Adenomatous Polyposis Coli Regulates Epithelial Membrane Protein 2 To

ADENOMATOUS POLYPOSIS COLI REGULATES EPITHELIAL MEMBRANE PROTEIN 2 TO

MEDIATE 3D MORPHOGENESIS AND APICAL-BASAL POLARITY

A Dissertation

Submitted to the Graduate School

of the University of Notre Dame

in Partial Fulfillment of the Requirements

for the Degree of

Doctor of Philosophy

Alyssa C. Lesko

Jenifer Prosperi, Director

Graduate Program in Biological Sciences

Notre Dame, Indiana

June 2018

ADENOMATOUS POLYPOSIS COLI REGULATES EPITHELIAL MEMBRANE PROTEIN 2 TO

MEDIATE 3D MORPHOGENESIS AND APICAL-BASAL POLARITY

Abstract

Alyssa C. Lesko

Adenomatous Polyposis Coli (APC) is lost in several epithelial cancers and regulates a variety of normal biological functions including cell migration and apical- basal polarity. Loss of apical-basal polarity disrupts several cellular processes including epithelial structure and intracellular signaling, and is an early marker for tumor development. We previously demonstrated that APC knockdown (APCKD) in Madin-

Darby Canine Kidney (MDCK) cells altered cyst size and inverted polarity in 3D culture.

Through microarray analysis we made the novel observation that APC loss increased

Epithelial Membrane Protein 2 (EMP2) expression. Here we discovered that EMP2 knockdown in APCKD cells decreased cyst size and restored apical polarity. These studies are the first to identify EMP2 as a regulator of apical-basal polarity though the mechanisms downstream of APC and EMP2 that influence polarity and the regulation of

EMP2 by APC are not yet fully understood. With these studies we determined that APC and EMP2 do not control β1 integrin/FAK/Src, Cav1/ERK/JNK, and Scrib/Hippo signaling suggesting a novel downstream mechanism by which APC and EMP2 regulate cyst size Alyssa C. Lesko and polarity. We identified filamin A or plectin isoform X2, histone H3.3, histone H2A type 1-E-like, and 3-hydroxyacyl-CoA dehydrogenase type-2 as possible candidates of

APC/EMP2-mediated polarity. Additionally, we investigated transcriptional regulation of

EMP2 by APC. In parallel studies we identified transcription factors that are predicted to bind the EMP2 promoter and transcription factors that are regulated by APC. Together these data suggested STAT-1 and STAT-3 as potential APC-mediated transcriptional regulators of EMP2. Overall, these studies identified a novel role for EMP2 in controlling apical-basal polarity and have begun to elucidate the molecular mechanisms by which

EMP2 influences tumorigenesis resulting from APC loss. Gaining a better understanding of how APC and EMP2 interact to regulate normal cellular functions will allow us to elucidate the mechanisms by which APC and EMP2 contribute to tumor progression and identify novel therapeutic targets for APC-mutant cancers.

This is for Papa Z, Grandma Barb, Papa Lesko, Grammie Lesko, Mom, Dad, Zane, Karee, and the rest of the Zurovchak and Lesko families who have provided unconditional love,

support, and encouragement.

CONTENTS

Figures ...... vi

Tables ...... viii

Acknowledgments...... ix

Abbreviations ...... xi

Chapter 1: Introduction ...... 1 1.1 Abstract ...... 1 1.2 Apical-basal polarity ...... 3 1.2.1 Apical-basal polarity is required for tissue maintenance and development ...... 3 1.2.2 Apical-basal polarity and disease ...... 11 1.2.3 Models to study apical-basal polarity ...... 14 1.3 Adenomatous Polyposis Coli: normal functions and role in tumorigenesis ... 16 1.3.1 Introduction ...... 16 1.3.2 Loss of functional APC in human cancers ...... 17 1.3.3 Normal functions of the APC tumor suppressor ...... 20 1.3.4 Conclusion ...... 35 1.4 The APC tumor suppressor is required for epithelial cell polarization and three-dimensional morphogenesis ...... 36 1.4.1 Introduction ...... 36 1.4.2 Results ...... 39 1.4.3 Discussion...... 50 1.5 Epithelial Membrane Protein 2 ...... 54

Chapter 2: Epithelial Membrane Protein 2 and β1 integrin signaling regulate APC- mediated processes ...... 56 2.1 Abstract ...... 56 2.2 Introduction ...... 57 2.3 Materials and methods ...... 61 2.3.1 Cell culture ...... 61 2.3.2 Morphological assay ...... 62 2.3.3 Immunofluorescence (IF) ...... 62

iii

2.3.4 Migration assay ...... 63 2.3.5 Western Blotting ...... 63 2.3.6 Real-time PCR ...... 64 2.3.7 Cell adhesion assay ...... 65 2.3.8 Statistics ...... 65 2.4 Results ...... 65 2.4.1 APC loss upregulates Epithelial Membrane Protein 2 and β1 integrin...... 65 2.4.2 APC regulates cyst size through EMP2 and β1 integrin signaling. ... 68 2.4.3 APC regulates apical-basal polarity through EMP2...... 74 2.4.4 APC loss increases cell motility through 1 integrin and Src...... 77 2.5 Discussion ...... 81

Chapter 3: Molecular Mechansims of APC/EMP2-mediated apical-basal polarity ...... 87 3.1 Abstract ...... 87 3.2 Introduction ...... 88 3.3 Materials and Methods ...... 90 3.3.1 Cell culture ...... 90 3.3.2 3D culture IF ...... 91 3.3.3 2D IF ...... 92 3.3.4 Western Blotting ...... 92 3.3.5 Statistics ...... 93 3.4 Results ...... 93 3.4.1 APC and EMP2 do not regulate known downstream signaling pathways of EMP2 ...... 93 3.4.2 Scribble and Hippo are not downstream of APC and EMP2 ...... 97 3.4.3 2D DIGE analysis identified several candidates by which APC and EMP2 may control polarity ...... 102 3.5 Discussion ...... 115

Chapter 4: APC Transcriptional Regulation Of EMP2 ...... 117 4.1 Abstract ...... 117 4.2 Introduction ...... 118 4.3 Materials and Methods ...... 119 4.3.1 Cell Culture ...... 119 4.3.2 ConTra bioinformatics...... 119 4.3.3 Protein/DNA array ...... 119 4.3.4 Firefly and renilla luciferase reporter assays ...... 120 4.3.5 Chromatin immunoprecipitation ...... 120 4.3.6 Statistics ...... 121 4.4 Results ...... 121 4.4.1 Predicted transcriptional regulators of EMP2 ...... 121

4.4.2 APC loss increases the expression and activity of several transcription factors including STAT-1 and STAT-3 ...... 127 4.4.3 Transcription factor binding in the canine EMP2 promoter ...... 129 4.5 Discussion ...... 133

Chapter 5: Conclusions and Future Perspectives ...... 137 5.1 EMP2 independent APC-mediated cell migration ...... 137 5.2 EMP2 as a novel regulator of polarity ...... 138 5.3 Regulation of EMP2 by APC ...... 140 5.4 Tissue specific functions of EMP2 ...... 142 5.5 Summary ...... 144

Bibliography ...... 145

FIGURES

Figure 1.1 APC protein structure...... 21

Figure 1.2 MDCK 3D morphogenesis is perturbed by APC knockdown and restored by reintroduction of APC...... 41

Figure 1.3 Reintroduced APC fragments are detected by immunofluorescence staining for GFP...... 43

Figure 1.4 APC knockdown disrupts MDCK cell polarity in cysts grown 3D culture...... 45

Figure 1.5 An APC-knockdown gene signature is associated with altered cell-cell and cell- matrix communication...... 49

Figure 2.1 APC loss upregulates Epithelial Membrane Protein 2 and β1 integrin...... 67

Figure 2.2 EMP2 knockdown and inhibition of β1 integrin signaling decreases size of APC shRNA cysts...... 70

Figure 2.3 Proliferation and apoptosis are not affected by APC loss...... 71

Figure 2.4 Chemical inhibitors target β1 integrin signaling pathway...... 72

Figure 2.5 Apical polarity in APC shRNA cysts is restored by EMP2 knockdown...... 76

Figure 2.6 Increased migration in APC shRNA cells is controlled by β1 integrin signaling...... 79

Figure 2.7 Schematic of APC-mediated mechanisms...... 82

Figure 3.1 APC and EMP2 do not regulate Cav1/ERK/JNK signaling...... 95

Figure 3.2 Scribble localization and expression is unchanged by APC loss...... 98

Figure 3.3 YAP is not localized to the nucleus of MDCK cells with or without APC loss. 100

Figure 3.4 Representative 2D DIGE images...... 104

vi Figure 3.5 APC loss does not increase filamin or plectin expression...... 112

Figure 4.1 Predicted transcription factor binding sites in the canine EMP2 promoter. 124

Figure 4.2 APC loss increases the expression and activation of several transcription factors...... 128

Figure 4.3 CREB binds the EMP2 promoter...... 131

Figure 4.4 EMP2 promoter driven luciferase reporter plasmid...... 136

vii

TABLES

Table 3.1 2D DIGE proteins with changed expression upon APC loss ...... 106

Table 3.2 2D DIGE proteins specific to APC regulation ...... 108

Table 3.3 2D DIGE proteins specific to APC and EMP2 regulation ...... 109

Table 3.4 2D DIGE candidates identified by mass spectrometry ...... 110

Table 3.5 Pathway Enrichent Analysis of 2D DIGE data ...... 114

Table 4.1 Predicted Binding Sequences in the EMP2 promoter of Several Transcription Factors ...... 126

Table 4.2 CREB ChIP experiment input ratios ...... 132

viii

ACKNOWLEDGMENTS

I would like to acknowledge my advisor Dr. Jeni Prosperi for giving me the opportunity to grow as a scientist during my time in her lab and for her support and guidance throughout this project and my graduate education. Thank you to all of the

Prosperi lab members past and present for creating a great lab environment, thoughtful scientific discussions, and experimental support. I would like to specifically thank

Carolyn Ahlers, an undergraduate student who worked closely with me for three years for her contributions to this project. Thank you to Dr. Monica VanKlompenberg and

Casey Stefanski for being wonderful lab mates and friends who provided thoughtful advice and many laughs during late nights in the lab.

Thank you to my dissertation committee members: Dr. Rebecca Wingert, Dr.

Holly Goodson, Dr. Kevin Vaughan, and Dr. Reginald Hill for improving my research as well as professional development through scientific feedback and support of fellowship applications. I would also like to thank our collaborator Dr. Madhuri Wadehra for the use of EMP2 reagents and sharing her advice and expertise on EMP2, and my co-advisor

Dr. Jeremy Zartman for his guidance especially on the migration and cytoskeletal aspects of this project. Thank you to the faculty, administration, staff, and members of

Harper Cancer Research Institute (HCRI) for providing a collaborative and engaging research environment. ix

My dissertation research was made possible by funding from the

Interdisciplinary Interface Training Program through HCRI.

I would also like to thank my friends and family who have made a great impact on my graduate education. Ian Guldner and Dr. Josh Mason thank you for being great friends and peer mentors throughout our time together at Notre Dame. Thank you to my grandparents for always supporting and encouraging me. Thank you Mom and Dad for always pushing me to be my best, providing me with every opportunity for success, and believing in me even when others did not. Without your love and unconditional support I would not be where I am today. Zane and Karee thank you for following me to the Midwest and for always being there to listen to my hardships and celebrate my triumphs. Kate and Mason I am grateful that we were able to complete our journeys together at the same university. Finally, I would like to thank the rest of the Zurovchak and Lesko families for their encouragement, inspiration, and visits especially in the fall.

ABBREVIATIONS

ADPKD Autosomal dominant polycystic kidney disease

AKT Protein kinase B

AMPK AMP-activated protein kinase

AP-1 Activator protein 1

APC Adenomatous polyposis coli aPKC Atypical protein kinase C

Cav1 Caveolin 1

ChIP Chromatin immunoprecipitation

Crb Crumbs

CREB cAMP responsive element binding protein 1

DIGE Difference gel electrophoresis (DIGE)

Dlg Discs large

E2F-1 E2F transcription factor 1

ECM Extracellular matrix

EGFR Epidermal growth factor receptor

EMP2 Epithelial membrane protein 2

EMT Epithelial to mesenchymal transition

ERBB2 Erythroblastic oncogene B xi

ERK Extracellular signal-regulated kinase

FAK Focal adhesion kinase

FAP Familial adenomatous polyposis

GAS Gamma interferon activation sites

GPI-APS Glycosylphosphatidyl inositol-anchored proteins

ISRE Interferon stimulated response element

JAM Junctional adhesion molecules

JNK Jun N-terminal kinase

Lats Large tumor suppressor kinase

Lgl Lethal giant larvae

LKB1 Liver kinase B 1

MDCK Madin darby canine kidney

MET Mesenchymal to epithelial transition

NFAT-1 Nuclear factor of activated T-cells

NF-κB Nuclear factor kappa-light-chain-enhancer of activated B cells

PALS1 Protein associated with Lin-7

Par Partition-defective

PATJ PALS1-associated tight junction protein

PAX2 Paired box gene 2

PDZ Postsynaptic density/discs Large/zonula occludens

PI3K Phosphoinositide 3-kinsase

PPAR Peroxisome proliferator-activated receptor xii

Sav Salvador

Scrib Scribble

Sp-1 Specificity protein 1

STAT Signal transducer and activator of transcription

TGFβ Transforming growth factor beta

YAP Yes-associated protein

YY1 Yin and yan 1

xiii

CHAPTER 1:

INTRODUCTION

This text appears in part in publication: Lesko AC, Goss KH, and Prosperi JR

(2014). Exploiting APC Function as a Novel Cancer Therapy. Current Drug Targets, 15:

90-102. And in part in publication: Lesko AC, Goss KH, Yang FF, Schwertner A, Hulur I,

Onel K, and Prosperi JR (2015). The APC tumor suppressor is required for epithelial cell polarization and three-dimensional morphogenesis. BBA-Molecular Cell Research (ESL),

1854(3): 711-723.

1.1 Abstract

Tissue structure is maintained by interactions between epithelial cells and the establishment of proper apical-basal polarity. Apical-basal polarity is regulated by three core polarity complexes: partition-defective (Par) 3/Par6/atypical protein kinase C

(aPKC), Crumbs (Crb)/protein associated with Lin-7 (PALS1)/PALS1-associated tight junction protein (PATJ), and Scribble (Scrib)/Lethal giant larvae (Lgl)/Discs large (Dlg).

Loss of these complexes and subsequently cell polarity disrupts normal epithelial structure and can lead to disease progression including chronic kidney disease and tumorigenesis. Therefore, it is critical to identify the molecular mechanisms underlying the maintenance of polarity to better understand how these processes are hijacked in 1

disease states. In this thesis, we investigate the role of the tumor suppressor

Adenomatous Polyposis Coli (APC) in the regulation of apical-basal polarity.

APC is most commonly mutated in colorectal cancers such as familial adenomatous polyposis (FAP); as well as many other epithelial cancers like breast, pancreatic, and lung cancer. APC mutations usually result in a truncated form of the protein lacking the Carboxy-terminal region resulting in loss of function. Mutations in

APC have been identified in early stages of cancer development making it a gatekeeper of tumor progression and therefore an ideal therapeutic target. APC is best known for its role as a negative regulator of the Wnt/β-catenin pathway. However, APC also mediates several other normal cell functions independently of Wnt/β-catenin signaling such as: apical-basal polarity, microtubule networks, cell cycle, DNA replication and repair, and cell migration. Our lab implicated APC in the control of apical-basal polarity by generating APC knockdown epithelial cell lines. APC depletion resulted in loss of polarity and enlarged, filled spheroids with disrupted polarity in 3D culture.

Interestingly, these effects of APC knockdown were rescued with either full-length or a

Carboxy (C)-terminal segment of APC. Moreover, we identified a gene expression signature associated with APC knockdown that points to several candidates including the tetraspan membrane protein Epithelial Membrane Protein 2 (EMP2). EMP2 functions to maintain plasma membrane composition and changes in EMP2 expression are associated with several epithelial cancers. These data suggest that the initiation of epithelial-derived tumors as a result of APC mutation or gene silencing may be driven by increased EMP2 expression, loss of polarity, and dysmorphogenesis. This chapter will 2

discuss apical-basal polarity, the normal functions of APC, the role of APC in mediating polarity, and an overview of EMP2.

1.2 Apical-basal polarity

1.2.1 Apical-basal polarity is required for tissue maintenance and development

1.2.1.1 Tissue structure and cell interactions

Epithelial cells form a uniform layer with an established asymmetrical apical- basal polarity that is essential for cell-cell interactions, signaling, and tissue organization

(O'Brien et al. 2002, Debnath and Brugge 2005, Datta et al. 2011, Pieczynski and

Margolis 2011, Bazzoun et al. 2013, Rodriguez-Boulan and Macara 2014). For example, apical-basal polarity is required for renal tubule formation (O'Brien et al. 2002, Sawyer et al. 2010), mammary duct formation (Debnath and Brugge 2005), gastrulation (Sawyer et al. 2010), the proper division of cells (Lechler and Fuchs 2005), cell movement and signaling though the proper orientation of cilia (Singla and Reiter 2006), polarization of the cytoskeleton to mediate intracellular signaling, cellular transport, cell adhesion and cell migration (Bazzoun et al. 2013), and epithelial to mesenchymal transition (EMT) and mesenchymal to epithelial transition (MET) which are required for development and wound-healing ((Savagner 2001, Mikawa et al. 2004, Nelson 2009, Lim and Thiery 2012) and reviewed (Rodriguez-Boulan and Macara 2014)). Cells interact with each other through junctions including gap junctions and the apical junctional complex, which is comprised of the tight junction and adherens junction (Bazzoun et al. 2013). These 3

junctional complexes are important for cell-cell adhesion and communication as well as cell environment interactions. Therefore many of the junctional proteins (described in detail below) are transmembrane proteins (Bazzoun et al. 2013). The adherens junction regulates signaling and cytoskeletal organization, and is the primary mediator of cell adhesion and the formation of other junctions (Capaldo and Macara 2007). Tight junctions control the diffusion of molecules between cells and act as the border by which the apical and basal surface are defined (Perez-Moreno et al. 2003, Tamura et al.

2008, Schluter and Margolis 2012, Bazzoun et al. 2013). Finally, gap junctions mediate action potential signaling, exchange of nutrients between cells and the environment, and second messenger uptake for intracellular signaling (Talhouk et al. 2005, Talhouk et al. 2008).

In mature epithelial organs, the apical surface lines the lumen while the basal surface faces outward toward the extracellular matrix (ECM) (Bornens 2008, Fedeles and Gallagher 2013). Lumen formation is dependent on apical-basal polarity as cues from the environment are needed to feed through a polarized cytoskeleton via tight junction complexes (Ivanov et al. 2005, Wu and Crabtree 2007, Datta et al. 2011, Terry et al. 2011). Given the importance of proper apical-basal polarity, this process is highly regulated and is maintained by several polarity and junctional protein complexes which are described in detail below.

1.2.1.2 Polarity and junctional complexes

Apical-basal polarity is regulated by three core polarity complexes,

Par3/Par6/aPKC, Crb/PALS1/PATJ, and Scrib/Lgl/Dlg. They act by antagonizing one another to mutually exclude and restrict proteins to the correct cellular surface

(reviewed in (Lee and Vasioukhin 2008, Martin-Belmonte and Perez-Moreno 2012,

Fedeles and Gallagher 2013, Rodriguez-Boulan and Macara 2014)).

1.2.1.2.1 Par3/Par6/aPKC

Par3 is localized to the apical tight junction where it binds directly to Par6 through its Postsynaptic Density/Discs Large/Zonula Occludens (PDZ) binding domain and subsequently recruits aPKC to the apical complex (Etemad-Moghadam et al. 1995,

Macara 2004, Lechler and Fuchs 2005, Suzuki and Ohno 2006). Upon phosphorylation of aPKC by the Rho GTPase cdc42, the tight junctions develop (Ebnet et al. 2004, Bradfield et al. 2007, Weber et al. 2007). aPKC is an apical kinase that excludes basolateral proteins by phosphorylation and therefore can phosphorylate Par3 to localize it to the lateral adherens junction freeing Par6 which can now bind to Crb or Lgl (Betschinger et al. 2003, Plant et al. 2003).

Par1 is another protein kinase however it is localized to the basolateral membrane and conversely to aPKC excludes apical proteins (Rodriguez-Boulan and

Macara 2014). In this way Par1 can phosphorylate Par3 to localize it to the lateral membrane where Par5, a cytoplasmic phospho-protein interactor, binds to Par3 to release it from the lateral membrane into the cytoplasm. Par3 can now be

dephosphorylated (Benton and St Johnston 2003). Par1 is regulated by aPKC as aPKC excludes Par1 from the apical surface by direct phosphorylation (Hurov et al. 2004,

Kusakabe and Nishida 2004, Suzuki et al. 2004). Similarly to Par3, once phosphorylated

Par1 can bind to Par5 and this complex dissociates form the membrane into the cytoplasm where Par1 is subsequently dephosphorylated (Rodriguez-Boulan and Macara

2014).

Par4, or the mammalian orthologue liver kinase B1 (LKB1), is a serine/threonine kinase whose primary target is AMP-activated protein kinase (AMPK) through which

LKB1 regulates cell metabolism (Williams and Brenman 2008). LKB1 was first identified as a tumor suppressor in Peutz-Jeghers syndrome which is characterized by tumors in the gastrointestinal tract (Hemminki et al. 1998), and is implicated in several other epithelial cancers including lung, cervical, ovarian, and breast (reviewed in (Martin-

Belmonte and Perez-Moreno 2012)). In Drosophila Par4 regulates the first asymmetric divisions during embryonic development and also phosphorylates Par1 to mediate cell polarity (Kemphues et al. 1988, Martin and St Johnston 2003, Mirouse and Billaud

2011). Furthermore intestinal epithelial cells lacking junctional cell interactions were able to polarize upon LKB1 activation (Baas et al. 2004). These interactions demonstrate the complex regulatory mechanisms underlying polarity maintenance.

1.2.1.2.2 Crb/PALS1/PATJ

The Crb/PALS1/PATJ complex is localized to the apical surface as well as cell junctions (Margolis and Borg 2005, Suzuki and Ohno 2006, Olsen et al. 2007). Crb is a

transmembrane protein, while PALS1 and PATJ are both cytoplasmic scaffolding proteins (Assemat et al. 2008). There are three mammalian homologues of Crb. Crb1 is expressed in retina and brain, Crb2 is expressed in the retina, brain and kidney, and Crb3 is the predominant form expressed in all epithelial tissues (den Hollander et al. 2001,

Makarova et al. 2003, Lemmers et al. 2004, Fogg et al. 2005, van den Hurk et al. 2005).

Crb3 defines the apical membrane and has a role in tight junction formation as MCF-10A cells, normal mammary cells which express low Crb levels, do not form tight junctions; however, overexpression of Crb is able to induce tight junction formation and polarization (Fogg et al. 2005). Additionally, overexpression of Crb3 in Madin Darby

Canine Kidney (MDCK) cells, normal kidney cells, delayed tight junction formation (Roh and Margolis 2003, Lemmers et al. 2004). Crb3 is also important for cilia development.

Crb3 can be found localized in puncta of cilia of MDCK cells and loss of Crb3 prevented cilia formation in these cells (Fan et al. 2004).

Crb directly binds PALS1 and PATJ through its PDZ domain (Makarova et al.

