The Role of Polarity Protein Angiomotin (AMOT) in the Human Placenta

Abby Patricia Farrell

A thesis submitted in conformity with the requirements for the degree of Master of Science Institute of Medical Science University of Toronto

The Role of Polarity Protein Angiomotin (AMOT) in the Human

Placenta

Abby Patricia Farrell

Master of Science

Institute of Medical Science University of Toronto

2018 Abstract

Angiomotin (AMOT) is a scaffolding protein involved in cell polarity regulation, cell migration and early embryo lineage differentiation, yet its biological significance in the human placenta remains unknown. I hypothesized that AMOT controls trophoblast cell polarity and migration, and is further regulated by transforming growth factor beta (TGF) signalling and upstream oxygen tension. AMOT localization to extravillous trophoblast (EVT) cells, corroborated by AMOT 80 overexpression increasing JEG3 cell migration rate, supports a role for AMOT in EVT migration.

TGF1/3 treatment decreased AMOT protein levels and redistributed AMOT from the tight- junction to cytoplasmic F-actin in JEG3 cells. TGF1/3 also prompted a novel association between

AMOT 80 and Partitioning Defective Protein-6. Similarly, low oxygen exposure negatively regulated AMOT levels and localization. Furthermore, Jumonji C Domain Containing Protein-6

(JMJD6), an oxygen sensor, was discovered to positively regulate AMOT via lysyl hydroxylation.

Finally, AMOT levels were found markedly reduced in preeclampsia, a disease characterized by aberrant TGF signalling and chronic hypoxia. In conclusion, this study reveals AMOT is a mediator of TGF and oxygen signalling to regulate trophoblast migration in the human placenta.

Table of Contents Table of Contents ...... iii Acknowledgments ...... vi List of Abbreviations ...... vii List of Tables ...... x List of Figures ...... xi Chapter 1 Introduction...... 1 Introduction ...... 1 1.1 Angiomotin (AMOT) ...... 2 1.1.1 Discovery and Structure ...... 2 1.1.2 Role of AMOT in Hippo Pathway Signalling...... 6 1.1.3 Role of AMOT in Cell Migration ...... 8 1.2 Cell Polarity Regulation ...... 9 1.2.1 Tight Junctions ...... 9 1.2.2 Polarity Protein Complexes ...... 10 1.2.3 Role of Cell Polarity in Proliferation, Migration and Invasion ...... 11 1.2.4 AMOT as a Novel Regulator of Cell Polarity ...... 12 1.3 Human Placenta Development...... 16 1.3.1 Trophoblast Differentiation ...... 17 1.3.2 TGF signalling pathways ...... 22 1.3.3 Role of TGFβ Signalling in Trophoblast Differentiation ...... 26 1.4 Preeclampsia ...... 29 1.4.1 Altered Trophoblast Differentiation in preeclampsia ...... 30 1.4.2 Impairments in oxygen sensing in preeclampsia ...... 32 1.4.3 JMJD6: a novel oxygen sensor and regulator in the placenta ...... 33 1.5 Rationale, Hypothesis and Objectives ...... 37 Chapter 2 Materials and Methods...... 39 Materials and Methods ...... 39 2.1 Human Placenta Tissue Collection ...... 39 2.2 JEG3 Human Choriocarcinoma Cell Culture ...... 41 2.3 In vitro treatments in JEG3 cells ...... 42 2.3.1 Transforming Growth Factor- (TGF) Treatment ...... 42 2.3.2 SB-431542 Treatment in JEG3 cells...... 42 iii

2.3.3 Minoxidil Treatment ...... 43

2.3.4 Low Oxygen (3% O2) Treatment in JEG3 cells...... 43 2.4 Plasmid DNA Constructs for Overexpression Studies ...... 43 2.4.1 Plasmid DNA Transfection ...... 45 2.5 siRNA Transfections ...... 45 2.6 Wound Healing Assay ...... 46 2.7 Time-Lapse Live Cell Imaging ...... 46 2.8 Western Blot Analysis ...... 47 2.9 Antibodies ...... 49 2.10 Immunoprecipitation (IP)...... 50 2.11 Immunohistochemistry (IHC) ...... 51 2.12 Immunofluorescence (IF) ...... 53 2.13 Proximity Ligation Assay ...... 56 2.14 RNA Isolation, cDNA conversion and Quantitative-PCR ...... 57 2.15 In vitro JMJD6 Hydroxylation Reaction...... 58 2.16 MALDI-TOF Mass Spectrometry...... 59 2.17 Statistical analysis ...... 60 Chapter 3 Results ...... 61 Results ...... 61 3.1 AMOT exhibits distinct temporal and spatial expression patterns during human placenta development...... 61 3.2 AMOT is regulated by TGF signalling pathway ...... 67 3.2.1 AMOT resides at tight junction, cytoplasm and protruding edge of JEG3 cells ...67 3.2.2 TGF1/3 ligand treatment reduces AMOT 130 and 80 protein levels ...... 68 3.2.3 TGF1/3 treatment promotes subcellular redistribution of AMOT ...... 68 3.3 AMOT 130 is regulated by Smad-dependent TGF pathway ...... 74 3.4 TGFβ promotes AMOT redistribution in migrating cells ...... 78 3.5 AMOT 80 promotes JEG3 cell migration ...... 79 3.6 Novel AMOT/Par6 interaction and its regulation by TGFβ ...... 84 3.7 PDZ and coiled-coil binding domains are important for AMOT and Par6 interaction .....89 3.8 AMOT promotes dissolution of RhoA at the tight junction ...... 93 3.9 AMOT protein levels and distribution is disrupted in preeclampsia ...... 96 3.9.1 AMOT and Par6 interaction is impaired in preeclampsia ...... 97

3.10 AMOT protein levels and localization is disrupted in low oxygen ...... 101 3.11 Oxygen sensor JMJD6 positively regulates AMOT ...... 104 3.11.1 AMOT is subject to lysyl hydroxylation by JMJD6 ...... 104 Chapter 4 Discussion ...... 112 Discussion and Conclusions ...... 112 4.1 General Discussion ...... 112 4.2 Conclusions ...... 126 Chapter 5 Future Directions ...... 129 Future Directions ...... 129 5.1 Using in vivo villous explants to assess AMOT’s role in trophoblast cell differentiation ...... 130 5.2 Deciphering the role of AMOT in placental mesenchymal cells ...... 132 5.3 Establishing a role for the AMOT/TAZ axis in trophoblast cell differentiation ...... 137 5.4 Investigating lysosomal degradation of AMOT ...... 141 References ...... 143 Appendix - Statement of Contributions ...... 158

Acknowledgments

The academic and personal growth I have experienced while working towards my Master’s degree is beyond what I could have imagined, and there are many people I would like to thank for their support along the way. First and foremost, I want to express my sincerest gratitude to my supervisor, Dr. Isabella Caniggia, for her steadfast commitment and unwavering guidance over these past two years. Thank you for providing me with an innovative project and incredible opportunities that have inspired me as a young scientist and aspiring clinician. Your passion for science, life and adventure is admirable and I feel incredibly fortunate to have been your student.

I would also like to thank my PAC members Dr. Kellie Murphy, Dr. Brian Cox and Dr. Theodore Brown for their attention to detail and guidance throughout the duration of my project.

In my short time with the Caniggia Lab, I have had the absolute pleasure of working alongside driven, brilliant and energetic people. Joelcio, Julien, Tyler, Leo, Andrea and Tingting- you all, in one form or another, have made invaluable contributions to my project and enriched my graduate student experience. Thank you for always supporting and caring about me, and of course for the many, many laughs we all shared together. To Taylor, thank you for being my best friend this past year, and empathizing with me every step of this journey. I am grateful to have found a life-long friend in you. To Sruthi, I can confidently say that my project would not have been as successful if it weren’t for you. Your daily academic and personal support has been irreplaceable and I can’t thank you enough for being such a great friend and mentor to me.

To my extraordinary friends, Jessie and Nicole, thank you for always checking up on me, asking me about my research and jumping through hoops to find time to see me.

To my boyfriend Brandon, thank you for your unconditional love and patience day in and day out. I am fortunate enough to celebrate milestones, such as this one, with someone as kind-hearted and sweet as you. Thank you for being so wonderful to me, always.

Lastly, I would like to thank my family, who collectively have shaped the young woman I am today. To my sister Jenna, living in Toronto with you this past year has been so much fun. Being able to actively support each other’s goals has been profound, and I will always look back fondly on this time we have spent together. To my guardian angel, Dad- you always told me I could achieve anything I set my mind to. Thank you for instilling perseverance in me to follow my dreams, even when it gets tough. I miss you, always. Finally, to my Mom- there will never be enough words to describe what you mean to me. Thank you for doing everything in your power to ensure I am healthy, happy and successful. But most importantly, thank you for constantly grounding me and reminding me what is truly important in life.

List of Abbreviations

AMOT Angiomotin ABC complex Activin-biotin complex ACOG American College of Obstetricians and Gynecologists Alk1-7 Activin receptor-like kinase 1-7 AMOT 130 Angiomotin 130kDa isoform AMOT 80 Angiomotin 80kDa isoform AMOTL1 Angiomotin-Like 1 AMOTL2 Angiomotin-Like 2 aPKC atypical protein kinase BSA Bovine serum albumin Cdc42 Cell division control protein 42 cDNA complementary DNA ChIP Chromatin Immunoprecipitation cm centimeter Co-IP Co-immunoprecipitation Coll-1 Collagen type 1 Crb Crumbs CT Cytotrophoblast DAB Diaminobenzidine tetraaminobiphenyl DAPI 4',6 Diamidino-2-phenylindole ddH2O Double distilled water DEPC Diethyl Pyrocarbonate Dlg1 Drosophila disc large tumour suppressor Dlg1 Discs large protein DMEM Dulbecco’s Modified Essential Medium DMSO Dimethyl sulfoxide DNA Deoxyribonucleic Acid E-PE Early-onset preeclampsia ECL Enhanced chemiluminescence ECM Extracellular matrix EMEM Eagle's Minimal Essential Medium EMT Epithelial-mesenchymal transition ERVW-1 Endogenous retrovirous group W member 1 EVT Extravillous trophoblast EV Empty Vector FBS Fetal bovine serum Fe2+ Ferrous iron FIH Factor inhibiting HIF1 Flt1 VEGF receptor 1 (aka VEGFR1) GAP GTPase activating protein GCM-1 Glial cells missing homolog 1 GTP Guanosine triphosphate H&E Hematoxylin & Eosin H2O2 Hydrogen peroxide

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HCl Hydrochloric acid HIF-1α Hypoxia inducible factor-1 alpha HRP Horseradish peroxidase Hz Hertz IF Immunofluorescence IHC Immunohistochemistry JAM Junctional adhesion molecules JmjC Jumonji C JMJD6 Jumonji C domain containing protein 6 KCl Potassium chloride L-PE Late-onset preeclampsia LAMP-1 Lysosomal associated membrane protein 1 LATS1/2 Large tumour suppressor kinase 1/2 Lgl1/2 Lethal giant larvae protein 1/2 MAE Mouse aortic endothelial cells MALDI-TOF Matrix-assisted laser desorption ionization time of flight MAP kinase Mitogen-activated protein kinase MDCK Madin-Darby Canine Kidney MgCl2 Magnesium chloride mL Millilitre mM Millimolar mm Hg Millimetres of mercury MMP Matrix metalloproteinases mRNA Messenger ribonucleic acid NH4Cl Ammonium Chloride O2 Molecular oxygen OE Overexpression Pals1 Protein associated with Lin-7 1 Par3 Partitioning defective protein-3 Par6 Partitioning defective protein-6 Patj Pals1 associated tight junction protein homolog PBS phosphate buffer saline PDZ PSD95, Dlg1, ZO-1 PE Preeclampsia PFA Paraformaldehyde PHD1-3 Prolyl hydroxylase domain (1-3) PLA Proximity ligation assay PLOD1 Procollagen-lysine 5-dioxygenase 1 pMSC Placental mesenchymal cells pO2 Partial pressure of oxygen PSD95 Post synaptic density protein pSMAD2-3 Phosphorylated SMAD 2-3 PTC Pre-term control qPCR Quantitative polymerase chain reaction Rac1 Ras related C3 Botulinum Toxin Substrate 1 RhoA Ras homolog gene family member A Rich1 Rho-type GTPase activating protein 17

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RIPA buffer Radioimmunoprecipitation buffer RNA Ribonucleic acid Scrib Scribble SDS Sodium dodecyl sulfate SDS-PAGE SDS polyacrylamide gel electrophoresis SEM Standard error of the mean siRNA Small interfering RNA Smad2,3,7 Small-Mothers Against Decapentaplegic 2,3,7 Smurf1 Smad ubiquitination regulatory factor 1 ss Scrambled sequence ST Syncytiotrophoblast Syx Rho guanine exchange factor TAZ Transcriptional coactivator with PDZ binding motif TBST Tris buffered saline +Tween TEAD1 TEA domain family member 1 TGFβRI-II Transforming growth factor beta receptor I-II TGFβ1-3 Transforming growth factor beta 1-3 TIMPS Tissue inhibitors of MMPs TJ Tight junction uM Micromolar VEGF Vascular endothelial growth factor VHL von Hippel-Lindau tumour suppressor protein WB Western blotting/blot YAP Yes-associated protein ZEB2 Zinc finger E-box binding homeobox 2 ZO-1 Zona occludens-1 ZONAB ZO- associated nucleic acid binding protein α-tubulin alpha tubulin β- actin Beta-actin µg Microgram µL Microliter

List of Tables

Table 2.1 Clinical features of patient population ...... 40

List of Figures

Figure 1.1 Protein structures of ‘motin’ family members ...... 5

Figure 1.2 Polarity complexes in migrating epithelial cells ...... 14

Figure 1.3 Human placenta development and trophoblast differentiation ...... 20

Figure 1.4 Role of Smad-dependent and Par6-mediated TGFβ signalling in trophoblast cell differentiation ...... 28

Figure 1.5 Molecular and phenotypic characteristics of preeclampsia ...... 35

Figure 1.6 Model of Hypothesis ...... 38

Figure 3.1 AMOT protein levels and mRNA expression during early placenta development ..... 64

Figure 3.2 Spatial localization of AMOT in floating villi in early placenta development ...... 65

Figure 3.3 Spatial localization of AMOT in anchoring villi in early placenta development ...... 66

Figure 3.4 Co-localization of AMOT with tight junction protein ZO-1 in JEG3 choriocarcinoma cells ...... 70

Figure 3.5 Effect of TGFβ1/3 on AMOT protein levels in JEG3 cells ...... 71

Figure 3.6 Effect of TGFβ1/3 on AMOT and ZO-1 co-localization in JEG3 cells ...... 72

Figure 3.7 Effect of TGFβ1/3 on AMOT and F-actin co-localization in JEG3 cells ...... 73

Figure 3.8 Contribution of Smad-dependent TGFβ signalling on AMOT localization ...... 76

Figure 3.9 Contribution of Smad-dependent TGFβ signalling on AMOT protein levels ...... 77

Figure 3.10 Effect of TGFβ on AMOT localization in migrating edge of JEG3 cells ...... 80

Figure 3.11 AMOT localization in live cell imaging of migrating JEG3 cells...... 82

Figure 3.12 Effect of AMOT 80 overexpression on migration rate of JEG3 cells during wound healing...... 83

Figure 3.13 Effect of TGFβ1/3 on AMOT and Par6 co-localization in JEG3 cells ...... 86

Figure 3.14 Effect of TGFβ1/3 on AMOT and Par6 interaction in JEG3 cells ...... 87

Figure 3.15 AMOT 80 and Par6 interaction in human placenta tissue ...... 88

Figure 3.16 Description and validation of AMOT 130 and AMOT 80 plasmid constructs ...... 91

Figure 3.17 Importance of PDZ and coiled-coil binding domains to AMOT-Par6 interaction .... 92

Figure 3.18 Effect of AMOT overexpression on protein levels of RhoA ...... 94

Figure 3.19 Effect of AMOT overexpression on RhoA localization ...... 95

Figure 3.20 AMOT protein levels in preeclamptic and normotensive pre-term control placentae

...... 98

Figure 3.21 AMOT localization in preeclamptic and pre-term control placenta tissue sections .. 99

Figure 3.22 AMOT 80 and Par6 association in preeclamptic and pre-term control placenta .... 100

Figure 3.23 Effect of low oxygen on AMOT protein levels in JEG3 cells ...... 102

Figure 3.24 Effect of low oxygen on AMOT localization in JEG3 cells ...... 103

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Figure 3.25 Effect of silencing and overexpressing JMJD6 on AMOT protein in JEG3 cells... 107

Figure 3.26 Effect of overexpressing JMJD6 on AMOT localization in JEG3 cells ...... 108

Figure 3.27 MALDI-Mass Spectrometry analysis of AMOT peptide mass profile following in vitro JMJD6 enzyme reaction ...... 109

Figure 3.28 Effect of minoxidil induced inhibition of lysyl hydroxylation on AMOT protein levels in JEG3 cells ...... 110

Figure 3.29 Effect of minoxidil induced inhibition of lysyl hydroxylation on AMOT localization in JEG3 cells ...... 111

Figure 4.1 Putative model depicting the role and regulation of AMOT in normal placentation and in preeclampsia...... 128

Figure 5.1 Investigating AMOT in placenta mesenchymal cells (pMSC) isolated from term placentae...... 135

Figure 5.2 Investigating AMOT localization term and preeclamptic placental mesenchymal cells

(pMSC) ...... 136

Figure 5.3 Investigating AMOT and TAZ spatial association in the human placenta ...... 139

Figure 5.4 Effect of TGFβ1/3 on AMOT and TAZ localization in JEG3 cells ...... 140

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Chapter 1

Introduction

Trophoblast proliferation, migration and invasion are critical events during human placenta development that contribute to the establishment of the feto-maternal interface in pregnancy.

