable to question whether these phenotypes resulted from FGFR2-deficiency, PTEN- deficiency or a LeCre toxicity affect. When comparing these mutants to another mutant hemizygous for cre and homozygous for loxP-flanked PTEN, which does not have reduced Aquaporin0, DnaseIIβ or nuclear retention, we can confidently conclude that the phenotype is resulting from FGFR2 deficiency. Additionally, we compared LeCre hemizygous (LeCre +/-) E12.5 lenses to Cre negative lenses for TUNEL (Fig. 4.12 compare B to A), PAX6 (Fig. 4.12 compare D to C), γ-crystallin (Fig. 4.12 compare F to E), and E15.5 LeCre +/- to control lenses for γ-crystallin and Aquaporin0 (Fig. 4.12 compare J to I) and did not detect any observable lens fiber cell differentiation defects resulting from LeCre+/-, although hemizygous for LeCre resulted in a higher apoptotic index. In conclusion, these data expands the known relationship of PTEN and FGFR signaling. In the context of the lens, PTEN and FGFR signaling act antagonistically, primarily at the level of cell survival, and to an extent, lens fiber cell differentiation. The elucidation of the interactions of FGFR signaling and PTEN is critical as the maintenance of these two pathways prove crucial during development and in disease. Achondroplasia, craniosynostosis, thanatophoric dysplasia, and cancer result from abnormal FGFR signaling. Mutations in Pten are correlated with Cowden’s Syndrome, Autism, and nearly every cancer type. This work provides for another system revealing the antagonism existing between these two pathways during development.

114 4.5 FIGURES

P< 0.0021 P< 0.00045 P< 0.035 A P< 0.003 B P< 0.00062

Δ/ Δ Δ/Δ Δ/Δ 0.02 Control Pten Fgfr2 (Pten/R2) 0.08 mRNA mRNA 0.015 0.06 GAPDH GAPDH 0.01 0.04 Fgfr2 to Pten to of

of 0.005 0.02 Ratio Ratio

0 0 Δ/ Δ Δ/Δ Δ/ Control Pten Fgfr2 (Pten/R2)

Figure 4.1 Le-Cre efficiently deletes LoxP-flanked Pten and Fgfr2.

RT-qPCR was used to detect Fgfr2 (left bar graph) and Pten (right bar graph) E15.5 Le-Cre negative (Control), PtenΔ/Δ, Fgfr2Δ/Δ, and Pten/R2Δ/Δ lenses to determine the efficiency of the deletion of LoxP-flanked alleles. Lenses with LoxP-flanked Pten and the Le-Cre transgene

(PtenΔ/Δ) displayed similar expression levels of Fgfr2 transcript as control lenses, but expressed very little Pten transcript. LoxP-Flanked Fgfr2 lenses containing the Le-Cre transgene

(Fgfr2Δ/Δ) displayed a significant reduction in Fgfr2 transcript and a significant increase in Pten transcript. Lenses containing both LoxP -flanked Pten and Fgfr2 and the Le-Cre transgene contained almost no Fgfr2 or Pten transcript.

115 Control Pten / Fgfr2 / (Pten/R2) / A B C D Le Le Le Le

Re Re Re Re

100⎧m

A’ C’ E12.5 B’ n D’ a s a l 20⎧m

E F G H

Le Le Le Le Re Re Re Re 50⎧m

E’ F’ G’ H’ E15.5

Fibers Fibers Fibers Fibers Bow

Bow Bow Bow 20⎧m

I J K L

Le Le Le Le

E18.5 Re Re Re Re 200⎧m

116 Figure 4.2 Pten deletion rescues the lens size and elongation defects in Fgfr2Δ/Δ lenses. Hematoxylin and Eosin staining was implemented to analyze the morphology and size of the four different genotypes being compared, Cre-negative controls (A, A’, E, E’, I), PtenΔ/Δ (B, B’, F, F’ J) Fgfr2Δ/Δ (C, C’, G, G’ K), and Pten/R2Δ/Δ (D, D’, H, H’ L) at E12.5 (A-D, A’-D’), E15.5 (E-H, E’-H’), and E18.5 (I-L). A’-D’ and E’-H’ are higher magnifications of the boxed in regions of A- D and G-H’ respectively. Lens planar area measurements were taken using an Nikon TI-80 microscope in conjunction with their advanced research software at E15.5 and E18.5 (M). Quantitative reverse transcriptase PCR was used to assess the efficiency of Cre- mediate deletion of LoxP-flanked alleles and were standardized to GAPDH mRNA levels (N- Fgfr2 O-Pten). At E12.5 the posterior cells of Fgfr2Δ/Δ lenses did not elongate (compare C to A, C’ to A’- white arrows). Deleting Pten restored the fiber cell elongation phenotype displayed in FGFR2- deficient lenses (compare D to C, D’ to C’-white arrows). At E15.5 and E18.5 Fgfr2Δ/Δ lenses were significantly smaller than control lenses (compare G to E, K to I, M). Pten deletion rescued lens size in FGFR2-deficient lenses at E15.5 and E18.5 (compare H to G, L to K), but did not restore the eyelid closure phenotype present in FGFR2 lenses (compare L and K to I). Pten deletion by itself did not visible alter the size or morphology of the lens at any of the 3 stages compared (compare B, B’, F, F’ J to A, A’ E, E’, I, M). Errors bars on the graphs represent s.e.m, with each bar representing a minimum of 9 measurements (3 sections from the lens center of 3 different embryos). Scale bars: 100µm in A-D; 20 µm in A’-D’ and E’-H’; 50 µm in E’H; 200 µm in (I-L).