2003). PALS1 contributes to tight junction formation as knockdown of PALS1 in MDCK cells disrupted tight junction formation and polarity and led to mistargeting of E- cadherin to the cell membrane (Straight et al. 2004, Wang et al. 2007). PALS1 and PATJ expression appear to be dependent on one another. Loss of PALS1 decreased PATJ expression (Straight et al. 2004, Wang et al. 2007), and PALS1 expression is dependent on its interactions with PatJ (Michel et al. 2005). The downregulation of PATJ leads to mislocalization of the tight junction proteins ZO1, ZO3, and occludin (Lemmers et al.

2002, Michel et al. 2005, Shin et al. 2005). Additionally, loss of PATJ resulted in loss of 7

both Crb3 and PALS1 from the apical membrane suggesting PATJ is required for the targeting of the Crb complex to the apical membrane through interactions and stability of tight junctions and for the maintenance of apical membrane identity (Roh et al. 2002,

Michel et al. 2005). Crb can interact with other polarity complexes by binding Par6 directly or indirectly via PALS1 (Hurd et al. 2003, Lemmers et al. 2004, Wang et al. 2004).

Overexpression of Par6 and Crb3 delayed tight junction formation in MDCK cells (Gao et al. 2002, Lemmers et al. 2004) suggesting this interaction is critical for tight junction assembly.

1.2.1.2.3 Scrib/Lgl/Dlg

The Scrib/Lgl/Dlg complex is localized to the basolateral membrane (Bilder 2004,

Assemat et al. 2008). The main goal of this complex is to establish basolateral identity by antagonizing the apical Par complex ((Humbert et al. 2006, Assemat et al. 2008) and reviewed in (Rodriguez-Boulan and Macara 2014)). Scribble is a cytoplasmic protein mostly associated with and targeted to the adherens junction (Navarro et al. 2005, Qin et al. 2005). The targeting of Scrib and E-cadherin to the adherens junction is dependent on their interaction with each other (Navarro et al. 2005, Qin et al. 2005). Furthermore, suppression of Scrib in MDCK cells disrupted adhesion by dissociating E-cadherin and catenin complexes causing these MDCK cells to acquire a mesenchymal phenotype and exhibit increased migration (Qin et al. 2005).

Par6 directly interacts with Lgl to promote phosphorylation of Lgl by aPKC. Upon phosphorylation by aPKC, Lgl is localized to lateral membrane right below the adherens

junction (Musch et al. 2002, Betschinger et al. 2003, Plant et al. 2003, Yamanaka et al.

2003, Kallay et al. 2006), and this localization is critical for the establishment of polarity during the early stages of polarization (Yamanaka et al. 2003). Additionally, studies have established a role for the Lgl3 isoform in exocytosis of core vesicles and the more prominent mammalian isoform Lgl1 can interact with syntaxin4, a protein that mediates vesicle fusion with the plasma membrane, suggesting Lgl can also mediate polarity through regulation of basolateral exocytosis (Musch et al. 2002).

Although there are five homologues of Dlg in mammalian cells, Dlg1 is most similar to Drosophila Dlg and is the most studied in epithelial tissues (McLaughlin et al.

2002). Dlg is essential for tight junction formation and is localized to the lateral membrane through several interactions (Bohl et al. 2007). Dlg can bind to matrix metallopeptidase 9 (MMP-9) and Lin7, involved in delivery and recycling of proteins to plasma membrane, to stabilize its localization at tight junctions (Bohl et al. 2007). Dlg can also interaction with Lin2/hCask, a calmodulin associated serine/threonine kinase, which distributes Dlg throughout the lateral membrane (Lee et al. 2002). Additionally,

Dlg can interact with 4.1, a cytoskeletal stabilizing protein (Lue et al. 1996), and loss of

Dlg prevents the recruitment of both PI3K to cell-cell contacts and disrupts cytoskeletal organization (Laprise et al. 2002). In opposing but complimentary functions to the Par proteins, the Scrib/Lgl/Dlg complex establishes the identity of the basal membrane.

1.2.1.2.4 Adherens junction complexes: cadherins and catenins

Adherens junction formation is dependent on E-cadherin which forms cadherin- catenin clusters (Gumbiner 2005). Cadherins are transmembrane proteins which form weak cell adhesions by the interaction of the cadherin extracellular domain with the extracellular domain of cadherin of an adjacent cell (Gumbiner 2000, Chen et al. 2005)

Stronger adhesions are formed by the lateral clustering of cadherins which is mediated by catenins and the cytoskeleton (Aberle et al. 1994, Huber and Weis 2001). Cadherins are linked indirectly to the cytoskeleton through interactions with β-catenin and α- catenin (Aberle et al. 1994, Huber and Weis 2001). E-cadherin also mediates intracellular signaling through activation of MAPK signaling (Pece and Gutkind 2000), and negative regulation of Wnt signaling by recruiting β-catenin to cell junctions and away from the nucleus (Wijnhoven et al. 2000). Furthermore, loss of E-cadherin mislocalizes Crb (Navarro et al. 2005) suggesting a role for cadherins in maintaining polarity in addition to maintaining tissue structure.

1.2.1.2.5 Tight junction complexes: claudins, occludins, junctional adhesion molecules

(JAM), and zonula occludens

Tight junctions are composed of claudins, occludins, and junctional adhesion molecules (JAM) which are all transmembrane proteins that interact with cytoskeletal connectors including zonula occludens (Chiba et al. 2008). Claudins function to control the ion permeability of the membrane and also recruit occludins to the tight junctions

(Wu et al. 1998). Occludins help regulate the movement of molecules between adjacent

cells and interact with several intracellular signaling proteins including transforming growth factor (TGFβ) (Barrios-Rodiles et al. 2005, Martin and Jiang 2009). JAMs regulate epithelial cell adhesion to the extracellular matrix (ECM) and morphology through β1- integrin and Rap1 GTPase activity (Mandell et al. 2005). Finally, zonula occludins have been shown to be a possible link between tight junctions and adherens junctions and serve as scaffolding proteins to connect other cytoplasmic junctional proteins

(Utepbergenov et al. 2006). Without tight junction proteins, cells are unable to receive cues from the environment in order to establish the correct apical-basal polarity.

1.2.2 Apical-basal polarity and disease

Epithelial to mesenchymal transition (EMT) is characterized by loss of both tight and apical junctions, loss of apical-basal polarity, and loss of expression of epithelial proteins such as E-cadherin, and increase in mesenchymal markers (Macara et al. 2014).

This transition is essential in normal development for cells to differentiate and migrate, and cells will undergo the reverse process of mesenchymal to epithelial transition (MET) in order to form the correct epithelial tissue structure. Unfortunately, the process of

EMT can be hijacked by mutant cells during disease states to disrupt normal epithelial morphology and promote malignant phenotypes such as kidney disease and tumorigenesis (Chandramouly et al. 2007, Wodarz and Nathke 2007, Lee and Vasioukhin

2008, Lim and Thiery 2012, Martin-Belmonte and Perez-Moreno 2012, Bazzoun et al.

2013).

1.2.2.1 Loss of polarity in kidney disease

Apical-basal polarity is important for normal kidney function as many transporters and aquaporin channels are normally apically localized and polarization is required for ciliogenesis ((Wilson et al. 1991, Orellana et al. 1995) and reviewed in

(Wilson 2011)). Loss of polarity can lead to kidney disease as a result of mislocalization of transporters and subsequent increased proliferation and secretion of fluid into the lumen leading to organ malfunction (Wilson 2011). Additionally, disruption of cilia function due to loss of apical-basal polarity can lead to formation of cysts, fibrosis, and subsequently kidney disease ((Wilson et al. 1991, Orellana et al. 1995) and reviewed in

(Wilson 2011)).

Polycystin-1, encoded by the PKD1 gene and commonly mutated in polycystic kidney disease (PKD), interacts with the Par complex and contributes to polarization and tubulogenesis (Castelli et al. 2013). Interestingly, mice with podocyte specific knockout of aPKC develop nephrotic syndrome and glomerular dysfunction resulting in death shortly after birth (Huber et al. 2009). Similarly, mice with deletion of one allele of

PALS1 develop cyst formation and lose kidney function resulting in death within 4-6 weeks of birth (Weide et al. 2017). Furthermore, Scrib overexpression in a zebrafish autosomal-dominant polycystic kidney disease (ADPKD) model reduced cyst formation that was induced by pkd morphants and Scrib is down regulated in ADPKD cell lines (Xu et al. 2018). Mice with knockout of MALS3 (mammalian homologue of Lin7), which binds both Crb and Dlg complexes to stabilize them at cell junctions and maintain polarity, develop cystic and fibrotic kidneys (Olsen et al. 2007). These studies provide a role for 12

polarity disruption in renal cyst formation and kidney disease progression and identify polarity complexes and their downstream mechanisms as possible therapeutic targets.

1.2.2.2 Loss of polarity in breast cancer

The structure of kidney tubules is very similar to that of mammary ducts as they both form hollow, polarized spherical structures. Given the physical similarities between kidney and mammary epithelial tissues, the mechanisms underlying the loss of polarity in kidney disease may be similar to those in breast cancer. Loss of polarity has been well studied in the context of breast cancer though polarity loss has also been implicated in several other epithelial cancers (reviewed in (Martin-Belmonte and Perez-Moreno

2012)). Loss of Par3 in conjunction with activation of Notch or hRAS induced mouse mammary tumors and increased invasion and metastasis through activation of aPKC

(McCaffrey et al. 2012). Additionally, Par6 is upregulated in breast cancer cell lines as well as precancerous and advanced stage breast cancer tissues. Interestingly, Par6 upregulation alone is sufficient to induce proliferation and hyperplasia in mammary epithelial cells grown in a 3D in vitro model system (Nolan et al. 2008). Activation of aPKCζ is expressed in more mesenchymal breast cancer cell lines as well as human breast cancer tissues. Furthermore, depletion of aPKCζ increased E-cadherin expression in the metastatic MDA-MB-231 breast cancer cell line and decreased metastasis in MDA-

MB-231 xenograft mouse models (Paul et al. 2015). LKB1, or Par4, is well known as a tumor suppressor and studies have shown that LKB1 is downregulated in several breast cancer cell lines leading to increased cell migration and invasion and disrupted polarity

in 3D culture (Li et al. 2014). Loss of LKB1 is also associated with poor prognosis in human breast cancer patients (Shen et al. 2002).

Like the Par complex the other apical complex Crb/PATJ/PALS1 and the lateral polarity complex Scrib/Lgl/Dlg have also been implicated in tumorigenesis. A multipotent cancer stem cell phenotype and induction of EMT was caused by knockdown of Crb3 in MCF-10A cells (Li et al. 2017). Loss or mislocalization of Scrib induced mammary tumor formation in mouse mammary fat pads (Zhan et al. 2008), and human tissue from lobular carcinoma exhibits downregulated Scrib expression (Navarro et al. 2005). Activated ERBB2 binds directly to Par6 and disrupts apical-basal polarity and promotes proliferation in breast tumors (Muthuswamy et al. 2001, Aranda et al. 2006,

Guo et al. 2006), and dissociates Par3 from Par6/aPKC complex to provide protection from apoptosis (Aranda et al. 2006). Together these studies show loss of polarity proteins leads to tumorigenic phenotypes such as disrupted morphology and increased mesenchymal characteristics in mammary tissue and identify polarity complexes as important tumor suppressors through the maintenance of epithelial structure.

1.2.3 Models to study apical-basal polarity

To identify novel therapeutic targets, it is important to understand the mechanisms by which cells normally regulate apical-basal polarity and how these mechanisms are disrupted in disease states. Several in vitro methods have been developed to model epithelial polarity to better understand the molecular mechanisms that mediate this process. Although 2D assays like calcium switch studies and transwell

filters provide a good platform to study cell-cell interactions and junction formation, there are many limitations and these assays are not representative of in vivo conditions.

When cultured on plastic or a transwell filter the apical (media) and basal membranes

(plastic) are already predefined (Schluter and Margolis 2009). 2D assays are also unable to replicate the 3D tissue structure of epithelial organs. Most epithelial tissues are made up of cysts, hollow spherical structures like mammary acini or ducts and lung alveoli, and tubules, hollow cylindrical structures like the tubules of the nephrons in the kidney

(O'Brien et al. 2002).

β1 integrin and ECM interactions are critical for the establishment of polarity and correct tissue structure in vivo based on signals from the environment (Klein et al. 1988,

Sorokin et al. 1990, Howlett et al. 1995, Weaver et al. 2002, Cohen et al. 2004, Yu et al.

2005, Lee and Vasioukhin 2008, Myllymaki et al. 2011, Pieczynski and Margolis 2011).

Therefore, two methods have been developed where cells are either embedded or overlayed in ECM depending on the cell type. For example, MDCK cells are often grown in collagen because they can secrete their own laminin while MCF-10A cells require exogenous laminin and therefore are grown in Matrigel (O'Brien et al. 2001, Debnath and Brugge 2005). The need for β1 integrin and ECM is demonstrated by growing MDCK cells in suspension. Although they will from cysts, the apical surface will face the outside in contact with media and lumens will not form. If these cells are put into collagen they will repolarize and form a lumen with the apical surface facing the inside (Chambard et al. 1984, Wang et al. 1990, Wang et al. 1990). Although these cysts form spontaneously when cells are cultured in ECM, tubulogenesis can also be induced in MDCK cells by 15

treating with HGF (Montesano et al. 1991, Soriano et al. 1995, Pollack et al. 1998,

O'Brien et al. 2002). Therefore 3D culture model systems have proven to be a very useful system to study the mechanisms that establish and maintain epithelial cell polarity that resembles in vivo tissue structures.

Not only do these models allow for studies to better understand the mechanisms that regulate normal cellular functions but they can be exploited to determine how these mechanisms are disrupted during disease. For example, understanding how cells invade collectively during tubulogenesis can lead to insights into EMT mechanisms as cells migrate in a similar manner (O'Brien et al. 2002, Debnath and Brugge 2005). In our studies, we utilize the MDCK 3D Matrigel culture cell model to understand how the tumor suppressor Adenomatous Polyposis Coli (APC) regulates apical-basal polarity and

3D morphogenesis.

1.3 Adenomatous Polyposis Coli: normal functions and role in tumorigenesis

1.3.1 Introduction

Mutations in the Adenomatous Polyposis Coli (APC) tumor suppressor gene are most commonly found in colorectal cancers such as the inherited syndrome familial adenomatous polyposis (FAP), but have also been identified in many epithelial cancers including breast cancer ((Herrera et al. 1986, Tanaka et al. 1989, Leppert et al. 1990) and reviewed in (Prosperi 2011)). Most APC mutations are nonsense mutations, frequently created by frame-shifts, resulting in premature stop codons and a truncated gene product lacking the Carboxy-terminus of the protein. Patients with FAP inherit one 16

germline mutation and develop tumors when another somatic mutation is received resulting in loss of the wild-type APC allele. In this way, APC mutation follows the classical ‘two-hit’ model. The disease presentation in patients with APC mutation often depends on the location of the mutation. For example, mutations at the 5’ and 3’ ends of the coding sequence are associated with a weakened FAP phenotype, while extracolonic phenotypes are associated with mutations in other regions of the sequence

((Kinzler and Vogelstein 1996) and reviewed in (Goss and Groden 2000)). Importantly,

APC mutations similar to those found in FAP have been identified in 50-80% of sporadic colon adenomas and adenocarcinomas (reviewed in (Segditsas and Tomlinson 2006,

Prosperi 2011)). Since APC has been identified in the earliest stages of tumor progression (Fishel et al. 1993), it has emerged as the ‘gatekeeper’ of colorectal cancer development. This section will describe the normal functions of APC as they relate to

APC-mutant tumors and will discuss the importance of APC in a variety of epithelial- derived tumors.

1.3.2 Loss of functional APC in human cancers

Modifications of APC are most commonly found in hereditary and sporadic colorectal cancers. Depending on the type of analysis performed or the subset of tumors investigated, 25%-88% of sporadic colorectal adenomas and 31%-83% of adenocarcinomas are a result of APC inactivation (Powell et al. 1992, Miyaki et al. 1994,

Yashima et al. 1994, Huang et al. 1996, Konishi et al. 1996, Yu and Wang 1998, Rowan et al. 2000, Fukushima et al. 2001, De Filippo et al. 2002, Miyoshi et al. 2002, Tsai et al.

2002, Diergaarde et al. 2003, Luchtenborg et al. 2005). The majority of somatic mutations of APC in FAP and sporadic colorectal cancers occur in the mutation cluster region (MCR) in the middle portion of APC where β-catenin binding and downregulation occurs (reviewed in (Miyoshi et al. 1992, Goss and Groden 2000).

Interestingly, mutations in the MCR causing allelic loss are often found in colorectal cancers; whereas mutations outside the MCR are commonly found in breast cancers

(Lamlum et al. 1999, Rowan et al. 2000). APC loss can also be the result of hypermethylation of the promoter and epigenetic silencing of APC, and has been observed in many gastrointestinal and non-gastrointestinal tumor types (Hiltunen et al.

1997, Esteller et al. 2000, Arnold et al. 2004, Lind et al. 2004, Segditsas et al. 2008).

Specifically in breast cancer, both epigenetic and genetic alterations in APC have also been identified. APC mutations that regulate Wnt signaling and result in the accumulation of cytosolic and nuclear -catenin have been identified in breast cancer

(Jonsson et al. 2000, Abraham et al. 2002, Ozaki et al. 2005); however, most APC mutations associated with sporadic human breast cancer occur outside the MCR and function independently of the Wnt pathway to cause cancer progression (Kashiwaba et al. 1994, Wada et al. 1997, Furuuchi et al. 2000, Abraham et al. 2002). While APC mutations occur in a subset of breast cancers, the most common method of APC inactivation in breast cancer is promoter methylation (Jin et al. 2001, Virmani et al.

2001, Sarrio et al. 2003, Dulaimi et al. 2004, Prasad et al. 2008, Van der Auwera et al.

2008). Recent studies demonstrated that alterations of APC, both by deletion or

methylation, occur more frequently in ER-/PR- breast cancers, and are associated with higher tumor grade and poor survival (Mukherjee et al. 2012).

Interestingly, APC inactivation occurs at different frequency in specific breast cancer subtypes. For example, 50% of lobular carcinomas (Sarrio et al. 2003), and 70% of inflammatory breast cancers exhibit promoter methylation and silencing of APC (Van der Auwera et al. 2008). Similarly, subtype specificity has been observed with -catenin cytosolic and nuclear accumulation (Khramtsov et al. 2010, Lopez-Knowles et al.); however, it is unclear whether the subtype specificity of APC inactivation and -catenin nuclear accumulation are related. Exploring this relationship could lead to potential therapeutic treatments because it appears that APC functions separately from the

Wnt/-catenin pathway to suppress tumor activity (Phelps et al. 2009 for review,

Prosperi et al. 2009). Not only does the frequency of APC inactivation help to determine the cancer phenotype, but it may also be associated with certain clinical parameters such as tumor stage and size, poor prognosis, and overall survival (Kashiwaba et al.

1994, Jin et al. 2001, Virmani et al. 2001, Prasad et al. 2008).

Although mutations and silencing of APC are most commonly found in colorectal cancer, loss of APC also occurs in several other epithelial cancers (reviewed in (Prosperi

2011)). Briefly, APC promoter methylation has been found in 95% of lung cancers

(Brabender et al. 2001, Usadel et al. 2002), and 90% of prostate cancers (Jeronimo et al.

2004, Yegnasubramanian et al. 2004, Bastian et al. 2005). APC mutations have been detected in 18% of pancreatic acinar cell carcinomas (Abraham et al. 2002), and have been associated with pancreatoblastomas (Abraham et al. 2001, Chetty et al. 2006). 19

Recently, APC loss by missense or frameshift mutations has been identified in 16% of invasive urothelial cancers (Kastritis et al. 2009). Biallelic deletion has been detected in

33% of ovarian cancers (Sarrio et al. 2006), and in 17% of melanoma tumors

(Korabiowska et al. 2004, Worm et al. 2004, Castiglia et al. 2008). These data collectively suggest that APC mutations likely contribute to the development of diverse types of cancer, suggesting that treatments that specifically target APC status may be applicable to a variety of patients.

1.3.3 Normal functions of the APC tumor suppressor

APC is a large multifunctional protein made up of 2843 amino acids and multiple binding domains. While the best-known role of APC is to act as a negative regulator of the Wnt/-catenin signaling pathway, there are many other normal functions of APC.

Based on the ability of APC to bind a variety of protein partners, other activities of APC include mediating cell migration, regulation of apical-basal polarity, microtubule networks, cell cycle, DNA replication and repair, and apoptosis (summarized in Figure

1.1). These functions of APC, including regulation of Wnt/β-catenin signaling and other non-Wnt related functions have been reviewed elsewhere (Prosperi and Goss 2011,

Nelson and Nathke 2013). Here we will briefly summarize some key functions of APC that may contribute to its tumor suppressive function.

20 21

Figure 1.1 APC protein structure. Specific APC functional domains are shown and labeled on top, and known binding partners are listed below the domain with which they interact. 1.3.3.1 Association of APC with polarity and junctional complexes

1.3.3.1.1 Role in apical-basal polarity of epithelial cells

Epithelial polarity is a highly regulated process and loss of polarity can contribute to disease progression. Some non-gastrointestinal model systems have provided evidence for the involvement of APC in regulating aspects of apical-basal polarity and epithelial morphogenesis. For example, a role for APC in overall epithelial organization is supported by the homozygous germline deletion of Apc preventing development past embryonic day 6 (Moser et al. 1995). Apc mutations in the mature cochlea result in a decreased number of parallel microtubule arrays, which are essential for epithelial polarization (Mogensen et al. 2002). Studies from our laboratory have also demonstrated a role for APC in mammary epithelial cell polarization and morphogenesis as ApcMin/+ female mice, containing a mutant Apc allele, exhibited altered mammary epithelial polarity and overall integrity (Prosperi et al. 2009). In addition, knockdown of

APC in the MDCK model, a standard model for epithelial polarity, resulted in altered epithelial morphogenesis and polarity defects in 3D culture (described in detail in section 1.4 and (Lesko et al. 2015)). In Drosophila neuroepithelial cells, APC is recruited to the adherens junction, and mediates symmetric cell division along the planar axis through its interactions with microtubules by positioning the mitotic spindle (Lu et al.

2001). Other roles for APC include the polarized distribution of MUC1, a transmembrane glycoprotein that is frequently mislocalized in cancers, to the apical membrane of differentiated mammary epithelial cells (Prosperi et al. 2009), and the binding of APC to

Striatin, a calmodulin-binding protein predominately expressed in the central nervous system, in cell-cell junctions (Breitman et al. 2008).

The role of APC in maintaining apical-basal polarity is also conveyed in its importance in epithelial cell differentiation, as establishment of polarity is required for differentiation. APC’s role in zebrafish intestinal differentiation involves the transcriptional co-repressor Carboxy-terminal binding protein 1 (CtBP1) which interacts with APC to drive differentiation independent of -catenin through controlling the expression of enzymes like intestinal retinal dehydrogenases (Hamada and Bienz 2004,

Nadauld et al. 2006). A recent study showed that APC is required for apical extrusion of human bronchial epithelial cells (Marshall et al. 2011). The investigators further determined that apical extrusion is not dependent on Wnt/-catenin signaling, but instead requires the Carboxy-terminus of APC and microtubule binding. In addition, the introduction of the EB1 binding domain of APC to DLD-1 colorectal cells was sufficient to shift cells from basal extrusion to apical extrusion, suggesting that APC requires microtubule binding to mediate cell differentiation and migration through apical-basal polarity (Marshall et al. 2011).