Changes in oxygen tension and downstream TGF signalling experienced by the developing placenta have been demonstrated to regulate these events. However, in preeclampsia (PE), a devastating pregnancy disorder associated with increased maternal and perinatal mortality worldwide, persistent chronic hypoxia and aberrant TGF signalling has been shown to impair trophoblast differentiation and function, ultimately leading to defects in spiral artery remodelling and vascularization of the fetus. A fundamental element in eukaryotic cells regulated by TGF signalling, that is also integral to cell migration processes, is apical-basolateral cell polarity. Yet, the contribution TGF to the regulation of a novel cell polarity protein, termed Angiomotin

(AMOT), remains to be established. Moreover, AMOTs role in the human placenta and in trophoblast cell events remains unknown. In this chapter, I will first introduce the current literature surrounding AMOT’s discovery and function, next elucidate the complex regulation of placenta development by upstream oxygen and TGF signalling, and finally how it all goes awry in PE.

1 2

1.1 Angiomotin (AMOT) 1.1.1 Discovery and Structure

Angiomotin (AMOT) was first discovered in 2001 for its ability to bind to and mediate the effects of angiostatin, a circulating inhibitor of endothelial cell migration and tube formation during angiogenesis (Troyanovsky et al., 2001). Comprised of 675 amino acid residues and with a molecular mass of 80kDa, AMOT was the founding member of the ‘motin’ family of proteins

(Bratt et al., 2002; Troyanovsky et al., 2001). Succeeding investigations identified a 130kDa

AMOT product, structurally identical to the 80 kDa AMOT protein with an additional 409 amino acid N-terminal extension that arises from alternative splicing of the AMOT gene between exons

2 and 3 (Ernkvist et al., 2006). Thus, it was determined that the AMOT gene encodes for two isoforms: AMOT 80 and AMOT 130. In pursuit of identifying conserved domains important for

AMOT function, two other motin family members with significant sequence homology to AMOT were also identified: angiomotin-like 1 (AMOTL-1), and angiomotin-like 2 (AMOTL-2) (Bratt et al., 2002). The motin family of proteins exhibit two conserved protein motifs, namely a coiled-coil and PDZ binding domain (PSD95 (post synaptic density protein), Dlg1 (drosophila disc large tumour suppressor), ZO-1 (zona occludens-1)) (Figure 1.1).

Coiled-coil domains, structurally characterized by two alpha helices woven around each other, are a principal protein motif permitting the oligomerization of proteins and mediating protein-protein interactions (Burkhard et al., 2001). PDZ domains are abundant, globular interaction motifs that facilitate protein-protein interactions during signal transduction events, and organize proteins to specific sites or membrane structures (Lee and Zheng, 2010). In particular, PDZ domains are common components of proteins that localize to tight junctions (Tsukita et al., 2001). In fact, studies in epithelial and endothelial cells have revealed all motin members to localize at the tight

junctions, where it is suggested they elicit an important role in apical-basolateral cell polarity and stability of the cytoskeleton (Moleirinho et al., 2014). The presence of these characterized protein- interaction domains underscores the capability of the motin family of proteins to be involved in cellular scaffolding and signal transduction events.

The structural similarities amongst AMOT (80kDa and 130kDa isoforms), AMOTL-1 and

AMOTL-2 foster a degree of functional redundancy; however, there is functional variability within the family. These functional differences are highlighted by the variable spatial and temporal expression patterns amongst members, and the tissue-and cell-dependent expression patterns

(Moleirinho et al., 2014). Further, a number of functional and mechanistic studies have been conducted in pursuit of distinguishing the relevance of individual motin family members in different organ systems and signalling pathways.

Early studies in mouse aortic endothelial (MAE) cells revealed that AMOT 80 expression increases endothelial cell migration and contributes to vessel tube formation (Troyanovsky et al., 2001).

Moreover, studies examining the functional necessity of PDZ binding domain on AMOT 80 revealed that mutations in its PDZ domain resulted in complete loss of migratory activity in MAE cells (Levchenko et al., 2003). Further, transgenic mice expressing AMOT with mutated PDZ lose response to growth factors and are embryonic lethal due to impaired vascularization. On the other hand, AMOT 130 expression did not promote migration or mediate a response to angiostatin but was found to induce changes in cell shape through binding and stabilization of the F-actin cytoskeleton (Ernkvist et al., 2006). This AMOT 130/ F-actin interaction was determined to be mediated by the N-terminal domain on AMOT 130 (Ernkvist et al., 2008). Thus, the distinction was made that AMOT 80 functions primarily in endothelial cell migration, whereas AMOT 130 primarily controls changes in cell shape and regulates cytoskeleton reorganization. With regards

to AMOTL1 and AMOTL2, both proteins have been shown to interact with tight junction residing transmembrane proteins (Nishimura et al., 2002a; Patrie, 2005; Sugihara-Mizuno et al., 2007).

However, AMOTL1 and AMOTL2 lack the hydrophobic angiostatin binding domain present in

AMOT 130 and 80 and thereby fail to mediate its effects during angiogenesis (Bratt et al., 2002).

Additionally, transcript expression analysis of motin family members at the time of AMOT discovery revealed that AMOT had highest mRNA expression in the human placenta, compared to AMOTL1 and AMOTL2 which had highest expression in lung and skeletal muscle (Bratt et al.,

2002). Although this study identified AMOT expression in the human placenta, there has been no further investigation into its role. Hence, the focus of the present study remained on AMOT protein, both 130kDa and 80kDa isoforms, and its undiscovered role in the human placenta.

These aforementioned studies act as an introduction to the breadth of work which has been subsequently conducted to distinguish the function of AMOT isoforms. These include investigations into regulation of the Hippo pathway, as well as studies in cell polarity regulation, which will be described in this chapter.

5 Angiomotin (AMOT)

2 7 9 4 8 Coiled coil Angiostatin PDZ 0 2 2 1 Y Y Y E E T P P binding binding binding P 9 P 4 P 6 L 3 8 1 0 2 2 429 689 721 751 867 1005 1081 1084 1 AMOT 130 130 kDa 1084 a.a

1 20 280 312 342 458 596 672 675 AMOT 80 80 kDa 675 a.a

AMOT Like-1 (AMOTL1) AMOT Like-2 (AMOTL2)

3 0 3 1 1 7 1 5 9 3 3 7 2 5 1 0 2 Y Y 1 Y F Y E E Y Q V T P P T P P P 0 P P L 1 7 P 0 P 2 P 1 8 3 6 4 L 1 5 8 3 438 694 729 762 953 956 1 0 2 2 308 581 777 780 1 106 kDa 1 86 kDa 780 a.a 956 a.a

Figure 1.1 Protein structures of ‘motin’ family members

The members of the motin family of proteins include: (1) Angiomotin (AMOT), (2) Angiomotin Like-1 (AMOTL1), and (3) Angiomotin Like-2 (AMOTL-2). Notably, AMOT is expressed as two isoforms due to alternative splicing, AMOT 130 and AMOT 80, which corresponds to their respective molecular weights. The motin protein members share distinct structural characteristics. AMOT 130 and AMOT 80 contain coiled coil regions and PDZ binding domains, motifs conserved across all motin members, as well as an angiostatin binding domains. However, AMOT 130 contains a 409 amino acid N-terminal extension comprising distinct PPxY binding motifs that promotes its interaction with Hippo Pathway effectors, YAP (Yes-associated protein) and TAZ (Transcriptional coactivator with PDZ binding motif), in addition to mediating the interaction with the F-actin cytoskeleton (Chan et al., 2011; Ernkvist et al., 2006). AMOTL-1 and AMOTL-2 exhibit similar sequence identity with AMOT 130, however they lack the angiostatin binding domain. AMOTL-1 was initially identified in a screen for novel tight-junction associated proteins (Nishimura et al., 2002b) and named JEAP (junction-enriched and-associated protein), but later discovered to share sequence homology with AMOT 130 and was subsequently named AMOTL- 1. AMOTL-2 is also referred to as MASCOT (MAGI-1-associated coiled-coil tight junction protein) due to its colocalization with MAGI-1 at epithelial tight junctions (Patrie, 2005). Despite these structural similarities, the individual motin proteins are functionally distinct, made evident by their differential expression in various tissues and cell types. PDZ- PSD95 (post synaptic density protein), Dlg1 (drosophila disc large tumour suppressor), ZO-1 (zona occludens-1).

1.1.2 Role of AMOT in Hippo Pathway Signalling

Yes-associated protein (YAP) and transcriptional coactivator with PDZ-binding motif (TAZ) are transcriptional co-activators of the Hippo Pathway, a signalling pathway which has over the years been regarded to have important regulatory control in several cellular processes including cell proliferation and differentiation, and in cancer cell progression (Moroishi et al., 2015). YAP and

TAZ are structural homologs and exhibit functional redundancy. Specifically, YAP and TAZ transcription factors shuttle between cytoplasm and the nucleus, where they are permitted to interact with TEA domain family members (TEAD) transcription factors to induce the transcription of genes controlling cell proliferation and epithelial-mesenchymal transition (Lei et al., 2008; Zhao et al., 2008). Tight regulation of YAP/TAZ activity is essential as hyperactivity of

YAP/TAZ has been implicated in a variety of malignant and metastatic cancers and associated with poor outcomes (Piccolo et al., 2014). YAP and TAZ activity is regulated by inhibitory Hippo kinases large tumour repressor 1 and 2 (LATS1 and LATS2), which directly phosphorylate YAP and TAZ at several serine residues, promoting the sequestration of YAP and TAZ in the cytoplasm

(Oh and Irvine, 2010), or alternatively targeted for ubiquitination and subsequent proteasomal degradation (Zhao et al., 2010). However, a third fate exists, which is the direct binding and regulation of YAP and TAZ by AMOT 130 (Chan et al., 2011; Zhao et al., 2011). AMOT 130 interaction with YAP/TAZ occurs between tryptophan binding domains (WW) on YAP/TAZ, and the PPxY binding motif within the N-terminal extension found on AMOT 130 (Chan et al., 2011).

However, the nature of AMOT’s regulation over YAP/TAZ remains a subject of controversy within the field of the Hippo pathway, as conflicting reports have outlined AMOT as both a negative and a positive regulator of YAP/TAZ activity.

AMOT 130 as a Negative Regulator of YAP/TAZ in the Hippo Pathway

The first evidence of AMOT 130 as a negative regulator of Hippo Pathway was demonstrated in a study by Zhao et al showing that AMOT 130 inhibits YAP activity in Madin-Darby Canine Kidney

(MDCK) cells by sequestering YAP to intracellular tight junctions, or to the F-actin cytoskeleton in the cytoplasm, thus effectively preventing YAP nuclear translocation and gene transcription

(Zhao et al., 2011). This mechanism is independent of LATS1/2 mediated phosphorylation.

Additionally, AMOT 130 can negatively regulate YAP/TAZ by promoting inhibitory phosphorylation of YAP and TAZ (Zhao et al., 2011). This apparent tumour suppressor role of

AMOT 130 on YAP/TAZ has been elucidated in a number of cancer studies (Moleirinho et al.,

2017; Moroishi et al., 2015). For example, expression of AMOT 130 is significantly decreased in clinical lung cancer specimens, and furthermore knockdown of AMOT 130 in lung adenocarcinoma cancer cell line was shown to initiate cancer proliferation, migration and invasion by promoting YAP/TAZ nuclear translocation (Hsu et al., 2015).

AMOT 130 as a Positive Regulator of YAP/TAZ in the Hippo Pathway

In stark contrast to these findings attributing an inhibitory role for AMOT 130 in Hippo signalling, reports in biliary epithelial cells have demonstrated AMOT 130 to function as a positive-regulator of YAP/TAZ (Hong, 2013). AMOT 130 can compete with LATS1 for binding to YAP, thereby preventing YAP phosphorylation, which facilitates YAP/TAZ nuclear translocation. Further, studies in hepatic epithelial cells have demonstrated AMOT 130 can also localize to the nucleus, where it forms a complex with YAP and TEAD1, thus contributing to transcription of YAP/TAZ target genes associated with tumorigenesis (Yi et al., 2013a). This positive interplay between

AMOT and YAP/TAZ has been observed in models of hepatic carcinoma, renal epithelial cells and renal cell cancers (Lv et al., 2016; Yi et al., 2013a).

Recent studies in embryonic kidney cells, endothelial cells, and mouse and zebrafish embryos suggest that the phosphorylation status of AMOT is important in distinguishing whether AMOT functions as a positive or negative regulator of YAP/TAZ mediated transcription in the Hippo pathway (Chan et al., 2013; Dai et al., 2013; Hirate and Sasaki, 2014). Phosphorylation of AMOT by LATS1/2 promotes AMOT mediated redistribution of YAP/TAZ from either the nucleus or cytoplasm, to the plasma membrane. As such, the ability of YAP/TAZ to promote cell proliferation and tumorigenesis is abrogated. On the other hand, when AMOT is in its unphosphorylated state,

AMOT is able to translocate into the nucleus and function as a positive cofactor in the transcription of YAP target genes.

1.1.3 Role of AMOT in Cell Migration

Independent of its role in mediating downstream Hippo pathway gene transcription, AMOT has also been shown to influence endothelial and epithelial cell migration. This was apparent in zebrafish studies which showed Amot knockdown reduced the number of filopodia in endothelial cells, severely impairing the migration of intersegmental vessels during embryogenesis (Aase et al., 2007). Total knockdown of Amot in mice resulted in embryonic lethality between E11 and

E11.5, as a result of severe vascular insufficiency in the intersomitic region and dilated vessels in the brain (Aase et al., 2007). Notably, the placenta was not thoroughly investigated. Nonetheless, this study suggested AMOTs role in cell migration could be attributed to novel function of AMOT in the regulation of cell polarity (Aase et al., 2007). Thereafter, studies have underscored how the intracellular scaffolding abilities of AMOT can promote the binding and shuttling of various cell polarity components, including tight junction components, polarity complex proteins and small G- proteins, in order to promote alterations in cell polarity.

1.2 Cell Polarity Regulation 1.2.1 Tight Junctions

Cell polarity is a fundamental element in all eukaryotic cells that controls changes in cell-shape, cell migration, cell fusion and epithelial-to-mesenchymal transition (Martin-Belmonte and

Mostov, 2008). Apical-basolateral cell polarity arises in epithelial cells as a result of asymmetric distribution of lipids and proteins to the cell surface on the apical or basolateral ends of the cell

(Assemat et al., 2008). Alongside the adherens junction, which provide structural support to the cells via linkage to cytoskeletal elements, the tight junctions (TJ) is a key intracellular junctional complex involved in the regulation of cell polarity (Shin et al., 2006). The TJ mediates adhesion between neighboring cells by forming a selectively permeable barrier to diffusion through intercellular space, a property referred to as the “barrier” function. However, in regards to its role in cell polarity, the TJ is able to restrict the intracellular localization of proteins and macromolecules between the apical and basolateral regions of the cell by delineating the boundary between these two domains, a characteristic referred to as the “fence function” (Shin et al., 2006;

Zihni et al., 2016). The TJ is comprised of transmembrane protein components; occludin, claudin and junctional adhesion molecules, all of which interact with underlying peripheral membrane proteins to form a complex protein network (Shin et al., 2006). Importantly, these underlying peripheral membrane proteins serve as a link to the actin cytoskeleton (Zihni et al., 2016). These underlying proteins, which typically contain PDZ and tryptophan (WW) binding domains, include scaffolding proteins, kinases, phosphatases, small GTPases and their activating proteins, actin binding proteins and F-actin itself (Quiros and Nusrat, 2014). For example, Zonula occludens-1

(ZO-1) is a peripheral scaffolding protein belonging to the membrane-associated guanylate kinase

(MAGUK) family of proteins, which connects tight junction proteins (occludin and claudin) to the actin cytoskeleton (Shin et al., 2006). Further, Ras homolog gene family member A (RhoA), a

small GTPase protein, is another peripheral membrane protein that has dual regulatory roles in tight junction assembly and actin cytoskeleton arrangement (Quiros and Nusrat, 2014). Thus, molecules involved in tight junction stability, are also involved in cytoskeleton reorganization and altogether contribute to the establishment of epithelial polarization.