117 TUNEL/DAPI Control Pten Δ/ Δ Fgfr2Δ/Δ (Pten/R2)Δ/Δ A B C D

Le Le Le Le E12.5

Re Re Re Re 100⎧m

E F G H

Fibers

E15.5 Fibers Fibers Fibers

100⎧m

P< 0.0261 I J P< 0.0009 P< 0.0001

P< 0.0002 P< 0.0132 P< 0.0053 P< 0.0066 P< 0.0006 0.14" 0.14"

Cell

0.12" 0.12" Lens 0.1" 0.1" Index

Index 0.08" 0.08" Epithelial Whole

0.06" 0.06"

0.04" 0.04" Lens Apoptotic

E12.5 0.02" Apoptotic 0.02"

0" Δ/Δ 0" Δ/ Δ Δ/Δ E15.5 Δ/ Δ Δ/Δ Δ/Δ CCoonnttrool"l PtePtne n" FgFgffrr22 " (PPtteenn"R/2R"2) CCoonnttrorol""l PPtteenn"L/L" FFggfrf2r"2L/L" Pte(nP"tLe/Ln"F/gRfr22L)/L"

118 Figure 4.3 Pten deletion restores lens cell survival in FGFR2-deficient lenses TUNEL analysis was implemented on E12.5 and E15.5 (E-F) lenses comparing Cre-negative controls (A, E), PtenΔ/Δ (B, F) Fgfr2Δ/Δ (C G), and Pten/R2Δ/Δ (D, H). I and J represent the apoptotic index at E12.5 and E15.5 respectively. At E12.5, the apoptotic index was calculated for the entire lens due to the apoptosis detected in both posterior and anterior lens cells, whereas the apoptotic index was only calculated for the epithelial cell layer at E15.5 as only apoptosis was detected in the epithelium. At E12.5 and E15.5 Fgfr2Δ/Δ lenses displayed a significant increase in TUNEL positive cells as compared to control lenses (compare C to A, G to E I), although the apoptotic index was higher at E12.5 than E15.5 (compare C to G). At both E12.5 and E15.5 deleting Pten rescued the cell survival phenotype associated with FGFR2 loss (compare D to C, H to G, I, J). Despite the rescue of apoptosis when Fgfr2Δ/Δ lenses are compared to Pten/R2Δ/Δ lenses, Pten/R2Δ/Δ lenses remained to exhibit higher levels of apoptosis at E12.5 and E15.5 (compare D to A, H to E, I, J). Moreover, the apoptotic index in PtenΔ/ lenses was increase at E12.5 in comparison to control lenses (compare B to A-white arrows; I). In part, the apoptosis could be the affect of the Cre-transgene, as Le-Cre transgenic mice have been reported to display a phenotype in the absence of floxed alleles (Dora et al. 2014). Errors bars on the graphs represent s.e.m, with each bar representing a minimum of 9 measurements (3 sections from the lens center of 3 different embryos). Scale bars: 100µm in A-H. White arrows are pointing to TUNEL positive foci.

119 BrdU/DAPI Control Pten / Fgfr2 / (Pten/R2) /

A B C D

Le Le Le Le

Re Re Re Re 100⎧m

E12.5 A’ B’ C’ D’

50⎧m

E F G H

Le Le Le Le E15.5

Re Re Re Re 200⎧m

E12.5 Whole Lens E15.5 Lens Epithelial Cell M Proliferation Index N Proliferation Index P< 0.0001 50 50

45 P< 0.0001 P< 0.0001 45 40 40 35 35 30 30 25 25

Incorporation 20 Incorporation 20 15 15 10

BrdU BrdU 10 5 5 % % 0 0 Control Pten / Fgfr2 / / / / / ()*+,)-" .+/*" 012,%" (P.+t/e*n"3/%R"2) (C)*o+n,)tr-"ol .P+t/e*n010" 2F3g4f,r%2 "010".+(/P*t"e2n3/4R,2%)"01

120 Figure 4.4 PtenΔ/Δ,Fgfr2Δ/Δ, and Pten/R2Δ/Δ lenses display early cell cycle withdrawal defects. None of the knockouts exhibited lens epithelial cell proliferation alterations later in development.

BrdU incorporation assay was used on Cre-negative controls (A, A’, E, I), PtenΔ/Δ (B, B’, F, J) Fgfr2Δ/Δ (C, C’, G, K), and Pten/R2Δ/Δ (D, D’, H, L) to assess the impact of deleting Pten, Fgfr2, and the combination of the two on lens epithelia cell proliferation and cell cycle withdraw. By E12.5, control lenses did not exhibit any BrdU incorporation in the differentiation fiber cells (A, A’). E12.5 PtenΔ/Δ (B, B’) ,Fgfr2Δ/ Δ (C, C’), and Pten/R2Δ/Δ (D, D’) lenses displayed retained nuclei in the fiber cell mass, although Fgfr2Δ/Δ lenses contained the most proliferation in their posterior cells (compare C’ to B’ and D’). Furthermore, overall proliferation was increased in E12.5 Fgfr2Δ/Δ lenses (M), yet was not significantly increased in E12.5 PtenΔ/Δ or Pten/R2Δ/Δ lenses (M). By E15.5 (E- H, N) and E18.5 (I-L) the lens epithelial proliferation was not altered in PtenΔ/Δ (F, J) Fgfr2Δ/Δ (G, K), and Pten/R2Δ/Δ (H, L). A’-D’ represent boxed regions of A-D selected for higher magnification. Dashed white lines outline the edges of the lens and white arrows point towards posterior lens cells remaining prolific (A’-D’). Errors bars on the graphs represent s.e.m, with each bar representing a minimum of 9 measurements (3 sections from the lens center of 3 different embryos). Scale bars: 100 µm in A-D; 50 µm in A’- D’; 200 µm in E-L.

121 Control Pten⊗/ Fgfr2⊗/⊗ (Pten/R2)Δ/Δ ⊗ A B C D

Le Le Le E12.5 Le

Re Re Re Re 100⎧m

E F G H γ - crystallin/ DAPI

Le Le Le E15.5 Le

Re Re Re Re 200⎧m

I J K L

E12.5 Le Le

Re Re Re Re 100⎧m

M N O P β - crystallin/ DAPI

E15.5 Le Le Le

200⎧m

Q 140.0000# R 0.3"

0.25"

120.0000# to

100.0000# 0.2"

mRNA 80.0000# maf maf CrygD mRNA 0.15" c- 60.0000# 0.1" 40.0000# GAPDH

Ratio of Ratio 0.05" 20.0000# GAPDH Ratio of Ratio

0" 0.0000# Control Pten Δ/ Δ Fgfr2Δ/Δ (Pten/R2)Δ/Δ Control Pten Δ/ Δ Fgfr2Δ/Δ (Pten/R2)Δ/Δ Control# PTEN# R2# PTEN/R2# Control" PTEN" R2" PTEN/R2"