Despite the increased understanding about the role of APC in regulating apical- basal polarity, it is still unclear how much of the tumor suppressive activity of APC can be attributed to this function; however, the loss of epithelial polarity and the disruption of tissue organization are normally associated with more aggressive cancers ((Thiery

2002, Huang and Muthuswamy 2010) for review). Moreover, disrupted localization of adherens junction proteins is frequent in cancers (Schmalhofer et al. 2009), and 23

components of the Scrib/Dlg/Lgl complex, protein partners of APC, have recently been implicated in several cancers such as cervical and colorectal cancer (Scheffner and

Whitaker 2003, Gardiol et al. 2006, Huang and Muthuswamy 2010). Collectively, these data support a role for APC in epithelial polarization that may be important for its function during tumor development.

1.3.3.1.2 Junctional complexes

A potential role of APC in mediating cell-cell interactions and cell migration emerged through the interaction of APC with -catenin at the adherens junction

(Rubinfeld et al. 1993), and plakoglobin (-catenin), at the desmosome (Shibata et al.

1994, Rubinfeld et al. 1995). However, the APC/-catenin and -catenin/E-cadherin complexes were observed to be mutually exclusive (Rubinfeld et al. 1993, Hulsken et al.

1994, Rubinfeld et al. 1995), and studies have largely focused on APC’s regulation of the

Wnt pathway through -catenin degradation. Although the role of APC in cellular adhesion, motility, polarization and morphogenesis has been overshadowed, there is evidence that unregulated APC levels are associated with altered cell migration and adhesion. Manipulation of APC levels in mouse intestinal epithelial cells resulted in altered enterocyte migration along intestinal villi in vivo (Wong et al. 1996, Mahmoud et al. 1997, Sansom et al. 2004). Reintroduction of APC in SW480 (APC-mutant) colorectal cancer cells caused changes in the localization of adherens junction proteins, tighter cell-cell contacts, and an epithelial phenotype (Faux et al. 2004). Localization of APC at cell-cell contacts has been shown to be dependent on -catenin and VE-cadherin, as

depletion of either -catenin or VE-cadherin resulted in the disruption of cell-cell adhesion and the loss of APC from the lateral membrane (Harris and Nelson 2010).

Studies using wound healing assays also showed that the inhibition of glycogen synthase kinase 3 (GSK3) and casein kinase I α (CK1α) decreased wound closure, suggesting that cell migration is promoted by a shift of APC to cell protrusions resulting from the phosphorylation by GSK3 and CKIα (Harris and Nelson 2010).

While APC likely mediates cell-cell interactions by regulating levels of -catenin or plakoglobin directly, and -catenin/TCF target genes specifically implicated in adhesion, such as E-cadherin (Jamora et al. 2003), other APC binding partners such as polarity complex proteins or the cytoskeleton may also play an important role. For example, Odenwald et al. showed that the localization of APC/-catenin at protrusion ends controls cell migration and the maintenance of the mesenchymal morphology

(Odenwald et al. 2013). APC knockdown prohibited β-catenin localization at the protrusions, but Wnt pathway activation was not observed. The formation of these complexes was dependent on the microtubule cytoskeleton suggesting that microtubules are required for APC’s role in cell-cell interactions, migration, and morphology (Odenwald et al. 2013).

1.3.3.2 APC interaction with the cytoskeleton

1.3.3.2.1 Actin

APC localized F-actin to the plasma membrane through its Carboxy-terminal region, which also directly interacts with microtubules (Rosin-Arbesfeld et al. 2001).

These data and the observation that APC binds to EB1 (a microtubule-binding protein that will be discussed in detail below) suggest that APC may be involved in regulating actin and microtubule dynamics (Moseley et al. 2007). Recent work demonstrates that

APC promotes nucleation by recruiting actin monomers, and its synergistic activity with the formin mDia is involved in the assembly of actin filaments (Okada et al. 2010).

Although some studies suggest that APC directly interacts with F-actin through the basic domain, it has been shown that APC localization to the plasma membrane is primarily mediated by the ARM domain in the Amino-terminus (Okada et al. 2010). These discrepancies warrant further investigation to understand the interactions between APC and actin.

1.3.3.2.2 Actin effectors

Through its armadillo repeats APC interacts directly with the APC-stimulated guanine nucleotide exchange factor (Asef)-1 and -2 to promote epithelial cell motility and maintain cell morphology (Kawasaki et al. 2000, Kawasaki et al. 2003). However, in

MDCK epithelial cells and colorectal tumor cells, cell migration was only induced by interaction between Asef-1 and the truncated form of APC (Kawasaki et al. 2003). Some

studies suggest that APC may be a scaffold for a complex containing APC/Asef-1/- catenin despite a lack of direct interaction between -catenin and Asef-1 (Kawasaki et al. 2000). Asef-1 and-2 both function as guanine nucleotide exchange factors (GEF) for

Rac (Kawasaki et al. 2000), and binding with APC activates the GEF activity of both Asef-

1 and -2 to promote tumor cell migration (Kawasaki et al. 2007). Unlike Asef-1, which only contains one binding domain for APC, Asef-2 contains two binding sites for APC, the

APC binding region and the Src-homology (SH) 3 domain. Interactions between APC and the SH3 domain of Asef-2 have been shown to activate the GEF activity of Asef-2

(Hamann et al. 2007).

APC also modulates the actin cytoskeleton through direct binding of the actin cross-linking protein IQ motif containing GTPase activating protein (IQGAP1), an effector of Rac and Cdc42, through its armadillo repeat domain (Watanabe et al. 2004). IQGAP1 recruits APC to membrane ruffles and is involved in maintaining the Rho GTPases in an active state (Tirnauer 2004). This complex promotes cell polarization and migration, and binds the microtubule stabilizing protein, CLIP-170 (Watanabe et al. 2004), indicating that APC/IQGAP1 could link actin and microtubule networks.

1.3.3.2.3 Microtubules and microtubule-binding partners

In addition to binding to the actin cytoskeleton, APC also interacts, through the

Carboxy terminus, with microtubules to promote assembly (Munemitsu et al. 1994) by increasing both the stability and the lifespan of the microtubules (Zumbrunn et al. 2001,

Kita et al. 2006). Phosphorylation of APC by GSK3 and Protein Kinase A (PKA) decreases

its interactions with microtubules and also strengthens binding with -catenin

(Zumbrunn et al. 2001). These observations suggest that these interactions are differentially regulated and provide evidence for mutually exclusive pools of APC with different cellular functions (Zumbrunn et al. 2001).

Several other binding partners have been found to link APC with microtubules.

Interaction with end-binding protein 1 (EB1), a plus-end binding protein that regulates microtubule polymerization through a Carboxy-terminal binding domain of APC, results in the localization of APC to microtubule distal ends (Askham et al. 2000, Mimori-

Kiyosue et al. 2000). In addition to EB1, Kap3 and Kif3, members of the kinesin superfamily, interact with APC in the armadillo repeat region to direct its transport to microtubule clusters at cell protrusions and the leading edge of migrating cells (Jimbo et al. 2002). mDia, a formin protein that stabilizes microtubules, binds the Carboxy- terminus of APC in a complex with EB1 to promote fibroblast motility through Rho- induced stabilization (Wen et al. 2004). KIF17, an anterograde kinesin recently shown to participate in localizing APC to the plus ends of microtubules, is required for proper epithelial polarization and morphogenesis (Jaulin and Kreitzer 2010). The nuclear pore complex protein importin-, interacts with the central region of APC where it can compete with -catenin, and also in two regions of the Carboxy-terminus, to regulate

APC-mediated microtubule assembly and spindle integrity (Dikovskaya et al. 2010). APC also interacts with the nuclear pore complex proteins, Nup153 and Nup358, to regulate cell migration and microtubule localization (Collin et al. 2008, Murawala et al. 2009).

Finally Dlg promotes APC accumulation at plus ends of microtubules and the interaction 28

of microtubules with the plasma membrane, suggesting it has an APC-mediated role in microtubule dynamics (Etienne-Manneville et al. 2005).

1.3.3.2.4 Role in front-rear polarity and directional cell migration

APC’s association with actin and the microtubule network is closely linked with its role in directional cell migration. APC expression is localized to puncta at the leading edge or ends of cell protrusions in central nervous system cells, as well as subconfluent epithelial cells and some tumor cells in culture (Nathke et al. 1996, Barth et al. 1997,

Odenwald et al. 2013). This localization depends on the microtubule cytoskeleton and is regulated by APC phosphorylation (Nathke et al. 1996, Barth et al. 1997). APC loss results in decreased microtubule stability, fewer cell protrusions and reduced cell migration (Kita et al. 2006, Kroboth et al. 2007); conversely, APC over-expression induces the formation of cell protrusions (Kroboth et al. 2007).

It has been shown that the regulation of APC by common cytoskeleton modulators is required for the polarization of migrating astrocytes (Etienne-Manneville and Hall 2003). The Rac effector Cdc42, a small Rho GTPase required to polarize the actin and microtubule networks during migration, phosphorylates and inactivates GSK3 promoting the association of APC with microtubule plus ends and the assembly of Dlg- containing puncta at the plasma membrane (Etienne-Manneville and Hall 2003, Etienne-

Manneville et al. 2005). Centrosome orientation and polarized motility was severely disrupted in APC-mutant cells lacking the Carboxy-terminus, which contains the PDZ- binding domain and the binding domains for microtubules, EB1, Dlg, and Scrib (Etienne-

Manneville and Hall 2003, Etienne-Manneville et al. 2005). These data indicate an important role for APC in mediating microtubule polarization and directed cell migration through its interactions with polarity proteins like Dlg and Scrib.

APC also interacts with activated focal adhesion kinase (FAK) at the leading edge in migrating cells to regulate focal adhesion turnover, suggesting that APC may also impact cell motility by directly controlling cell-matrix interactions (Matsumoto et al.

2010). The association of APC with FAK is biologically important as FAK activity regulates proliferation and is required for tumorigenesis in the intestinal epithelium of Apc- mutant mice (Ashton et al. 2010). In the intestinal epithelium, APC-mediated FAK expression is Wnt-dependent (Ashton et al. 2010). Mammary tumors from MMTV-

PyMT;ApcMin/+ mice demonstrated increased phosphorylated FAK, and increased signaling through Src and JNK (Prosperi et al. 2011). Although FAK signaling was enhanced, β-catenin was not localized to the nucleus and TCF activity was not detected, suggesting that the activation of FAK in this model is independent of Wnt signaling

(Prosperi et al. 2011). Combined, these studies support a model in which the interactions between APC and FAK are important for the role of APC as a tumor suppressor; however, the mechanisms of FAK activation may be context-specific.

1.3.3.3 Cell cycle control and DNA replication

During mitosis, APC localizes near the centrosomes and is associated with the microtubule-organizing center to maintain chromatin structure (Olmeda et al. 2003).

Although it is expressed and phosphorylated throughout the cell cycle, APC is transiently

hyperphosphorylated at the Carboxy-terminus in M-phase by the cyclin-dependent kinase (CDK) complex, cyclinB/cdk1, which regulates its interaction with EB1

(Bhattacharjee et al. 1996, Trzepacz et al. 1997). Similarly the cyclin A/cdk2 complex is required for mitotic entry, and also associates with APC in G2/M (Beamish et al. 2009).

The APC/EB1 complex, along with the phosphorylation of APC by the Bub1 and Bub3 mitotic checkpoint kinases, provide stable kinetochore microtubule attachment for proper chromosome alignment (Green et al. 2005, Zhang et al. 2007).

There is evidence that APC plays a direct role in mitosis through its association with the centrosome and microtubules at the mitotic spindle to mediate all phases of the cell cycle. APC functions in the cell cycle partially through the activation of the

Wnt/-catenin pathway. Studies have shown that loss of APC in colorectal cancer cells can modulate cell cycle progression; however, overexpressing APC results in cell cycle arrest (Baeg et al. 1995), and introduction of -catenin only partially rescued progression (Heinen et al. 2002, Carson et al. 2004). In fibroblasts, the interaction of APC with Dlg is required for the APC-induced G1/S arrest independent of mediating - catenin degradation (Ishidate et al. 2000). Further studies showed that the direct binding of APC to A/T rich sequences of DNA through the Carboxy-terminal region of

APC inhibits DNA replication in the G1/S progression (Deka et al. 1998); however phosphorylation of APC by CDK1 and 2 allows cells to progress though the G1/S phase

(Qian et al. 2008).

The role of APC in chromosome segregation and cytokinesis is critical, as embryonic stem cells from Apc-mutant mice and cells with APC depletion suffer from 31

severe chromosomal and mitotic spindle defects (Fodde et al. 2001). This is most likely an early consequence of APC loss since normal tissues isolated from Apc-mutant mouse models exhibit aneuploidy and severe mitotic defects, and the misorientation of spindles worsens as intestinal tumor development progresses (Caldwell et al. 2007,

Dikovskaya et al. 2007). APC’s Amino-terminus may be involved in mitosis as introduction of Amino-terminal APC mutants into HCT-116 colorectal cancer cells inhibits proliferation and mitosis by impacting chromosomal stability through weakening the microtubule and kinetochore interactions (Tighe et al. 2004). The role of APC in chromosomal instability has also been attributed to -catenin/TCF-mediated transcription as shown in studies with APC mutant embryonic stem cells and DLD-1 colorectal cancer cells (Aoki and Taketo 2007).

Additional regulation of APC during proliferation comes from casein kinase 2

(CK2), which interacts with APC at the Amino-terminus to promote nuclear translocation of APC by phosphorylation (Zhang et al. 2001). Interestingly, it is the Carboxy-terminal region that inhibits CK2 and promotes proliferation (Homma et al. 2002). These findings suggest that full-length APC is needed for the APC/CK2 complex activity, although there is evidence that shows no changes in proliferation in cells with loss of APC (Kroboth et al. 2007).

APC also binds proliferating cell nuclear antigen (PCNA), a marker for the G1/S phase and a co-factor of DNA polymerase hence it has roles in both DNA replication and repair (Narayan et al. 2005, Jaiswal and Narayan 2008). The APC binding site for PCNA occurs in the 15-amino acid repeat region of APC that contributes to binding -catenin 32

(Wang et al. 2008). The binding site for topoisomerase II to regulate cell cycle progression also involves the 15-amino acid repeat region. Functionally, cells overexpressing the APC fragment responsible for binding topoisomerase II exhibit a G2 cell cycle arrest (Wang et al. 2008).

Targeting the mitotic spindle checkpoint may present an important therapeutic approach for APC-mutant cancers. Some chemotherapeutic agents, such as taxanes and vinca alkaloids, work as spindle poisons by disrupting microtubules. Recent work demonstrated that low levels of taxol caused APC-deficient cells in vivo to resist arrest due to Wnt-independent microtubule destabilization. However, high taxol concentrations failed to prevent arrest (Radulescu et al. 2010). Another in vitro study implicated a role for APC in response to low levels of mitotic arrest caused by nocodazole treatment. However, similar to previous findings, APC loss had no impact on high levels of arrest (Draviam et al. 2006, Dikovskaya et al. 2007). These data indicate that APC status may influence therapeutic efficacy of specific chemotherapies.

1.3.3.4 Role in DNA repair

APC is involved in the inhibition of multiple DNA repair pathways, such as single nucleotide base nucleotide repair (SN-BER) where it interacts with DNA polymerase 

(Pol-), and long-patch base excision repair (LP-BER) where it interacts with flap endonuclease 1 (Fen-1) (Narayan et al. 2005, Jaiswal et al. 2006, Balusu et al. 2007). APC is also recruited to the site of DNA damage during double-stranded DNA break repair where it induces early response to the damage by interacting with the DNA-dependent 33

protein kinase catalytic subunit (Kouzmenko et al. 2008). This activity is consistent with

APC’s role as the ‘gatekeeper’, as its mutation may slow DNA repair and increase mutations in surrounding cells to promote tumor progression (Kinzler and Vogelstein

1996, Kouzmenko et al. 2008).

The loss or mutation of APC has also been associated with increased ability to repair mutated DNA (Narayan et al. 2005). Specifically, APC interacts with Pol- and Fen-

1 to block LP-BER, and treatment with a DNA alkylating agent methylmethane sulfonate

(MMS) did not damage cells with mutated APC (Jaiswal et al. 2006, Balusu et al. 2007).

This observation is important as APC status could impact treatment of tumor cells with chemotherapeutic alkylating agents such that cells with APC mutations may be more resistant than those with wild-type APC. Interestingly, APC expression is regulated by a variety of alkylating agents, including MMS (Narayan and Jaiswal 1997), and carcinogens

(Jaiswal and Narayan 1998, Jaiswal et al. 2004, Jaiswal et al. 2006) suggesting that there may be a feedback loop that regulates APC levels and activity.

1.3.3.5 Association of APC with apoptosis regulators

APC mutation has long been associated with changes in cellular apoptosis

(Browne et al. 1994, Browne et al. 1998, Venesio et al. 2003). In fact, reintroduction of full-length APC to colorectal cancer cells with truncated APC enhanced both basal apoptotic activity and drug-induced apoptosis (Morin et al. 1996). APC can induce apoptosis through regulation of the Wnt target gene survivin, a member of the inhibitor of apoptosis family of pro-survival factors (Zhang et al. 2001); however, control of Wnt

signaling by APC may not be sufficient to account for APC’s role in apoptosis. For example, an increase in apoptosis was observed during lactation in the mammary gland of ApcMin/+ mice in the absence of Wnt pathway activation (Prosperi et al. 2009) and APC modulated apoptosis in a transcription-free Xenopus extract assay (Steigerwald et al.

2005).

The context-dependent manner in which APC induces apoptosis with both APC mutation or depletion and APC over-expression presents a challenge in identifying the role of APC in regulating apoptosis. As an example, apoptosis in mammary epithelial in

ApcMin/+ mice may be caused by a disruption in epithelial polarization (Prosperi et al.

2009); while apoptosis in Apc-mutant adult small intestine epithelium of mice may be a result of perturbed migration and differentiation (Clarke 2005). Because APC’s regulation of apoptosis may be dependent on cancer type, methods triggering apoptosis in APC-deficient cancers by introducing APC domains or other approaches might prove to be a promising therapeutic strategy, but still need to be thoroughly examined for context-dependence (Zhang et al. 2010).

1.3.4 Conclusion

Loss of function mutations and silencing of the APC tumor suppressor gene are commonly observed in several epithelial cancers such as colorectal, breast, ovarian, pancreatic, and lung cancer. APC is best known for regulating the Wnt/β-catenin pathway; however, APC has many Wnt-independent roles which influence its tumor suppressor activity. APC is associated with polarity and junctional complexes and has

been shown to regulate apical-basal polarity in a variety of epithelial tissues.

Additionally, APC interacts and stabilizes the cytoskeleton through both direct and indirect interactions. Nuclear functions of APC have been identified as APC interacts with several proteins involved in DNA replication and repair. Finally, APC influences the cell cycle by mediating proteins involved in both proliferation and apoptosis. Gaining a better understanding of the normal functions of APC allow us to glean important insights into how the loss of APC and disruption of these cellular processes drive tumor progression.

1.4 The APC tumor suppressor is required for epithelial cell polarization and three- dimensional morphogenesis

1.4.1 Introduction

Epithelial morphogenesis is a tightly coordinated process that requires extrinsic and intrinsic cues to couple cell-cell and cell-matrix interactions, polarity, proliferation, cell death, and differentiation. In contrast to traditional two-dimensional (2D) culture on glass or plastic, the organotypic 3D culture of epithelial cells in extracellular matrix

(ECM), such as the reconstituted basement membrane Matrigel or collagen, is a powerful in vitro model that recapitulates many of the features of tissue polarity and architecture (reviewed in (Zegers et al. 2003)). Common features of these organoid or spheroid models (conventionally referred to as “acini” for mammary cells and “cysts” for kidney cells) are that after a couple of cell divisions of plated single cells, they polarize to form a basal surface that contacts the ECM, a lateral surface between cells, and an 36

apical surface which faces the lumen. Apoptosis will occur in those cells that do not contact the ECM, and cells that do not yet have an apical surface will generally form a lumen at the point of contact with other cells (reviewed in (Bryant and Mostov 2008)).

Recent insights into the molecular mechanisms that guide polarization and lumen formation, for example, have supported the importance of junction and polarity complexes, laminins, integrins, phosphoinositides and Rho GTPases family members in these processes (O'Brien et al. 2001, Yu et al. 2005, Yu et al. 2008, Zhan et al. 2008, Kim and Giardiello 2011). Importantly, these polarity and morphogenesis programs are often disrupted or hijacked in pathological conditions such as chronic wounds, kidney fibrosis and cancer; therefore, a more complete understanding of the pathways and critical players involved has significant clinical relevance.

Previous work from our laboratory and others (described in section 1.3.3) has established a link of APC to polarity and tissue architecture. Endogenous and exogenous

APC is concentrated in puncta at the ends of cell protrusions in motile cells, such as astrocytes, radial glia or subconfluent epithelial cells, where it associates with the microtubules and is required for front-rear polarity downstream of the Cdc42

RhoGTPase (Nathke et al. 1996, Barth et al. 2002, Etienne-Manneville and Hall 2003,

Etienne-Manneville et al. 2005, Reilein and Nelson 2005, Kita et al. 2006, Yokota et al.

2009). In polarized epithelia in vitro and in vivo, APC localizes to the basal plasma membrane (Rosin-Arbesfeld et al. 2001, Mogensen et al. 2002, Prosperi et al. 2009) in an actin-dependent fashion (Rosin-Arbesfeld et al. 2001), where it controls the establishment of parallel arrays of microtubule bundles (Mogensen et al. 2002). 37

However, there are also reports of APC localization to the apical membrane of polarized epithelial cell types, including in the differentiated mammary epithelium (Reinacher-

Schick and Gumbiner 2001, Prosperi et al. 2009). Our laboratory has demonstrated that heterozygous Apc mutation abrogates mammary lobuloalveologenesis by inhibiting proliferation during pregnancy, inducing apoptosis during lactation and severely altering epithelial integrity, including cell-cell interactions and polarity (Prosperi et al. 2009).

Furthermore, knockdown of APC in MDCK cells perturbs mitotic spindle orientation (den

Elzen et al. 2009) that can lead to monolayer disruption, and APC expression in EpH4 mammary epithelial cells was required for normal monolayer formation (Prosperi et al.

2009). APC also mediates directionality of cell extrusion from an epithelial monolayer through its control of microtubule dynamics (Marshall et al. 2011). However, key questions regarding the role of APC in epithelial morphogenesis and the mechanisms by which APC mediates these behaviors remain unanswered, and, importantly, it has not been established whether this is one of the essential ways in which APC acts as a tumor suppressor.

In the following study by Lesko et. al., we tested the hypothesis that APC function is required for normal epithelial polarity and 3D morphogenesis. By establishing an in vitro model of stable APC knockdown in MDCK cells, we found that APC is required for normal spheroid morphogenesis in 3D culture. The effects of APC depletion were rescued with overexpression of either full-length or a Carboxy (C)-terminal fragment of

APC, but not by a central region containing the -catenin-binding domain. Our results are consistent with the interactions between APC and cytoskeletal and/or polarity 38

complex proteins being required for normal polarity and morphogenesis programs, but the phenotypes associated with APC knockdown do not involve activation of the Wnt signaling pathway. These studies highlight the importance of APC as a regulator of epithelial behavior and tissue architecture, and suggest that tumor initiation as a result of APC mutation or inactivation may be driven by loss of proper apical-basal polarity and dysmorphogenesis.

1.4.2 Results

1.4.2.1 APC knockdown disrupts cyst morphology and polarity in MDCK cells.

We sought to address the role for APC in establishing or maintaining polarity, and identify the molecular mechanisms responsible, in the MDCK cell line because it is a well studied model system for investigating epithelial polarity and morphogenesis.