1.2.2 Polarity Protein Complexes

There are three main protein complexes involved in the maintenance and loss of apical-basolateral polarity: Par/aPKC complex, composed of Par3, Par6, and atypical protein kinase C (aPKC);

Crumbs complex, composed of crumbs, Pals1 (Protein associated with Lin-7), Patj (Pals1 associated tight junction protein homolog); and Scribble complex, comprised of scribble, discs large (Dlg) and lethal giant larvae (Lgl) (Assemat et al 2007). Protein subunits within these complexes are able to interact with one another through shared PDZ-binding domain motifs and localize certain cellular components to poles of the cell, promoting apical-basolateral membrane identity and cell polarization (Assemat et al., 2008; Martin-Belmonte and Mostov, 2008).

The Par complex defines the apical region of the cell, and is involved in the early events of cell- cell adhesion and tight junction formation (Henrique and Schweisguth, 2003). Specifically, Par3 binds to transmembrane junctional adhesion molecules (JAM) at the site of cell-cell contact, while

Par6 and aPKC initially reside with Lgl component of the scribble complex in the cytoplasm. Next, cell division control protein 42 homolog (Cdc42), a master regulator of actin cytoskeleton rearrangements explained in detail below, becomes activated and promotes Par6 and aPKC to associate with Par3 at the tight junction to complete the Par complex at the apical/basolateral membrane (Yamanaka et al., 2001). Simultaneously, the Lgl component of the scribble complex is displaced to the basolateral membrane where it is found associate to other scribble proteins and defines the basolateral domain. The crumbs complex resides in the apical region alongside the Par

complex, and collectively they identify the apical-membrane (Figure 1.2A). Loss of any one of these polarity complex proteins can lead to the disruption of cell polarity in mammalian epithelial cells (Assemat et al., 2008; Macara, 2004; Martin-Belmonte and Mostov, 2008).

1.2.3 Role of Cell Polarity in Proliferation, Migration and Invasion

It is evident that tight junction proteins are not only permeability barriers between cells, but are sensors involved in a variety of cellular processes, such as cell proliferation. In conditions of increasing cell density within epithelial sheets, tight junction associated mechanisms sense this, and inhibit further cell proliferation by impeding transcription of proliferative related genes.

Specifically, tight junction protein ZO-1 increases expression in conditions of high cell density

(Balda and Matter, 2000), binds to ZO-1-associated nucleic acid binding protein (ZONAB), and sequesters it to the cytoplasm to prevent its mediated gene transcription and ultimately reduce proliferation (Balda et al., 2003). Another example is the previously outlined role of AMOT 130 in recruiting Hippo pathway kinases to phosphorylate and inactivate transcription activators

YAP/TAZ, or sequestering YAP/TAZ out of the nucleus (Zhao et al., 2011).

Tight junction proteins are also important when applied in the context of epithelial cell migration.

Importantly, a series of tightly coordinated steps are required for proper directional cell migration.

Cell migration begins in response to external stimuli such as growth factors or extracellular matrix

(ECM) molecules, where the driving force is the polarized extension of a leading edge protrusion, or “lamellipodium” in response to direction of movement. Following the formation of the lamellipodium, new adhesion sites are established at the leading edge, the cell undergoes actin/myosin contraction, and previous adhesion sites located at the tail of the cell are detached to permit cell movement. One family of proteins that plays a pivotal role in these steps of cell migration is the tight junction residing Rho family of GTPases, namely Cdc42, Rac1 (Ras related

C3 Botulinum Toxin Substrate-1) and RhoA (Ras homolog gene family member A). Cdc42 responds to external stimuli and regulates the direction of cell movement, Rac1 stimulates actin polymerization and integrin adhesion complexes to the lamellipodium, and RhoA promotes actin: myosin contraction to complete the cycle (Raftopoulou and Hall, 2004). Studies aimed at investigating the mechanism in which polarity proteins assemble at the leading edge of migrating epithelial cells revealed that tight junction component occludin localized to the leading edge of migrating cells, and regulated directional cell migration by binding to and localizing polarity complex aPKC-Par3 and Patj to the leading edge. In turn, this promoted cell protrusions by activating Rac1 to direct lamellipodia formation (Du et al., 2010) (Figure 1.2B).

1.2.4 AMOT as a Novel Regulator of Cell Polarity

Studies conducted in endothelial and epithelial cells have characterized AMOT localization at the tight junction and determined a role for AMOT in apical-basolateral cell polarity and cytoskeletal stabilization (Lv et al., 2017). A study investigating polarity complexes in MDCK cells determined that the coiled-coil domain on AMOT 80 binds to Rich1, a GTPase activating protein necessary to tight junction formation and stabilization, and collectively the two proteins localize to the tight junction (Wells et al., 2006). Here, AMOT 80 was demonstrated to directly interact with Par and

Crumbs Complex proteins Pals1, Patj/Mupp1, and Par3 via their individual PDZ-binding motifs

(Wells et al., 2006). However, MDCK cells stably overexpressing AMOT 80 exhibited tight junction dissolution. A potential explanation is that AMOT 80 overexpression above a certain threshold can promote the selective redistribution of Par and Crumbs complex components alongside scaffolding protein AMOT into punctuate structures within the cytoplasm, and away from the tight junction (Wells et al., 2006). Resulting from the sequestration of essential tight junction proteins away from cell boundaries to the cytoplasm include disruptions in tight junction

integrity, alterations in apical-basolateral cell polarity and promotion of cell movement. The roles of AMOT 80 in cell migration and cell polarity were linked in a study by Aase et al, where isolated and immortalized embryonic stem cells from an Amot deficient mouse model (Amot - EC) exhibited defects in cell migration, as wild type EC migrated five times longer than Amot – EC in two independent migration assays (Boyden chamber and time-lapse wound healing) (Aase et al., 2007).

AMOT – EC also displayed defects in cell polarization, as shown by failure of the Golgi apparatus and GTPase Rich1 to localize to the lamellipodia. Further, AMOT – EC exhibited defects in actin cytoskeleton organization and changes in cell shape, as elucidated by disorganized pattern of actin fibers and shorter focal adhesions in Amot – EC compared to wildtype EC (Aase et al., 2007). In another study, Ernkvist et al found that AMOT 80 binds to RhoA GTPase exchange factor ‘Syx’ via its PDZ binding motif, which forms a ternary protein complex with Patj (AMOT/Patj/Syx) and regulates activity of RhoA GTPase in lamellipodium of migrating cells (Ernkvist et al., 2009).

These findings provided further evidence that AMOT’s role as a cell polarity regulator is intricately connected to its role in promoting cell migration.

The role of cell polarity, and the signalling pathways that regulate it, has been a subject of intense investigation in the field of cancer due to the strong correlation between malignancy of epithelial cancer and loss of epithelial organization attributed to loss of polarity (Bilder, 2004). Further, loss or deregulation of epithelial cell polarity processes has been characterized as a hallmark of cancer, as it plays a role in the initiation of tumorigenesis and later stages of tumour development (Royer and Lu, 2011). Interestingly, comparisons have been drawn between cancer cell differentiation in the tumour microenvironment, and the differentiation of trophoblast cells at the fetomaternal interface (Holtan et al., 2009). More specifically, the parallel lies in the proliferation, migration and invasion of tumour cells during cancer, and trophoblast cells during placental development.

Figure 1.2 Polarity complexes in migrating epithelial cells

(A) The Crumbs, Par and Scribble protein complexes tightly regulate cell polarity in epithelial cells. The localization of these complexes and their complex protein-protein interactions define the apical and basolateral regions within epithelial cells. The Par and Crumbs complex define the apical domain of epithelial cells, whereas the Scribble complex defines the basolateral domain. During the early events of tight junction formation, Par6 and aPKC components of the Par complex are found bound to Lgl component of the scribble complex within the cytoplasmic region of the

cell. Upon sensing of extracellular cues to establish apical basolateral cell polarity, Cdc42 is activated and promotes Par6 and aPKC to redistribute to the tight junction to join Par3 component which is bound to transmembrane protein JAM, and ultimately complete the Par complex. Simultaneously, Lgl component is displaced to basolateral membrane and joins to other scribble complex proteins(Chatterjee and McCaffrey, 2014). The complex interactions amongst these protein complexes, as well as interactions between individual proteins and cellular scaffolding proteins, occur via shared putative PDZ binding domains. The known interactions between polarity proteins and scaffolding protein AMOT are noted by the blue asterisk (Sugihara-Mizuno et al., 2007; Wells et al., 2006). (B) The distinct, step-wise events during epithelial cell migration require precise regulation by Rho family members (Cdc42, Rac1 and RhoA), as well as require changes in polarity complex localization and their complex interactions (1) Cdc42, a master regulator of actin cytoskeletal rearrangements, controls the direction of cell migration in response to external growth factors or ECM signals by promoting the protrusion of the lamellipodia (cells leading edge). Rac1 stimulating actin polymerization at the cell’s leading edge, as well as the localization of apical polarity complex proteins to the leading edge mediate the formation of this protrusion. (2) New cell adhesions at the lamellipodia are generated via Rac1 promoting the formation integrin adhesion complexes. (3) Rho A promotes actin-myosin contraction of the epithelial cell body, (4) simultaneously promoting the dissolution of old, rear adhesions and retraction of the trailing ‘tail’ of the epithelial to promote cell migration in the respective direction (Raftopoulou and Hall, 2004). TJ (Tight junction) AJ (Adherens junction), Par6 (Partitioning defective protein 6), Par3 (Partitioning defective protein 3), Patj (Pals1 associated tight junction protein), Pals1 (Protein associated with Lin-7 1), Crb, (Crumbs) Cdc42 (Cell division control protein 42), aPKC (atypical protein kinase 1), Scrib (Scribble), Lgl1/2 (Lethal giant larvae protein homolog1/2), Dlg (Discs large protein), RhoA (Ras homolog gene family member A), Rac1 (Ras related C3 Botulinum Toxin Substrate 1).

1.3 Human Placenta Development

Successful pregnancy is fundamentally dependent on placental function, and thereby hinges on proper placenta development. The human placenta is vital to the proper growth and development of the fetus, as elucidated by its multi-functionality in a variety of critical functions including gas exchange, transportation of nutrients, elimination of waste, and production of hormones (Costa,

2016). Development of the human placenta begins immediately following fertilization and is a continuous process that occurs alongside development of fetus. Specifically, the formation of the blastocyst on the fifth day after fertilization delineates two critical structures: (1) the inner cell mass, which subsequently forms the embryo and fluid-filled blastocoel, and (2) the outer layer of cells referred to as trophectoderm, which continue on to form the placenta and fetal membranes.

Initial nourishment of the developing embryo is provided through uterine secretions containing oxygen and metabolic substrates, and subsequent uptake by the trophoblast layer (Burton et al.,

2001). However, to provide sustainable nutrient and oxygen support necessary for further development, access to maternal decidua is required. Specifically, the blastocyst implants into the uterine lining (referred to as the ‘decidua’) via invasion of the outer trophoblast cells into the endometrial layer of the uterus. Initial trophoblast invasion into the decidua leads to erosion of uterine tissue, and consequently generates a network of lacunae which become filled with maternal blood for nutrition of the fetus. This blood-filled lacuna network is referred to as the ‘intervillous space’. Following implantation of the blastocyst, progenitor trophoblast cells differentiate into distinct trophoblast subtypes which are vital to the development of a fully functioning placenta.

These trophoblast differentiation events, and their regulation, will be outlined in this next section.

1.3.1 Trophoblast Differentiation

Progenitor trophoblast cells, referred to as cytotrophoblast (CT), are proliferative cells with two fates during development. One is fusion of CT to form the multi-nucleated syncytiotrophoblast cell layer (ST), which functions as the site of gas and nutrient exchange between the mother and fetus in the intervillous space. These two cellular layers, along with a mesenchymal core of connective tissues and blood vessels, comprise a villous structure directly bathed in maternal blood referred to as the floating villus. Concurrently, CT also undergo transformation to become extravillous trophoblast cells (EVT). Here, the EVT’s erupt through the overlaying syncytium at

4-5 weeks of gestation to form cellular columns, which anchor the villi to the maternal decidua, and are known as the anchoring villi. This transformation of CT to EVT and subsequent attachment to uterine tissue involves cellular differentiation of CT from a proliferative, to a migratory, and finally invasive phenotype moving from proximal to distal ends of the anchoring villi. Once EVTs acquire invasive capabilities, they first undergo interstitial invasion and infiltrate the decidua. At the decidua, the EVTs begin endovascular invasion of the maternal spiral arteries by displacing vascular smooth muscle and endothelial cells. This effectively transforms the spiral arteries from narrow, high-resistance vessels, into wide, low resistance conduits, facilitating the increase of oxygenated blood flow into the lacunar vascular system and establishment of uteroplacental circulation. It is at this point in development where important physiologic changes in oxygen tension occur (Figure 1.3).

Distinct molecular changes occur within epithelial CT as they transform into EVTs and begin their quest towards invading the maternal spiral arteries. One such change includes the increased secretion of metalloproteinases, which breaks down extracellular matrix proteins to facilitate EVT migration and invasion through the endometrium (Fisher et al., 1989). Another important

molecular alteration involves the upregulation of 5 and 1 integrins in differentiating and invasive EVTs, which has proven to be critical for EVT invasion (Damsky et al., 1994).

Interestingly, this process of EVT differentiation in the human placenta has been compared to the process of epithelial-mesenchymal transition (EMT) often seen during embryonic development or cancer metastasis (Kalluri and Weinberg, 2009). During EMT, epithelial cells lose expression of cell-cell junction molecules (i.e. E-cadherin, ZO-1, and desmoplakin), adhere to ECM via integrin binding, and shift cell polarity from apical-basolateral to “front-back” orientation to facilitate migratory and invasive potential (Hay, 1995). Recently, gene expression analysis by qPCR revealed that EVT cells isolated from first trimester placentae exhibited downregulated expression of epithelial markers such as E-cadherin and occludin, and upregulated expression of mesenchymal markers such as vimentin, fibronectin and extracellular matrix integrins 5 and 1, when compared to first-trimester isolated CT (DaSilva-Arnold et al., 2015). Additionally, EVT cells had upregulated matrix metalloproteinases MMP2 and MMP9 that are necessary for invasion of ECM, as well as a robust increase in EMT regulator ZEB2 (Zinc finger E-box binding homeobox 2)

(Gheldof et al., 2012). EMT processes require strict regulation by growth factors and other molecules, or else pathologies may arise in which cells have aberrant abilities to grow, proliferate, migrate or invade (i.e. carcinoma progression). EMT-like trophoblast differentiation, and resultant invasion, is tightly regulated during human placenta development by a variety of growth factors, cytokines and most evidently, changes in oxygen tension (Knofler, 2010).

Prior to endovascular EVT invasion of the spiral arteries, the early human placenta must develop in a hypoxic environment to protect the embryo against oxygen free radical mediated damage

(Burton et al., 2003). The low oxygen environment is established as a result of EVTs forming endovascular ‘plugs’ along the lumen of spiral arteries, which restricts maternal blood flow into

the intervillous space. Development in low oxygen is required not only to protect the developing embryo, but also functions to maintain progenitor trophoblast cells in a proliferative, non-invasive phenotype (Genbacev et al., 1996). Later, coinciding with the timing of spiral artery remodeling, steep changes in placental oxygen tension occur; oxygen levels measured to be <20mmHg at 8- weeks of gestation, rise to >50mmHg by 12- weeks of gestation (Jauniaux et al., 2000). Recent studies have, however, suggested increases in oxygen tension may occur even earlier; contrast enhanced ultrasound imaging showed detectable increases in blood flow into intervillous space as early as 6-weeks of gestation due to progressive disintegration of uteroplacental ‘plugs’ (Roberts et al., 2017). Nonetheless, these alterations in oxygen tension experienced by the placenta have been shown to regulate the aforementioned trophoblast differentiation events via transcription factor hypoxia-inducible factor 1 (HIF-1) (Caniggia et al., 2000). Specifically, in low oxygen conditions such as those seen in early placenta development, HIF-1 positively regulates levels of transforming growth factor 3 (TGF3) to maintain trophoblast cells in a proliferative, non- invasive phenotype. Increased exposure to oxygen reduces HIF-1 and levels of TGF3, allowing

EVT differentiation and invasion into the decidua to continue (Caniggia et al., 1999; Caniggia et al., 2000; Ietta et al., 2006).