122 Figure 4.5 Pten deletion restores E12.5 γ-crystallin expression. Later in development Fgfr2Δ/Δ, and Pten/R2Δ/Δ lenses display normal transcript levels of γ-crystallin and c-maf. Cre-negative control (A, E, I), PtenΔ/Δ (B, F, J) , Fgfr2Δ/Δ (C, G, K), and Pten/R2Δ/Δ (D, H, L) lenses were analyzed by immunohistochemistry at E12.5 (Carrara-de Angelis et al.), E15.5 (Muise-Helmericks et al.), E18.5 (I-L) to determine the expression of γ-crystallin. At E12.5 γ- crystallin was present in very few posterior cells of Fgfr2Δ/Δ lenses (compare C to A). E12.5 Pten/R2Δ/Δ lenses displayed an increase in γ-crystallin expression in relation to Fgfr2Δ/Δ lenses (compare D to C), but remained reduced in comparison to Control lenses (compare D to A). By E15.5, Fgfr2Δ/Δ lenses increased its γ-crystallin protein (compare G to C). Quantitative RT-PCR using whole E15.5 Fgfr2Δ/Δ and Pten/R2Δ/Δ lenses indicated that the γ-crystallin transcript levels in both Fgfr2Δ/Δ and Pten/R2Δ/Δ lenses remained comparable to control lenses (M). Furthermore, the regulatory transcription factor of γ- crystallin, c-maf, did not display reduced transcript levels in Fgfr2Δ/Δ and Pten/R2Δ/Δ lenses (R). Pten deletion by itself did not alter γ-crystallin protein at E12.5 (compare B to A), E15.5 (compare F to E) or E18.5 (compare J to I) or E15.5 transcript levels of γ-crystallin (M) and c-maf (N). The quantification of γ-crystallin and c-maf were standardized to gapdh mRNA. Errors bars on the graphs represent s.e.m, with each bar representing a minimum of 9 measurements (3 sections from the lens center of 3 different embryos). Scale bars: 100 µm in A-D; 200 µm in E- H; 200 µm in I-L.

123 Control Pten⊗/ Fgfr2⊗/ (Pten/R2)Δ/Δ ⊗ ⊗

A B C D DAPI

Le Le Le E15.5 Le

Re Aquaporin0/ Re Re Re 200⎧m

E F G H

Lens Center Lens Center Lens Center Lens Center DAPI E18.5 TUNEL/ 100⎧m

P< 0.021 I P< 0.009 J 0.025"

to 0.02"

0.015" mRNA Dlad Dlad

0.01" Aquaporin0 GAPDH Ratio of Ratio 0.005"

0" Δ/ Δ Fgfr2Δ/Δ Δ/Δ GAPDH Control Pten Control" PTEN" R2" (PPTtEeNn/R/R2"2)

4 P< 0.0157 P< 0.0384

2 Aquaporin0 to to Aquaporin0

GAPDH mRNA GAPDH 1 Ratio of Ratio

0 Control Pten Δ/ Δ (Pten/R2)Δ/Δ Fgfr2Δ/Δ

124 Figure 4.6 Fgfr2Δ/Δ lenses exhibit reduced Aquaporin0 and a nuclear retention phenotype that is not rescued by Pten deletion.

Cre-negative control (A, E, I), PtenΔ/Δ (B, F, J) , Fgfr2Δ/Δ (C, G, K), and Pten/R2Δ/Δ (D, H, L) lenses were analyzed for Aquaporin0 protein expression at E15.5 (Idrovo Espin et al.), and TUNEL at E18.5 in the lens center-indicated denucleating cells (Muise-Helmericks et al.). Both Fgfr2Δ/Δ (C) and Pten/R2Δ/Δ (D) lenses experienced a reduced Aquaporin0 expression level (compare D and C to A; J). Additionally, Fgfr2Δ/Δ (C) and Pten/R2Δ/Δ (D) contained several heightened areas of Aquaporin0 expression in a circular pattern (C and D white arrows). PtenΔ/Δ lenses did not display an altered expression of Aquaporin0 (compare B to A; J). Western blot analysis was performed on E15.5 to confirm the reduced immunofluorescent detection of Aquaporin0 observed with FGFR2 deficiency (J). The western blot quantification was standardized to GAPDH (K). TUNEL analysis was implemented to detect DNA degradation that occurs in control lenses, in that, TUNEL foci can be found in cells removing their nuclei (E). Both Fgfr2Δ/Δ (G) and Pten/R2Δ/Δ (H) lenses displayed very few TUNEL positive foci (compare G and H to E). Quantitative RT-PCR was performed on E16.5 lenses using primers to detect the nuclease, DnaseIIβ (Dlad) (I) to further characterize the nuclear retention phenotype present in the FGFR2-deficient lenses. The quantification of DnaseIIβ was standardized to GADPH mRNA. Reduced transcript levels of DnaseIIβ were observed in Fgfr2Δ/Δ and Pten/R2Δ/Δ (I). PtenΔ/Δ lenses did not have defects in Aquaporin0 protein expression (J)/localization (B), nuclear removal (F), or DnaseIIβ expression (I). White arrows point towards heightened and abnormal expression of Aquaporin0 in C and D, and TUNEL foci in E, F, and H. Errors bars on the graphs represent s.e.m, with each bar representing a minimum of 9 measurements (3 sections from the lens center of 3 different embryos). Scale bars: 200 µm in A-D; 100 µm in E-H.