MDCK cells with stable expression of APC shRNA were generated using transduction by lentiviral shRNA. Western blot analysis confirmed that APC protein levels were markedly decreased in the APC shRNA cells compared to the controls (Figure 1.2 A,B). APC knockdown in MDCK cells plated in 3D Matrigel cultures exhibited very large and highly disorganized cysts in the APC shRNA cells compared to the control cells by phase- contrast microscopy (Figure 1.2 E). These qualitative differences were confirmed by morphometric analysis of the phase-contrast images and demonstrated that the cysts were larger in the APC shRNA cells compared to the control cells (Figure 1.2 F). The stable introduction of full-length APC (hAPC-FL) was able to rescue this phenotype

(Figure 1.2 E,F) confirming that normal MDCK 3D morphogenesis is dependent on APC. 39

For the rescue experiments, it is important to note that the construct contains the human APC cDNA and is not targeted by the APC shRNA.

To dissect the region of APC required for the altered cyst size, we generated cell lines stably expressing a large central segment of APC tagged with GFP (APCmid; residues

220-2164) and the C-terminus of APC tagged with GFP (APCC-ter; residues 2165-2843) in

APC-knockdown and control cells (Figure 1.2 C), and compared them to expression of the vector only and full-length APC (as a positive control for phenotype rescue as shown in Figure 1.2 E,F). Expression of these fragments was confirmed through western blots of APC (Figure 1.2 D), and immunofluorescence staining of GFP (Figure 1.3). Notably, this central segment is necessary and sufficient for -catenin binding and downregulation (Rubinfeld et al. 1993, Munemitsu et al. 1995), and the C-terminal fragment associates with the microtubule and actin cytoskeleton, and Dlg and Scrib polarity proteins (Munemitsu et al. 1994, Matsumine et al. 1996, Askham et al. 2000, Takizawa et al. 2006, Moseley et al. 2007). While the APCmid fragment had minimal effect on cyst size, the APCC-ter significantly decreased the cyst size of the APC-knockdown cells compared to APC shRNA cells stably expressing the vector control (Figure 1.2 E,F).

Figure 1.2 MDCK 3D morphogenesis is perturbed by APC knockdown and restored by re-introduction of APC. A) APC protein expression in MDCK cells was analyzed by Western blot. B) APC protein expression in MDCK cells was quantified and normalized to actin expression. C) Schematic of the full-length APC protein including functional domains (top) and binding partners (bottom) and APCmid and APCC-ter fragments. D) APC protein expression in MDCK cells with expression of human full- length APC (hAPC-FL), the central fragment of APC (APCmid), and the C-terminus of APC (APCC-ter) was analyzed by Western Blot and normalized to actin expression. E) Phase-contrast images of MDCK and APC-knockdown MDCK cysts show an increase in cyst size and altered morphology in APC-knockdown cell compared to controls (left panels). The 2nd-4th columns show the phenotype with ectopic expression of human full-length APC (hAPC-FL), APCmid, or APCC-ter. Scale bars, 100 m. F) Cyst size was increased in APC- knockdown cells compared to controls, and this size difference was reversed with the introduction of hAPC-FL. The enhanced cyst size was significantly abrogated by introduction of the APCC-ter construct. Shown is a bar graph with average cyst size. A one-way ANOVA was performed to determine significance, p <0.05 . Representative images are shown of experiments, which were all performed three times.

Figure 1.3 Reintroduced APC fragments are detected by immunofluorescence staining for GFP. Control and APC knockdown MDCK cells transfected with either the vector alone, hAPC-FL, APCmid, or APCC-ter fragment were stained for GFP (green) and phalloidin (red).

To further assess the impact of APC depletion on MDCK 3D morphogenesis and polarization, MDCK control and APC shRNA cells were plated in Matrigel over a time course of 7 days and analyzed by immunofluorescence and confocal microscopy for the localization of the apical marker gp135 (also referred to as podocalyxin) and phalloidin to stain F-actin. While the lumens observed at 3 days post-plating in the MDCK control cells were quite small, gp135 was localized to apical cell surface adjacent to these early lumens (Figure 1.4 A), which is consistent with previous analysis of MDCK 3D morphogenesis in Matrigel (Engelberg et al. 2011, Kim and Giardiello 2011). After 7 days in culture, the lumens of MDCK cell cysts were generally hollow, and most cysts exhibited only a single lumen. In striking contrast, the APC shRNA cells formed many cysts without lumens and gp135 was frequently localized to basal surface (Figure 1.4 A), an effect that was observed as early as 3 days post-plating and very pronounced by day

7. Like the gross cyst morphological defects observed in APC-knockdown cells (i.e. increased size and non-spherical shape), the lack of discernible lumens and mislocalization of gp135 were abrogated by introduction of full-length APC (Figure 1.4

B,C). Similar to the effect of the APC fragments on cyst size, the APCC-ter fragment, but not the APCmid fragment, partially rescued the polarized expression of membrane markers in the APC-knockdown cells (Figure 1.4 B,C). These data demonstrate that the

C-terminus of APC mediates epithelial polarization and morphogenesis, and suggest that one consequence of APC mutation and deletion of the C-terminus during tumorigenesis is loss of polarity and tissue architecture.

Figure 1.4 APC knockdown disrupts MDCK cell polarity in cysts grown 3D culture. A) Control and APC-knockdown MDCK cells were grown in Matrigel for 3 or 7 days and stained for the apical marker gp135 (red) and phalloidin (green). Control MDCK cells show apical localization of gp135 over the time course, but basal localization (white arrows) of gp135 is observed as early as 3 days post-plating in the APC-knockdown MDCK cells and the cysts are generally larger and have a less spherical morphology. Insets are higher magnification images of interest. Scale bars, 20 m. B) Cysts from APC rescue cells (using hAPC-FL, APCC-ter, and APCmid) were grown for 7 days and stained for the apical marker, gp135, which as shown in Figure 3A is inverted in the APC-knockdown cells. With re-introduction of full-length APC or the APCC-ter construct, gp135 localization is restored to the apical surface in knockdown cells. Scale bars, 20 m. Shown is a bar graph (C) quantifying the polarity phenotypes as percent apical (normal), and basal or mixed (abnormal). 100 cysts were counted for each cell line with the various APC constructs re-introduced. Significance was determined using Fisher’s exact test (* p < .05 compared to APC shRNA vector cells). Representative images are shown of experiments, which were all performed three times.

1.4.2.2 Gene profiling experiments identify candidates for mediating APC-regulated epithelial morphogenesis.

Given that the effects of APC-knockdown in MDCK cells could not be fully explained by Wnt/-catenin activation (Lesko et al. 2015), we addressed other potential molecular mechanisms responsible by gene expression profiling to identify candidate pathways or molecules dysregulated upon APC knockdown. In addition, we sought to address whether the APC-knockdown cells could be used as a tool to identify a gene signature associated with APC deficiency and epithelial polarity and morphogenesis.

Gene expression microarray analysis was performed on RNA from control shRNA and

APC shRNA MDCK cells grown in 3D Matrigel culture for 5 days, a time point when the polarity inversion and dysmorphogenesis phenotypes are pronounced in APC- knockdown cells (n=3 per cell line). Surprisingly, comparison of APC shRNA to control shRNA MDCK cells at this time point demonstrated that only 125 genes were significantly differentially expressed (>1.2-fold change; 75 up-regulated and 50 downregulated). A representative heat-map of the signature is shown (Figure 1.5 A), and the differential expression of multiple genes was validated by real-time RT-PCR (Figure 1.5

B). Consistent with previous data (Lesko et al. 2015), no characterized -catenin/TCF target genes were contained in this APC-knockdown signature. Some of the genes in the signature had been previously linked to APC loss-of-function, and are generally implicated in mediating epithelial cell-cell or cell-matrix interactions. For example, epithelial membrane protein 2 (EMP2), which has been shown to regulate the activity and phosphorylation of focal adhesion kinase (FAK) through interaction with 1 integrin 47

(Morales et al. 2009, Morales et al. 2009, Fu et al. 2011) was significantly up-regulated in the APC shRNA MDCK cells. This alteration in gene expression is of particular interest given our previous observations that mutation or loss of APC controls FAK activity in breast tumor cells (Prosperi et al. 2011), and that inhibition of 1 integrin signaling, like

APC depletion, confers an inverted polarity phenotype in MDCK cells (Ojakian and

Schwimmer 1994, Yu et al. 2005). The transmembrane glycoprotein MUC16 was also up-regulated in the APC shRNA MDCK cells. MUC16 has been implicated in early tumor development in both pancreatic and ovarian models (Bast et al. 1983, Chauhan et al.

2006, Haridas et al. 2011), and we have previously shown that APC is required for the localization of a related membrane-associated mucin, MUC1 (Prosperi et al. 2009).

Interestingly, down-regulation of Lipocalin2 (LCN2), a marker for kidney injury (Viau et al. 2010), implicated in kidney epithelial cell morphogenesis (Gwira et al. 2005) and previously shown to be dysregulated in ApcMin/+ intestinal adenomas (Reichling et al.

2005), was observed in the APC shRNA MDCK cells. Other dysregulated genes including the alpha 1 subunit of type IV collagen (COL4A1, a basement membrane component), C-

X-C chemokine receptor type 7 (CXCR7), and ADAM metallopeptidase with thrombospondin type 1, motif 6 (ADAMTS6), are implicated in controlling cell-cell or cell-matrix interactions (Kuhn 1995, Bevitt et al. 2005, Hou et al. 2010, Aikio et al. 2012).

These data collectively support a model in which APC loss-of-function in epithelial cells

(through mutation and deletion of its C-terminus or gene silencing) leads to loss of polarity and tissue architecture, and subsequent tumor initiation, via altered communication between neighboring cells and the substratum. 48

Figure 1.5 An APC-knockdown gene signature is associated with altered cell-cell and cell-matrix communication. A) Microarray analysis was performed on RNA from cells grown in 3D Matrigel culture for 5 days. Four wells were pooled to generate a single RNA sample, and 3 independent RNA samples per cell line were utilized in these studies. Global gene expression changes are shown in the heat map where overexpression is shown in red and underexpression is blue. B) Real-time PCR was used to validate some key gene expression differences, including CXCR7, ADAMTS6, LCN2, COL4A1, MUC16, and EMP2. Normalized expression relative to 18S rRNA is shown for real-time data. 49

1.4.3 Discussion

1.4.3.1 Summary

The data presented here underscore the importance of APC in controlling key components of the normal epithelial morphogenesis program, including polarity and cell-cell and cell-matrix communication. Down-regulation of APC MDCK epithelial cell lines significantly perturbed acinar/cyst formation in 3D culture conditions. The 3D dysmorphogenesis phenotype of APC-knockdown cells, namely large, non-spherical structures with abnormal polarity, was rescued by introduction of exogenous human

APC and its C-terminal end but not a central segment containing the -catenin binding and down-regulation domain. These data, combined with previous observations support a model in which the control of polarity and morphogenesis by APC is independent of canonical Wnt pathway regulation (Lesko et al. 2015). We identified an APC-knockdown gene signature characterized by several genes involved in cell-cell and cell-matrix interactions and tumor initiation.

It was striking that APC-knockdown epithelial cells did not have any overt morphological defects or alterations in growth properties when plated on solid substrata (e.g glass or tissue culture plastic) but had very dramatic phenotypes when cultured on permeable supports or in 3D ECM, respectively. A similar phenotype in epithelial morphogenesis has recently been observed with loss of the tumor suppressor gene, ductal epithelium-associated ring chromosome 1 (DEAR1). In human mammary epithelial cells, loss of DEAR1 causes no change in 2D growth; however, these cells

exhibit irregular acini morphogenesis and loss of polarity in 3D culture (Chen et al.

2014). These differences presumably occur because on plastic or glass, the cells are not receiving asymmetric polarization cues that are necessary for orienting their axis of polarity in 3D (Bryant and Mostov 2008). APC may be required for receiving or integrating these cues, such as those derived from cell-matrix interactions.

1.4.3.2 Potential molecular mechanisms downstream of APC in regulating polarity and morphogenesis

A multi-layering phenotype specifically and loss of contact inhibition has been observed in MDCK cells in which a kinase-dead or constitutively activated Rac1 effector p21-activated kinase (PAK1) was introduced (Zegers et al. 2003). Consistent with these data, the expression of a constitutively active PAK1 in MDCK cells misorients the apical surface and induces a multi-lumen phenotype, identical to the effect of 1 integrin inhibition (deLeon et al. 2012). In fact, Yu et al. (Yu et al. 2005) showed that 1 integrin orients epithelial polarity in the 3D MDCK model through Rac1 and laminin signaling, an effect that is likely mediated by PAK1. In addition, APC loss has been shown to regulate directionality of cell extrusion (Marshall et al. 2011) or apical constriction through RhoI and Myosin II in Drosophila (Zimmerman et al. 2010). We have previously shown that

Apc mutation in PyMT-driven mammary tumor cells results in hyperactivation of focal adhesion kinase signaling (Prosperi et al. 2011). Consistently, enhanced FAK phosphorylation and activation is also observed in intestinal tumor models from Apc- knockout mice, and FAK activity is required for tumorigenesis in these animals (Ashton

et al. 2010). It is possible that APC regulates the signaling pathways downstream of cell-

ECM interactions and its loss uncouples integrin/ECM-mediated polarization and morphogenesis signaling pathways. Further support for this hypothesis is provided by the gene expression changes observed in APC-knockdown cells grown in 3D culture conditions. EMP2 is an intriguing candidate given its role in mediating 1 integrin signaling (Morales et al. 2009, Fu et al. 2011), previously described as a critical component of MDCK cell polarity (Ojakian and Schwimmer 1994, Yu et al. 2005), and the

APC/FAK crosstalk identified in Apc-mutant tumors by our laboratory (Prosperi et al.

2011) and others (Ashton et al. 2010). Furthermore, EMP2 and MUC16 are both early markers of tumor development (Fu et al. 2011, Haridas et al. 2011), consistent with the model of APC loss resulting in dysregulation of epithelial polarity preceding tumorigenesis. The specific molecular mechanism by which APC controls these pathways is the focus of current study but may involve interactions of APC with actin itself or actin remodeling proteins such as the Rho GTPase effector IQGAP (Watanabe et al. 2004) or the Rac and Cdc42 guanine-exchange factors Asef1 and 2 (Kawasaki et al.

2000, Kawasaki et al. 2003, Kawasaki et al. 2007).

Another attractive possibility is that APC elicits many of its control of epithelial polarization and morphogenesis through its direct and indirect interactions with the plus-ends of microtubules, particularly during mitosis. APC localizes to the kinetochores and centrosomes in mitotic cells, and its mutation is associated with defects in chromosome segregation/cytokinesis, and genomic instability (Fodde et al. 2001, Kaplan et al. 2001). Mitotic spindle orientation is disrupted in intestinal crypts heterozygous for 52

an Apc mutation and even more dramatically in tumors from Apc-mutant mice that demonstrate loss of heterozygosity (LOH) (Fleming et al. 2009). Recent work has shown that APC is necessary for proper spindle orientation perpendicular to the apical surface in the stem cell compartment of human and mouse gastrointestinal epithelium, and that this alignment is required for asymmetric stem cell division (Quyn et al. 2010). Studies by den Elzen et al. (den Elzen et al. 2009) demonstrated that adherens junctions provide a necessary cue for proper planar spindle orientation in MDCK monolayers, and that E- cadherin disruption mislocalizes APC. Furthermore, APC is required for spindle alignment during symmetric cell division in this model (den Elzen et al. 2009). The APC partner IQGAP also has been implicated in cytokinesis (Shannon 2012). Additionally, the kinesin KIF17 helps to localize APC to the plus ends of a subset of microtubules, and

KIF17 depletion results in aberrant 3D epithelial cysts that lack both a central lumen and polarized apical markers (Jaulin and Kreitzer 2010). A recently characterized integrin- linked kinase (ILK)-microtubule pathway to regulate the delivery of apical cargo to the correct membrane domain (Akhtar and Streuli 2013) raises the attractive possibility that

APC provides a link between integrin signaling and microtubules in epithelia.

1.4.3.3 Conclusion

APC is commonly mutated or down-regulated in epithelial cancers, it is important to consider how loss of APC-mediated control of morphogenesis and polarization may contribute to tumor initiation. Our previous work (Prosperi et al. 2009,

Lesko et al. 2015) demonstrates that heterozygous APC-mutant tissues recapitulate

many of the architectural abnormalities modeled by APC-knockdown in epithelial cell lines. These data are consistent with other studies illustrating that Apc-knockout colonic mucosa and tumors have defective barrier function (Jamora et al. 2003, Segditsas and

Tomlinson 2006) and that intestinal epithelial cells from histologically normal heterozygous Apc transgenic and mutant tissues had altered migration and patterns of gene expression along the crypt-villus axis (Wong et al. 1996, Homma et al. 2002).

While kidney tumors have not been identified in the ApcMin/+ mice or FAP patients, there is a connection between Gardner’s syndrome (the non-intestinal tumors of FAP patients) and cilia (Gomez Garcia and Knoers 2009). Further, homozygous deletion of

Apc in the kidney predisposes it to tumorigenesis (Sansom et al. 2005), suggesting that

APC plays a critical tumor suppressive role in the kidney. These findings collectively support a model in which mutation of one APC allele, or decreased APC expression by gene methylation, is sufficient to perturb epithelial polarization and architecture so as to uncouple growth and survival cues and promote genomic instability to promote carcinogenesis.

1.5 Epithelial Membrane Protein 2

From microarray analysis we previously discovered APC loss increases the mRNA expression of Epithelial Membrane Protein 2 (EMP2). EMP2 is a tetraspan membrane protein which regulates several cellular functions including lipid raft formation and apical membrane recycling, proliferation and apoptosis, and cell migration and adhesion

(reviewed in (Wang et al. 2017)). EMP2 expression is very heterogeneous in different

organs and depending on tissue type, EMP2 has been implicated as both an oncogene and tumor suppressor (reviewed in (Wang et al. 2017)). EMP2 was first identified as a tumor suppressor. Low expression of EMP2 in lung cancer cells promotes cell migration through Caveolin 1 (Cav1)/extracellular signal-regulated kinase (ERK)/jun N-terminal kinase (JNK) signaling (Lee et al. 2016). Conversely, EMP2 is overexpressed in breast and endometrial cancer cell lines and promotes invasive phenotypes through β1 integrin/focal adhesion kinase (FAK)/Src signaling ((Fu et al. 2011, Fu et al. 2014) and reviewed in (Wang et al. 2017)). Although not much is known about the regulation of

EMP2, studies have shown EMP2 expression is influenced by hormonal regulation. EMP2 expression can be induced in endometrial cancer cells by treatment with estrogen and progesterone and EMP2 expression in the endometrium varies during different phases of the uterine cycle (Wadehra et al. 2008). The normal functions of EMP2 in epithelial cells remain largely unknown and warrant further investigation to identify the mechanisms by which EMP2 contributes to tumorigenesis.

CHAPTER 2:

EPITHELIAL MEMBRANE PROTEIN 2 AND Β1 INTEGRIN SIGNALING REGULATE APC-

MEDIATED PROCESSES

This text can be seen in publication: Lesko AC, and Prosperi JR (2017). Epithelial Membrane Protein 2 and β1 integrin signaling regulate APC-mediated processes. Experimental Cell Research, 350(1): 190-198.

2.1 Abstract

Adenomatous Polyposis Coli (APC) plays a critical role in cell motility, maintenance of apical-basal polarity, and epithelial morphogenesis. We previously demonstrated that APC loss in Madin Darby Canine Kidney (MDCK) cells increases cyst size and inverts polarity independent of Wnt signaling, and upregulates the tetraspan protein, Epithelial Membrane Protein 2 (EMP2). Herein, we show that APC loss increases

β1 integrin expression and migration of MDCK cells. Through 3D in vitro model systems and 2D migration analysis, we have depicted the molecular mechanism(s) by which APC influences polarity and cell motility. EMP2 knockdown in APC shRNA cells revealed that

APC regulates apical-basal polarity and cyst size through EMP2. Chemical inhibition of

β1 integrin and its signaling components, FAK and Src, indicated that APC controls cyst size and migration, but not polarity, through β1 integrin and its downstream targets.

Combined, the current studies have identified two distinct and novel mechanisms required for APC to regulate polarity, cyst size, and cell migration independent of Wnt signaling. 56

2.2 Introduction

Epithelial cells normally form a uniform layer maintained through the regulation of cell-cell interactions and apical-basal polarity (Bazzoun et al. 2013) by several complexes including cell junctions and polarity proteins such as Par6, atypical Protein

Kinase C (aPKC), Scribble (Scrib), and Discs large (Dlg) (Wodarz and Näthke 2007, Lee and Vasioukhin 2008). The loss of these complexes, and subsequently apical-basal polarity, disrupts normal developmental processes including epithelial structure, migration, and intracellular signaling (Martin-Belmonte and Perez-Moreno 2012), predisposing cells to tumorigenesis ((Shin et al. 2006, Chandramouly et al. 2007) and reviewed in (McCaffrey and Macara 2011, Macara and McCaffrey 2013)). Several studies have shown that the loss of junctional complexes and polarity complexes leads to disrupted morphogenesis and polarity. Down regulation of Junctional Adhesion

Molecule A (Jam-A), Par6, aPKC, or lethal giant larvae (Lgl) in Madin Darby Canine

Kidney (MDCK) cysts prevented lumen formation and disrupted apical-basal polarity

(Rehder et al. 2006, Yamanaka et al. 2006, Kim et al. 2007, Karp et al. 2008, Feigin and

Muthuswamy 2009, Archibald et al. 2015). Similar phenotypes are seen in acini formed by MCF-10A mammary epithelial cells with the loss of Scrib (Zhan et al. 2008), and the mislocalization of Scrib from cell junctions promotes mammary tumorigenesis through

PTEN and Akt signaling (Feigin et al. 2014). Furthermore, studies have shown that changes in mammary cell adhesion, signaling, and polarity can lead to development of breast cancer ((Feigin et al. 2014, Mojallal et al. 2014) and reviewed in (Bazzoun et al.

2013, Chatterjee and McCaffrey 2014)). For example, loss of the polarity complex 57

protein Par3 in conjunction with activation of Notch signaling or the expression of the oncogene h-RAS increases mammary tumor growth (McCaffrey et al. 2012). These studies stress the importance of regulating cellular processes like cell polarity, adhesion, and motility to prevent tumorigenesis.

It has recently been appreciated that tumor suppressors play an important role in regulating polarity to maintain tissue structure. One example of this is Liver Kinase B1

(LKB1), a tumor suppressor known to regulate cell polarity proteins and complexes, such as the Par proteins ((Goransson et al. 2006) and reviewed in (Martin-Belmonte and

Perez-Moreno 2012)). In breast epithelial cells, LKB1 loss increased cell migration and invasion in 2D, and disrupted morphology and polarity in 3D culture (Li et al. 2014).

Similarly, loss of a novel tumor suppressor ductal epithelium associated ring chromosome 1 (DEAR1) in human mammary epithelial cells resulted in acini with disrupted polarity and filled lumens (Chen et al. 2014). Studies from our laboratory provide evidence for a role for Adenomatous Polyposis Coli (APC) in regulating apical- basal polarity and morphogenesis as mammary glands of Apc-mutant mice exhibit alterations in polarity and epithelial structure during pregnancy and lactation (Prosperi et al. 2009), and the loss of APC in MDCK cells resulted in larger cysts with filled lumens and the mislocalization of the apical marker gp135 to the basal membrane (Lesko et al.