TGFs are a subgroup of growth factors within the TGF superfamily that regulate multiple biological processes including cell proliferation, differentiation, migration, EMT and apoptosis

(Massague, 2008). Interestingly, the human placenta is a major tissue source of TGF (Jones et al., 2006), where it has been reported to regulate trophoblast cells during development. The signalling events activated by the TGF family of growth factors, and how this impacts trophoblast cell function is outlined next.

20 A Umbilical Intervillous Umbilical space veins arteries

Chorionic Villi

ST CT EVT Maternal spiral artery

Fetal Side Maternal Side Decidua Myometrium B Fetal Blood Migratory ST CT Vessel Stroma EVT Spiral artery

Invasive pMSC EVT Anchoring villi column

Proximal Intermediate Distal Floating villi 5-9 weeks 10-12 weeks

~20 mmHg Oxygen Gradient ~55 mmHg

~2-3% O2 ~6-8 % O2 Figure 1.3 Human placenta development and trophoblast differentiation

(A) General architecture and vascularization of the mature human placenta. The fetal side of the placenta (chorion) is comprised of umbilical veins and arteries, chorionic villi (cytotrophoblast (CT) and syncytiotrophoblast (ST) layers), and amnion. The maternal side of the placenta attached to the uterine wall, is comprised of uterine vessels and decidua basalis. Upon remodeling of the maternal spiral arteries by distal, endovascular extravillous trophoblast cells (EVT), the maternal

blood pools into the intervillous space and comes in direct contact with the chorionic villi structure, achieving uteroplacental vascularization. (B) Structure of floating villi and anchoring villi at the fetomaternal interface. In the floating villi, underlying proliferative CT fuse to form the overlying, multinucleated ST that facilitates gas and nutrient exchange. Placenta mesenchymal cells (pMSC) and fetal blood vessels and capillaries are found within the stromal core of the chorionic villi. During the formation of the anchoring villi, trophoblast cells break through the syncytium and develop into EVT to form an anchoring column. In the anchoring column, EVTs differentiate through proliferative, migratory and invasive phenotypes as they move from proximal to distal ends of the column. Distal EVTs establish fetomaternal blood flow by invading through the maternal decidua (interstitial EVTs) to ultimately reach and remodel the spiral arteries within the myometrium layer (endovascular EVTs). Levels of oxygen regulate these trophoblast differentiation events. During early gestation (5-9 weeks), low oxygen levels retain trophoblast cells in a proliferative, undifferentiated state; however, following a rise in oxygen tension around 10-12 weeks, CT differentiate into migratory and invasive EVTs required for spiral artery remodeling and increase in maternal blood flow to the intervillous space and ultimately the fetus (Simon and Keith, 2008).

1.3.2 TGF signalling pathways

The TGF superfamily is the largest family of secreted morphogens and its members are highly conserved across animals (Wrana, 2013). In addition to the three TGF isoforms (TGF1, TGF2 and TGF3), this superfamily of growth factors also includes activins, bone morphogenic proteins

(BMP), and growth differentiating factors (GDF). While all three TGF isoforms are expressed in the human placenta, TGF3 is the only isoform demonstrated to exhibit a temporal expression pattern, peaking at 7/8 weeks of gestation, and dropping thereafter. This is unlike TGF1 and

TGF2, which remained consistently expressed across gestation (Caniggia et al., 1999). As mentioned, this temporal peak in TGF3 is attributed to the hypoxic environment during early placenta development, where HIF-1 levels are abundant and positively regulate levels of TGF3

(Caniggia et al., 2000; Ietta et al., 2006). The drop in TGF3 levels coincide with an increase in oxygen tension and a reduction of HIF-1 stability at 10-12 weeks of gestation. Aside from its differential expression pattern, TGF3 has proven to be an inhibitor of cytotrophoblast outgrowth and invasion. Specifically, in vivo studies of placental explants showed that inhibition of TGF3, but not TGF1 or TGF2, restored the invasive capabilities of trophoblast cells, and increased both MMP production and increase fibronectin deposition (Caniggia et al., 1999). Moreover, only the TGF3 isoform was found elevated in placentae complicated with preeclampsia. Altogether these data suggest a role for TGF3 in regulating trophoblast differentiation events during human placenta development. However, investigation into the soluble factors within the decidua that regulate trophoblast invasion revealed that decidual derived TGF1 plays a role in the inhibition of trophoblast outgrowth and invasion (Graham and Lala, 1991; Lala and Graham, 1990).

Mechanistically, in vitro studies in first trimester derived trophoblast cells demonstrated TGF1 to supress trophoblast invasion by: (1) upregulating tissue inhibitors of metalloproteases (TIMPs),

which inhibit trophoblast-derived matrix proteases required for invasion (Graham and Lala, 1991);

(2) increasing the fusion of trophoblast cells into non-invasive multinucleated cells (Graham et al.,

1992); (3) upregulating cell surface integrins that result in elevated trophoblast adherence to ECM that impedes cell migration (Irving and Lala, 1995); (4) downregulation of urokinase type plasminogen activator (uPA), which is also required for invasion (Graham and Lala, 1992).

Investigation into the expression of TGF in maternal decidua and placenta revealed TGF2 to be particularly expressed in the ECM of first-trimester decidua and cytoplasm of term decidua cells

(Graham et al., 1992; Lysiak et al., 1995).

TGF signalling begins with generation of mature homo-or heterodimer TGF ligands. Following proteolytic cleavage of dormant, precursor TGF proteins within the extracellular matrix (ECM), the mature cleaved segments actively dimerize through disulfide links (Budi et al., 2017). The resultant dimerized TGF ligands are now primed to bind to a heteromeric transmembrane complex of serine/threonine receptors and activate downstream TGF signalling. This heteromeric complex is composed of two major types of transmembrane receptors: TGF receptor type I

(TGFRI), also referred to as activin receptor-like kinase (alk); and TGF receptor type II

(TGFRII), both of which possess intrinsic serine-threonine kinase activity (Attisano and Wrana,

2002; Wrana, 2013). While seven types of TGFRI, commonly referred to as alk1-7, have been identified, TGF signalling occurs primarily via alk5. In the instance of endothelial cells, however, signalling also occurs through alk1 (Piek et al., 1999). Following TGF ligand binding, downstream signalling occurs via Smad-dependent and Smad-independent pathways.

In Smad-dependent TGF signalling, also referred to as canonical TGF signalling, the TGF ligand binds to the constitutively active TGFRII, and promotes trans-phosphorylation and

activation of TGFRI (alk5) at a glycine-serine (GS) rich region. Activation of alk5 kinase activity, in turn, promotes the phosphorylation of Smad2 and Smad3 at C-terminal serine residues, which form a trimeric complex with Smad4 that translocates into to the nucleus (Wrana, 2013). Resident in the nucleus, this Smad complex interacts with other transcription factors to either activate or repress the expression of select genes, such as those involved in cell proliferation, migration and invasion. For example, in oral squamous cancer cells, TGFβ signalling via Smad3 upregulated the expression of microRNA miR-455-5p which promoted cancer cell proliferation (Cheng et al.,

2016). Inhibitory Smad proteins including Smad7 can repress TGF signalling by directly binding to phosphorylated TGFRI (Wrana, 2013). This effectively prevents the phosphorylation of

Smad2/3 while simultaneously targeting the receptor for degradation. Although Smad proteins are recognized as the main mediators of TGF signalling, TGF signalling can also occur independent of Smad activation, commonly referred to as Smad-independent or non-canonical TGF signalling

(Moustakas and Heldin, 2005). These include activation of MAP kinase pathway, as well as phosphatidylinositol-3-kinase/protein kinase B pathway (AKT) (Zhang, 2009). Notably, another non-canonical TGF pathway that has been implicated in TGF-mediated loss of tight junctions and loss of cell polarity during EMT, is the Par6/Smurf1 polarity pathway (Ozdamar et al., 2005).

In the TGF-Par6 polarity pathway, TGF ligand binding to the type II receptor induces the direct association and subsequent phosphorylation of polarity protein Par6 at the intracellular tight junctions (Bose and Wrana, 2006; Ozdamar et al., 2005). As mentioned previously, Par6 is a regulator of epithelial cell polarity and tight junction integrity (Assemat et al., 2008). Following the direct phosphorylation of Par6 by TGFRII, Smad ubiquitination regulatory factor 1 (Smurf1), an E3 ubiquitin ligase, is recruited to interact with Par6. In turn, Smurf1 ubiquitinates the tight junction stabilizer protein GTPase RhoA, resulting in its targeted proteasomal degradation.

Considering RhoA plays a fundamental role in tight junction stabilization and formation, its degradation leads to disassembly of the actin cytoskeleton, dissolution of tight junctions and loss of apical-basolateral cell polarity, all events which typify EMT (Ozdamar et al., 2005). In addition to EMT, phosphorylation of Par6 via non-canonical TGF signalling has proven to be essential for cell migration and invasion, key characteristics of cancer progression. Particular studies in prostate cancer showed phosphorylated Par6 to form a complex with aPKC polarity component at the leading edge of membrane ruffles. Further, the use of PKC specific inhibitors interfered with the formation of this polarity complex, and prevented prostate cancer cell invasion (Mu et al., 2015).

Congruently, studies in breast cancer showed the TGFβ-Par6 polarity pathway to regulate cancer metastasis; interference with Par6 signalling prevented TGFβ induced loss of polarity in mammary cells grown in 3D structures (Viloria-Petit et al., 2009). In addition, suppression of Par6 in an in vivo orthotopic mouse model induced formation of ZO-1 positive epithelium within the tumour whilst supressing lung metastasis (Viloria-Petit et al., 2009). This further highlights the involvement of non-canonical TGF signalling via polarity protein Par6, and its importance not only to EMT but to other cellular processes regulated by TGF including cell migration and invasion.

It is evident that both canonical and non-canonical TGFβ signalling pathways are involved in

TGFβ regulation over multiple cellular processes. In some cases, activation of two different TGFβ signalling arms can result in the same effect. For instance, induction of EMT that is achieved by the non-canonical TGFβ/Par6 polarity pathway via RhoA degradation, can also be the result of canonical TGFβ signalling where Smad2/3 promotes the transcription of genes involved in EMT, such as Snail (Peinado et al., 2003). On the other hand, TGFβ ligands have the ability to induce different TGFβ signalling arms, and thus variable downstream pathways, which can account for

the vast multifunctionality of these growth factors, as well as explain the conflicting reports which show TGFβ to promote opposing effects in the same tissues/cells. This is particularly relevant to the human placenta, where different TGFβ isoforms have been implicated as negative regulators of trophoblast cell migration and invasion (Caniggia et al., 1999; Graham and Lala, 1991, 1992;

Karmakar and Das, 2002; Tse et al., 2002), as well as positive regulators of trophoblast migration and proliferation (Xu et al., 2016). This latter role of TGF is in line with the positive effect of

TGFβ on cell migration and invasion observed in many cancers.

1.3.3 Role of TGFβ Signalling in Trophoblast Differentiation

TGFβ regulation of trophoblast cell differentiation in the human placenta occurs via both the canonical (Smad mediated) and non-canonical signalling (Par6/Smurf1 mediated) (Xu J et al 2016,

Sivasubramaniyam et al 2013) (Figure 1.4). Furthermore, this regulation during placentation occurs in a temporal and spatial manner. In regards to the canonical TGFβ signalling, receptor activated Smad2 is found particularly expressed in ST, where studies using explants and BeWo cells showed it to negatively regulate trophoblast cell fusion via downregulation of fusion regulators GCM-1 and ERVW-1 (Xu et al., 2016). Receptor activated Smad2 is also found in proliferating EVTs, and in vitro studies using JEG3 cells showed it to positively regulate trophoblast cell proliferation as elucidated by the upregulated expression of cell cycle regulators

CCNE1 and CDK4 (Xu et al., 2016). As gestation progressed, ST levels of pSmad2 decreased, and levels of inhibitory Smad7 increased, consistent with increased rates of cell fusion observed with advancing gestation. Concurrently, decreasing pSMAD2 and increasing SMAD7 levels were seen toward the distal end of the EVT column (Xu et al., 2016). This is consistent with the differentiation of EVTs from a proliferative to a migratory and invasive phenotype with increasing

gestation. The differential Smad expression patterns seen here highlight the fact that canonical

TGFβ signalling plays a distinct role in different trophoblast subtypes.

Concerning the non-canonical TGFβ pathway, Par6/Smurf1 signalling was observed to be activated at 10-12 weeks of gestation in EVTs located in the intermediate region of anchoring column (Xu et al., 2016). Studies using JEG3 cells revealed that TGFβ treatment increases the association of Par6 and Smurf1 and promotes trophoblast cell migration in vitro via dissolution of tight junctions (Xu et al., 2016). This prompted the conclusion that trophoblast cell migration is regulated by the TGFβ-Par6 polarity pathway. Additionally, this study also observed Par6/Smurf1 association in CT at 10-12 weeks of gestation, a time when CT are known to undergo fusion into

ST, suggesting that TGFβ-Par6 signalling also mediates trophoblast fusion. These findings underscore the fundamental role that cell polarity plays in the human placenta, particularly in the proper functioning and differentiation of trophoblast cells. However, aside from the aforementioned findings on polarity protein Par6 from our group, no studies have investigated the role of other cell polarity proteins and signalling events on trophoblast cell differentiation. Further, no studies have looked at the impact of TGFβ on the functionality of other polarity proteins, or scaffolding proteins, in the placenta.

This section has elucidated to the complexity of trophoblast differentiation during human placenta development, and thus has underscored the grave importance of its precise regulation. In fact, dysregulation of these differentiation processes is implicated in the pathogenesis of placenta related diseases such as preeclampsia.

Figure 1.4 Role of Smad-dependent and Par6-mediated TGFβ signalling in trophoblast cell differentiation

TGFβ regulates trophoblast cell fate via canonical (Smad-dependent) and non-canonical (Smad- independent) signalling pathways. (A) In the Smad-dependent TGFβ pathway, TGFβ ligand binding to its tetrameric receptor complex promotes the phosphorylation and activation of Smad2, which complexes together with Smad4 to collectively translocate into the nucleus and may activate/repress target gene transcription that favor trophoblast cell proliferation over cell fusion. (B) In the Par6 mediated TGFβ pathway, TGFβRII can directly promote the phosphorylation of polarity protein Par6, which recruits Smad ubiquitination regulator factor (Smurf1) to selectively ubiquitinate (u) small GTPase protein RhoA for targeted proteasomal degradation. RhoA degradation results in tight junction dissolution leading to a loss in apical-basolateral cell polarity and promotion of cell motility events, such as trophoblast cell migration (Bose and Wrana, 2006; Ozdamar et al., 2005; Wrana, 2013).

1.4 Preeclampsia

Preeclampsia (PE) is a serious hypertensive disorder that affects 5-8% of all pregnancies, and is associated with increased maternal and perinatal mortality worldwide (American College of et al.,

2013). According to the current American College of Obstetricians and Gynecologists (ACOG) guidelines, a diagnosis of PE is given when the blood pressure of a previously normotensive mother is greater than or equal to 140 mmHg (systolic) and 90mmHg (diastolic) and there is new onset of one of the following events: proteinuria, thrombocytopenia, renal insufficiency, impaired liver function, pulmonary edema or visual disturbances (American College of et al., 2013). Any of these conditions pose a great deal of stress on both the mother and offspring, and due to the lack of treatment for PE, the solution is typically pre-term delivery. Although the clinical symptoms of

PE typically resolve upon delivery, there are often significant long-term health consequences for both mother and offspring. Women who develop PE are at double the risk of developing cardiovascular or cerebrovascular diseases, and three times the risk of chronic hypertension

(Goffin et al., 2018). Offspring of preeclamptic mothers also possess an increased risk for stroke later in life (Kajantie et al., 2009), and show significantly increased blood pressure in childhood and adulthood (Davis et al., 2012). In serious cases, immediate fetal complications include cognitive and physical impairments such as cerebral palsy, epilepsy and blindness. PE is subclassified into two distinct disorders depending on when clinical symptoms manifest during pregnancy: early-onset preeclampsia (E-PE), manifested prior to 34 weeks of gestation; and late- onset (L-PE), manifested after 34 weeks (Raymond and Peterson, 2011). Evidence suggests that the E-PE and L-PE have unique biological profiles and pathogenesis: E-PE originates from the placenta, and exhibits more serious clinical symptoms, whereas L-PE is maternal in origin, and results from an enhanced susceptibility of the maternal endothelium to react to pro-inflammatory factors and elicit an abnormal maternal response (Huppertz, 2008; Redman and Sargent, 2005). In

the present study, PE and E-PE are used interchangeably, unless otherwise stated. The placenta is considered an integral figure in the etiology of PE since in most cases, removal of the placenta is required for symptoms to regress. In pursuit of understanding how the placenta contributes to development of PE, several studies have showed that impaired trophoblast differentiation events during early development are central to the pathogenesis of this disease.