125 PAX6/DAPI Control Pten Fgfr2 (Pten/R2) /

A B C D

Le Le Le Le

Re Re Re Re 100⎧m

E12.5 A’ B’ C’ D’

50⎧m

E F G H

Le Le Le Le E15.5

Re Re Re Re 200⎧m

126 Figure 4.7 E12.5 Fgfr2Δ/Δ lenses maintain PAX6 expression in the posterior fiber cells and Pten deletion restores the normal removal of PAX6. Immunological detection of PAX6 was implemented at E12.5 (A-D; A’-D’) and E15.5 (Muise- Helmericks et al.) comparing Cre-negative control (A, A’, E), PtenΔ/Δ (B, B’, F) , Fgfr2Δ/Δ (C, C’, G), and Pten/R2Δ/Δ (D, D’, H) lenses. Control lenses remove PAX6 from differentiating, posterior fiber cells by E12.5 (A, A’), and further to E15.5, mature fiber cells do not possess PAX6 expression (E). Fgfr2Δ/Δ lenses display abnormal retention of PAX6 throughout all the nuclei in the posterior cells of the lens (compare C to A and A’ to C’-white brackets), and contain removed “islands” of PAX6 expression in mature fiber cells at E15.5 (G-inset, white arrows). Pten deletion, in the presence of FGFR2 removal, restores the normal removal of PAX6 in differentiating fiber cells (compare D to C, D’ to C’-white brackets. E15.5 Pten/R2Δ/Δ lenses, like control lenses, remove their nuclei from the lens center (H). Pten deletion on its own did not disrupt the normal removal of PAX6 in either E12.5 (B, B’) or E15.5 (F) lenses. A’-D’ are higher magnifications of the boxed in regions of A-D. Brackets in A’-D’ indicate posterior lens cells that should not be expressing PAX6. Scale bars: 100 µm in A-D; 50 µm in A’-D’; 200 µm in E-H.

127 A P< 0.0048 P< 0.02 B P< 0.0046 C P< 0.017 P< 0.03 P< 0.041 P< 0.0054 P< 0.0041 P< 0.0031 p-AKT

Total AKT

p-ERK P< 0.0075 D P< 0.0147 E P< 0.0325 P< 0.0042 P< 0.0014

Total ERK

GAPDH

Figure 4.8 Deleting Pten restores pAKT and pERK1/2 in FGFR2-deficient lenses. E15.5 Cre-negative control, PtenΔ/Δ , Fgfr2Δ/Δ , and Pten/R2Δ/Δ lenses were analyzed by western blot analysis to determine the impact of deleting Pten, Fgfr2, or the double knockout on MAPK/ERK1/2 and PI3K/AKT activation. Total levels of pAKT were significantly reduced Fgfr2Δ/Δ lenses (A, B). Additionally deleting Pten with the Fgfr2 deletion brought the levels of pAKT well beyond the levels of control lenses (A, B). As expected, PtenΔ/Δ lenses experienced very high levels of pAKT (A, B). Interestingly, the total amount of AKT was reduced in both PtenΔ/Δ lenses and Pten/R2Δ/Δ lenses (A, C). Fgfr2Δ/Δ lenses experienced a significant reduction in p-ERK (A, D), and additionally deleting Pten brought the levels of pERK1/2 above the levels of control lenses (A, D). PtenΔ/Δ lenses experienced very high levels of pERK1/2 (A, D). Total ERK levels were unaltered by any of the conditional knockouts (A, E). Total ERK1/2 and AKT and pERK1/2 and pAKT were standardized to GAPDH for quantification. Errors bars on the graphs represent s.e.m. Scale bars: 50 µm in A-H.

128 P< 0.036 A P< 0.043 P< 0.023 B to

p53

p-p53 GADPH - Phospho

Ratio

GAPDH Control Pten Δ/ Δ Fgfr2Δ/Δ (Pten/R2)Δ/Δ C P< 0.015 - P< 0.018 P< 0.027 p-cJun GAPDH

phosphoSer63/73

Jun

GAPDH of

c- Ratio Control Pten Δ/ Δ Fgfr2Δ/Δ (Pten/R2)Δ/Δ

Figure 4.9 FGFR2 deficiency leads to increased activation of C-JUN and p53 E15.5 whole lenses of Cre-negative control, PtenΔ/Δ , Fgfr2Δ/Δ , and Pten/R2Δ/Δ were analyzed by western blot analysis using antibodies to p-p53 and p-cJun. GAPDH was used as a loading control. Fgfr2Δ/Δ lenses increased both phospho-p53 (A-top panel, B) and p-cJun (A-3rd panel, C). Pten deletion reduced the level of p53 activation (A-top panel, B) below the levels of the control when Fgfr2 was deleted from the lens. Deleting Pten also brought back C-JUN activation to control levels (A-3rd panel, C). Errors bars on the graphs represent +/- s.e.m. p-p53 and p-cJun were standardized to GAPDH for quantification.

129 E18.5 Control Pten / Fgfr2 (Pten/R2)

A Epi B C D Epi Epi

Epi Fibers Fibers DAPI Fibers Fibers BrdU/

E F G H

DAPI

Le Le Le Le crystallin/

I J K L

DAPI Le Le Le Le crystallin/

200⎧m

M N O P

DAPI

/ Le Le Le Le KIP1 p27

Re Re Re Re

Q R S T DAPI Le Le Le Le PROX1/

Re 200⎧m

E18.5 Lens Epithelial Cell Proliferation Index

!#!'$" !#'" mRNA mRNA !#!'" !#&$"

!#!&$" !#&" GAPDH

!#!&" GAPDH to

to !#%$"

KIP1 !#!%$" Incorporation

!#%"

p27 !#!%"

Prox1

of !#!$"

!#!!$" BrdU

% !" !" Ratio

/ / / Ratio / 2&" / ./0132&" / Control Pten / Fgfr2 / (Pten/R2) / C()o*+n,t)-r"ol P.t/e01n" Fg2f&r2" (P./t0e13n2/&R"2) Co()*n+t,)r-"ol Pt./e0n1" Fgfr2 (Pten/R2)

130 Figure 4.10 Deletions in Pten, Fgfr2, or both did not alter E18.5 lens fiber cell differentiation or proliferation.

Primary antibodies to BrdU (A-D), β-crystallin (E-H), γ-crystallin (I-L), p27KIP1 (M-P), and Prox1 (Q-T) were used on E18.5 control (A, E, I, M, Q), PtenΔ/Δ (B, F, J, N, R), Fgfr2Δ/Δ(C, G, K, O, S), and Pten/R2Δ/Δ (D, H, L, P, T) paraffin lens sections. At E18.5, there were no differences observed between the 3 different mutant lines compared in regards to the proliferation or cell cycle withdraw (BrdU A-D), or the fiber cell differentiation markers β-crystallin (A-D), γ- crystallin (I-L). It did not appear that there were “islands” of cells that appeared to have experience an upregulation of p27KIP1 (M-P) and Prox1 (Q-T) separated from the bow region of the lens and in closest proximity to the neural retina in Fgfr2Δ/Δ lenses (O, S-white dashed region). The BrdU index was calculated from the E18.5 lens sections, which represents the total number of BrdU positive cells (green) over the total epithelail cell nuclei (U). RT-qPCR was performed on E18.5 control, PtenΔ/Δ , Fgfr2Δ/Δ, and Pten/R2Δ/Δ whole lens rlysates fo Prox1 and p27KIP1 to quantify these transcripts. Fgfr2Δ/Δ lenses experience a slight upregulation of Prox1 and p27KIP1 transcript levels. Error bars represents S.E.M. Scale bar 200µm A-T.