2015). Furthermore, heterozygous mutation of Apc altered polarity and tissue structure in human familial adenomatous polyposis (FAP) colonic tissue and mouse kidney and colonic tissue (Lesko et al. 2015). The maintenance of apical-basal polarity by tumor suppressors contributes to their role in preventing disease progression. For example, 58

ERBB-2 mediated mammary tumorigenesis was accelerated upon loss of the 14-3-3σ tumor suppressor resulting in tumor cell lines with disrupted junctional complexes and mislocalization of Par3 (Ling et al. 2012). Together these studies suggest that maintenance of apical-basal polarity is a critical function by which APC suppresses tumor development.

Although APC is most known for its role in the regulation of Wnt/β-catenin signaling, it has many Wnt-independent functions including regulating apical-basal polarity, microtubule and actin networks, and cell migration (reviewed in (Prosperi and

Goss 2011, Nelson and Nathke 2013)). Consistent with previous results showing Wnt- independent functions of APC (Prosperi et al. 2009, Prosperi et al. 2011, Cai et al. 2015), we recently demonstrated that APC regulates polarity and cyst size through a Wnt- independent mechanism as TCF/LEF transcriptional activity was not increased upon APC loss and stabilized β-catenin did not increase cyst size or mislocalize gp135 in MDCK cysts (Lesko et al. 2015). Furthermore, the reintroduction of the β-catenin binding domain of APC was unable to restore cyst size and apical polarity in MDCK cysts with

APC loss (Lesko et al. 2015). Interestingly, a C-terminal fragment of APC, important for binding polarity complexes, decreased cyst size and reversed the polarity phenotypes caused by APC loss (Lesko et al. 2015). The C-terminus of APC also contains binding domains for actin, microtubules, and associated cytoskeletal proteins suggesting a role for APC in cell motility. This role has been supported by studies in which Apc-mutant mice displayed altered enterocyte migration in the intestine (Wong et al. 1996,

Mahmoud et al. 1997, Sansom et al. 2004), mammary tumor cell migration was 59

controlled by APC localization at cell protrusions (Odenwald et al. 2013), and the C- terminus of APC promoted the apical extrusion of human bronchial cells (Marshall et al.

2011). Furthermore truncation mutations in APC in MDCK cells and colorectal tumors cells increased cell migration (Kawasaki et al. 2000, Kawasaki et al. 2003).

We identified that APC knockdown in MDCK cells does not activate the Wnt/- catenin pathway, but results in elevated levels of Epithelial Membrane Protein 2 (EMP2)

(Lesko et al. 2015). Previous studies demonstrated that overexpression of EMP2 increases endometrial cell migration in wound healing assays (Fu et al. 2011), upregulates expression of β1 integrin, and activates Focal Adhesion Kinase (FAK)/Src signaling (Fu et al. 2011, Fu et al. 2014). β1 integrin has been shown to regulate polarity and cyst size in MDCK cells (Yu et al. 2005). Furthermore FAK and Src have been identified as downstream targets of β1 integrin ((Cary et al. 1996, Parsons 2003) and reviewed in (Israeli et al. 2010, Vachon 2011)). Given the interaction of EMP2 and β1 integrin with FAK and the role of FAK/Src signaling in migration and tumorigenesis, FAK and Src are promising candidates as downstream effectors of APC. Several studies have shown that FAK has a role in regulating cell motility. APC interacts with FAK at the leading edge of migrating cells (Matsumoto et al. 2010). Additionally, phosphorylated

FAK and Src are increased in mammary tumors from Apc-mutant mice (Prosperi et al.

2011), and correlate with enhanced cell migration of breast cancer cells in wound healing assays (Fang et al. 2014). These data provide a possible mechanism by which

APC mediates cyst size, polarity, and migration through upregulation of EMP2 and subsequent activation of β1 integrin and downstream signaling. 60

The current studies test the hypothesis that APC mediates epithelial polarity, cyst size, and migration through EMP2 and β1 integrin signaling. Given the known interactions between EMP2, β1 integrin, FAK and Src, and their roles in regulating cell growth, adhesion, motility, and tumorigenesis these pathways present a possible mechanism by which APC regulates these processes. The current studies show for the first time that EMP2 regulates cyst size and apical-basal polarity, while β1 integrin signaling mediates cyst size and migration, but not apical-basal polarity. Combined, we have identified two novel molecular mechanisms by which APC influences epithelial morphogenesis, polarity, and cell motility.

2.3 Materials and methods

2.3.1 Cell culture

Madin Darby Canine Kidney (MDCK) cells were obtained from K. Matlin

(University of Chicago), tested for contamination, and maintained in MEM media with

5% FBS, 2mM L-glutamine, 10mM HEPES, and 1% Penicillin/Streptomycin. ctl shRNA and

APC shRNA cells were established previously (Lesko et al. 2015), and were grown in

MEM media with 5% FBS, 2mM L-glutamine, 10mM HEPES, 1% Penicillin/Streptomycin, and 2µg/ml Puromycin. EMP2 was knocked down in APC shRNA cells using ribozymes targeting EMP2 (a kind gift from M. Wadehra, UCLA) (Wadehra et al. 2005), and cells were selected using 600µg/ml G418. Inhibition was confirmed using rt-PCR as described below.

2.3.2 Morphological assay

For 3D cultures, 5,000 cells per well were overlaid in single cell suspension of 2%

Matrigel media over a solid bed of 50 µl Matrigel (BD Biosciences; San Jose, CA, USA) in an 8 well chamber slide (protocol modified from Debnath et al., 2003). Cysts were grown for 7 days as previously described (Lesko et al. 2015). Cysts were imaged on an

EVOS fl microscope at room temperature with 20x objective and Sony ICX285AL CCD camera, size was assessed using ImageJ software, and significance was determined using a one-way ANOVA. Adobe Photoshop and Illustrator were used to generate figures. For inhibitor treatments, 8µg/ml AIIB2 (Developmental Studies Hybridoma Bank; Iowa City,

Iowa, USA) (Zawistowski et al. 2013) was applied at Day 1, and 50µM PP2 (Sigma-

Aldrich; St. Louis, MO, USA) or 2µM PF-04554878 (Cayman; Ann Arbor, MI, USA) were applied at Day 4. A two-way ANOVA was used to determine statistical significance.

2.3.3 Immunofluorescence (IF)

MDCK cysts were fixed in 4% paraformaldehyde at days specified. Cysts were permeabilized with 0.5% Triton-X-100, blocked in IF buffer (0.1% BSA, 0.2% Triton X-100,

0.05% Tween 20, and 0.1% goat serum in PBS) and incubated with anti-gp135 (1:100;

Developmental Studies Hybridoma Bank) (Mao et al. 2011), phospho-histone H3 (Cell signaling), or cleaved caspase 3 (Cell signaling) diluted in IF buffer overnight at room temperature. To visualize the staining, cysts were incubated in Alexa-conjugated secondary antibodies diluted in IF buffer (1:1000; Invitrogen A21422, lot:1180091;

Carlsbad, CA, USA) (Ausubel and al. 1992) for 1 hour at room temperature. Actin was

visualized using Alexa conjugated phalloidin (1:200; Invitrogen A-12379, lot: 1217967)

(Bubb et al. 1994). Slides were mounted with Fluoromount G, Hoescht dye, and TOPRO-

3, and imaged on a Zeiss 710 confocal at room temperature with Pin Apo 63x objective.

Images were analyzed with ZEN software and figures were generated with Adobe

Photoshop and Illustrator. At least 100 cysts were classified as either normal, exhibiting apical localization of gp135, or abnormal, exhibiting basal or mixed localization of gp135 and statistical analysis was done using Fisher’s exact test.

2.3.4 Migration assay

1 x 105 cells were plated per well in a 6 well plate. Once cells reached confluence, cells were treated with 2.5µg/ml mitomycin (Sigma-Aldrich) to inhibit proliferation and scratched with a 1-10µl pipette tip. For inhibitor treatments, cells were treated with 2.5µg/ml mitomycin and 6.25µM PP2, 8µg/ml AIIB2, or 2µM PF-04554878.

Wounds were imaged at 0, 24, and 48 hours post scratching with phase-contrast microscope (Evos fl) at room temperature with 20x objective and Sony ICX285AL CCD camera. Analysis was completed using TScratch software (developed by the

Koumoutsakos group (CSE Lab), at ETH Zürich (Gebäck 2008) and significance was determined using a one-way ANOVA. Adobe Photoshop and Illustrator were used to make figures.

2.3.5 Western Blotting

Protein lysates were isolated using 20mM Tris-HCL, 150mM NaCl, 1% Triton x-

100, 0.5% IGEPAL, 50mM NaF, 1mM Na3VO4, 5mM Sodium Pyrophosphate, phosphatase

inhibitors and protease inhibitors. 30µg of lysate were separated by an 8, 10, or 12% acrylamide gel and transferred to an Immobilon P membrane. Membranes were probed with the following primary antibodies diluted in 1% BSA in TBST: anti-phospho-FAK

(2D11) (1:1000; Santa Cruz sc-81493, lot:180091 ; Dallas, TX, USA) (Schlaepfer and

Hunter 1996), anti-FAK (1:1000; BD Biosciences 6180087, lot:40498) (Zeng et al. 2003), anti-phospho-Src (Tyr416) (1:1000; Cell Signaling 2101, lot:20; Danvers, MA, USA)

(Thomas and Brugge 1997), anti-Src (1:1000; Cell Signaling 2108, lot:8) (Thomas and

Brugge 1997), anti-CD29 (1:1000; 1 integrin, BD Biosciences 610467, clone 18) (Ivaska et al. 2002), and anti-β-Actin (1:25,000; Sigma-Aldrich A1978, clone AC-15) (Sawyer et al.

2003). Blots were analyzed with Image J software and a one-way or two-way ANOVA for inhibitor treatments was used to determine statistical significance. Adobe Photoshop and Illustrator were used to create figures.

2.3.6 Real-time PCR

RNA was isolated and cDNA was synthesized using iScript Reverse Transcriptase

(Biorad; Hercules, CA, USA). Real-time PCR was performed using SsoAdvanced universal supermix (Bio-Rad) and PrimePCR SYBR Green Assay: EMP2, Dog (Bio-Rad). Gene expression changes were quantified using the ΔΔC(t) method using transferrin receptor

(TFRC) as the housekeeping gene with the Bio-Rad FX Manager 3.0 Software. Statistics for experimental replicates were performed using a one-way ANOVA. Figures were constructed using Adobe Photoshop and Illustrator.

2.3.7 Cell adhesion assay

A 96 well plate was coated with Collagen I (Upstate; Charlottesville, VA, USA) overnight at 4˚C, and incubated with 1% BSA in PBS for 1 hour at 37˚C. 8 x 104 cells/well were plated and incubated for 1 hour at 37˚C. Adherent cells were stained with 0.1% crystal violet in 20% methanol and absorbance at 590nm was read. Statistical significance was determined with two-way ANOVA. Figures were created using Adobe

Photoshop and Illustrator.

2.3.8 Statistics

For one-way and two-way ANOVAs a Tukey post hoc test was preformed. For all data analysis a p-value < 0.05. was determined to be significant.

2.4 Results

2.4.1 APC loss upregulates Epithelial Membrane Protein 2 and β1 integrin.

To investigate the molecular mechanisms downstream of APC loss responsible for mediating the cyst size and polarity phenotypes, we turned to our previously published array data (Lesko et al. 2015). EMP2, which interacts with β1 integrin to induce signaling cascades (Fu et al. 2011, Fu et al. 2014), was upregulated in 3D cysts from APC shRNA cells (Lesko et al. 2015). Therefore, we sought to understand if EMP2 and β1 integrin signaling are responsible for APC regulation of morphogenesis and polarity. To evaluate if EMP2 upregulation caused by APC loss increases cyst size and inverts polarity, we utilized a ribozyme targeting EMP2 (Wadehra et al. 2005) to create

APC shRNA cell lines with stable EMP2 knockdown (APC shRNA EMP2KD), and confirmed knockdown with rt-PCR (Figure 2.1 A). Next we evaluated β1 integrin protein expression in MDCK, ctl shRNA, APC shRNA, and APC shRNA EMP2KD cells (Figure 2.1 B). Image J analysis and quantification displayed that β1 integrin is significantly upregulated in APC shRNA cells compared to controls (Figure 2.1 B,C). Although EMP2 has been shown to upregulate β1 integrin (Fu et al. 2011, Fu et al. 2014), EMP2 knockdown in APC shRNA cells did not significantly decrease β1 integrin expression (Figure 2.1 B,C). These data suggest that upregulation of EMP2 and β1 integrin in APC shRNA cells are two distinct possible mechanisms by which APC loss increases cyst size accompanied by inversion of polarity.

Figure 2.1 APC loss upregulates Epithelial Membrane Protein 2 and β1 integrin. A) rt-PCR data confirms that EMP2 is upregulated in APC shRNA cells, and that hEMP2 ribozyme knocks down EMP2 expression in MDCK APC shRNA cells. Expression is normalized to TRFC expression and is plotted as fold change expression. B) Western blot showing β1 integrin protein expression in MDCK, ctl shRNA, APC shRNA, and APC shRNA EMP2KD cells. Actin is utilized as the loading control. C) Quantification using Image J shows that β1 integrin is upregulated in MDCK cells upon APC loss. However, EMP2 knockdown in APC shRNA cells does not decrease β1 integrin expression. Experiments were replicated three times and data are presented as mean ± s.d. and a one-way ANOVA was utilized to determine significance, * p < 0.05.

2.4.2 APC regulates cyst size through EMP2 and β1 integrin signaling.

Given that APC loss increases EMP2 and β1 integrin, and β1 integrin has been implicated in polarity (Ojakian and Schwimmer 1994, Yu et al. 2005, Zovein et al. 2010), we next evaluated the role of EMP2 and β1 integrin in APC-mediated cyst size. To assess the effect of EMP2 knockdown in APC shRNA cysts, MDCK control, APC shRNA, and APC shRNA EMP2KD cells were grown in 3D Matrigel culture for 7 days (Figure 2.2 A). As we previously described, the APC shRNA cysts, which arise from a population of single cells, were larger (Lesko et al. 2015), with no change in cell proliferation or apoptosis as measured by phospho-histone H3 or cleaved caspase 3 respectively (Figure 2.3). Image J analysis confirmed that EMP2 knockdown decreased cyst size of APC shRNA EMP2KD cells compared to APC shRNA cells, and that the double knockdown resulted in areas similar to MDCK control cells (Figure 2.2 B). To test whether APC mediates cyst size through β1 integrin, we blocked 1 integrin with the AIIB2 antibody, which inhibits the interaction of 1 integrin with Collagen I. To measure the functionality of AIIB2 in the

APC shRNA cells, we assessed cell adhesion to Collagen I, and found that AIIB2 significantly reduces adhesion (Figure 2.4 A). MDCK, ctl shRNA, and APC shRNA cells were plated in 3D Matrigel culture, treated with AIIB2 on Day 1 and grown to Day 7

(Figure 2.2 C). Image J analysis was used to quantify area and indicated that inhibition of

β1 integrin significantly decreased cyst size in APC shRNA cysts (Figure 2.2 D). AIIB2 treatment significantly increased the area of MDCK and ctl shRNA cysts Figure 2.2 D) consistent with previous studies showing AIIB2 treatment of MDCK cells increased cyst size due to aberrant intracellular signaling caused by inversion of polarity (Yu et al. 68

2005)¸ Given that EMP2 and β1 integrin activate FAK/Src signaling, we assessed the effects of inhibiting FAK/Src signaling on cyst size using the following chemical inhibitors: PF-04554878 (targeting FAK) or PP2 (targeting Src). Inhibition of phosphorylated protein expression by PF-04554878 and PP2 was measured using western blots of phospho-FAK and phospho-Src (Figure 2.4 B-D). MDCK, ctl shRNA, and

APC shRNA cells were plated in 3D Matrigel culture and treated with inhibitors on Day 4 to avoid long-term toxicity to the cysts. Cysts were grown to Day 7 and size was assessed using Image J analysis (Figure 2.2 C). Inhibiting FAK and Src decreased APC shRNA cyst size to areas comparable to MDCK and ctl shRNA cysts (Figure 2.2 D).

Together, these data suggest that EMP2, β1 integrin, and FAK/Src signaling impact APC’s regulation of 3D cyst size.

Figure 2.2 EMP2 knockdown and inhibition of β1 integrin signaling decreases size of APC shRNA cysts. A) Phase contrast images on Day 7 of MDCK, APC shRNA, and APC shRNA EMP2KD cysts. Scale bar = 200μm. B) Quantification of area using Image J reveals that EMP2 knockdown in APC shRNA cells significantly decreases cyst size. C) Phase contrast images on Day 7 of MDCK, ctl shRNA, and APC shRNA cells in 3D culture treated with the following inhibitors: AIIB2 (targeting β1 integrin), PF-04554878 (targeting FAK), and PP2 (targeting Src). Scale bar = 200μm. D) Quantification using Image J shows inhibition of β1 integrin, FAK, and Src significantly decreases APC shRNA cyst size. Data from three replicates are presented as mean ± s.d. and a one-way (B) or two-way (D) ANOVA determined significance, * p < 0.05.

Figure 2.3 Proliferation and apoptosis are not affected by APC loss. A) Quantification of phospho-histone H3 staining showed no significant changes in proliferation between APC shRNA cysts compared to control cysts at Day 3, 5, or 7. B) No significant changes were observed in cleaved caspase 3 staining in cysts with APC loss compared to MDCK controls at Day 7. Data from three replicates are presented as mean ± s.d. and a one-way ANOVA determined significance, * p < 0.05.

Figure 2.4 Chemical inhibitors target β1 integrin signaling pathway. A) AIIB2 functions to inhibit 1 integrin by blocking its interaction with Collagen I. Absorbance of crystal violet in cell adhesion assays was utilized to confirm the activity of AIIB2 to inhibit 1 integrin mediated cell adhesion to Collagen I. APC shRNA cells treated with AIIB2 exhibited decreased absorbance of crystal violet indicative of inhibited 1 integrin/Collagen I interactions. B) Western blot analysis for phospho-FAK after PF- 04554878 treatment. C) Phospho-FAK protein expression is normalized to DMSO treatment within each cell line, and is significantly decreased in APC shRNA cells treated with PF- 04554878 as shown by quantification using Image J. D) Western bot analysis for phospho-Src after PP2 treatment. E) Phospho-Src expression is normalized to DMSO treatment within each cell line. Analysis using Image J shows PP2 significantly inhibits phospho- Src expression in APC shRNA cells. Data of four experimental replicates are presented as mean ± s.d. and a two-way ANOVA was performed to determine significance, * p < 0.05.

2.4.3 APC regulates apical-basal polarity through EMP2.

We next investigated the mechanisms by which APC mediates apical-basal polarity. MDCK, APC shRNA, and APC shRNA EMP2KD cells were grown in 3D Matrigel culture for 7 days and immunofluorescence and confocal microscopy for the apical marker gp135 was utilized to evaluate polarity in cysts with APC loss and EMP2KD (Figure

2.5 A). Quantification of the cysts with normal (apical) or abnormal (basal or mixed) gp135 localization demonstrated that EMP2KD in APC shRNA cells restores gp135 to the apical membrane (Figure 2.5 B), suggesting that APC mediates epithelial polarity through EMP2. Given that β1 integrin protein expression is increased in APC shRNA cells, we next evaluated the role of β1 integrin, FAK, and Src in APC-mediated apical- basal polarity. Although β1 integrin signaling has been shown to be necessary for the establishment of polarity in MDCK cells (Yu et al. 2005), we hypothesize that this process must be tightly regulated and that aberrant signaling may also invert apical-basal polarity. MDCK, ctl shRNA, and APC shRNA cells were treated with AIIB2, PF-04554878, or PP2 and grown in 3D Matrigel culture for 7 days, and confocal microscopy analysis for the apical marker gp135 was utilized to evaluate epithelial polarity (Figure 2.5 C).

Quantitative analysis showed that inhibiting β1 integrin, FAK, or Src did not restore the inverted polarity of APC shRNA cysts from the basal to the apical membrane (Figure 2.5

D). In control cysts, inhibition of β1 integrin mislocalized gp135 from the apical membrane to the basal membrane (Figure 2.5 D), which is consistent with previous studies (Yu et al. 2005). Additionally, inhibition of Src inverted polarity in 90% of the

MDCK and 70% of the ctl shRNA cysts, however this effect was suppressed in APC shRNA 74

cells which may be attributed to increased EMP2 expression and levels of Src of activation (Figure 2.5 D). Therefore, APC regulates apical-basal polarity through EMP2 independent of β1 integrin and downstream signaling.

Figure 2.5 Apical polarity in APC shRNA cysts is restored by EMP2 knockdown. A) Immunofluorescence confocal images of MDCK, APC shRNA, and APC shRNA EMP2KD for gp135 (red, indicated by white arrows) and nuclei, (TOPRO-3, blue). Scale bar = 50μm. B) Quantification of polarity as normal (apical) or mislocalized (basal and mix) reveals EMP2 knockdown restores gp135 localization to the apical membrane. C) Immunofluorescence confocal images of MDCK, ctl shRNA, and APC shRNA cells treated with β1 integrin signaling inhibitors AIIB2 (targeting β1 integrin), PF-04554878 (targeting FAK), and PP2 (targeting Src) for gp135 (green, indicated by white arrows), actin via phalloidin (red), and nuclei (TOPRO-3, blue). Scale bar = 50μm. D) Quantification of polarity as normal (apical) or mislocalized (basal or mix) suggests inhibition of β1 integrin signaling does not restore apical polarity. Data are presented as mean percentage and Fisher’s exact test was utilized to determine significance, * p < 0.05, compared to control MDCK cells. Experiments were replicated three times.

2.4.4 APC loss increases cell motility through 1 integrin and Src.

The larger and more disorganized cysts that develop upon APC loss (Lesko et al.

2015) led us to investigate whether the APC shRNA MDCK cells were more migratory. To evaluate the role of APC in regulating cell motility we utilized wound-healing assays.

MDCK and APC shRNA cells were grown to confluence, treated with mitomycin C to inhibit proliferation, scratched, and imaged over 48 hours (Figure 2.6 A). Using the

TScratch program developed by the Koumoutsakos group (CSE Lab), at ETH Zürich

(Gebäck 2008), we quantified the percent filled area of the scratches and found that at

48 hours APC shRNA cells filled 90% of the wound area while MDCK cells filled 45% of the wound area (Figure 2.6 B), indicating that the APC shRNA cells migrate significantly faster than control cells. Since EMP2 upregulation and activation of FAK/Src signaling increase epithelial cell migration (Fu et al. 2011, Fu et al. 2014), we next sought to test if these signaling molecules were responsible for the increased cell migration in APC shRNA cells. Wound-healing assays were utilized as above for MDCK, APC shRNA, and

APC shRNA EMP2KD cells (Figure 2.6 C). Analysis with TScratch software revealed that

APC shRNA EMP2KD cells filled in 90% of the wound area at 48 hours and therefore

EMP2 knockdown was unable to alleviate the increased migration caused by APC loss

(Figure 2.6 D). Next we assessed whether chemical inhibition of 1 integrin, FAK, or Src decreases migration of APC shRNA cells (Figure 2.6 C). APC shRNA cells treated with

AIIB2, PF-04554878, or PP2 only filled in 40%, 50%, and 10% of the wound area, respectively (Figure 2.6 E), indicating that inhibition of 1 integrin and downstream signaling alleviated increased cell motility upon APC loss. Although other studies provide 77

evidence for EMP2, β1 integrin, FAK, and Src in mediating epithelial cell migration, our studies demonstrate that APC regulates cell motility through β1 integrin independent of

EMP2.