1.4.1 Altered Trophoblast Differentiation in preeclampsia

In PE placentae, extravillous trophoblast cells are arrested in an immature, proliferative phenotype

(Redline and Patterson, 1995), and further exhibit shallow cell migration and invasion into the maternal decidua (Naicker et al., 2003; Robertson et al., 1985). Proliferative cytotrophoblast cells at the base of the anchoring column within the uterine wall retain their original “epithelial” like phenotype, and neglect to undergo the differentiation process which is required to invade the decidua (Fisher, 2015). This phenotype is demonstrated in one early study, where EVT from PE placentae failed to undergo integrin switching to express the invasive 51 integrin that is involved in EVT invasion in normal pregnancy (Zhou et al., 1993). Further, impaired ECM degradation by lowered levels and activity of MMP-9 in PE trophoblasts contributes to defective invasion (Lim et al., 1997). As a result, there is insufficient remodelling of the maternal spiral arteries, leading to a marked reduction in uteroplacental blood flow to the fetus. This reduction in placental blood flow, and increase in uteroplacental vascular resistance, in PE pregnancies can be observed via doppler ultrasound of the uterine arties (Harrington et al., 1996). Higher sensitivity analysis using MRI imaging has since confirmed this reduction in placental perfusion in E-PE placentae compared to normal pregnancies, which did not experience decreases in placental perfusion until later in gestation (Sohlberg et al., 2014). Reduced placental perfusion into the intervillous space ultimately leads to persistent uteroplacental hypoxia, a defining feature of PE (Soleymanlou et al., 2005).

Work from our lab implicated chronic hypoxia as a trademark of PE through microarray analyses, which revealed gene expression profiles of placental villous explants exposed to 3% O2, placentae from high-altitude residence, and PE placentae were strikingly similar (Soleymanlou et al., 2005).

In turn, this resultant uteroplacental hypoxia can further exacerbate impaired trophoblast invasion and contribute to PE pathogenesis (Caniggia and Winter, 2002; Roberts and Cooper, 2001).

To date, it is still not entirely clear what the molecular aberration is that leads to this impaired trophoblast differentiation and invasion seen in PE and thus investigations aimed at understanding this are ongoing. However, studies from our lab have attributed elevated TGFβ3 signalling as a major contributing factor in the pathogenesis of PE (Caniggia et al., 1999; Caniggia et al., 2000).

Mechanistically, elevated levels of HIF-1 maintain high TGFβ3 expression and thereby arrest trophoblast cells in a proliferative, non-invasive phenotype (Caniggia et al., 2000). The role of

TGFβ3 in maintaining trophoblasts in a proliferative, non-invasive state was elucidated by inhibition of TGFβ3 in PE placental explants restoring their invasive capability (Caniggia et al.,

1999). More recently, studies from our lab have demonstrated elevated pSMAD2 levels in PE placentae compared to gestational age-matched controls, particularly in the extravillous trophoblast cell population, providing further evidence that active Smad-dependent TGFβ signalling is likely contributing to the proliferative PE phenotype (Xu et al., 2016). Smad- independent TGFβ signalling via Par6 polarity protein was also elevated in PE placentae as indicated by an increase in Par6/Smurf1 association (Xu et al., 2016). This suggested that the non- invasive phenotype of PE could in part be due to Smad-independent TGFβ signalling maintaining trophoblast cells in an intermediate migratory phenotype that does not reach invasive potential.

1.4.2 Impairments in oxygen sensing in preeclampsia

Upstream of the aberrant TGFβ signalling seen in PE are notable disruptions in placental oxygen sensing mechanisms. This has been regarded as the underlying mechanism for the increased HIF-

1 and associated signalling that typifies PE (Rolfo et al., 2010). Canonical oxygen sensors implicated in regulation of HIF-1 in the human placenta include prolyl hydroxylase domain proteins (PHD1, PHD2, PHD3) and factor inhibiting HIF (FIH), both of which regulate HIF-1 on the basis of oxygen availability (Semenza, 2004). In normoxic conditions, PHD2 regulates HIF-

1 protein stability by hydroxylating HIF-1 at proline 402 and 564 residues, which targets it for proteasomal degradation via von Hippel Lindau tumour suppressor protein, a component of the E3 ubiquitin ligase complex (Maxwell et al., 1999). In hypoxia, PHD activity is inhibited by lack of molecular oxygen, therefore HIF-1 protein remains stable (Semenza, 2004). All three PHDs exhibit HIF-1 hydroxylation capabilities; however, PHD2 is regarded as the primary regulator of

HIF-1 prolyl hydroxylation, while PHD1 is less involved in this process, and PHD3 is a HIF-1 transcription target and involved in a feedback loop of autoregulation (Appelhoff et al., 2004;

Aprelikova et al., 2004). In parallel, FIH proteins regulate HIF-1 at the level of its transcriptional activity. In normoxic conditions, FIH prevents the association of HIF-1 and other transcription co-activators which repress HIF-1 transcriptional activation whereas in hypoxia, FIH is functionally inactive and thus HIF-1 is free to translocate into the nucleus and induce gene transcription (Lando et al., 2002). In PE, dysregulation of PHD1/2 and FIH is demonstrated by a reduction in their placental protein levels, and by decreased HIF-1 hydroxylation by PHD2 at target prolines (Rolfo et al., 2010). Furthermore, placental explants from PE pregnancies cultured in low oxygen (3%) showed no changes in PHD mRNA expression, yet explants from pre-term control placentae experienced significant increases following low oxygen exposure (Rolfo et al.,

2010). This revealed that in PE, placentae fail to sense changes in oxygenation, contributing to the overall stabilization and persistence of HIF-1. The molecular and phenotypic characteristics of

PE discussed in this section are depicted in Figure 1.5. In addition to the mentioned canonical oxygen sensors, ongoing work has revealed a novel group of oxygen sensors and mediators of hypoxic gene expression: the family of Jumonji C domain containing proteins.

1.4.3 JMJD6: a novel oxygen sensor and regulator in the placenta In the human placenta, work from our lab has discovered a role for Jumonji C domain containing protein 6 (JMJD6), a novel oxygen sensor belonging to a superfamily of JmjC domain containing proteins that, similarly to PHDs and FIH, require molecular O2, -keto-glutarate, and ferrous iron

(Fe2+) to execute their enzymatic functions, despite being paradoxically upregulated in conditions of hypoxia (Alahari et al., 2015; Beyer et al., 2008). In particular, studies have shown that JMJD6 uniquely functions as a histone arginine demethylase (Chang et al., 2007), as well as a lysyl hydroxylase (Mantri et al., 2011; Webby et al., 2009). The latter marked a unique function for

JMJD6 as a specialized lysyl hydroxylase, in addition to the only other known lysyl hydroxylases, the procollagen-lysine 5-dioxygenase (PLOD1). Considering this, studies have only recently started to uncover the novel substrates for JMJD6-mediated lysine hydroxylation and implicating its role in distinct biological and pathological processes (Unoki et al., 2013; Wang et al., 2014).

In the human placenta, JMJD6 was found to be a key regulator of HIF-1 signalling. Specifically,

JMJD6 indirectly regulates HIF-1 transcription factor stability by regulating the expression and stability of von Hippel-Lindau (VHL), a protein critical to the proteasomal degradation of HIF-1.

JMJD6 was also found to directly regulate VHL protein stability via oxygen-dependent lysyl hydroxylation, and this in turn promoted HIF-1 proteasomal degradation (Alahari et al., 2015).

More recently, Alahari et al revealed that JMJD6 regulates VHL gene expression also by targeted

histone arginine demethylation (Alahari et al., 2018). However, in PE, the persistence of low oxygen renders the JMJD6 enzyme compromised, and as a result, lysyl hydroxylation and histone demethylation of VHL is impaired and HIF-1 levels remain high due to its impaired degradation

(Alahari et al., 2015; Alahari et al., 2018).

There are a number of other features that make JMJD6 an oxygen sensor of particular interest.

Firstly, it is known to be essential for cellular differentiation and normal embryonic development, as Jmjd6-/- mice exhibit a plethora of defects throughout various stages in gestation, namely; embryonic growth retardation at E12.5, stunted differentiation of lungs, kidneys and intestines and complete peri-natal lethality as a result of cardiac malformations (Bose et al., 2004). In the field of cancer, JMJD6 has been found to exhibit oncogenic properties and has been implicated as a biomarker for tumorigenesis. This is shown in breast cancer studies where JMJD6 promoted the transcription of genes involved in cell proliferation (Lee et al., 2012), as well as regulated cell migration and invasion (Poulard et al., 2015), and in colon cancer studies, where JMJD6 inhibits p53-mediated suppression of cell proliferation by repressing its transcriptional activity by lysyl hydroxylation (Wang et al., 2014). A role for JMJD6 in angiogenesis has also been established, where studies in endothelial cells revealed JMJD6 regulates the splicing of VEGF-receptor 1 (Flt1) that is involved in regulation of angiogenic sprouting (Boeckel et al., 2011). Finally, a novel localization and function of JMJD6 has been revealed in the ECM, where it is secreted as a soluble protein and functionally interacts with ECM component collagen type 1 (Coll-I) (Miotti et al.,

2017). Furthermore, this study showed JMJD6 to interfere with cell adhesion of breast cancer cells to Coll-I, a prerequisite action in epithelial cell migration. Overall, the dependence of JMJD6 on oxygen and its established role in the migration and invasion in cancer highlight the prospective capabilities of JMJD6 as a regulator of trophoblast differentiation in the human placenta.

Preeclampsia (PE)

Chronic placental hypoxia Preeclamptic EVT A

Impaired oxygen sensing

B ↓ EVT cell ↓ EVT ↑ Trophoblast migration invasion cell proliferation

Impaired spiral artery remodeling

↓ Uteroplacental vascularization

Figure 1.5 Molecular and phenotypic characteristics of preeclampsia

Chronic hypoxia is a key feature of preeclampsia (PE) (Soleymanlou et al., 2005), resulting in (A) impaired molecular signalling pathways and (B) altered trophoblast cell phenotype. Persistent hypoxia directly upregulates levels of HIF-1 and also reduces enzymatic activity of JMJD6 oxygen sensor, as JMJD6 requires molecular oxygen to elicit its lysyl hydroxylation (OH) or histone arginine demethylation enzymatic functions. Considering JMJD6 negatively regulates

HIF-1, impaired JMJD6 activity by hypoxia further contributes a rise in HIF-1 levels (Alahari et al., 2015; Alahari et al., 2018). Additionally, increased HIF-1 leads to increased levels of transforming growth factor beta-3 (TGFβ3) (Caniggia et al., 2000). Through Smad-dependent TGFβ signalling, elevated TGFβ3 levels contributes to impaired trophoblast differentiation by arresting trophoblast cells in a proliferative, non-migratory/invasive phenotype (Caniggia et al., 1999). Resultantly, the anchoring villi column does not effectively invade the maternal decidua, and there is poor spiral artery remodeling and reduced uteroplacental vascularization.

1.5 Rationale, Hypothesis and Objectives

Loss of cell polarity orchestrates cell migration, and AMOT appears to be an integral part of this process. Considering the requirement of proper trophoblast cell migration and invasion during placenta development, it is intuitive that cell polarity changes are impactful on the differentiation of these cells. However, aside from polarity protein Par6, no studies have investigated the role of cell polarity regulators in downstream trophoblast differentiation, or the impact of TGFβ on their functionality. Studies in mouse and zebrafish highlight AMOT’s importance in development and embryogenesis; however, the focus of these studies has remained primarily on the embryo/fetus.

A recent study has identified AMOT as a key regulator in the mammalian pre-implantation embryo for its specific role in cell-lineage specification (delineating inner cell mass and trophectoderm).

Within the inner cell mass, AMOT maintained pluripotency and prevented inappropriate cell fate by sequestering YAP from the nucleus and preventing expression of trophectoderm markers

(Leung and Zernicka-Goetz, 2013). Contrastingly, in the polarized trophectoderm cells, AMOT was sequestered to the apical domain, where it was hypothesized to bind to F-actin and/polarity factors that shield its ability to bind to YAP, which is then able to localize to the cell nucleus and promote transcription of genes promoting trophectoderm differentiation. Although focused on the very early stages of embryogenesis, this study reiterates how cell polarity and cell scaffolding, two features of AMOT, are essential for cellular differentiation. To our knowledge, there have been no investigations into the role of AMOT in downstream trophoblast differentiation in the human placenta, or in preeclampsia. This underscores the importance of deciphering how AMOT contributes to the proper development and functioning of the human placenta.

I hypothesized that during placental development, AMOT is regulated by TGFβ signalling pathway and upstream oxygen tension to control trophoblast cell polarity and orchestrate trophoblast cell migration.

Hence, my objectives were to:

1. Characterize the spatial and temporal distribution of AMOT in the developing placenta

2. Determine the involvement of TGFβ signalling, and oxygen, to AMOT expression,

localization and function in trophoblast cells

3. Establish the expression and localization of AMOT in preeclampsia

Figure 1.6 Model of Hypothesis

Chapter 2

Materials and Methods

Materials and Methods 2.1 Human Placenta Tissue Collection

Placentae from early gestation (5-16 weeks, n= 49) were obtained following elective termination of pregnancy by suction evacuation, dilatation or curettage from the Toronto Morgentaler Clinic.

Date of last menstruation and ultrasound measurements were used to determine age of gestation.

Pre-term (n= 15), and early-onset preeclamptic (n= 19) human placenta tissue was obtained through The Research Centre for Women’s and Infant’s Health (RCWIH) BioBank at Mount Sinai

Hospital in Toronto. Informed consent was given by all participants according to University of

Toronto Faculty of Medicine Ethics guidelines, and the Research Ethics Board of Mount Sinai

Hospital (Toronto, ON, Canada) prior to tissue collection. To ensure diversity in the sampling, different areas of the placenta (central and peripheral) were obtained, and tissue with any indications of calcification, ischemia or necrosis were excluded. Selection of PE placenta was made using the guidelines from the American College of Obstetricians and Gynecologists

(ACOG). Patients with chronic hypertension, diabetes, infection and kidney disease were excluded. Definition of early-onset PE included delivery prior to 34 weeks of gestation.

Normotensive, age matched placenta from pre-term (PTC) exhibiting no signs of disease were used as control. Following collection, placenta samples were either snap frozen for RNA and protein analysis or fixed in paraformaldehyde (PFA) for subsequent histological and immunohistochemical or immunofluorescence imaging and analyses. Clinical features of PE and

E-PE are summarized in Table 2.1.

39 40

Table 2.1 Clinical features of patient population

Clinical Parameter PTC E-PE p-value

n= 15 n= 19

Gestational Age at 30.2  0.9 29.0  0.6 ns Delivery (weeks)

Fetal Birth Weight 1558.9  340.2 1093.6  128.0 * p<0.05 (grams)

Fetal Sex 42.85% F 40% F ns

57.14% M 60% M

Blood Pressure S: 107.2  3.6 S: 159.6  5.5 *** S: p<0.001 (mm Hg, S/D) D: 66.6 2.3 D: 100  3.8 * D: p<0.05

Proteinuria Absent 3.7  0.2 n/a (grams/day)

Mode of Delivery 62.5% VD, 25% VD, ns

(%) 37.5% CS 75% CS

Exact p-values: Gestational age: p=0.460; Fetal birth weight: p=0.022; Fetal sex: p=0.899; Blood pressure systolic: p=0.0002; Blood pressure diastolic: p=0.010; Mode of delivery: p=0.073. ns=non-significant

Data are presented as mean ± standard error of the mean. Statistical significance was determined as *p<0.05 or ***p<0.001. A non-parametric Mann-Whitney test was used when comparing gestational age at delivery, fetal birth weight and blood pressure in E-PE to PTC, and a chi-square test was used when comparing fetal sex and mode of delivery in E-PE and PTC.