131 TUNEL/DAPI Control Pten Δ/ Δ Fgfr2Δ/Δ (Pten/R2)Δ/Δ

A B C D

100⎧m

E10.5 E F G H

50⎧m

Figure 4.11 PTEN deletion did not inhibit apoptosis at E10.5

TUNEL analysis was implemented on E10.5 lens sections comparing Cre-negative controls (A, E), PtenΔ/Δ (B, F) Fgfr2Δ/Δ (C G), and Pten/R2Δ/Δ (D, H). E-H represent higher magnification of A-D. TUNEL positive foci were mainly localized to the anterior edges of the lens pit (white arrows) on control (A, E), PtenΔ/Δ (B, F), and Pten/R2Δ/Δ (D, H). The apoptosis appeared to be spread throughout the lens pit of Fgfr2Δ/Δ mice (C, G). Scale bars: 100 µm (Carrara-de Angelis et al.), 50µm (Muise- Helmericks et al.).

132 Cre Negative LeCre +/-

A B 10 cells

positive 6 DAPI 4 2 TUNEL/

Percent TUNEL Percent 0 100⎧m Control LeCre mi

C D

AX6 P E12.5

100⎧m

E F

DAPI 100⎧m

200⎧m

E15.5 I J

DAPI Aquaporin0/ 200⎧m

133 Figure 4.12 Le-Cre Hemizygosity did not alter fiber cell differentiation, but did increase apoptosis.

Control lenses (A, C, E, G, I) were compared to lenses that were hemizygous for Le-Cre without any loxP-flanked alleles (LeCre +/- B, D, F, H, J) at E12.5 (A-F) and E15.5 (G-J). TUNEL assay was used to determine if LeCre+/- resulted in increased apoptosis at E12.5 (A,B, C). LeCre hemizygosity resulted in an increase in TUNEL positive foci (compare B to A, C). LeCre hemizygosity did not alter PAX6 expression at E12.5 (compare D to C), γ-crystallin expression at E12.5 (compare F to E) or E15.5 (compare H to G) or Aquaporin0 expression at E15.5 (comare J to I). Scale Bars: 100µm A-F, 200 µm G-J

134 A B 2.5 C 0.3

GAPDH 2 0.25 to

p-AKT GAPDH 0.2 to

1.5 AKT 0.15 Total AKT AKT 1 Total p- 0.1 of of

GAPDH 0.5 0.05

Ratio 0 Ratio 0 Control Lecre hemi Control Lecre Hemi

D E F 1 1.2

GAPDH 1 0.8 to

p-ERK GAPDH 0.8

to 0.6

ERK 0.6

Total ERK AL ERK 0.4 0.4 p- TOT

of 0.2 of

0.2 GAPDH Ratio 0 0 Ratio Control Lecre hemi Control Lecre Hemi

4.13 LeCre Hemizygosity did not alter downstream PI3K/AKT or MAPK/ERK1/2 Protein from E18.5 lenses, either negative for Le-Cre (Lecre -/-) or hemizygous for Lecre (Lecre het or Lecre +/-) that did not cotain LoxP flanked alleles, was isolated and subjected to western blot analysis to determine if being hemizygous for Lecre altered downstream PI3K/AKT signaling (A, B, C) or MAPK/ERK1/2 (D, E, F) signaling. The protein levels was quantified and standardized to GAPDH expression level. LeCre hemizygosity did not alter p-AKT (A, B) or total AKT (A, C). Moreover Le-Cre hemizygosity did not alter p-ERK (D, E) or total ERK (D, F) expression. Error bars represent S.E.M.

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142 CHAPTER 5

SUMMARY OF CONCLUSIONS AND FUTURE DIRECTIONS

Prior to this work, the role of CDK1 primarily focused on the role of CDK1 in mitotic progression. Using the ocular lens as a model of development, we established the first post mitotic mechanism of CDK1. Furthermore, as the process of nuclear removal during terminal lens fiber cell differentiation remained enigmatic for more than a century, we revealed a complete mechanism elucidating the way DNaseIIβ gains accesses to the nuclear content. As mitotis requires CDK1, Diril et al reported in 2012 that one of the requirements of CDK1 in mitosis is in preventing endoreduplication of the DNA in cycling cells. Additionally, Diril demonstrated how CDK1 was required for mitotic entry. The work in this dissertation provided yet another system to confirm the necessity of CDK1 in preventing endoreduplication, and the critical nature of CDK1 during mitotic entry. Regarding PTEN and FGFR2, the relationship of these two molecules have recently been appreciated in several cell contexts, with particular interest in cancer therapeutics. In skin tumorigenesis, deletion of just Fgfr2 restored the tumor phenotype resulting from Pten deletion (Hertzler-Schaefer et al., 2014). Interestingly, this work demonstrated that Pten deletion restored the lens cell survival phenotype associated with FGFR2 loss. Moreover, the skin tumorigenesis relationship represented, a near perfect, antagonistic relationship between Pten and Fgfr2 in cellular proliferation (Hertzler-Schaefer et al., 2014). In contrast, the work in this dissertation displayed a near perfect antagonistic relationship of Pten and Fgfr2 in lens cell survival, while; their impact on lens epithelial proliferation was negligible.