Figure 2.6 Increased migration in APC shRNA cells is controlled by β1 integrin signaling. A) Phase contrast images of MDCK and APC shRNA cells during wound-healing assays at 0, 24, and 48 hours. Scale bar = 400μm. B) Analysis using TScratch software shows that APC shRNA cells fill a significantly larger wound area compared to MDCK controls at 48 hours. C) Phase contrast images of APC shRNA cells with EMP2 knockdown or treated with the following inhibitors: AIIB2 (targeting 1 integrin), PF-04554878 (targeting FAK), or PP2 (targeting Src) during wound-healing assays at 0, 24, and 48 hours. Scale bar = 400μm. D) TScratch quantification of percent filled area at 48 hours indicates rapid migration in APC shRNA cells was not alleviated by EMP2 knockdown. E) Quantification of percent filled area at 48 hours shows that inhibition of β1 integrin, FAK, and Src decreases migration in APC shRNA cells. Experiments were replicated three times and data are presented as mean ± s.d. and a student’s t-test (B and D) or a two-way ANOVA (E) was performed to determine significance, * p < 0.05.

2.5 Discussion

APC loss increases cyst size accompanied by inverted apical-basal polarity (Lesko et al. 2015), suggesting a necessary role for APC in mediating 3D morphology and epithelial polarity. We showed that these phenotypes are independent of Wnt signaling as the Wnt pathway was not activated and the C-terminal fragment of APC, lacking the

-catenin binding domain, partially restored cyst size and polarity (Lesko et al. 2015).

While we previously demonstrated that APC loss upregulates EMP2 mRNA expression

(Lesko et al. 2015), the precise molecular mechanisms of APC-mediated polarity remain unknown. With the data presented here, we show for the first time that APC loss upregulates β1 integrin protein expression and increases migration during wound- healing assays. Through analysis of cyst size, apical-basal polarity, and migration in APC shRNA cells with genetic or chemical manipulation of EMP2 or β1 integrin, we identified two novel mechanisms by which APC regulates these functions Figure 2.7.

Figure 2.7 Schematic of APC-mediated mechanisms. APC loss increases migration in wound-healing assays and increases cyst size accompanied by inverted polarity in 3D culture. The data presented here identify the molecular mechanisms by which APC regulates these processes. Knockdown of EMP2 reduces cyst size and restores apical polarity of APC shRNA cysts, but has no impact on the enhanced migration. Inhibition of β1 integrin signaling alleviates increased cyst size and migration due to APC loss, with no effect on the inversion of polarity.

Through knockdown of EMP2 or inhibition of 1 integrin signaling, APC was found to mediate cyst size via both molecular pathways. Although previous studies indicate that EMP2 upregulates α61 integrin expression at the plasma membrane

(Wadehra et al. 2002, Wadehra et al. 2005), and that EMP2 and 1 integrin both increase phosphorylation of FAK and Src (Cary et al. 1996, Parsons 2003, Morales et al.

2009, Fu et al. 2011), we have shown EMP2 knockdown does not affect 1 integrin expression (Figure 2.1 B,C) suggesting APC is mediating cyst size through two distinct mechanisms. Beyond cyst size, we demonstrated that the molecular mechanisms differ depending on phenotype (i.e., polarity and migration). Knockdown of EMP2, and not inhibition of β1 integrin signaling, restored apical localization of gp135 in APC shRNA cysts. Previous studies (Yu et al. 2005) suggest that β1 integrin signaling is necessary for the establishment of apical-basal polarity. However our data demonstrate that APC regulates polarity through an EMP2-mediated mechanism independent of β1 integrin.

Although EMP2 interacts with β1 integrin, only approximately 60% of EMP2 co- immunoprecipitates with β1 integrin in fibroblasts (Wadehra et al. 2002). The existence of two distinct pools of EMP2 supports our model by which EMP2 regulates APC- mediated cyst size and apical-basal polarity through two distinct mechanisms (Figure

2.7).

We show here that APC loss in MDCK normal epithelial cells increased migration during wound healing assays. This suggests that a part of the tumor suppressive function of APC may be through controlling cell motility. Our findings in normal epithelial cells are contradictory to previous studies (Sansom et al. 2004, Kroboth et al. 83

2007, Odenwald et al. 2013) showing that APC loss in multiple fibroblastic or malignant cell types decreases cell migration. Therefore APC may mediate cell motility depending on both the tissue type and stage of tumorigenesis. Despite the role of EMP2 in mediating cyst size and epithelial polarity, EMP2 knockdown had no effect on cell motility in APC shRNA cells. However, inhibition of β1 integrin, FAK, and Src selectively decreased migration in APC shRNA cells at the doses used. While these signaling pathways have been extensively studied in cell motility ((Matsumoto et al. 2010, Fang et al. 2014) and reviewed in (Hood and Cheresh 2002)), the present study is the first to link

APC with this Wnt-independent 1 integrin-mediated cell migration. An additional mechanism by which APC may regulate β1 integrin-mediated cell motility is through cytoskeletal interactions. APC interacts with the cytoskeleton through multiple binding domains; most notably through direct binding of actin and microtubules in the C- terminal domains (reviewed in (Prosperi and Goss 2011)). Studies have shown that APC promotes nucleation and actin assembly (Okada et al. 2010) and associates with the plus ends of microtubules to stabilize and promote assembly (Munemitsu et al. 1994).

These interactions may be important for APC’s regulation of cell motility as the APC C- terminus is required for apical extrusion of human bronchial epithelial cells (Marshall et al. 2011). Furthermore, β1 integrin is linked to actin at focal adhesions (Critchley 2000), and β1 integrin signaling is required for the localization of microtubule plus ends to focal adhesion (Krylyshkina et al. 2003, Akhtar and Streuli 2013), suggesting that APC may mediate cell migration by regulating β1 integrin signaling and cytoskeletal interactions.

Future studies will test the hypothesis that loss of APC and upregulation of β1 integrin promote microtubule and focal adhesion interactions to increase cell migration.

The data presented here suggest a role of EMP2 in mediating cyst size and apical-basal polarity; however the mechanism by which APC regulates EMP2 remains unknown. APC may indirectly regulate EMP2 by mediating transcription factors. Studies have shown EMP2 mRNA expression is induced by the cAMP responsive element binding protein 1 (CREB1) (Li et al. 2015) or the combination of estradiol and progesterone (Wadehra et al. 2008). Other candidate transcription factors that have predicted binding sites in the EMP2 promoter and may be mediated by APC to upregulate EMP2 include activator protein 1 (AP-1), a downstream target of β1 integrin

(Guo and Giancotti 2004), signal transducer and activator of transcription 3 (STAT3), activated by loss of Par3 and polarity (Guyer and Macara 2014), and paired box gene 2

(PAX2), an important regulator of kidney morphogenesis (Winyard et al. 1996). It is also possible that APC may directly interact with EMP2 through the APC C-terminus since

EMP2 knockdown or the reintroduction of the APC C-terminus in APC shRNA cells decreased cyst size and restored apical-basal polarity (Figure 2.5 A,B and (Lesko et al.

2015)). Current studies (discussed in Chapter 4) are ongoing in the laboratory to assess the precise mechanisms by which APC regulates EMP2.

As a multifunctional scaffolding protein, APC regulates several biological processes including 3D morphogenesis, apical-basal polarity, and cell motility that are critical for cellular maintenance and development. Loss of APC and subsequent loss of these functions promotes the progression of several diseases such as cancer and chronic 85

kidney disease ((Qian et al. 2005) and reviewed in (Lesko et al. 2014)). Therefore, it is important to understand the molecular mechanisms by which APC controls these processes. The current studies reveal that both knockdown of EMP2 and inhibition of β1 integrin restored increased cyst size due to APC loss, while inhibition of β1 integrin signaling alone alleviated increased cell migration in APC shRNA cells (Figure 2.7).

Surprisingly, EMP2 regulates APC-mediated apical-basal polarity independent of β1 integrin signaling (Figure 2.7). Therefore the signaling pathways downstream of EMP2 that influence polarity remain unknown. We have previously shown that the C-terminus of APC restores apical polarity in APC shRNA cells (Lesko et al. 2015). Investigating APC

C-terminus protein interactions may provide insight into the downstream mechanisms of EMP2-mediated polarity. The APC C-terminus contains a PDZ binding domain that interacts with the polarity proteins Dlg and Scrib which form a basolateral polarity complex with Lethal giant larvae (Lgl) (Matsumine et al. 1996, Takizawa et al. 2006). The loss of these proteins has been shown to result in loss of polarity and tissue overgrowth in Drosophila ((Bilder 2004) and reviewed in (Schluter and Margolis 2012)), and the loss of Scrib promotes mammary tumorigenesis (Zhan et al. 2008). Further studies will be needed to determine a role for APC C-terminus and Scrib interactions in regulating apical-basal polarity. With the data presented here, we identify a model in which APC mediates cyst size, epithelial polarity, and cell motility through distinct molecular mechanisms. Although these findings warrant further investigation into the details of downstream signaling mechanisms, here we have identified novel therapeutic targets, including EMP2 and β1 integrin, for the treatment of APC-mediated diseases. 86

CHAPTER 3:

MOLECULAR MECHANSIMS OF APC/EMP2-MEDIATED APICAL-BASAL POLARITY

3.1 Abstract

Adenomatous Polyposis Coli (APC) is lost in several epithelial cancers and is an important regulator of numerous normal cellular processes including maintenance of apical-basal polarity, cell migration, and intracellular signaling. We previously demonstrated that loss of APC in Madin Darby Canine Kidney (MDCK) cells upregulated

Epithelial Membrane Protein 2 (EMP2) expression, increased cyst size, and disrupted polarity. Our lab was the first to identify a role for EMP2 in controlling polarity and this regulation was independent of β1 integrin signaling. However, the molecular mechanisms downstream of APC and EMP2 that influence polarity remain unknown.

With these studies we determined that Caveolin 1 (Cav1)/extracellular signal-regulated kinase (ERK)/jun N-terminal kinase (JNK) and Scribble (Scrib)/Hippo pathways are not downstream mechanisms of APC/EMP2-mediated polarity. By utilizing 2D Difference gel electrophoresis (DIGE) gel analysis we identified several other possible downstream effectors of polarity including filamin A or plectin isoform X2, histone H3.3, histone H2A type 1-E-like, and 3-hydroxyacyl-CoA dehydrogenase type-2. Furthermore, pathway analysis revealed calcium/phospho-lipid binding and genes associated with DNA-binding and nucleosomes are enriched upon APC loss. Together these studies explored known signaling modalities as downstream effectors of APC and EMP2 mediated polarity and identified several novel proteins that may be regulated by APC and EMP2. Further 87

experiments are needed to better understand how these candidates contribute to APC and EMP2 mediated apical-basal polarity and cyst size.

3.2 Introduction

Epithelial Membrane Protein 2 (EMP2) is a tetraspan membrane protein which regulates several cellular functions including lipid raft formation and apical membrane recycling, proliferation and apoptosis, and cell migration and adhesion (reviewed in

(Wang et al. 2017)). EMP2 mediates the formation of lipid rafts by specifically recruiting glycosylphosphatidyl inositol-anchored proteins and negative regulating Cav1 (Wadehra et al. 2003, Wadehra et al. 2004). Normally highly expressed in lung tissue, EMP2 is downregulated in lung cancer cells causing the upregulation of Cav1 and activation of

ERK and JNK pathways (Lee et al. 2016). Additionally, knockout of EMP2 in zebrafish kidney upregulated Cav1 expression (Wan et al. 2016) suggesting a conserved role for

EMP2 in controlling Cav1 expression and lipid raft composition. In addition to regulating lipid composition, EMP2 also recruits integrins to the plasma membrane to regulate adhesion and cell migration (Wang et al. 2013). Furthermore, β1 integrin/FAK/Src signaling is activated in endometrial and breast cancer cell lines to increase migratory and invasive phenotypes ((Fu et al. 2011, Fu et al. 2014) and reviewed in (Wang et al.

2017)).

Depending on tissue type, EMP2 has been implicated as both an oncogene and tumor suppressor (reviewed in (Wang et al. 2017)). For example, EMP2 is overexpressed in ovarian, endometrial, and breast cancers as well as glioblastomas (Fu et al. 2010,

Habeeb et al. 2010, Fu et al. 2014), while EMP2 is lost in upper urinary tract, nasopharyngeal, and lung cancers (Wang et al. 2013, Ahmed et al. 2015, Lee et al. 2016).

Studies from our lab determined EMP2 was upregulated upon APC loss and identified a novel role for EMP2 in mediating cell polarity (Lesko and Prosperi 2017). MDCK cells grown in 3D Matrigel culture with APC knockdown (APCKD) exhibit increased cyst size and disrupted polarity (Lesko et al. 2015). These early tumorigenic phenotypes were caused by increased EMP2 expression or activation of β1 integrin signaling although these two pathways acted independently (Lesko and Prosperi 2017). Therefore the mechanisms that influence polarity and cyst size downstream of APC and EMP2 remain unknown.

Recent studies by Cai et. al. provided evidence for a role of Hippo signaling in

APC-mutant intestinal tumorigenesis (Cai et al. 2015). Hippo signaling, which controls organ growth and size, is active when Salvador (Sav) and Large tumor suppressor kinase

(Lats) are phosphorylated and subsequently phosphorylate yes-associated protein (YAP) to target it for degradation (Zhao et al. 2010). When Hippo is inactive YAP is no longer phosphorylated, translocates into the nucleus, and drives the transcription of several proliferative targets (Zhao et al. 2010). Although the mechanisms are not yet fully understood, there is evidence that polarity proteins are involved in the Hippo signaling pathway. For example, in Drosophila loss of Scrib or Lgl results in the upregulation of the

YAP homologue Yorki (Chen et al. 2012). APC interactions with Scrib (Takizawa et al.

2006) and the Sav/Lats complex (Cai et al. 2015) as well as the requirement for YAP

activation in APC-mutant adenocarcinomas suggest the Hippo pathway a possible mechanism by which APC and EMP2 may control apical-basal polarity and cyst size.

The current studies assess the Cav1/ERK/JNK and Scrib/Hippo pathways as possible downstream mechanisms of APC/EMP2-mediated polarity and cyst size.

However, we show that APC and EMP2 do not regulate either of these signaling modalities, and that the mechanisms downstream of APC/EMP2 that influence polarity are novel. We have identified filamin A or plectin isoform X2, histone H3.3, histone H2A type 1-E-like, and 3-hydroxyacyl-CoA dehydrogenase type-2 as possible downstream effectors of polarity through 2D DIGE gel analysis. Furthermore, pathway analysis revealed calcium/phospho-lipid binding and genes associated with DNA-binding and nucleosomes are enriched upon APC loss. Future studies are needed to validate these candidates and determine if they contribute to APC and EMP2 mediated polarity and cyst size.

3.3 Materials and Methods

3.3.1 Cell culture

Madin Darby Canine Kidney (MDCK) cells were obtained from K. Matlin

(University of Chicago), tested for contamination, and maintained in DMEM media with

5% FBS, 2mM L-glutamine, 10mM HEPES, and 1% Penicillin/Streptomycin. APCKD cells were established previously (Lesko et al. 2015), and were grown in DMEM media with

5% FBS, 2mM L-glutamine, 10mM HEPES, 1% Penicillin/Streptomycin, and 2µg/ml

Puromycin. APCKD EMP2KD cells were established previously (Lesko and Prosperi 2017) 90

and cells were grown in DMEM media with 5% FBS, 2mM L-glutamine, 10mM HEPES, 1%

Penicillin/Streptomycin, 2µg/ml Puromycin and 600µg/ml G418. EMP2 was overexpressed (EMP2OE) using the mEMP2 pEGFP-N3 plasmid (a kind gift from M.

Wadehra, UCLA) described in (Wadehra et al. 2005).MDCK EMP2OE cells were grown in

DMEM media with 5% FBS, 2mM L-glutamine, 10mM HEPES, 1%

Penicillin/Streptomycin, 2µg/ml Puromycin and 600µg/ml G418. MDA-MB-231 cells were grown in DMEM media with 5% FBS and 1% Penicillin/Streptomycin.

3.3.2 3D culture IF

For 3D cultures, 5,000 cells per well were overlaid in single cell suspension of 2%

Matrigel media over a solid bed of 50 µl Matrigel (BD Biosciences) in an 8 well chamber slide (protocol modified from (Debnath et al. 2003)). Cysts were grown for 7 days as previously described (Lesko et al. 2015) and fixed in 4% paraformaldehyde. Cysts were permeabilized with 0.5% Triton-X-100, blocked in IF buffer (0.1% BSA, 0.2% Triton X-100,

0.05% Tween 20, and 0.1% goat serum in PBS) and incubated with Scrib (C-20)(1:200;

Santa Cruz sc-11049) diluted in IF buffer overnight at room temperature. To visualize the staining, cysts were incubated in Alexa-conjugated secondary antibodies diluted in IF buffer (1:1000; Invitrogen A21422) for 1 hour at room temperature. Actin was visualized using Alexa conjugated phalloidin (1:200; Invitrogen A-12379). Slides were mounted with Fluoromount G, Hoescht dye, and TOPRO-3, and imaged on a Zeiss 710 confocal at room temperature with Pin Apo 63x objective. Images were analyzed with ZEN software and figures were generated with Adobe Photoshop and Illustrator.

3.3.3 2D IF

Cells were serum starved for 24 hours and serum was reintroduced into positive control wells for 1 hour before cells were fixed in 3.7% formaldehyde. Cells were permeabilized with 0.3% Triton-X-100 and incubated with Anti-YAP (1:400; Abnova

H00010413-M01) diluted in IF buffer for 1 hour at room temperature. To visualize the staining, cysts were incubated in Alexa-conjugated secondary antibodies diluted in IF buffer (1:1000; Invitrogen A21422) for 1 hour at room temperature. Actin was visualized using Alexa conjugated phalloidin (1:200; Invitrogen A-12379). Slides were mounted with Fluoromount G, Hoescht dye, and imaged on a Zeiss Axio Imager A.1 microscope at room temperature with 63x objective.

3.3.4 Western Blotting

Protein lysates were isolated using 20mM Tris-HCL, 150mM NaCl, 1% Triton x-

100, 0.5% IGEPAL, 50mM NaF, 1mM Na3VO4, 5mM Sodium Pyrophosphate, phosphatase inhibitors and protease inhibitors or fractionation was performed using a Nuclear

Extraction Kit (Affymetrix). 30µg of lysate were separated by a 7, 10, or 12% acrylamide gel and transferred to an Immobilon P membrane. Membranes were probed with the following primary antibodies diluted in either 5% NFDM or 1% BSA in TBST: Caveolin-1

(1:1000, CST 3238), p44/43 MAPK (ERK 1/2) (1375) (1:1000, CST 4695), filamin-A

(1:1000; CST #4762), anti-human JNK (1:1000, BD 551197), plectin-1 (D6A11) (1:1000,

CST 12254), phospho-SAPK/JNK (Thr183/Tyr185) (81E11) (1:1000, CST #4668), phospho- p44/42MAPK (Thr202/Tyr204) (20G11) (1:1000; CST #4276), Scrib (C-20) (1:200, Santa

Cruz sc-11049), Anti-YAP1 (1:1000 Abnova H00010412-M01), and anti-β-Actin (1:25,000;

Sigma-Aldrich A1978). Blots were analyzed with Image J software and a one-way ANOVA was used to determine statistical significance. Adobe Photoshop and Illustrator were used to create figures.

3.3.5 Statistics

For one-way and two-way ANOVAs a Tukey post hoc test was preformed. For all data analysis a p-value < 0.05. was determined to be significant.

3.4 Results

3.4.1 APC and EMP2 do not regulate known downstream signaling pathways of EMP2

Very few molecular signaling pathways have been identified downstream of

EMP2. Therefore we initially assessed pathways evaluated in the literature including β1 integrin/FAK/SRC (Fu et al. 2011, Fu et al. 2014) and Cav1/ERK/JNK (Lee et al. 2016, Wan et al. 2016) to test if these pathways were downstream of APC/EMP2 interactions and influence polarity in our MDCK 3D culture model. Although β1 integrin is upregulated in

MDCK cells with APC loss and inhibition of β1 integrin is able to rescue the cyst size phenotype we observe upon APC loss, we have previously shown that β1 integrin acts independently from EMP2 (Lesko and Prosperi 2017). Given that EMP2 downregulation in lung cancer cells increases Cav1 expression and activates ERK/JNK signaling (Lee et al.

2016), we hypothesized that EMP2 upregulation caused by APC loss would decrease

Cav1 expression and inhibit the activation of ERK/JNK signaling to influence polarity. To

test this hypothesis we performed western blots for Cav1, p-ERK, ERK, p-JNK, and JNK in

APCKD cells compared to control MDCK cells. Additionally, we tested if knockdown of

EMP2 in APCKD cells (APCKD EMP2KD) would restore any changes in expression and conversely, if overexpression of EMP2 (MDCK EMP2OE) would mimic changes in APCKD protein expression (Figure 3.1 A). Quantification revealed no significant changes in any proteins from the Cav1/ERK/JNK pathway between all of the cell lines (Figure 3.1 B) suggesting EMP2 does not mediate Cav1/ERK/JNK signaling in our MDCK model system.

These experiments as well as our previous findings from Lesko et. al., 2017 demonstrate that APC/EMP2 interactions do not function through any known downstream pathways of EMP2 to control cyst size and polarity.

Figure 3.1 APC and EMP2 do not regulate Cav1/ERK/JNK signaling. A) Western blotting for targets in the Cav1/ERK/JNK signaling pathways known to be mediated by EMP2 was performed to determine if these pathways are downstream of APC/EMP2 interactions. B) No significant changes in expression were observed in any of the cell lines upon quantification suggesting EMP2 regulates APC-mediated polarity through a novel mechanism in MDCK cells. Data from four replicates are presented as mean ± s.d. and a one-way ANOVA was used to determine significance.

3.4.2 Scribble and Hippo are not downstream of APC and EMP2

Because APC binds Scribble (Takizawa et al. 2006) and Scrib can mediate Hippo signaling which controls tissue growth and polarity (Chen et al. 2012), we assessed Scrib and Hippo as possible downstream regulators of APC/EMP2-mediated cyst size and polarity. First we assessed Scrib localization in both 2D and 3D culture using IF.

Fluorescent images revealed that in cells grown on plastic (2D) Scrib localization is unchanged in APCKD cells from its normal localization in the basolateral membrane in control cells (Figure 3.2 A). Similarly, when grown in 3D Matrigel culture, Scrib is still localized to the basolateral membrane in both MDCK and APCKD cysts (Figure 3.2 B).

Next we assessed whether Scrib expression was changed upon APC loss utilizing western blots. No significant changes were observed in Scrib expression between MDCK control and APCKD cells (Figure 3.2 C). Together these studies indicate Scrib localization and expression remain unchanged with loss of APC and therefore Scrib is not a downstream target of APC/EMP2 interactions.

Figure 3.2 Scribble localization and expression is unchanged by APC loss. IF for Scribble (red), phalloidin (green), and Hoescht (blue) in 2D (A) and TOPRO-3 (blue) in 3D Matrigel culture (B) demonstrates APC loss does not mislocalize Scrib from the basolateral membrane. C) APCKD cells do not exhibit changes in Scrib expression compared to controls in western blot analysis. Data from three replicates are presented as mean ± s.d. and a one-way ANOVA was utilized to determine significance.