PTC = Preterm Controls, E-PE = Early-Onset Preeclampsia, F = Female, M = Male, S = Systolic, D = Diastolic, CS = Caesarian Section, VD = Vaginal delivery

2.2 JEG3 Human Choriocarcinoma Cell Culture

JEG3 human choriocarcinoma cells (ATCC, Manassas, VA, USA) were cultured in 75 cm2 flasks containing Eagle’s minimal essential medium (EMEM). Cells were maintained at 37 C under standard culture conditions (5% carbon dioxide, 21% oxygen). EMEM was supplemented with

10% fetal bovine serum (FBS) and 1% of penicillin-streptomycin (10,000 units/mL). Once JEG3 cells reached 100% confluency, the cells were rinsed with 10 mL PBS, trypsinized with 0.25%

Trypsin-EDTA (Thermo Fisher Scientific, Waltham, MA, USA) and passaged forward at a 1:5 dilution in EMEM in a new 75 cm2 flask. Cells were seeded at appropriate cell densities corresponding to the size of culture plate and experiment. Cells were stained with trypan blue dye

(Invitrogen, Carlsbad, CA, USA) and counted using a hematocytometer. For 35 mm 6-well plates

(Corning Inc. Corning, NY, USA), JEG3 cells were seeded overnight at 2.0 x 105 cells/well before respective treatments. For 0.7 cm2 8-well chamber slides (Thermo Scientific™ Nunc™ Lab-Tek™

II, Waltham, MA, USA) JEG3 cells were seeded overnight at 1.5 x 104 cells/well before respective treatments. For 1.0 cm2 8-well -slide (ibidi Inc, Fitchburg, WI, USA) cells were seeded overnight at 9.0 x 104 cells/well before respective treatments and live-cell imaging.

JEG3 cells are a trophoblast derived choriocarcinoma cell line that shares key phenotypic characteristics with villous and extravillous trophoblast cells. Given their phenotype, they are a sufficient model to study events linked with proliferation, migration and invasion (Hannan et al.,

2010; King et al., 2000; Shiverick et al., 2001; Sullivan, 2004). Further, JEG3 cells display a stable phenotype and can be maintained and passed in culture, and are highly amenable to transection.

These mentioned features make JEG3 cells an advantageous in vitro model for this study.

However, it is acknowledged that JEG3 are derived from human choriocarcinoma, thus their cancerous and tumorigenic properties may impact the molecular cellular machinery of these cells.

2.3 In vitro treatments in JEG3 cells

2.3.1 Transforming Growth Factor- (TGF) Treatment

Transforming growth factor 1 and 3 (TGF1 and TGF3) (R&D Systems, Minneapolis, MN,

USA) were reconstituted in 4 mM HCl containing 1mg/ml bovine serum albumin (BSA) for a final stock concentration of 20 g/mL and stored in -20 °C. Once JEG3 cells reached 70% confluency in 6-well plate, 1 L of stock TGF1 or TGF3, or 1L of 4 mM HCl + 1mg/mL BSA (vehicle), was added to 2 mL EMEM for a final concentration of 10 ng/mL TGF. After 24 hours, cells were either collected for protein extraction by adding 60 L RIPA lysis buffer containing protease inhibitor and scraping the cells, or fixed for immunofluorescence staining in 6-well plates by adding 4% formaldehyde for 15 minutes at room temperature.

2.3.2 SB-431542 Treatment in JEG3 cells

SB-431542, inhibitor of TGFR1/ alk5 receptor and downstream Smad-signalling (Sigma-Aldrich

Corporation, St. Louis, MO, USA), was reconstituted in Dimethyl sulfoxide (DMSO) for a final stock concentration of 13 mM. Once cells reached 70% confluency in 6-well plate, SB-431542 were added to 2 mL EMEM for a final concentration of 5 M. For SB-431542 + TGF treatments,

SB-431542 inhibitor was added for 30 minutes prior to TGF ligand treatment to ensure sufficient time for inhibition. After 24 hours, cells were either collected for protein extraction by adding 60

L RIPA lysis buffer containing protease inhibitor (referred to as RIPA +) and scraping the cells, or fixed for immunofluorescence staining in 6-well plates by adding 4% formaldehyde for 15 minutes at room temperature.

2.3.3 Minoxidil Treatment

Minoxidil, an inhibitor of lysyl hydroxylase activity (Sigma-Aldrich Corporation, St. Louis, MO,

USA), was reconstituted in 95% (v/v) ethanol for a final stock concentration of 1 mM. Once cells reached 70% confluency in a 6-well plate, 40 L of stock minoxidil, and 40 L of 95% (v/v) ethanol (vehicle), were added to 2 mL EMEM for final concentration of 20 M. After 48 hours, cells were collected for protein extraction by adding 60L RIPA lysis buffer containing protease inhibitor and scraping the cells, or fixed for immunofluorescence staining by adding 4% formaldehyde for 15 minutes at room temperature.

2.3.4 Low Oxygen (3% O2) Treatment in JEG3 cells

JEG3 cells were grown to 70% confluency in their normoxic conditions of 21% O2, prior to low oxygen exposure in which JEG3 cells were incubated in 3% O2 (5% CO2, 92% N2) for 24 hours.

After 24 hours, cells were either collected for protein extraction by adding 60 L RIPA lysis buffer containing protease inhibitor and scraping the cells, or fixed for immunofluorescence staining by adding 4% formaldehyde for 15 minutes at room temperature.

2.4 Plasmid DNA Constructs for Overexpression Studies

The following plasmids were obtained from Addgene (Cambridge, MA, USA): AMOT 130 (HA-

AMOT p130, Addgene plasmid #32821), AMOT 130PDZ (HA-AMOT p130 deltaEYLI,

Addgene plasmid #32826), and AMOT 130 coiled-coil (HA-AMOT p130 deltaCC, Addgene plasmid #32827) (Zhao et al., 2011). AMOT 130PDZ was generated by deletion of the last 4 amino acids (EYLI) at the c-terminal end of AMOT p130 where the PDZ binding domain is located. AMOT 130 coiled-coil was generated by deletion of amino acid residues at position 410 through 751 in the coiled-coil region of AMOT 130. These plasmids contained a pcDNA3-HA

vector backbone and bacterial resistance against ampicillin. Plasmids were sent in DH5alpha E. coli bacteria contained in agar stabs. Bacteria were streaked and grown overnight at 37 C on

Luria’s broth (LB) agar plates containing ampicillin antibiotic. Selected colonies were inoculated overnight at 37 C in 5 mL LB broth containing 100 g/mL of ampicillin. QIAGEN Plasmid

Maxi Kit was used to purify plasmid DNA according to manufacturer’s instructions (Qiagen,

Hilden, Germany). Quantification of plasmid DNA yield was analyzed using Nanodrop 1000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA).

The following plasmids were obtained through OpenFreezer Laboratory Reagent Tracking and

Workflow Management System, a reagent repository at Mount Sinai Hospital (Toronto, ON,

Canada): AMOT 80 (Angiomotin/Kiaa1071/FL, pLP Triple Flag SD, ID #V988-Pawson Lab)

AMOT 80-YFP (Angiomotin FL, pLP EYFP C1, ID #V1489-Clontech) and AMOT 80 PDZ

(Angiomotin/Kiaa1071/-YLI, pLP Triple Flag SD, ID #V1145-Pawson Lab) (Wells et al., 2006).

AMOT 80 PDZ was generated by deletion of the last 3 amino acids (YLI) in the c-terminal region of AMOT 80 where the PDZ binding domain is located. These three plasmids contained bacterial resistance against kanamycin/neomycin, and as such were prepared and purified as outlined above, except using 100g/mL of kanamycin antibiotic. pcDNA3 (c-Flag pcDNA3, Addgene plasmid #

20011) was used as empty vector control for all AMOT plasmids (Sanjabi et al., 2005).

JMJD6 plasmid (p6352 MSCV-CMV-Flag-HA-JMJD6, Addgene plasmid #31358) (Rahman et al., 2011), Par6 plasmid (pCMV5 T&-Par6 WT T7 tagged, Addgene plasmid #24649) (Ozdamar et al., 2005) were both obtained from Addgene. MSCV PIG (Puro IRES GFP, Addgene plasmid

#18751) served as the empty vector control for these plasmids and was also obtained from

Addgene.

2.4.1 Plasmid DNA Transfection

JEG3 cells were grown to 60-80% confluency in EMEM prior to plasmid DNA transfection using jetPRIME transfection protocol (Polyplus transfection, Illkirch, France). For transfections in

6-well plates (Corning Inc. Corning, NY, USA), one g of plasmid DNA (AMOT 130, AMOT

130PDZ, AMOT 130coiled-coil, AMOT 80, AMOT 80-YFP, AMOT 80PDZ, Par6, JMJD6 or empty vectors) was added to 200L of jetPRIME buffer, vortexed for 10 seconds and centrifuged. Next, 4L of jetPRIME reagent were added to the mixture, vortexed for 10 seconds and centrifuged. The transfection mixture was incubated at room temperature for 10 minutes and before addition to EMEM media of JEG3 cells, containing 10% FBS and 1% pen/strep. Media was replaced 6 hours following transfection. After 24-48 hours, transfected cells were collected for protein extraction by adding 60 L RIPA lysis buffer containing protease inhibitor and scraping the cells, or fixed for immunofluorescence staining by adding 4% formaldehyde for 15 minutes at room temperature.

In 8-well chamber slides used for proximity ligation assay (Thermo Scientific™ Nunc™ Lab-

Tek™ II, Waltham, MA, USA), and in 8-well ibidi -slide used for time-lapse imaging in live cells (ibidi Inc, Fitchburg, WI, USA), 0.5 g of plasmid DNA was added to 25 L of jetPRIME buffer and 1L of jetPRIME reagent. Aside from these volume adjustments, the transfection protocol remained identical.

2.5 siRNA Transfections

JEG3 cells were grown to 50-60% confluency in EMEM prior to addition of siRNA using the jetPRIME transfection protocol (Polyplus transfection, Illkirch, France). 30 nM of Silencer select siRNA targeted against JMJD6, or scrambled siRNA sequences (control) were added to 200

L of jetPRIME buffer, vortexed for 10 seconds and centrifuged. Next, 4 L of jetPRIME reagent were added to the mixture, vortexed for 10 seconds and centrifuged. The transfection mixture was incubated at room temperature for 10 minutes and before addition to EMEM media of JEG3 cells. After 24-48 hours, transfected cells were collected for protein extraction by adding

60 L RIPA lysis buffer containing protease inhibitor and scraping the cells.

2.6 Wound Healing Assay

JEG3 cells plated on coverslips in 6-well plates, or 8-well ibidi -slides, were grown until confluency. Next, a sterile 200L pipet tip was used to make a linear scratch that spanned the length of the coverslip or slide. Fresh tips were used in each well to ensure optimal wounds.

Following initiation of the wound, cells were gently rinsed 2X with PBS and 2X with EMEM to remove cell debris, and fresh 2 mL of EMEM media were added each well. In 6-well plates, JEG3 cells were fixed for immunofluorescence immediately following wound initiation (0 hours), as well as 3 hours and 24 hours after the scratch by addition of 3.7% formaldehyde for 15 minutes at room temperature. In 8-well ibidi -slides, cells were placed under the time-lapse microscope immediately following wound initiation and imaged for 24 hours. In the case of TGF1/3 treatment during wound healing assay, 10 ng/mL TGF1/3 were added to fresh media following scratch/rinse steps. AMOT 80-YFP was transfected 24-48 hours prior to establish fluorescence and confluency prior to wound initiation. Transfection efficiency for AMOT 80 YFP was 70%, as established by counting YFP positive cells vs non-YFP cells.

2.7 Time-Lapse Live Cell Imaging

Following overexpression of the AMOT 80-YFP construct, or pcDNA3 empty vector control, and subsequent wound initiation (as described above), JEG3 cells seeded on 8-well ibidi -slides were

placed in a temperature and CO2 regulated incubation chamber (37 °C, 5% CO2, 21% O2) and inserted into the microscope stage. Images were acquired from 3 separate wells with a 10x or 20x air immersion Leica objective, and captured every 15 minutes for 24 hours. Linear migration rates for AMOT-80 YFP overexpressing JEG3 cells were determined by measuring the area of the scratch at various time points (0 hr, 4 hrs, 8 hrs, 12 hrs, 16 hrs) using Volocity Imaging software and plotting the area values on a scatter plot in Microsoft Excel. A trendline was applied to the scatter plot, and the slope of the line was used as the linear migration rate. Three independent scratches from three different wells was assessed in three replicate experiments. Migration rates from overexpression AMOT 80-YFP JEG3 cells were compared to migration rates of JEG3 cells transfected with pcDNA3 empty vector control and expressed as a fold change.

2.8 Western Blot Analysis

To obtain placental tissue lysates, frozen placenta samples stored in -80 oC freezer were broken into smaller, more manageable chunks, and crushed in liquid nitrogen with a mortar and pestle to generate a fine powder. As outlined in section 2.1, multiple sampling of central and peripheral regions was performed to account for tissue heterogeneity in the placenta. Tissue powder was dissolved in 0.2-1mL RIPA plus proteasome inhibitor, depending on amount of placenta crushed, and homogenized (Ultra-Turrax T25 basic, IKA, Wilmington, NC, USA) 3 x 30 seconds, and stored at -80 oC until use. For JEG3 cell lysates, 65 L of RIPA plus proteasome inhibitor was used to collect cells off of 6-well culture plates with a 25 cm cell scraper (Sarstedt. Inc, Newton,

NC, USA) and transferred to 1.5 mL Eppendorf tube to be vortexed and subject to one freeze-thaw step. Both tissue and cell lysates were centrifuged at 16, 3000g for 10 mins. The supernatant containing protein was transferred to a fresh tube for further steps and the remaining pellet containing cell debris was discarded.

Protein concentration of tissue and cell lysates was quantified using a colorimetric Bradford protein assay (Bio-Rad, Mississauga, Ontario). Volume of lysate required for 30-50 µg of protein was calculated and added to 8µL of 4X sodium dodecyl sulfate (SDS) sample buffer containing

250 mM Tris-HCl (pH 6.8), 40% (v/v) glycerol, 8% (w/v) SDS, 0.2% (w/v) bromophenol blue and

20% (v/v) -mercaptoethanol. RIPA+ proteasome inhibitor buffer was also added to the sample mixture to maintain equal volumes amongst samples. Samples were vortexed for 10 seconds, heated at 95°C for 5 minutes, and centrifuged. Samples were then loaded and run on 6%, 7.5%,

10% or 12% sodium dodecyl polyacrylamide gel (SDS-PAGE) in running buffer (25 mM Tris base, 192 mM glycine, 0.1% SDS), ensuring sufficient separation at target protein molecular weight. Gels were then transferred onto methanol-activated polyvinylidene fluoride (PVDF) membrane (Trans-Blot Turbo Mini size PVDF, BioRad) in ice-cold transfer buffer (25 mM

Tris base, 192 mM glycine, 20% methanol), or using Turbo BioRad Trans-Blot Turbo System.

Membranes were blocked with 5% powdered non-fat milk dissolved in Tris-buffered saline containing 0.1% (v/v) Tween 20 (TBST) for 1 hour to reduce unspecific binding of antibodies.

Following blocking, membranes were incubated overnight with the appropriate primary antibody diluted in 5% non-fat milk in a 4 C cold room with gentle rocking. The next day, membranes were washed 3x10 mins in TBST, and horseradish peroxidase (HRP)-conjugated secondary antibody corresponding to the species of the primary antibody was diluted in 5% non-fat milk and added to the membrane for 1 hour at room temperature with gentle rocking. Membranes were washed again

3x10 mins in TBST before visualizing immunoreactive bands by addition of chemiluminescence

ECL-plus reagent (Western LightningTM Chemiluminescence Reagent Plus, Perkin Elmer,

Waltham, Massachusetts) and exposure to film (GE Healthcare, Amersham, Hyperfilm ECL).

Western blots were scanned using a Epson Perfection V39 scanner (Epson, Suwa, Japan), and densiometric analysis using Image J software.

Membranes were stripped with stripping buffer (0.2M glycine, ddHH2O, pH2.2) for 40 minutes, and blocked in 5% non-fat milk dissolved in TBST for 1 hour before overnight incubation with loading control primary antibodies, -actin or -Tubulin. Membranes were imaged using the aforementioned protocol. Direct band visualization of entire lanes following staining of membranes with 0.1% Ponceau S solution was also used as a loading control, particularly in cases where analysis of higher molecular weight proteins forces lower molecular weight proteins to run off, or when the experimental condition affected levels of -actin and -Tubulin cytoskeletal proteins.