5.1 CDK1 is required for lens fiber cell denucleation

Chapter 3 established a post mitotic role of CDK1 in fiber cell denucleation. Several prior studies implicated a role of CDK1 in fiber cell denucleation through the phosphorylation of nuclear lamins. The lab of Peggy Zelenka established the expression patterns of Cyclin B/CDK1 in the

143 fiber cell compartment of rat lenses (Gao et al., 1995). Moreover, in their following study, they established that CDK1/Cyclin B activity peaked at E18 in the lens fiber cell mass (He et al., 1998). Later, Dr. Allen Taylor demonstrated that p27KIP1 retention in the fiber cell mass was a result of mutating the sixth lysine 6 to tryptophan on ubiquitin (K6W-ubiquitin) (Caceres et al., 2010). The K6W-ubiquitin mutant mice also experienced nuclear retention resulting from low levels of Lamin A/C phosphorylation, and thus, suggesting a role of CDK1 in lamin phosphorylation leading to nuclear retention (Caceres et al., 2010). In collaboration with Dr. Allen Taylor, we deleted CDK1 with MLR10 Cre, which directs Cre mediated deletion at E10.5 in the mouse lens. MLR10 Cre mediated deletion of loxP-flanked CDK1 resulted in a mosaic expression of CDK1 protein in the entire lens epithelium and fiber cells by E15.5 (Fig. 3.2 compare E to D), and nearly a complete abrogation of CDK1 protein by E17.5 (Fig. 3.3 compare B to A) As a result of the CDK1 deletion, several targets of CDK1 experienced a reduction in phosphorylation levels specifically in the fiber cell mass, including NuMA, Histone H1, and pLamin A/C. Moreover, the nuclease required for the denucleating fiber cells, DLAD (also called DNaseIIβ), was incapable of penetrating the nucleus of CDK1-deficient fiber cells and their nuclei were retained within the lens center at E17.5 (Chaffee et al., 2014). In conclusion, this study demonstrated that CDK1 was required in a post mitotic context for the nuclear structural changes required for DLAD entry and denucleation (Chaffee et al., 2014).

5.2 S-phase entry occurs in the absence of CDK1, yet CDK1 is indispensible for mitosis.

In 2012, Diril reported that CDK1 was not required for S-phase entry, yet required for the prevention of re-replication of the DNA by competitively inhibiting CDK2, which mediates DNA replication (Diril et al., 2012). Furthermore, CDK1 deficiency resulted in “swollen” nuclei, as CDK1 was required for karyo- and cytokinesis, but not for S-phase entry, as the nuclei continually replicated their DNA without resulting in two daughter cells (Diril et al., 2012). The MLR10 Cre mediated deletion of CDK1 resulted in a depletion of CDK1 in the epithelial layer. Like the Diril 2012 publication, the lens epithelial cell nuclei experienced a 265% increase in nuclear cross-sectional area. The CDK1 deficient lens epithelial cells were capable of entering S- phase, and remained in late G2, (as recognized by the late G2 marker pH3) (Chaffee et al., 2014).

144 5.3 CDK1 is required for apoptosis?

Prior to this work, evidence existed that CDK1 can either inhibit or promote apoptosis. Not only is the role of CDK1 in apoptosis contradictory, but the understanding of the role of CDK1 is important in a clinical sense. Clinical researchers often use microtubule inhibitors to create mitotic catastrophe in cancer cells, and ultimately leading to the death of cancer cells. An example of this is with taxol, a common microtubule inhibitor. Interestingly, taxol increases cell death in cancer cells, yet requires CDK1 activity to do so (Shen et al., 1998). The predicted mechanism of CDK1 in apoptosis is through phosphorylating the pro-apoptotic protein BAD. Once phosphorylated BAD, enters the nucleus and antagonizes Bcl-2 like proteins, which promote cellular survival. Additionally, phosphorylated BAD activates BAX-like proteins which lead to the permeablilization of the mitochondrial membrane and cell death (Konishi et al. 2002). Contradictory to the role of CDK1 in promoting cell death, CDK1 can phosphorylate Survivin. Phosphorylated Survivin forms a complex with Caspase-9, preventing Caspase-9 cleavage, and therefore promoting survival (O’Connor et al., 2002). When we deleted CDK1 from the lens, we detected almost no cell death in the lens epithelium or the lens fiber cells. As normal lens development does not require high levels of apoptosis in the later stages of lens development (E12.5 onwards), we cannot definitely conclude that CDK1 is required for apoptosis in this context. Despite the results being inconclusive, it is interesting that CDK1 deletion does not generate massive amounts of cell death. In general, if a cycling cell does not enter mitosis following G2, it instead goes through apoptosis. Perhaps, apoptosis during mitosis or as a result of mitotic catastrophe requires CDK1.

5.4 Active CDK1 in the lens fiber cell mass opens a new door for future investigations of the role of CDK1 outside of the cell cycle.

As the kinase activity of CDK1 remains active in differentiating lens fiber cells, and the main targets of CDK1 reside inside the nucleus in contact with the chromatin, CDK1 may carry a potential role in gene expression changes accompanying lens fiber cell differentiation. We reported, the phosphorylation of both NuMA, and Lamin A/C require the kinase activity of CDK1 in the lens fiber cells. Additionally, NuMA and Lamin A/C are capable of contacting the

145 chromatin, and changes in their localization are associated with epigenetic changes during differentiation. Using the CDK1-deficient lenses, the overall impact of CDK1 on gene expression could be examined by a high-throughput RNA-sequencing effort comparing the MLR10; Cdk1L/L lenses to Cre-negative control lenses. As the phosphorylation of NuMA, Lamin A/C and Histone H1 required CDK1 in the lens fiber cell mass, and all three of these proteins contact the chromatin, an experiment investigating the gene expression changes in a CDK1 deficient lens would not provide sufficient evidence to determine the specific role of phosphorylating each individual protein on gene expression or nuclear architecture changes. To elucidate a the precise mechanism by which CDK1 alters gene expression, a different approach would need to be taken. It remains to be determined if NuMA phosphorylation in the nucleus alters chromatin organization in lens fiber cells, and whether or not NuMA phosphorylation directed fiber cell differentiation or gene regulation. Our studies observed a decrease in the phosphorylation of T2055 on NuMA in the lens fiber cell mass of CDK1-deficient lenses, and observed the localization of NuMA change throughout the differentiation process in control lenses. Despite these observations, the requirement of this phosphorylation on denucleation and fiber cell differentiation remains elusive. Due to the ease and efficiency of CAS9 mediated genome editing, it would be valuable to mutate T2055 to alanine in order to prevent NuMA phosphorylation at this site. Preventing NuMA phosphorylation at this site could provide evidence for a role in NuMA phosphorylation on gene expression, which as been suggested in the differentiation of human mammary epithelial cells (HMECs). In the context of HMECs, reminiscent of our observation of NuMA localization changes, undifferentiated HMECs exhibited a diffuse distribution of NuMA within the nucleus and upon further morphogenesis; NuMA distribution became concentrated into large foci (Lelievre et al., 1998). Furthermore, using an antibody to target the C terminus of NuMA on mammary acini cells grown in 3D culture shifted NuMA localization from being aggregated to diffuse NuMA across the nuclei (Lelievre et al 1998). The shift in NuMA localization generated a redistribution of acetyl-Histone 4 (acetyl H4) and H4K20 methylation, and an inhibition of acinar differentiation (Abad et al., 2007; Lelievre et al., 1998). As lens fiber cell differentiation is referred as “attenuated apoptosis” the NuMA phosphorylation and localization changes may be mimicking an apoptotic cell. An investagation by Weaver et al. in 1996 reported that NuMA becomes phosphorylated and cleaved prior to