To assess Hippo pathway inactivation we measured translocation of YAP to the nucleus utilizing two methods, IF to determine nuclear YAP localization and western blotting with fractionated lysates to identify nuclear YAP expression. As a positive control, nuclear YAP localization was induced in serum starved MDA-MB-231 breast cancer cells by treating with serum for 1 hour before fixation (Yu et al. 2012). After serum induction almost all YAP staining is localized to the nucleus of MDA-MB-231 cells compared to controls (Figure 3.3 A). However, very little nuclear YAP was observed in the nucleus of both MDCK and APCKD cells in either control or serum starved conditions

(Figure 3.3 A), suggesting the Hippo pathway is active and YAP is phosphorylated and degraded in the cytoplasm. Next, we used fractionated lysates to measure YAP expression in cytoplasmic and nuclear cell lysates. Normalization of YAP western blots to either HDAC or GAPDH, nuclear and cytoplasmic control genes respectively, revealed no significant changes in YAP expression in either the nucleus or cytoplasm of MDCK cells versus APCKD cells (Figure 3.3 B). Together, these data show that APC does not regulate both Scrib and Hippo pathway and have determined the polarity phenotypes observed upon APC loss are not driven by disruption of Hippo signaling.

Figure 3.3 YAP is not localized to the nucleus of MDCK cells with or without APC loss. A) IF for YAP (green) and phalloidin (red) was performed on cells that had been serum starved for 24 hours or cells that were serum starved for 23 hours and serum induced for 1 hour. YAP localizes to the nucleus of MDA-MD-231 cells upon serum induction; however, neither MDCK control cells nor APCKD cells showed nuclear localization of YAP. Images were taken at 63x and scale bar = 20μm B) Nuclear fractionation determined that YAP expression is unchanged between MDCK controls and APCKD cells in both the nucleus and cytoplasm suggesting Hippo signaling is actively degraded YAP. Experiments were all replicated three times and data are presented as mean ± s.d. A two-way ANOVA determined significance.

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101

3.4.3 2D DIGE analysis identified several candidates by which APC and EMP2 may control polarity

Since we determined that the known mechanisms downstream of EMP2, β1 integrin/FAK/Src and Cav1/ERK/JNK, and the Hippo pathway are not implicated in our system we decided to take a broader approach to identify possible downstream candidates. Therefore, we preformed 2D DIGE analysis (Applied Biomics) which separates protein lysates based on both size and pH to assess changes in protein expression. Lysates from several pairs of cell lines were compared: MDCK vs APCKD;

Reintroduction of full-length APC (APCFL) vs APCKD, APCKD vs APCKD EMP2KD, MDCK vs

APCFL, and MDCK vs APCKD EMP2KD and two representative gels are shown here (Figure

3.4). Protein expression ratios were calculated for each of the 1754 original spots and changes were defined as being upregulated if the ratio is > 1.3 and downregulated if the ratio is <-1.3. No change was define as an expression ratio between -1.3 and 1.3. 59 spots had changed expression upon APC loss (MDCK vs APCKD) (Table 3.1). Of the 59 spots affected APC loss, 14 were rescued in APCFL cells and 9 exhibited no change between APCFL and MDCK controls suggesting these 9 proteins were specific to APC knockdown and rescued to levels of controls by full-length APC (Table 3.2). Additionally,

10 of the 14 spots rescued in APCFL cells were also rescued by EMP2 knockdown in

APCKD cells (Table 3.3). Interestingly, 4 of the proteins that were rescued by EMP2 knockdown were restored to levels of control cells identifying these spots as the best candidates of APC/EMP2 regulation (Table 3.3). Curation of the spots was necessary to preform mass spectrometry to identify the proteins. Spots that passed curation where 102

assigned new spot numbers and are represented by the circles in Figure 3.4. Mass spectrometry identified the 4 best candidates (* in Table 3.3) as filamin A or plectin isoform X2, histone H3.3, histone H2A type 1-E-like, and 3-hydroxyacyl-CoA dehydrogenase type-2 (Table 3.4). We next sought to validate the 2D DIGE results starting with these 4 candidates.

103

Figure 3.4 Representative 2D DIGE images. 2D DIGE analysis was utilized to identify proteins that may be mediated by APC and EMP2. Several comparative gels were run to evaluate proteins that exhibited changes in expression upon APC and rescue of the expression by EMP2 knockdown in APCKD cells. Representative gels are shown here comparing MDCK (green) and APCKD (red) lysates (top) and APCKD (green) and APCKD EMP2KD (red) (bottom). Proteins exhibiting similar expression between the two cell lines will appear yellow, while differences in expression between the two cell lines will appear either green or red. Curated spots chosen for mass spectrometry are circled. White circles represent spots changed in both APCKD cell lines compared to controls, pink circles signify spots changed in only one of the two APCKD cell lines compared to controls, and red circles show spots that did not pass curation. Gels were performed twice with one biological replicate and changes in expression and curation limits were based on recommendations from Applied Biomics.

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105

TABLE 3.1

2D DIGE PROTEINS WITH CHANGED EXPRESSION UPON APC LOSS

Original No. Protein Expression Ratio

APCKD / MDCK

6 -1.4 175 -1.3 179 -1.3 272 1.4 369 -1.3 372 -1.3 457 -1.3 556 1.4 592 1.5 601 -1.7 605 -1.3 607 1.3 677 -1.3 697 1.3 698 1.3 703 -1.3 745 1.3 813 1.3 834 1.3 894 -1.3 896 1.3 912 -1.3 921 -1.3 965 -1.8 978 -1.4 1018 1.4 1030 1.3 1058 1.3 1167 1.3 106 TABLE 3.1 (CONTINUED)

Original No. Protein Expression Ratio

APCKD / MDCK

1237 1.3 1309 -1.5 1326 -1.3 1430 -1.3 1442 1.3 1448 1.4 1449 1.5 1483 1.3 1487 1.3 1497 1.6 1514 1.3 1536 1.3 1539 1.5 1551 1.5 1553 1.4 1573 -1.9 1588 1.3 1605 -1.7 1617 1.3 1619 -1.5 1633 -1.7 1661 -1.3 1679 1.4 1684 1.4 1698 1.5 1699 1.3 1700 1.3 1738 1.3 1740 1.3 1751 1.6

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TABLE 3.2

2D DIGE PROTEINS SPECIFIC TO APC REGULATION

Original No. Protein Expression Ratio

APCKD / APCFL / APCFL /

MDCK APCKD MDCK 6* -1.4 1.4 1.0 698* 1.3 -1.4 -1.0 894 -1.3 2.2 1.7 1309* -1.5 1.8 1.2 1430 -1.3 1.7 1.3 1448* 1.4 -1.6 -1.2 1487 1.3 -2.9 -2.3 1539 1.5 -2.2 -1.5 1573* -1.9 2.1 1.1 1605* -1.7 1.8 1.1 1619* -1.5 1.7 1.2 1633 -1.7 2.3 1.4 1740* 1.3 -1.3 1.0 1751* 1.6 -1.5 1.1 Note: *signifies 9 spots rescued to levels of control expression by reintroduction of full-length APC

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TABLE 3.3

2D DIGE PROTEINS SPECIFIC TO APC AND EMP2 REGULATION

Original No. Protein Expression Ratio

APCKD / APCFL / APCKD EMP2KD / APCFL / APCKD EMP2KD /

MDCK APCKD APCKD MDCK MDCK 6* -1.4 1.4 1.5 1.0 1.1 894 -1.3 2.2 2.3 1.7 1.8 1309* -1.5 1.8 1.5 1.2 1.0 1448 1.4 -1.6 -1.9 -1.2 -1.4 1487 1.3 -2.9 -1.8 -2.3 -1.4 1539 1.5 -2.2 -2.1 -1.5 -1.4 1573* -1.9 2.1 2.1 1.1 1.1 1619* -1.5 1.7 1.7 1.2 1.1 1633 -1.7 2.3 2.3 1.4 1.4 1751 1.6 -1.5 -2.1 1.1 -1.3 Note: *signifies 4 spots rescued to levels of MDCK controls by EMP2 knockdown

109 TABLE 3.4

2D DIGE CANDIDATES IDENTIFIED BY MASS SPECTROMETRY

New Spot Original No. Top Ranked Protein Name [Species] Accession No. Protein MW number

PREDICTED: filamin-A [Canis lupus familiaris] or plectin isoform X2 6 1 gi|74008809 280,450 [Canis lupus familiaris]

PREDICTED: 3-hydroxyacyl-CoA dehydrogenase type-2 [Canis lupus 1309 31 gi|74006997 27,138 110 familiaris]

1573 45 PREDICTED: histone H3.3 [Canis lupus familiaris] gi|57089073 15,319

1619 49 PREDICTED: histone H2A type 1-E-like [Canis lupus familiaris] gi|57110463 13,926

Because APC can bind the cytoskeleton both directly and indirectly (Munemitsu et al. 1994, Askham et al. 2000, Mimori-Kiyosue et al. 2000, Rosin-Arbesfeld et al. 2001,

Zumbrunn et al. 2001, Watanabe et al. 2004, Okada et al. 2010) and filamin and plectin are cytoskeleton bind proteins (Foisner et al. 1988, Yue et al. 2013), we attempted to validate the expression of filamin and plectin with western blotting first (Figure 3.5 A).

Quantification of filamin A and plectin expression in MDCK, APCKD, and APCKD EMP2KD cells revealed no significant changes in expression of either protein in all of the cell lines

(Figure 3.5 B). Therefore, data from western blots are inconsistent with the 2D DIGE results. Because proteins were separated by size and pH it is possible that changes seen in analysis are due to mislocalization or phosphorylation of proteins. Future experiments will assess localization of filamin and plectin to determine if these proteins are mislocalized in APCKD cells. Future studies will also seek to validate the other 3 hits histone H3.3, histone H2A type 1-E-like, and 3-hydroxyacyl-CoA dehydrogenase type-2 with western blotting.

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Figure 3.5 APC loss does not increase filamin or plectin expression. Western blot for filamin-A and plectin (A) and quantification (B) demonstrated neither protein is changed upon APC loss suggesting filamin and plectin are not downstream of APC and EMP2. Data from three replicates are presented as mean ± s.d. and a one-way ANOVA was utilized to determine significance, * p < 0.05.

112

Pathway enrichment analysis was performed on the proteins identified with mass spectrometry in the 2D DIGE experiments. Two functional groups were enriched calcium/phospholipid binding and chromosome associating genes including DNA binding, histone folding, and nucleosome (Table 3.5). Enrichment for calcium and phospholipid binding proteins is consistent with EMP2’s role as a tetraspan membrane protein where it is associated with glycosylphosphatidyl inositol-anchored proteins (GPI-

APs) in lipid rafts (Wadehra et al. 2004). Although APC has nuclear functions such as regulation of cell cycle progression and DNA repair and binds several proteins involved in DNA replication and repair (reviewed in (Lesko et al. 2014)), enrichment for chromosome associating genes was surprising. Because pathway analysis was completed on all proteins with changed expression in APCKD cells compared to MDCK controls, proteins related to this functional group may be specific to APC loss and not involved in signaling modalities downstream of EMP2. Nevertheless, pathway analysis identified two functional groups that may be important in mediating mechanisms downstream of APC and EMP2 and warrant further investigation.

113 TABLE 3.5

PATHWAY ENRICHENT ANALYSIS OF 2D DIGE DATA

Functional Group 1 Enrichment Score: 2.98 Category Term Fold Enrichment UP_KEYWORDS Calcium/phospholipid-binding 117.0214286 UP_KEYWORDS Annexin 106.3831169 INTERPRO IPR001464:Annexin 100.012987 INTERPRO IPR018502:Annexin repeat 100.012987

114 INTERPRO IPR018252:Annexin repeat, conserved site 100.012987

SMART SM00335:ANX 85.72027972 GOTERM_MF_DIRECT GO:0005544~calcium-dependent phospholipid binding 27.66190476 UP_KEYWORDS Calcium 4.875892857 Functional Group 2 Enrichment Score: 2.77 Category Term Fold Enrichment GOTERM_CC_DIRECT GO:0000786~nucleosome 31.63409091 UP_KEYWORDS Nucleosome core 28.89417989 INTERPRO IPR009072:Histone-fold 19.30075188 UP_KEYWORDS Chromosome 19.02787456 UP_KEYWORDS DNA-binding 2.916421896

3.5 Discussion

Although previously identified as downstream mechanisms of either APC or

EMP2, these studies determined neither Scrib/Hippo nor Cav1/ERK/JNK signaling are downstream of APC and EMP2 interactions. Several other downstream candidates were found through 2D DIGE experiments including filamin A or plectin isoform X2, histone

H3.3, histone H2A type 1-E-like, and 3-hydroxyacyl-CoA dehydrogenase type-2. All of these proteins were downregulated upon APC loss and expression was rescued in APCKD

EMP2KD double knockdown cell lines. We pursued filamin and plectin first because they act as links for cytoskeletal proteins (Foisner et al. 1988, Yue et al. 2013) and APC also binds both actin and microtubules to stabilize polymerization (Munemitsu et al. 1994,

Okada et al. 2010). Interestingly filamin A helps anchor membrane proteins by acting as a link with the actin cytoskeleton (Wang et al. 2015) providing evidence for a role for filamin as a linker for a complex consisting of APC, EMP2, filamin A, and actin. Future studies are needed to evaluate such a complex and would include IF for protein colocalization of filamin A and APC, EMP2, and actin as well as immunoprecipitation experiments to identify direct interactions between the proteins.

Future studies are also needed to validate the remaining candidates. The most promising is 3-hydroxyacyl-CoA dehydrogenase type-2 which is part of mitochondrial ribonuclease P, an enzyme which catalyzes the beta-oxidation of androgens and estrogens. EMP2 is implicated in several hormone driven cancers like ovarian, endometrial, and breast cancer (Fu et al. 2010, Habeeb et al. 2010, Fu et al. 2014). 115

Furthermore, other members of the EMP protein family EMP1 and EMP3 interact with epidermal growth factor receptor (EGFR) and erythroblastic oncogene B (ERBB2) respectively (reviewed in (Wang et al. 2017)) EMP1 has been identified as a biomarker for gefitinib, an EGFR inhibitor, resistance in non-small cell lung carcinoma (Jain et al.

2005) and EMP1 expression promotes tumorigenesis through activation of phosphoinositide 3-kinsase (PI3K)/protein kinase B (AKT) signaling (Lai et al. 2012).

Additionally, EMP3 increases urothelial carcinoma cell proliferation through upregulation of ERRB2 and activation of the PI3K and AKT pathway (Wang et al. 2014).

Given the role for 3-hydroxyacyl-CoA dehydrogenase type-2 in hormone synthesis and evidence supporting the EMP family of proteins interact with hormone receptors, it is possible that 3-hydroxyacyl-CoA dehydrogenase type-2 acts downstream of APC and

EMP2 interactions to mediate hormone receptors to drive PI3K/AKT signaling.

With this study we found that APC and EMP2 do not regulate known downstream mechanisms of EMP2 Cav1/ERK/JNK or Scrib and Hippo signaling. Although the mechanisms by which APC and EMP2 control cyst size and polarity remain unknown; we have identified several possible candidates through 2D DIGE studies. Future studies are needed to determine if these candidates do act downstream of both APC and EMP2 and influence polarity. Because we have ruled out all known mechanisms downstream of EMP2, the signaling pathways involved in APC/EMP2-mediated polarity are novel and could present new possibilities for targeted therapies against APC-mutant cancers.

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CHAPTER 4:

APC TRANSCRIPTIONAL REGULATION OF EMP2

This chapter was completed in collaboration with Carolyn Ahlers.

4.1 Abstract

The tumor suppressor Adenomatous Polyposis Coli (APC) is an important regulator of cell polarity and our laboratory previously demonstrated that loss of APC in

Madin Darby Canine Kidney Cells (MDCK) in 3D Matrigel culture increased cyst size, disrupted polarity, and upregulated gene expression of Epithelial Membrane Protein 2

(EMP2). Furthermore, we identified a novel role for EMP2 in mediating polarity as knockdown of EMP2 in cells with APC loss restored cyst size and polarity. However, how

APC increases EMP2 expression has not been elucidated. Because APC loss increased

EMP2 expression at the mRNA level in DNA microarrays and rt-PCR we hypothesize that

APC transcriptionally regulates EMP2. Here we identified several transcription factors with predicted binding sites in the EMP2 promoter and in parallel determined transcription factors regulated by APC. Together these studies suggest signal transducer and activator of transcription 1 (STAT-1) and STAT-3 as possible novel APC-mediated transcriptional regulators of EMP2. Furthermore, we have optimized ChIP assays and developed an EMP2 promoter driven luciferase construct to confirm a role for STAT-1 and STAT-3 in EMP2 transcriptional regulation with future studies.

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4.2 Introduction

Our lab has previously shown that APC loss in MDCK cells in 3D Matrigel cultures leads to early tumorigenic markers such as increased cyst size and disrupted polarity

(Lesko et al. 2015). Furthermore, a DNA microarray and rt-PCR validation showed that

APC loss increased EMP2 gene expression. Knockdown of EMP2 in cells with APC loss decreased cyst size and restored polarity, establishing a novel role for EMP2 in mediating epithelial polarity (Lesko et al. 2015, Lesko and Prosperi 2017). However, the mechanism(s) by which APC loss increases EMP2 expression remain unknown. Because

EMP2 expression is changed at the mRNA level, we hypothesize that APC transcriptionally regulates EMP2.

Few studies have investigated the mechanisms that drive EMP2 expression. In urinary bladder urothelial carcinoma cells, EMP2 promoter activity was decreased by mutating the cAMP responsive elements, binding site for cAMP responsive element binding protein 1 (CREB1) transcription factor (Li et al. 2015). A correlation between

CREB and EMP2 was shown by treatment with genistein, an angiogenesis inhibitor, which increased CREB1 transcriptional activity and EMP2 expression in in vivo xenograft models (Li et al. 2015). Furthermore, the combination of estradiol and progesterone increased EMP2 mRNA expression in human endometrial cancer cells; however, the mechanism of this hormone regulation is not yet fully understood (Wadehra et al.

2008). In the current studies we investigate how APC transcriptionally regulates EMP2.

We first identified transcription factors that are predicted to bind the EMP2 promoter.

Separately we assessed transcription factors that are regulated by APC. From these two 118

studies we identified STAT-3 and STAT-1 as possible transcriptional regulators of EMP2 by APC. Additionally we optimized ChIP assays and developed an EMP2 promoter driven luciferase reporter construct for use in further studies to confirm this mechanism.

4.3 Materials and Methods

4.3.1 Cell Culture

Madin Darby Canine Kidney (MDCK) cells were obtained from K. Matlin

(University of Chicago), tested for contamination, and maintained in DMEM media with

5% FBS, 2 mM L-glutamine, 10 mM HEPES, and 1% Penicillin/Streptomycin. APCKD cells were established previously (Lesko et al. 2015), and were grown in DMEM media with

5% FBS, 2 mM L-glutamine, 10 mM HEPES, 1% Penicillin/Streptomycin, and 2 µg/ml

Puromycin.

4.3.2 ConTra bioinformatics

The visualization tool on the ConTRA v3 webserver (Kreft et al. 2017) was utilized to identify a list of proteins predicted to bind the human EMP2 promoter (1500bp upstream). The visualization tool was used to determine predicted sites of specific transcription factors in the canine EMP2 promoter (1500bp upstream). Utilization of the cow genome was necessary to evaluate the canine genome and assess homology.

4.3.3 Protein/DNA array

Nuclear lysates from MDCK and APCKD cells were isolated utilizing the Nuclear

Extraction Kit (Affymetrix) and 7.5ug of nuclear protein was run on the TranSignal 119

Protein/DNA Array 1 (Affymetrix) with 56 transcription factors. Image J was used to quantify the blots. Arrays were replicated three times and data are plotted as normalized average relative density. Expression was considered changed if the fold change was greater than 0.5.

4.3.4 Firefly and renilla luciferase reporter assays

MDCK and APCKD cells were transfected with firefly luciferase plasmids pGL4.47[luc2P/SIE/Hygro] plasmid (STAT-3 activity) (Promega), AP-1 luc (Agilent

Technologies), or NF-κB and NF-κB mut plasmids (a kind gift from D. Guttridge OSUMC) for 48 hours. The pRL-TK plasmid (Promega) was used as the renilla luciferase transfection control. Additionally, Cignal Reporter Assay Kits (Qiagen) were utilized to assess Cre (CREB) and GAS (STAT-1) transcriptional activity following manufacturer instructions. For GAS assays, cells were treated with 100ng/ml canine interferon-γ (R&D systems, 781-CG) for 18 hours. Luciferase expression was measured using Dual-Glo

Luciferase assay (Promega) and the Dual-Glo Luciferase protocol on the SpectraMax M3 plate reader (Molecular Devices). Luciferase reporter assays are graphed as normalized mean ± standard deviation (n=3). Statistics were calculated using a one-way ANOVA with significance * = p<0.05.

4.3.5 Chromatin immunoprecipitation

ChIP experiments were performed using the SimpleChIP Enzymatic Chromatin

IP Kit with Magnetic Beads (Cell Signaling). 90 million cells were used for each chromatin prep, digested with 1.5µL of Micrococcal Nuclease and sonicated with 9 cycles of 20 120

seconds pulse and 59 seconds off. 10µg of digested chromatin was utilized for each immunoprecipitation (IP). CREB (48H2) Rabbit mAb (1:25) antibody (Cell Signaling,

#9197) was added to 500µL of each IP. Histone H3 (D2B12) XP Rabbit mAb (1:50) (Cell

Signaling, #4620) was used as a positive control IP, and respective concentration of

Normal Rabbit IgG (Cell Signaling, #2729P) was used as a negative control for each IP.

Chromatin was eluted by placing samples in a heat block for 30 minutes at 65˚C with gentle vortexing every 5 minutes. ChIP experiments were quantified using rt-PCR using the following primers: RPL30 F: 5’ GCACAGCATGTGGGAAATACTAC 3’, R: 5’

ACCTAGGAACCGAAGACATTGCTA 3’; NR4A3 F: 5’ TCCACCTTCCATCATCGACA 3’, R: 5’

TAGGTGTCCAGACCCATTCCTT 3’; EMP2 F: 5’ CTCCCTAAGCCACCCTACCT 3’, R: 5’

ATGCCTTGATGGTCCTGTGG 3’. Quantification shows mean percent input calculated as percent input = 2% x 2(Cq 2%Input Sample - Cq IP Sample) ± standard deviation (n=3).

Statistics were calculated using a two-way ANOVA with significance * = p<0.05.

4.3.6 Statistics

For one-way and two-way ANOVAs a Tukey post hoc test was preformed. For all data analysis a p-value < 0.05. was determined to be significant.

4.4 Results

4.4.1 Predicted transcriptional regulators of EMP2

The mechanisms underlying the transcriptional regulation of EMP2 remain largely unknown. Therefore, the ConTra v3 webserver (Kreft et al. 2017) was utilized to

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identify transcription factors that are predicted to bind the canine EMP2 promoter

(1500bp upstream of transcription initiation site). Due to limitations of exploration of the canine genome within the webserver, we first searched the human EMP2 promoter and found several transcription factors with predicted binding sites (Figure 4.1 A). Some of these transcription factors included activator protein 1 (AP-1), E2F transcription factor 1 (E2F-1), nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB),

STAT-1, and STAT-3. We also identified predicted binding sites for CREB, which is a known transcriptional regulator of EMP2 (Li et al. 2015). Next, we determined the predicted ability of these transcription factors to bind the canine promoter utilizing the alignment data from the cow genome. Representative alignment schematics show the number, location, and conservation of the predicted sites (Figure 4.1 B). Table 4.1 lists the specific predicted sequence at each site. Not surprisingly, CREB is predicted to bind the most sites, 19, in the canine EMP2 promoter and is well conserved with other species. AP-1 is predicted to bind 3 sites, STAT-1 is predicted to bind 6 sites, and STAT-3 is predicted to bind 4 sites. While AP-1, STAT-1, and STAT-3 are predicted to bind less sites than CREB they are still generally well conserved across the mouse and cow genomes. Though NF-κB is predicted to bind 6 sites these sites are not well conserved.