2.9 Antibodies

Primary antibodies used include: AMOT (Santa Cruz Biotechnology, Santa Cruz, CA, USA, L-

16 sc-82493, goat polyclonal [IF 1:1000]), AMOT (Thermo Scientific, Rockford, IL, USA, PA5-

31196, rabbit polyclonal [WB 1:1000]), AMOT (Cell Signalling, Danvers, MA, USA, #431305

D204H, rabbit polyclonal [IF/PLA 1:100]), -tubulin (Santa Cruz Biotechnology, P16 sc-31779, goat polyclonal [WB 1:500]), -actin (Santa Cruz Biotechnology, C-4 sc-47778, mouse monoclonal [WB 1:2000]), JMJD6 (Abcam Inc, Cambridge, MA, USA, #ab64575, goat polyclonal [WB 1:500]), Par6A (Santa Cruz Biotechnology, C-3 sc-365323, mouse monoclonal

[WB 1:500, IF/PLA: 1:100]), Par6A (Aviva Systems Biology, San Diego, CA, USA

ARP42883_P050, rabbit polyclonal [WB 1:600]), pSMAD2 (Abcam Inc, #ab53100, rabbit polyclonal [WB 1:800]), RhoA (Abcam Inc, #ab54835, mouse monoclonal [WB 1:500, IF 1:200]),

SMAD2 (Cell Signalling, #3122 86F7, rabbit monoclonal [WB 1:1000]), TAZ (Santa Cruz

Biotechnology, 1F1 sc-293183, mouse monoclonal [IF 1:300]), ZO-1 (Santa Cruz Biotechnology,

H-300 sc-10804, rabbit polyclonal [IF 1:200]). High affinity phalloidin probe conjugated to photostable, red fluorescent Alexa Flour 594 dye was used to selectively stain against the F-actin cytoskeleton. Of note, AMOT L-16 antibody validation for IHC was performed by a predecessor in the lab using AMOT competing peptide.

Secondary antibodies used in WB analyses were obtained from Santa Cruz Biotechnology and include horseradish peroxidase (HRP) conjugated rabbit anti-mouse (1:2000), HRP goat anti- rabbit (1:1000), HRP donkey anti-goat (1:1000). Secondary antibodies used in IF analyses were obtained from Thermo Scientific, used at a dilution of 1:300 and include: Alexa Fluor® 488 donkey anti-rabbit (A21206), Alexa Fluor® 594 donkey anti-rabbit (A-21207), Alexa Fluor® 488 donkey anti-goat (A-11055), Alexa Fluor® 594 donkey anti-goat (A-11058), Alexa Fluor® 488 donkey anti-mouse (A-21202), and Alexa Fluor® 594 donkey anti-mouse (A-21203).

2.10 Immunoprecipitation (IP)

Extracted protein from JEG3 cell or tissue lysates as described for western blot analysis was used to prepare IP samples. In IP samples, 300 µg of extracted protein were diluted in RIPA+ buffer to generate a final protein concentration of 1 µg/µL. To block nonspecific binding, IP samples were pre-cleared by adding 20 µL of Protein A or G agarose bead slurry (Santa Cruz, CA, USA) for 1 hour at 4 ºC on an orbital rocker. Two µg of appropriate primary antibody or IgG control were added to pre-cleared IP samples and incubated overnight at 4 ºC on an orbital rocker. The next day, 30 µL of Protein A or G agarose bead slurry were added to IP samples for 2-3 hours at 4 ºC on an orbital rocker. IP samples were centrifuged, supernatant discarded and the resultant protein- bead pellet was washed and re-suspended in 500 µL of RIPA+ buffer. IP samples were centrifuged, supernatant was discarded and the pellet was washed and re-suspended in 1XPBS for two rounds.

After washes, 40 µL of 2X SDS sample buffer (100 mM Tris-HCl (pH 6.8), 4% (w/v) SDS, 0.25%

(w/v) bromophenol blue, 20% (v/v) glycerol, 200 mM -mercaptoethanol) were added to the pellet to elute protein from the beads. The mixture was heated at 95 ºC for 5 minutes and centrifuged to pellet the disassociated beads. The resultant supernatant containing the immunoprecipitated protein of interest was loaded on SDS-PAGE and incubated with target and control antibodies in western blot analysis as previously outlined. For control blots, 15 µL of sample was loaded whereas for target protein blots, 35 µL of sample was loaded.

2.11 Immunohistochemistry (IHC)

Preparation of paraffin embedded placenta tissue sections (~7m thickness) on glass slides were performed either by a lab technician in our lab or by the pathology department at the Centre for

Phenogenomics (Toronto, ON, CAN). Placentae samples were obtained from the RCWIH Biobank at Mount Sinai as described above. Hematoxylin and eosin (H&E) staining was performed on every 10th or 20th section to establish tissue morphology prior to immunostaining. Tissue sections were selected for immunostaining on the basis that the both floating and anchoring placental villi were present, and the tissue was intact. Following this selection process, slide containing placenta sections were heated at 65ºC for 10 minutes prior to a series of deparaffinization and rehydration steps. Heated slides were deparaffinized by incubating 3x5 minutes in 100% xylene. Slides were rehydrated by incubating 3x2 minutes in 100% ethanol, and single incubations for 2 minutes in

95%, 90%, 85%, 80%, 75%, 70% and 50% ethanol. Rehydrated slides were then washed for 2 minutes in ddH2O, and 5 minutes in 1XPBS. Tissue antigen retrieval was conducted by immersing slides in 250L of 10mM sodium citrate buffer (pH 6.0) and heating in an 800-watt microwave for 5 minutes on power 6. The container was cooled for 15 minutes with the lid on, and re-heated for 3 minutes at power 6. The container was then cooled with the lid off until slides reached room temperature (~20 minutes) and then slides were washed 3x5 minutes in 1XPBS on a gentle rocker.

Endogenous peroxidase activity was inhibited by immersing slides in 250 mL of 3% (v/v) hydrogen peroxidase (H2O2) in methanol for 30 minutes at room temperature, and slides were then washed 3x10 minutes in 1XPBS on a gentle rocker. Tissue sections were circled using an

Immunopen (Calbiochem, Millipore Sigma, Burlington, MA, USA), and blocked by adding

100-200 L of 5% normal horse serum (5% NHS) (Sigma®, St. Louis, Missouri) containing 1%

BSA inside the circled tissue area for 1 hour at room temperature. Primary antibody (AMOT) was diluted in equal parts blocking solution (5% NHS) and antibody diluent (0.4% sodium azide,

0.625% gelatin), and 100-200 L was added to tissue sections overnight in 4ºC cold room. Placenta tissue sections on slides were traced using an Immunopen (Calbiochem, Millipore Sigma,

Burlington, MA, USA) which formed a hydrophobic barrier to aid in blocking and antibody incubation steps. For negative controls, normal goat-IgG (sc-2028) was used, corresponding to species and titer of primary antibody (AMOT). Blocking and antibody incubation steps were performed in a moist-humidified chamber to prevent evaporation of solutions or drying out of tissue sections. The following day, tissue sections were incubated with biotinylated secondary antibody diluted 1:200 in blocking solution for 2 hours at room temperature. Slides were washed

3x10 minutes in PBS, prior to incubating sections with an avidin-biotin-HRP complex from

VectaStain Standard Kit (Vector Laboratorie Inc., Burlingame, CA, USA). Slides were washed

2x5 minutes, followed by 1x10 minutes in 1XPBS, and incubated with DAB solution (0.075%

(w/v) 3,3-diaminobenzidine tetraaminobiphenyl (DAB) in 1XPBS solution containing 0.002%

(v/v) H2O2) to create positive brown staining. DAB reaction was stopped once positive brown staining was detected, approximately 2 minutes, by draining off DAB solution droplets, placing slides in container of water and washing slides with running tap water for 5 minutes. Tissue sections were counterstained by immersing slides in hematoxylin for 30 seconds to dye nuclei and washed with running tap water for 10 minutes. Slides were immersed in acid ethanol (70% ethanol

with 250 L HCl) for one second to reduce background and improve contrast and washed with running tap water for 10 minutes. Slides were dehydrated by immersing slides in reverse order in the ethanol-xylene gradient. Finally, coverslips were mounted to slides using Surgipath micromount medium (Leica, Wetzlar, Germany). IHC images were captured using Olympus BX61 motorized light microscope system (Olympus, Shinjuku, Tokyo, Japan).

2.12 Immunofluorescence (IF)

Tissue

Deparaffinization and rehydration steps were conducted as previously described in immunohistochemistry. Tissue antigen retrieval was conducted by immersing slides in 250 L of

10 mM sodium citrate buffer (pH 6.0) and heating in an 800-watt microwave for 5 minutes on power 4. The container was cooled for 15 minutes with the lid on, and re-heated for 3 minutes at power 4. The container was then cooled with the lid off until slides reached room temperature (~20 minutes) and then slides were washed 3x5 minutes in PBS on a gentle rocker. Next, slides were incubated in 0.4% Triton X-100 (Bioshop Canada Inc., Burlington, ON, CAN) for 5 minutes to permeabilize cells. Auto-fluorescence was quenched by incubating slides in a container filled with

0.3% (v/v) Sudan Black in 70% ethanol for 30 minutes (Erben et al., 2016; Qi et al., 2017). Slides were then washed 3x5 minutes in PBS with gentle rocking. Placenta tissue sections on slides were traced using an Immunopen (Calbiochem, Millipore Sigma, Burlington, MA, USA) which formed a hydrophobic barrier to aid in blocking and antibody incubation steps. Tissue sections were blocked by adding 100-200 L of 5% normal horse serum (5% NHS) (Sigma®, St. Louis,

Missouri) inside the circled tissue area for 1 hour at room temperature. Next, primary antibody/antibodies were diluted in equal parts blocking solution (5% NHS) and antibody diluent

(0.4% sodium azide, 0.625% gelatin), and 100-200 L was added to tissue sections overnight in

4ºC cold room. In dual-staining experiments, both primary antibodies were incubated overnight together. For negative controls, normal goat-IgG (sc-2028) or mouse IgG (sc-2025) was used, depending on the primary antibody. For dual staining, IgG controls for both primary antibodies were included. Blocking and antibody incubation steps were performed in moist-humidified chamber to prevent evaporation of solutions or drying out of tissue sections.

The following day, tissue sections were washed 3x10 minutes in PBS and incubated with 100-200

L of Alexa Flour®-conjugated secondary antibody (Invitrogen, Carlsbad, CA, USA) at 1:300 dilution in antibody diluent. From this point onwards, all incubations were performed in the dark using a container wrapped in tin foil. In dual-staining experiments, green Alexa Flour®-conjugated secondary antibody was added first for 1 hour, tissue section was then washed 3x5 minutes with

PBS, and red Alexa Flour®-conjugated secondary antibody was added second for 1 hour. Next, slides were finally washed 3x5 minutes in PBS. Tissue sections were stained with 0.5 g/mL of

DAPI nuclear staining (4’,6-diamino-2-phenylindole) for 5 minutes. Lastly, coverslips were mounted to slides by adding a drop of Immuno-Mount™ solution to tissue sections (Thermo Fisher

Scientific®, Waltham, MA). Fluorescence images were captured using Leica SD6000 spinning disk confocal microscope (Leica Camera, Wetzlar, Germany) and Volocity Imaging software.

Cells

Following appropriate treatments, JEG3 cells were fixed on coverslips in 6-well plates by adding

1mL of 4% formaldehyde for 15 minutes at room temperature, and rinsing 3x in 1XPBS thereafter.

Fixed cells were immediately used, or were stored in PBS in 4ºC cold room for later use. Cells were permeabilized with 0.2% TritonX-100 (Bioshop Canada Inc., Burlington, ON, CAN) for 5

minutes at room temperature, rinsed with PBS and blocked via addition of 100 L of 5% NHS to coverslips for 1 hour at room temperature, ensuring liquid covers entire coverslip surface area.

Next, primary antibody/antibodies were diluted in equal parts blocking solution (5% NHS) and antibody diluent, and 100 L was added to coverslips overnight in 4ºC cold room. For dual- staining experiments, cells were incubated overnight with both antibodies together. The following day, cells were washed 3x10 minutes with PBS, and incubated with 100 L of Alexa Flour®- conjugated secondary antibody (Invitrogen, Carlsbad, CA, USA) at 1:300 dilution in only antibody diluent. From this point forward, all incubations were done in the dark in containers wrapped in tin foil. For dual- staining experiments, green Alexa Flour®-conjugated secondary antibody was added first for 1 hour, cells were washed 3x5 minutes with PBS, and red Alexa Flour®-conjugated secondary antibody were added second for 1 hour. Cells were finally washed 3x5 minutes in

1XPBS. Cells were stained with 0.5 g/mL of DAPI nuclear staining (4’,6-diamino-2- phenylindole) for 5 minutes. Lastly, coverslips with immunostained cells were mounted onto

25x75x1 mm glass slides using Immuno-Mount™ solution (Thermo Fisher Scientific®, Waltham,

MA). Fluorescence images were captured using Leica SD6000 spinning disk confocal microscope

(Leica Camera, Wetzlar, Germany) and Volocity Imaging software. Colocalization between two fluorophores was quantified using Pearson’s correlation coefficient (PCC) through fluorescence analysis in Volocity Imaging software. An automated threshold as described by Costes et al

(Costes et al., 2004) was set to minimize background signals and noise. For each independent experiment (n=3), colocalization was quantified blindly in three regions from three different images for each treatment group. Guidelines for interpreting Pearson’s correlation coefficient describe PCC > 0.5 to have a large correlation 0.3

PCC<0.3 indicate a small correlation.

2.13 Proximity Ligation Assay

JEG3 cells were cultured on 8-well chamber slides (Thermo Scientific™ Nunc™ Lab-Tek™ II,

Waltham, MA, USA) in standard culture conditions and treated with vehicle and 10 ng/mL of

TGF1/3 for 24 hours, or transfected with AMOT plasmid constructs for 24-48 hours. JEG3 cells were rinsed 3x in 1XPBS, fixed with ice-cold methanol and acetone (1:1) for 3 minutes, and permeabilized with 0.2% TritonX-100 for 5 minutes. In the subsequent steps, Duolink PLA

Fluorescence Protocol was followed using Duolink PLA Reagents (Sigma-Aldrich Corporation,

St. Louis, MO, USA). All incubations were performed in a sterile moisture chamber at the appropriate temperature. For blocking, cells were treated with one drop (~40 L) of Duolink blocking solution for 30 minutes at 37 ºC, before incubating cells with AMOT and Par6 antibodies diluted in Duolink antibody diluent overnight at 4 ºC. The following day, cells were washed 2x5 minutes with Duolink Wash Buffer A with gentle rocking at room temperature and incubated with species-specific plus and minus PLA oligonucleotide probes diluted 1:5 in Duolink antibody diluent for 1 hour at 37 ºC. This step promoted hybridization of probes to their species corresponding primary antibody. Next, cells were washed 2x5 minutes with Duolink Wash

Buffer A with gentle rocking at room temperature and incubated with Ligase enzyme diluted 1:5 in 5X ligation buffer containing oligonucleotides and ddH20 for 30 minutes at 37 ºC. This step promoted ligation of oligonucleotide arms of PLA probes in close proximity (<40 nm) to form a closed oligonucleotide circle. Cells were then washed 2x2 minutes in Duolink Wash Buffer A with gentle rocking at room temperature and incubated with polymerase enzyme diluted 1:5 in 5X amplification buffer containing nucleotides and red fluorescently labelled oligonucleotides and ddH20 for 100 minutes at 37 ºC in the dark. This final step promotes the one PLA oligonucleotide probe to act as a primer for rolling circle amplification that generates a concatemeric product in

which fluorescently labelled oligonucleotides hybridize onto. All remaining steps were performed in the dark. Cells were then washed 2x10 minutes with Duolink Wash Buffer B, 1 minute with

0.01X Wash Buffer and mounted with coverslips using in situ mounting medium with DAPI

(Sigma-Aldrich Corporation, St. Louis, MO, USA). Three negative controls were used: PLA performing in the absence of either the AMOT or Par6 antibody, in the absence of plus-PLA probe, and in the absence of both plus and minus-PLA probes. Fluorescence indicating binding of AMOT and Par6 was visualized and captured using Leica SD6000 spinning disk confocal microscope

(Leica Camera, Wetzlar, Germany) and Volocity Imaging software. PLA quantification was performed by two independent assessors who were blinded to the treatment groups.