146 DNA degradation during apoptosis (Weaver et al., 1996). Perhaps the NuMA phosphorylation and cleavage primes the DNA for degredation, and NuMA participates in the process of nuclear removal during terminal lens fiber cell differentation. In the lens fiber cell mass, Lamin A/C is not phosphorylated until the nuclei have presumably shut down their transcriptional machinery, whereas the decrease in NuMA takes place in transcriptionally active lens fiber cells. Therefore, studying the impact of lamin A/C phosphorylation on gene expression changes in fiber cell differentiation does not carry the potential as investigating NuMA on gene regulatory mechanisms. Despite this, lamin A/C is implicated in epigenetic mechanisms during cellular differentiation. For example, patients with Huchinson-Gilford Progeria syndrome (HGPS), a well established laminopathy resulting from mutations in the gene encoding lamin A (LMNA), exhibit reduced trimethylation of both Histone H3 lysine 9 (H3K9me3) and Histone H3 lysine 27 (H2K27me3) as well as increased Histone H4 lysine 20 trimethylation (H4K20me3) (Columbaro et al., 2005; Scaffidi and Misteli, 2005; Shumaker et al., 2006). Although the “laminopathies” focus on mutations on genes responsible for lamin production, our discovery of a role of CDK1 in a post mitotic setting may bring more appreciation and interest in investigating the role of CDK1 in disease-related laminopathies. Emery-Dreifuss muscular dystrophy, dilated cardiomyopathy, limb-girdle muscular dystrophy, and Hutchinson-Gilford progeria syndrome primarily affect non-proliferation cells, and it is possible that CDK1 activity plays underappreciated roles in these laminopathies. 5.5 FGFR2 is required for lens cell survival and lens differentiation

Historically, the lens served as a premier tool to investigate Fgfr signaling due to the importance of Fgfr signaling in lens epithelial proliferation, survival and differentiation. Despite numerous publications focusing on Fgfr signaling in lens development, the question still remains, how does Fgfr signaling contribute to these different cellular responses, and whether or not Fgfr signaling directly influences fiber cell differentiation in vivo. One of the main questions centered on the impact on Fgfr2 and whether or not it was required for lens fiber cell differentiation. Previous reports stated that the deletion of Fgfr2 led to both survival and lens fiber cell differentiation defects, yet it still was not clear whether the differentiation defects were secondary to cell death present in Fgfr2 deficient lenses (Garcia et al. 2005). This dissertation reexamined the LeCre mediated deletion of Fgfr2. With this investigation, we reported that

147 Fgfr2-deficiency resulted in lens fiber cell differentiation defects at E12.5, including, lens fiber cells not fully elongating, reduced −γ and βcrystallin expression, cell cycle withdraw failure, and PAX6 retention in the nuclei of posterior lens cells. The reduced −γ and βcrystallin expression, cell cycle withdrawal failure, and PAX6 retention was restored later in development. This suggested that, perhaps much of the fiber cell differentiation defects present with FGFR2- deficiency was a result of delayed differentiation opposed to impaired differentiation. Despite the restoration of differentiation in later stages of FGFR2 deficient lenses, FGFR2 deficient lenses still displayed nuclear retention, low levels of DLAD transcript and Aquaporin0 protein expression. In addition to these fiber cell differentiation defects, FGFR2-deficient lenses displayed enhanced cellular apoptosis throughout lens development. This apoptotic phenotype generated the initial difficulty when concluding that FGFR2 is essential for fiber cell differentiation. Critics claimed that the fiber cell differentiation phenotype may be a secondary effect of apoptosis. Lastly, we examined the impact of deleting Fgfr2 downstream Fgfr signaling, specifically, the phosphorylation of ERK, AKT, and cJun. As expected deleting Fgfr2 resulted in significantly reduced levels of both p-ERK and p-AKT, but also led to an increase in p-cJun.

5.6 Pten deletion restores lens cell survival in FGFR2-deficient lenses.

As Fgfr2 deletion by Lecre results in both apoptosis and fiber cell differentiation defects, and a significant reduction in both AKT and ERK phosphorylation, our initial goal was to additionally delete Pten. By deleting Pten, we hoped to decouple the two main arms of FGFR signaling, PI3K/AKT and MAPK/ERK as the most established role of PTEN is through the inhibition of AKT activation. Moreover, AKT activation primarily results in cell survival pathways and inhibiting apoptotic pathways, and therefore, we hypothesized that deleting Pten in the presence of an Fgfr2 deletion, would permit the isolation of both arms of the Fgfr signaling pathway, and isolate the cell death phenotype away from the differentiation phenotype of FGFR2-deficient lenses. Although our double knockout of Pten and Fgfr2 did not permit the isolation of the FGFR downstream signaling pathways, we did successfully decouple the apoptotic phenotype from the differentiation phenotype. Deleting Pten in the presence of the Fgfr2 deletion resulted in high levels of phosphorylated AKT and ERK, a rescue of cellular survival and size, but it did

148 not restore Aquaporin0 expression, DLAD expression, or the nuclear retention phenotype displayed by FGFR2-deficient lenses. This provides support that signaling through FGFR2 is crucial in the fiber cell differentiation response, which cannot be supplemented by just high AKT and ERK activation levels.