Interestingly, the canine EMP2 promoter only contains 1 predicted binding site for E2F-1 which is not conserved among other species (Figure 4.1 B, Table 4.1). Together these data reveal predicted binding sites for several transcription factors including AP-1, CREB,

E2F-1, NF-κB, STAT-1, and STAT-3 in the canine EMP2 promoter. Based on the number of

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predicted sites and the conservation among other species, AP-1, CREB, STAT-1, and

STAT-3 are promising candidates for APC-mediated transcriptional regulators of EMP2.

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Figure 4.1 Predicted transcription factor binding sites in the canine EMP2 promoter. A) The ConTra v3 webserver was utilized to explore transcription factors with predicted binding sites in the canine EMP2 promoter (1500bp upstream of the transcriptional initiation site). Transcriptional factors with higher binding affinity and biological relevance are shown here. B) The visualization tool in the ConTra v3 webserver shows an example of predicted binding sites for AP-1, NF-κB, CREB, E2F-1, STAT-1, and STAT-3. Yellow highlighted sites in scale (top) demonstrate the number of predicted sites for each transcription factor, while an example of a predicted sequence is highlighted in the alignment data (bottom) for each transcription factor. Conservation between cow and mouse genomes can be assessed with plots of alignment data.

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TABLE 4.1

PREDICTED BINDING SEQUENCES IN THE EMP2 PROMOTER OF SEVERAL TRANSCRIPTION

FACTORS

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4.4.2 APC loss increases the expression and activity of several transcription factors including STAT-1 and STAT-3

Next, we sought to gather an unbiased compilation of transcription factors that demonstrate altered activation upon APC loss using a DNA/protein binding array. This transcription factor array containing 56 transcription factors demonstrated that APC loss upregulates expression by a factor of 0.5 or higher fold of multiple transcription factors including nuclear factor of activated T-cells (NFAT-1), Yin and Yan 1 (YY1), STAT-1, gamma interferon activation sites (GAS)/interferon stimulated response element (ISRE),

E2F-1, peroxisome proliferator-activated receptor (PPAR), and specificity protein 1 (Sp-

1) (Figure 4.2 A, B). Additionally, the expression of AP-1, STAT-3 and NF-κB are decreased (more than 0.5 fold lower) in APC knockdown (APCKD) cells compared to controls (Figure 4.2 A, B). Interestingly, expression of CREB and estradiol receptor, known regulators of EMP2, was unchanged or did not bind in either MDCK or APCKD cells in DNA/Protein arrays (Figure 4.2 A, B). These data suggest that the transcriptional regulation of EMP2 downstream of APC is through a novel mechanisms. Of the transcription factors with increased expression upon APC loss, YY1, STAT-1, E2F-1, and

Sp-1 are predicted to bind the canine EMP2 promoter (Figure 4.1 and data not shown).

However, the YY1 and E2F-1 binding sequences are not conserved among mouse and cow genomes (Figure 4.1 B and data not shown).

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Figure 4.2 APC loss increases the expression and activation of several transcription factors. A) A transcription factor DNA/protein array (Affymetrix) identified several transcription factors that exhibited changed expression upon APC loss. B) Interestingly, quantification of DNA/protein arrays revealed E2F-1. STAT-1, YY1, NFAT-1, GAS/ISRE, PPAR, and Sp-1 have increased expression in APCKD cells compared to MDCK control cells. Data are presented as the average normalized fold change of three experimental replicates. C) Data from the DNA/protein array was validated using luciferase reporter assays and indicated STAT-3 activity is increased in APCKD cells compared to MDCK controls. Data from three replicates are presented as mean ± s.d. A two- way ANOVA was used to determine significance for both experiments, * p < 0.05.

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Because the DNA/protein array only measures transcription factor expression and not transcriptional activity, we performed luciferase reporter assays to validate the functional relevance of some of the transcription factors identified in the DNA/protein array. Given their known role in regulating EMP2 (Li et al. 2015), tumor initiation and progression (Biswas et al. 2004, Shen et al. 2008, Ndlovu et al. 2009, Shostak and

Chariot 2011, Smith et al. 2014), or mediating apical-basal polarity (McCaffrey et al.

2012) we chose to validate AP-1, NF-κB, CREB, STAT-1 and STAT-3 with reporter assays.

Consistent with the DNA/protein array there are no changes in the transcriptional activity of AP-1, NF-κB, and CREB in APCKD cells compared to control MDCK cells (Figure

4.2 C). Interestingly, STAT-3 activity was significantly increased upon APC loss; however, in disagreement with the DNA/protein array STAT-1 activity, as measured by GAS reporter activity, remained unchanged (Figure 4.2 C). Identification of transcription factors regulated by APC, along with bioinformatics analysis predicting a binding site in the canine EMP2 promoter, suggest STAT-1 and STAT-3 as potential APC-mediated regulators of EMP2.

4.4.3 Transcription factor binding in the canine EMP2 promoter

Since STAT-1 and STAT-3 are predicted to bind the EMP2 promoter, exhibit increased expression or activity upon APC loss, and are associated with loss of polarity

(McCaffrey et al. 2012, Guyer and Macara 2014), we next sought to determine whether they actually bind to the canine EMP2 promoter utilizing Chromatin

Immunoprecipitation (ChIP) assays. As the only known transcriptional regulator of

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EMP2, CREB was utilized as a positive control for EMP2 promoter binding in ChIP assays.

Histone H3 was used in all ChIP assays as a positive control for the assay. In CREB ChIP experiments histone H3 bound the Ribosomal Protein L30 (RPL30) locus, known histone enrichment, in both MDCK and APCKD cells as shown by greater percent input of the histone H3 antibody than the IgG control (Figure 4.3 A), and an input ratio (percent input IP/percent input IgG) greater than 1 (Table 4.2). CREB is known to highly bind the

Nuclear Receptor Subfamily 4 Group A Member 3 (NR4A3) gene and therefore NR4A3 was utilized as a positive control for the CREB IP. NR4A3 was enriched in DNA pulled down by CREB in both MDCK and APCKD cells although the input ratio was lower in APCKD cells suggesting binding of CREB to NR4A3, was weaker upon APC loss (Figure 4.3 B,

Table 4.2). We next assessed CREB binding to the EMP2 promoter and determined that

CREB binds the EMP2 promoter in both MDCK and APCKD cells (Figure 4.3 C), but similar to NR4A3 binding to the EMP2 promoter was weaker in APCKD cells (Table 4.2). Together these data are consistent with previous studies showing that CREB binds the EMP2 promoter; however CREB binding seems to be decreased overall in cells with APC loss.

With ChIP assays optimized for EMP2 promoter studies in our lab, future studies will seek to determine if STAT-1 and STAT-3 bind the EMP2 promoter.

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Figure 4.3 CREB binds the EMP2 promoter. A) Histone H3 (H3) was used as a positive control assay in Chromatin Immunoprecipitation (ChIP) experiments. H3 IP was able to pull down the RPL30 gene (known H3 enrichment). B) The NR4A3 gene (known CREB binding) was used as a positive control for CREB IP. C) CREB IP samples were analyzed with rt-PCR for EMP2 gene expression. Percent input analysis of CREB IP and IgG control samples revealed that CREB binds the canine EMP2 promoter in both MDCK and APCKD cells, though there is less binding in APCKD cells. Experiments were replicated three times and data are presented as mean ± s.d. A two-way ANOVA was utililzed to determine significance.

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TABLE 4.2

CREB CHIP EXPERIMENT INPUT RATIOS

Input Ratio H3/IgG CREB/IgG CREB/IgG (RPL30) (NR4A3) (EMP2)

MDCK 8.791435 6.120454 5.747002 APCKD 8.716536 2.58091 2.289389

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4.5 Discussion

These studies glean novel insights into the transcriptional regulation of EMP2 by

APC. We identified several transcription factors with predicted binding sites in the EMP2 promoter including AP-1, NF-κB, E2F-1, STAT-1, and STAT-3. Through transcription factor binding arrays we determined several transcription factors with increased expression upon APC loss such as E2F-1, STAT-1, YY1, and Sp-1. Although E2F-1, STAT-1, YY1, and

Sp-1 are all predicted to bind the canine EMP2 promoter, only the STAT-1 and Sp-1 sites are conserved in cow and mouse genomes. Therefore, regulation by Sp-1 is an additional transcriptional mechanism to be assessed in future studies. Luciferase reporter assays revealed STAT-3 is activated in APCKD cells. Interestingly, the known transcriptional regulators of EMP2 CREB and estrogen and progesterone either showed no change in expression upon APC loss or did not bind the array in either MDCK or APCKD cells respectively. Additionally, CREB was not activated upon APC loss in luciferase reporter assays and exhibited decreased binding in APCKD cells to both the control

NR4A3 gene and EMP2. This overall decrease in DNA binding could be due to the upregulation of corepressors by the loss of APC. For instance, increased STAT-3 activation has been shown to negatively regulate CREB transcriptional activity

(Ramadoss et al. 2009). Overall these studies suggest APC regulates EMP2 through a novel mechanism.

Based on the number and homology of predicted binding sites, transcription factor expression in DNA/protein array, and transcriptional activity we believe that

STAT-1 and STAT-3 are promising candidates as APC-mediated transcriptional regulators 133

of EMP2. Though STAT-3 expression was not increased in DNA/binding arrays, STAT-3 activity was increased in luciferase reporter assays. This could be attributed to the STAT-

3 binding site on the DNA/protein array being different from the actual binding site and to the higher specificity of the luciferase reporter. We also observed increased STAT-1 expression in APCKD cells; however STAT-1 activity was unchanged in GAS luciferase reporter assays which measures transcriptional activation of STAT-1 homodimers. This contradiction could be caused by STAT-1 and STAT-3 forming heterodimers to drive

EMP2 expression instead of STAT-1 homodimers. STAT-3 has been shown to recruit

STAT-1 into heterodimers leaving less STAT-1 protein available to form STAT-1 homodimers (Delgoffe and Vignali 2013). Further studies are needed to confirm a STAT-

1:STAT-3 heterodimer complex.

Although we have shown that STAT-1 expression and STAT-3 transcriptional activity is increased in APCKD cells, it remains to be elucidated whether these transcription factors do increase EMP2 expression. To assess this we developed a luciferase reporter construct driven by the EMP2 promoter. 1700bp upstream of the transcriptional start site was cloned into the pGL4 promoterless luciferase plasmid

(Figure 4.4). The pGL4 plasmid alone will be used as a negative control and treatment with estradiol and/or progesterone will be as a positive control to confirm the functionality of this plasmid. Future studies will assess if EMP2 expression is increased in

APCKD cells compared to MDCK controls. Site-directed mutagenesis will be utilized to mutate the binding sites for STAT-1 and STAT-3 to create mutant reporter constructs which we predict will decrease EMP2 expression in APCKD cells. Finally, a STAT-3 134

overexpression line will be created by transfecting STAT3-flag-pcDNA vector into MDCK control cells and STAT-3 will be inhibited in APCKD cells using a small molecular inhibitor

A69 which inhibits STAT-3 binding to DNA (Huang et al. 2014, Huang et al. 2016). Then we will assess EMP2 expression by rt-PCR and cyst size and polarity in cells grown in 3D

Matrigel culture. These studies will identify a novel mechanism of APC transcriptional regulation of EMP2 through STAT-1 and STAT-3.

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Figure 4.4 EMP2 promoter driven luciferase reporter plasmid. The EMP2 promoter, 1700bp upstream of the initiation site, was cloned into the pGL4.10 [luc2] vector (Promega) utilizing the following primers: F- 5’ CATGCTCGAGGATGCAGGCGTAAA 3’ and R- 5’ CGAAGCTTTTTCACGGGGCA 3’. The plasmid was confirmed with restriction digest using XhoI and HindIII enzymes as well as Sanger sequencing through the Notre Dame Genomics & Bioinformatics core.

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CHAPTER 5:

CONCLUSIONS AND FUTURE PERSPECTIVES

Several studies have established a role for the APC tumor suppressor in regulating epithelial polarity including data from our lab showing that loss of APC in

MDCK cells increases cyst size and disrupts polarity (Lesko et al. 2015). However, the molecular mechanisms by which APC controls these processes to attenuate tumor progression remain largely unknown. Here we investigated the role of the tetraspan protein EMP2 in influencing APC-mediated polarity and gained important insight into the downstream mechanisms of APC tumor suppression.

5.1 EMP2 independent APC-mediated cell migration

In Chapter 2, we showed that APC loss in parental MDCK epithelial cells increased migration during wound healing assays. While inhibition of β1 integrin/FAK/Src signaling decreased migration, EMP2 knockdown had no effect on cell motility in APCKD cells. These data suggest APC mediates cell migration through β1 integrin signaling independent of EMP2, and were our first indication of EMP2 not signaling through β1 integrin and FAK in the MDCK cells. It has been well established that integrins act as links between the cytoskeleton and ECM at focal adhesions to regulate cell motility ((Critchley 2000, Krylyshkina et al. 2003, Akhtar and Streuli 2013) and reviewed in (Huttenlocher and Horwitz 2011)). Additionally, APC promotes actin and microtubule assembly through direct and indirect interactions ((Munemitsu et al. 137

1994, Okada et al. 2010) and reviewed in (Prosperi and Goss 2011)). Therefore, APC may mediate interactions between the cytoskeleton and β1 integrin to regulate cell migration. Future studies investigating cytoskeletal dynamics and adhesion during wound-healing assays are needed to further understand the mechanisms by which APC loss increases cell motility.

5.2 EMP2 as a novel regulator of polarity

EMP2 has been implicated in several cancers, however the normal functions of

EMP2 remain largely unknown. EMP2 is known to be involved in regulating the lipid composition of the plasma membrane (Wadehra et al. 2003, Wadehra et al. 2004) as well as recruitment of integrins to the membrane in order to regulate cell adhesion and migration (Wang et al. 2013). Here our lab identified a novel role for EMP2 in controlling polarity. Knockdown of EMP2 in APCKD cells rescued increased cyst size and disrupted polarity caused by APC loss. These studies were the first to implicate EMP2 as a downstream regulator of APC-mediated early tumorigenic phenotypes and identified

EMP2 as a potential therapeutic target in APC-mutant cancers.

Very few studies have explored therapies targeting EMP2; however the use of anti-EMP2 antibodies has proven promising in several cancers. An anti-EMP2 diabody that targets the extracellular domains of EMP2 was successful in treating human endometrial adenocarcinoma cell lines by inducing cell death and in in vivo mouse xenografts by significantly decreasing tumor growth (Shimazaki et al. 2008). Similar results were observed in the treatment of glioblastoma. Cell death was induced in 138

glioblastoma cell lines treated with the EMP2 diabody and tumors resulting from subcutaneous injections exhibited decreased growth upon treatment (Qin et al. 2014).

Additionally, a monoclonal IgG1 antibody was developed and tested in breast cancer models. Treatment of breast cancer cells with the monoclonal EMP2 antibody inhibited

EMP2 signaling and invasion and promoted apoptosis (Fu et al. 2014). These results were confirmed in vivo as monoclonal antibody treatment reduced tumor growth in mouse models (Fu et al. 2014). This monoclonal antibody had similar effects as the diabody in the treatment of endometrial cancer models as it decreased tumor burden and increased overall survival of mice in study (Gordon et al. 2013). Together these studies demonstrate the efficacy and low toxicity of EMP2 targeted antibodies; however, the mechanisms by which these antibodies work remain unknown and the use of EMP2 antibodies is still far from clinical use. Although promising, further in vivo studies are needed to assess the use of the anti-EMP2 antibodies in treating APC-mutant cancers.

Though we identified EMP2 as an important regulator of APC-mediated polarity, the molecular mechanisms downstream of APC and EMP2 remain unknown.

Understanding these mechanisms could provide insight into novel therapeutic targets or identify already existing therapies that can be repurposed as therapies for the treatment of APC-mutant cancer. EMP2 has been shown to mediate both β1 integrin/FAK/Src signaling as well as Cav1/ERK/JNK signaling (Fu et al. 2011, Fu et al. 2014, Lee et al.

2016); however in these studies we determined EMP2 does not regulate these molecular pathways in the MDCK APCKD model. We also investigated Scrib and Hippo 139

signaling, known to regulate growth and polarity, as downstream targets of APC and

EMP2 and again found no changes in expression upon APC loss or reintroduction of

EMP2 suggesting APC and EMP2 do not regulate Scrib/Hippo signaling. Together these studies suggested that APC and EMP2 influence a novel pathway to control polarity.

Since we eliminated all known downstream targets, we performed 2D DIGE analysis and identified several candidates that may be regulated by APC and EMP2 including filamin A or plectin isoform X2, histone H3.3, histone H2A type 1-E-like, and 3- hydroxyacyl-CoA dehydrogenase type-2. Furthermore, pathway analysis revealed calcium/phospho-lipid binding and genes associated with DNA-binding and nucleosomes are enriched upon APC loss. Further studies are needed to assess whether these candidates and pathways are downstream of APC and EMP2 and will provide novel insight into the molecular pathways underlying APC and EMP2-mediated polarity.

Overall, these studies identified a novel role for EMP2 in regulating APC-mediated polarity. Although the downstream mechanisms still remain unknown, we have identified several proteins and functional pathways that may be downstream of APC and

EMP2 interactions that could provide possible novel therapeutic targets for APC-mutant cancers.

5.3 Regulation of EMP2 by APC

We have previously shown that APC loss increases EMP2 gene expression through DNA microarrays and rt-PCR (Lesko et al. 2015). However, how APC loss upregulates EMP2 expression remains unknown. In these studies we present evidence 140

that APC transcriptionally regulates EMP2. We identified several transcription factors that are predicted to bind the EMP2 promoter with the ConTra v3 webserver.

Simultaneously we determined several transcription factors affected by APC loss through DNA/protein binding arrays and luciferase reporter assays. Based on the number and homology between species of predicted binding sites as well as the ability of APC loss to upregulate expression and transcriptional activity, we identified STAT-1 and STAT-3 as possible APC-mediated transcriptional regulators of EMP2. Future studies are needed to evaluate the ability of STAT-1 and STAT-3 to bind the EMP2 promoter, drive EMP2 expression, increase cyst size and disrupt polarity.

Although our data suggest transcriptional regulation of EMP2, it is possible that

APC may mediate EMP2 through direct interactions. APC and EMP2 are both localized to the basal membrane of 3D spheres grown in Matrigel and to the protrusions of human breast cancer cell lines (data not shown). Additionally, although the structural domains of EMP2 are well conserved with other EMP family members, EMP2 does not contain a myrstoylation domain which is important for membrane protein targeting and signal transduction (Wang et al. 2017). Therefore, it is possible that EMP2 requires another scaffolding protein such as APC to recruit proteins and form complexes required for the normal functions of EMP2. Further localization and immunoprecipitation experiments are needed to elucidate whether APC and EMP2 form a complex through either indirect or direct interactions.

Another possible way APC may mediate EMP2 is through post-translational modifications. A recent study in nasopharyngeal cancer cells showed that miR-101-3p 141

expression decreased EMP2 expression leading to increased cell proliferation and migration (Wang et al. 2017). Furthermore, miR-101-3p suppression of EMP2 was negatively regulated by the long non-coding RNA nuclear enriched abundant transcript 1

(NEAT-1) (Wang et al. 2017). This study by Wang et. al. is the first to identify EMP2 regulation by miRNA and long non-coding RNAs and presents another possible mechanism of EMP2 regulation by APC. Taken together our studies present evidence for

STAT-1 and STAT-3 mediated transcriptional regulation of EMP2 by APC; however, other mechanisms of regulation such as direct interactions and post-translational modifications are still possible. Further studies are needed to confirm the role of STAT-1 and STAT-3 in EMP2 upregulation and to assess other forms of EMP2 regulation by APC.

5.4 Tissue specific functions of EMP2

EMP2 expression is very heterogeneous throughout different tissue and organ types. High expression of EMP2 can be found in the lung, while the heart, uterus, eyes, breast, ovary, and thyroid exhibit moderate amounts of expression ((Fu et al. 2011, Fu et al. 2014) and reviewed in (Chung et al. 2017)). Kidney, liver, skeletal muscle, prostate, and epididymis tissue show very low levels EMP2 expression (reviewed in (Chung et al.

2017)). Furthermore, EMP2 expression in the endometrium is dependent on hormone regulation and therefore EMP2 expression is higher during secretory phases compared to proliferative stages (Wadehra et al. 2008). Additionally, although minimally expressed in the kidney overall, EMP2 is highly localized to the glomeruli (Gee et al. 2014).

Similarly, high expression of EMP2 can be found in the eye, however, this expression is 142

specific to the cornea and ciliary body (Wadehra et al. 2003). Finally, locations of ocular expression of EMP2 differed in murine and human samples. Although both mouse and human tissues exhibited low EMP2 expression in the lens and retina, EMP2 can be found expressed in the human iris while expression is low in the murine iris (Wadehra et al.

2003). These studies reveal the heterogeneity of EMP2 expression in different organs, tissue types, and species and highlight the importance of understanding the function of

EMP2 in a variety of tissues, stages, and species.

Given that the expression of EMP2 is highly differentiated among organs, EMP2 has been identified as both an oncogene and tumor suppressor. Low expression in lung cancer cells, nasopharyngeal cancer, and urinary bladder urethral carcinoma has been correlated with tumor growth, higher tumor grade and poor prognosis establishing

EMP2 as a tumor suppressor in these tissues ((Lee et al. 2016) and reviewed in (Chung et al. 2017)). In contrast, high levels of EMP2 expression have been shown to drive proliferation, migration, invasion, tumor growth, and have been associated with more aggressive tumors in ovarian, breast, and endometrial cancers (Fu et al. 2010, Fu et al.

2011, Fu et al. 2014). In this thesis we show that APC loss in MDCK cells increases EMP2 expression to increase cyst size and disrupt polarity suggesting an oncogenic function of

EMP2 in canine kidney cells with APC loss. However, it has yet to be elucidated how APC loss affects EMP2 expression in other organs and species. Future studies in the lab seek to determine if these molecular mechanisms are similar in the mammary gland and conserved among species by establishing a model with primary mammary epithelial cells isolated from mice with Apc loss, as well as developing a human mammary in vitro 143

model system by knocking down APC in the normal human mammary epithelial cell line

MCF-10A. These studies will glean important insight into molecular mechanisms of APC and EMP2 in other epithelial tissues and are critical to evaluate the use of EMP2 targeted therapies in APC-mutant cancers.

5.5 Summary

Here we identified a novel a role for EMP2 in regulating APC-mediated epithelial polarity, but not epithelial cell migration. Though the mechanisms downstream of APC and EMP2 remain unknown, we have determined that APC and EMP2 do not influence polarity through β1 integrin/FAK/Src, Cav1/ERK/JNK, or Scrib/Hippo signaling suggesting a novel signaling modality downstream of EMP2. We have also evaluated transcriptional regulation of EMP2 by APC and provide evidence that suggests STAT-1 and STAT-3 as possible novel APC-mediated transcriptional regulators of APC. Further studies are needed to fully understand the molecular mechanisms by which APC upregulates EMP2 expression and the signaling pathways downstream of APC and EMP2 that control cyst size and apical basal polarity. Furthermore, additional studies are needed to evaluate the role of APC and EMP2 in mediating polarity in other organs and tissue types.

Determining the molecular mechanisms underlying the regulation of apical-basal polarity by APC and EMP2 will contribute to a better understanding of the mechanisms that drive tumor progression upon APC loss and will identify novel therapeutic targets for the treatment of a variety of APC-mutant cancers.

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