5.7 Lens development does not require PTEN

While characterizing the double deletion of Pten and Fgfr2 by LeCre, we needed to run the experiments alongside an LeCre Pten deletion to avoid the possibility that any lens fiber cell differentiation defects present in the double knockouts was not a result of PTEN loss, but a result of FGFR2 loss. Furthermore, although the physiological function of PTEN in the ocular lens was previously described (Sellitto et al., 2013), the developmental role of PTEN remained elusive. LeCre mediated deletion of Pten resulted in lenses that were indistinguishable from control lenses. Deleting Pten resulted in significant increases in both AKT and ERK activation, yet did not alter lens epithelial cell proliferation nor lens fiber cell differentiation. Over activation of Frs2α by transgenic expression of the TrkC and NT3, lead to enhanced epithelial cell AKT and ERK activation and ectopic fiber cell differentiation in the lens epithelial cell layer (Madakashira et al. 2012). Additionally, transgenic lenses expressing Fgf1, Fgf3, Fgf4, Fgf7, Fgf8, and Fgf9 undergo ectopic differentiation in the lens epithelium (Lovicu and Overbeek, 1998; Robinson et al., 1998; Robinson et al., 1995). Provided the numerous examples of over activation of FGFR signaling leading to ectopic differentiation, and the two main downstream pathways promote AKT and ERK activation, it is surprising that Pten deletion does not promote ectopic differentiation in the epithelial cell layer. Taken together, these results suggest that specifically FGFR signaling through Frs2α leads to fiber cell differentiation, and not just heightened AKT and ERK activation. Another interesting aspect of the PTEN deficient lenses is that Pten deletion did not lead to lens vesicle defects, as it did not prevent the normal apoptosis occurring in between the lens pit and lens vesicle stage. This is in contrast to the role of PTEN in involuting mammary glands and developing lung epithelial cells as PTEN-deficiency enhances cell survival promoting enlarged tissue (Li et al. 2002; Tiozzo 2009). Even within our lens studies, Pten deletion

149 restored cell survival in Fgfr2-deficient lenses, but did not disrupt the normal apoptosis that occurs during the formation of the lens vesicle. Lastly, deleting Pten had a neglible effect on lens epithelial cell proliferation. Deleting Pten, and/or enhancing ERK and AKT activation frequently results in tumor formation and enhanced proliferation (Hertzler-Schaefer et al. 2014). It would be reasonable to hypothesize that Pten deletion would result in high levels of AKT and ERK, and these enhanced levels of AKT and ERK activation would lead to either enhanced proliferation or fiber cell differentiation. In contrast, deleting Pten did not alter lens epithelial cell proliferation or lens fiber cell differentiation. As lens epithelial cells represent one of the only epithelial cell tissue that does not exhibit spontaneous tumor formation, and high levels of AKT and ERK does not lead to enhanced cellular proliferation, the lens epithelium must have several mechanisms to prevent over-proliferation.

5.8 Does lens fiber cell differentiation require Fgfr signaling outside of AKT and ERK?

Increased levels of AKT and ERK1/2 activation did not result in ectopic fiber cell differentiation in the lens epithelium in PTEN-deficient lenses. Moreover, enhancing AKT and ERK activation above the levels of control lenses in the double knockout of Pten and Fgfr2 did not restore DLAD transcript, Aquaporin0 protein, and the nuclear retention phenotype. These data suggests a role of FGFR2 in lens fiber cell differentiation away from AKT and ERK1/2 activation. In 2013, the lab of Ales Cvekl reported 131 FGF responsive miRNAs, which target over 3000 transcripts (Wolf et al., 2013). Additionally, the conditional knockout of Dicer1, which is responsible for miRNA production, initiated the interest in miRNAs in lens development (Li and Piatigorsky, 2009; Wolf et al., 2013). The MLR10 Cre –mediated deletion of Dicer1 resulted in fiber cell elongation defects, fiber cell organization defects, and nuclear retention, which is a phenotype similar of our Le-Cre mediated deletion of Fgfr2 (Wolf et al., 2013). The components of fiber cell differentiation not restored by Pten deletion in FGFR2- deficient lenses may be due to the dependency of differentiation on drastic changes of the miRNA environment stimulated by FGFR signaling. In particular, the FGF-dependent miRNAs may be responsible for Aquaporin0 or expression. To date, only miR-204 has been

150 functionally characterized in the lens (Conte et al., 2010). The precise function of miRNAs in lens and their endogenous control through FGFR signaling remains an important avenue yet to be investigated. It is important to note that the miRNA environment changes in response to FGF2 addition may largely lie downstream of the two traditional arms of FGFR signaling, AKT and ERK. It would interesting to repeat the experiment done in Wolf 2013, and compare the miRNA environment of rLECs following FGF-2 addition to rLECs receiving FGF-2 and AKT and ERK inhibitors. In that way, we could determine which miRNAs change in response to FGF-2 addition, but do not depend on AKT and ERK. I would predict that upon FGF-2 addition while inhibiting both AKT and ERK activation, Aquaporin0 and DLAD expression would remain low. Several of the fiber cell differentiation defects in FGFR2-deficient lenses were restored by Pten deletion-including fiber cell elongation, and early (E12.5) PAX6 removal from differentiating posterior fiber cells and γ-crystallin expression. Presumably, these rescued aspects of fiber cell differentiation are responsive to increased pAKT and pERK1/2 levels. Of particular interest, PAX6 may be FGF-responsive in an ERK1/2 or AKT-dependent manner. Inhibition of FGF signaling by retinoic acid in explanted caudal regions decompacts the chromatin near the PAX6 locus; contrarily, FGF signaling compacts the PAX6 locus inhibiting its expression (Patel et al. 2014). On a protein level of control, PAX6 is a phospho-protein with 4 highly conserved phosphorylation sites, which can be phosphorylated by ERK1/2-which promotes the transactivation of PAX6 (Mikkola et al., 1999). As PAX6 is essential for early lens development, and can act as both an inhibitor and activator of genes associated with lens differentiation, it will would be interesting to see what the requirement of PAX6 phosphorylation has on lens development. Furthermore, as both FGFR signaling and PAX6 regulate lens, retinal, pancreas, and brain development it is crucial to further understand, in an in vivo context, the relationship PAX6 has with FGFR signaling on both its activation through phosphorylation and the transcriptional activation of the Pax6 gene.

151 5.9 References

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