2.14 RNA Isolation, cDNA conversion and Quantitative-PCR

RNA isolation: Following collection of human placenta tissue samples from early development

(5-15 weeks), as previously described, total RNA was isolated by adding 1mL of TRIzol reagent

(Invitrogen, Carlsbad, CA, USA) to crushed placenta. The tissue/TRIzol mixture was homogenized on ice, 200 µL of chloroform were added, and the sample mixture was shaken vigorously for 15 seconds prior to storing on ice for 5 minutes. Samples were centrifuged at 8900 g for 15 minutes at 4 ºC, supernatant was collected and added to 500 µL of isopropanol, and the mixture was incubated at 20 ºC for 1 hour. Samples were then centrifuged at 8900 g for 15 minutes at 4ºC, supernatant discarded and the resultant pellet was resuspended in 500 µL of ice cold 70% ethanol in Diethyl pyrocarbonate (DEPC)-treated dH2O. The tissue resuspension was vortexed, centrifuged at 3150 g for 10 minutes at 4 ºC, and supernatant was removed to promote air-drying of the RNA pellet. The dried pellet was dissolved in 50 µL of DEPC- dH2O and RNA yield was quantified and assessed for purity using Nanodrop 1000 spectrophotometer. RNA samples were stored in -80 ºC.

cDNA conversion: Isolated RNA was reverse transcribed into cDNA using qScript cDNA

SuperMix following the manufacturer’s protocol (Quanta Biosciences, Beverly, MA, USA). For each reverse transcription reaction, 4 µL of 5X qScript cDNA SuperMix (containing optimized concentrations of MgCl2, dNTPs (A, C, G, T), recombinant RNase inhibitor protein, qScript reverse transcriptase, random primers, oligo (dT) primer and stabilizers) were added to 1 µg of isolated RNA, and the reaction volume was brought to 20 µL with RNase/DNase- free water.

Samples were incubated at 25 ºC for 5 minutes, 42 ºC for 30 minutes, 85 ºC for 5 minutes and held at 4ºC until stored at -20 ºC.

Quantitative polymerase chain reaction (qPCR): PCR amplification of cDNA was conducted using the DNA Engine Opticon® 2 System (MJ Research, Waltham, MA) following addition of 1

µL AMOT probe (Hs00611096_m1 AMOT TaqMan® Assays-on-DemandTM from Applied

Biosystems (ABI), Foster City, CA, USA), 10 µL of TaqMan® Universal PCR Master Mix (ABI) and 6 µL of DEPC H2O to the converted cDNA (3 µL). Ribosomal 18S RNA was used as a normalization gene and all AMOT Ct values were normalized to 18S using comparative Ct method

(Livak and Schmittgen, 2001).

2.15 In vitro JMJD6 Hydroxylation Reaction

Protein sequences for AMOT 130 (NCBI Ref Seq: NP_001106962.1) and AMOT 80 (NCBI Ref

Seq: NP_573572.1) were input into in silico RF-Hydroxysite Prediction software in FASTA format (Ismail et al., 2016), and the probability of lysine hydroxylation of AMOT was assessed.

The following parameters were set in the search: Residue=Lysine (K);Window size=15; Score threshold: 0.6. Under these parameters, one-hundred percent probability of hydroxylation was predicted for a shared lysine residue on AMOT 130 and 80 (position 758 and 349, respectively).

It is noted that several other high probability lysine residues in other amino acids windows were

identified with >90% probability. Using the protein sequence information for AMOT 130/80, a

10-amino acid peptide containing this 100% probability lysine residue and flanking residues was synthesized (CanPeptide Inc, Point-Claire, QC, CA). The sequence of the synthesized AMOT peptide was AQIIEKDAMI, and its molecular weight was determined to be 1131.35 Daltons (Da).

For in vitro enzyme studies, 7.5 g of AMOT peptide was incubated with 100 or 200 ng of purified human recombinant JMJD6 enzyme (BPS Bioscience, San Diego, CA, USA) in the presence of hydroxylation buffer (50mM Tris pH 7.9, 50 mM KCl, 10 mM MgCl2, 2 mM ascorbate, 1 mM -

2+ ketoglutarate, 50 M Fe in dH2O) at 37 ºC for 2 hours. Reactions were performed in a 24-well plate, with a final volume of 100 L. Reactions performed in the absence of JMJD6 enzyme served as a negative control. Following a 2 hour enzymatic reaction, reaction volumes were collected and stored at -20 ºC.

2.16 MALDI-TOF Mass Spectrometry

The mass profile of AMOT peptide incubated in the presence or absence of JMJD6 enzyme (100 and 200 ng) was analyzed at the Analytical Facility for Bioactive Molecules at the Hospital for

Sick Children (Toronto, ON, CAN) using matrix-assisted laser desorption ionization time of flight mass spectrometry (MALDI-TOF MS). Four L of enzyme reaction mixture were mixed with 6

L of 15 mg/mL 9-aminoacridine matrix in methanol (Sigma-Aldrich, St. Louis, MO, USA) to promote isolation of the peptide analyte and prevent its clustering into aggregates. One L of analyte/matrix mixture was spotted on a MALDI plate and allowed to dry at room temperature to create a crystalized bed of analyte/matrix. The MALDI plate containing the crystalized analyte was then exposed to Nd:YAG laser (337 nm) at 3 nanosecond pulse width and 200 Hz firing rate, and the mass of peptide was analyzed by time-of-flight tandem mass spectrometer (SCIEX

TOF/TOFTM 5800 System; SCIEX, Concord, ON, CAN) and TOF/TOF Series Explorer Software

(SCIEX, Concord, ON, CAN). Negative-ion reflector mode was used to acquire the spectra data within the 0.5-2 kDa range. The resultant mass spectra are derived from 200 individual laser shots from different positions on the spotted sample. Viewing and processing of spectra files for the

16OH (hydroxyl) shift was performed using Data Explorer Software (SCIEX).

2.17 Statistical analysis

GraphPad Prism 7 software (San Diego, CA, USA) was used for statistical analysis. For the comparison of two groups, a non-parametric Mann Whitney test was used. For the comparison of three or more groups, pairwise Mann Whitney tests between individual groups was used. Data are presented as means +/- SEM (standard error of the mean). N values are provided in the figure legends for each experiment. Statistical significance was accepted at *p<0.05, **p<0.01 and

***p<0.001.

Chapter 3

Results Results

During human placenta development, trophoblast cell differentiation events, in particular migration, requires tight control of cell polarity and tight junction stability. Alterations in apical- basolateral epithelial cell polarity via reorganization of necessary tight junction proteins can disrupt intracellular tight junctions, and in turn promote cell movement. Angiomotin (AMOT) is one such scaffolding protein that has been implicated in the regulation of cell polarity (Wells et al., 2006). However, the role of scaffolding polarity protein AMOT in trophoblast cells during placental development remains unknown. Hence, I sought to first characterize the temporal expression pattern of AMOT in first-trimester placentae to establish a developmental profile.

Considering the fundamental role for AMOT in intracellular scaffolding, analyzing changes in protein levels alone is not sufficient. Thus, it is imperative to also investigate the spatial localization of AMOT within the placenta tissue. This concept is reiterated throughout this thesis, as a large focus is placed on AMOT subcellular localization in subsequent in vitro experiments.

3.1 AMOT exhibits distinct temporal and spatial expression patterns during human placenta development

I first assessed the temporal expression pattern of both AMOT isoforms in the developing placentae. AMOT 130, the isoform demonstrated to play a role in changes in cell shape through interaction with the F-actin cytoskeleton (Ernkvist et al., 2006), is present in the developing placentae as early as 5-6 weeks of gestation, as assessed by Western blot (WB) analysis (Figure

3.1A). AMOT 80, known for its role primarily in cell migration (Ernkvist et al., 2008), is present in lower amounts during early first trimester but increases at 10 weeks of gestation (Figure 3.1A).

61 62

Densitometry analysis revealed that both AMOT 130 and AMOT 80 protein levels are significantly increased in placentae from 10-15 weeks of gestation compared to 5-9 weeks of gestation (p<0.01 and p<0.001, respectively). Using a quantitative-PCR TaqMan AMOT probe, which does not discriminate between AMOT isoforms due to the probe spanning exons common to both the 80 and 130 kDa AMOT transcripts, total AMOT mRNA levels in the developing placentae were assessed. In line with protein analysis, qPCR findings revealed AMOT mRNA levels in the placentae are significantly increased at 10-15 weeks compared to 5-9 weeks of gestation (p< 0.05)

(Figure 3.1B). The 5-9 and 10-12-week gestational age windows were selected because significant changes in oxygen tension and levels of spiral artery remodeling occur at 10-12 weeks of gestation, and thus I wanted to see if AMOT expression patterns changed at this critical time point. However, the gestational ages of developmental placentae samples are an estimate based upon the date of last menstruation and ultrasound imaging, and could potentially account for the wide distribution of data points in these windows. Interestingly however, the higher data points indicating highest

AMOT expression and lower data points indicating lower AMOT expression corresponded to later and earlier gestation points, respectively.

Utilizing tissue sections from human placentae ranging from 5-13 weeks of gestation, I examined

AMOT spatial distribution in floating villi (Figure 3.2) and anchoring villi (Figure 3.3). The

AMOT antibody stains both AMOT 80 and AMOT 130; thus, the following findings represent the collective localization of both isoforms. In the floating villi at 6 weeks of gestation, AMOT primarily localizes to underlying cytotrophoblasts and to a lesser extent, the overlying syncytiotrophoblasts (Figure 3.2A). At 8 weeks of gestation, AMOT localization to cytotrophoblast is maintained and also appears in the overlying syncytiotrophoblast layer (Figure

3.2B). At 13 weeks of gestation, AMOT localization shifts primarily to the apical villi of the

syncytiotrophoblast layer, and to a lesser extent in the cytotrophoblast cells (Figure 3.2C).

Additionally, AMOT localization to mesenchymal cells within the core of the floating villi is more abundant in 8 and 13-week placenta than 6-week placentae. Examination of the anchoring villi in placentae from 6, 8, and 13 weeks of gestation revealed striking localization of AMOT to the extravillous trophoblast cells (EVTs) (Figure 3.3A-C). In particular, AMOT was found to localize to EVTs within the anchoring column, particularly at cell boundaries. This is unlike the cytoplasmic and nuclear distribution of AMOT observed in cytotrophoblasts and syncytiotrophoblasts of the floating villi. Intriguingly, AMOT localization to EVTs is restricted to the intermediate and distal regions of the anchoring column, and absent in the proximal region.

Furthermore, AMOT staining is stronger in anchoring villi of 13-week placentae compared to both

6 week and 8-week placenta sections. These findings are consistent with WB analysis showing an increase in both AMOT 130 and 80 protein levels in 10-15-week placentae vs 5-9-week placentae.

64 A 6 7 8 9 10 10 12 14 15 AMOT 130 kDa

80 kDa

Ponceau

Weeks of gestation

8 8 *** **

6 6

4 4

2 2 Fold Change FoldChange 0 Change Fold 0 5-9 10-15 5-9 AMOT 80 Protein 80 Protein AMOT AMOT 130 Protein Protein 130 AMOT 10-15 Weeks of gestation Weeks of gestation

4 * B

3 mRNA mRNA 2

Fold Change FoldChange AMOT 0 5-9 10-15 Weeks of gestation Figure 3.1 AMOT protein levels and mRNA expression during early placenta development

(A) Representative Western blot showing developmental profile of AMOT 130 and AMOT 80 protein levels in human placenta tissue from 5-15 weeks of gestation. Densitometry analysis of AMOT 130 and AMOT 80 protein levels normalized to entire lane following Ponceau staining and expressed as fold change relative to 5-9 weeks. AMOT 130 and AMOT 80 protein levels are significantly increased in placentae from 10-15 weeks of gestation vs 5-9 weeks of gestation (5-9 weeks: n=9, 10-15 weeks: n=10). Statistical significance was determined as **p<0.01 or ***p<0.001 using non-parametric Mann-Whitney test. (B) qPCR analysis for AMOT mRNA expression in human placentae tissue. Data are expressed as fold change relative to 5-9 weeks of gestation. qPCR probe does not discern AMOT 80 and 130 transcripts thus represents mRNA expression of both isoforms. Total AMOT mRNA levels are significantly increased in placentae from 10-15 weeks of gestation vs 5- 9 weeks of gestation (5-9 weeks: n=10, 10-15 weeks: n=10). Statistical significance was determined as *p<0.05 using non-parametric Mann-Whitney test.

6 Weeks A 20x IgG control

20x

40x ST CT

8 Weeks 13 Weeks C B 20x 20x

40x 40x ST CT

CT ST

Figure 3.2 Spatial localization of AMOT in floating villi in early placenta development

Representative images illustrating AMOT localization in placenta tissue sections from; (A) 5-6 weeks (B) 8 weeks and (C) 13 weeks of gestation using immunohistochemistry (IHC) staining and imaging with stereology microscope (n=11). Magnification is indicated in the top left corner of the images. Normal Goat IgG was used as a negative control. Scale bars for 10X, 20X and 40X images are 100m, 50m and 20m, respectively. CT (cytotrophoblast) ST (syncytiotrophoblast).

66 6 Weeks A 20x PC IC 20x ST IC DC IgG control 10x CT

40x PC 40x IC/DC 20x

EVT EVT

B 8 Weeks C 13 Weeks 10x10x 10x DC DC

DC IC DC IC PC

20x 20x 20x DC DC

Figure 3.3 Spatial localization of AMOT in anchoring villi in early placenta development

3.2 AMOT is regulated by TGF signalling pathway

Studies from our lab and others have revealed the role that TGF signalling plays in mediating the differentiation events that shape early placenta development in humans (Caniggia et al., 1999;

Caniggia et al., 2000; Lala and Chakraborty, 2003; Xu et al., 2015). Further, our lab has demonstrated a TGF-dependent regulation of polarity protein Par6 in trophoblast cell migration

(Xu et al., 2015). Considering that AMOT is a scaffolding protein involved in cell polarity regulation, as well as its temporal and spatial distribution pattern during early placenta development where TGF signalling is known to be active, I next investigated whether there was a TGF-dependent regulation of AMOT. Before TGF ligand treatment, I first characterized the endogenous subcellular localization of AMOT in JEG3 cells. Despite being choriocarcinoma,

JEG3 cells are widely used in the field of placenta biology to study trophoblast events linked to proliferation, migration and invasion (Rothbauer et al., 2017; Sivasubramaniyam et al., 2013; Xu et al., 2016).

3.2.1 AMOT resides at tight junction, cytoplasm and protruding edge of JEG3 cells

As previously discussed, the cellular localization of AMOT is constantly in flux. AMOT can be found at the plasma membrane, where it is permitted to interact with tight junction residing polarity proteins; however, it has also been located within the nucleus, and in the cytoplasm where it is able to interact with the F-actin cytoskeleton (Ernkvist et al., 2006; Ernkvist et al., 2008). To first assess endogenous AMOT localization in JEG3 cells, I performed dual-staining immunofluorescence (IF) of AMOT with tight junction marker ZO-1. In JEG3 cells cultured under standard conditions,

AMOT co-localization with ZO-1 was observed at intracellular tight junctions (Figure 3.4). Strong

AMOT localization was also observed at the protruding edges (‘leading edge’) of JEG3 cells.

3.2.2 TGF1/3 ligand treatment reduces AMOT 130 and 80 protein levels

Next, I treated JEG3 cells with 10 ng/mL TGF1 or TGF3 ligand for 24 hours and examined protein levels and cellular localization of AMOT by WB and IF analyses, respectively. This particular concentration and time of TGFβ exposure was selected following a time and dose course of TGFβ treatment in these cells. Specifically, JEG3 cells were treated with 1, 5 and 10 ng/mL of

TGFβ for 3, 6, 12, 24 and 48 hours and it was established that 10ng/mL for 24 hours produced a prominent effect. Activation of TGF signalling following addition of ligand was confirmed by an increase in pSMAD2 levels (Figure 3.5). Following TGF1 and TGF3 treatment, protein levels of both AMOT 130 and AMOT 80 decreased significantly compared to vehicle controls

(p<0.0001, p<0.01, p<0.001, p<0.05) (Figure 3.5). Notably, no differences in AMOT 130 and 80 protein levels were observed between TGF1 and TGF3.

3.2.3 TGF1/3 treatment promotes subcellular redistribution of AMOT

TGF1/3 ligand treatment resulted in distinct changes in the subcellular localization of AMOT, particularly with respect to the tight junction and F-actin cytoskeleton (Figure 3.6, Figure 3.7). In vehicle control, AMOT displayed primary localization to the tight junction. Quantification of

AMOT and ZO-1 colocalization by Pearson’s correlation confirmed strong AMOT colocalization at the tight junction (Pearson’s correlation coefficient (PCC)=0.56). Following TGF1 and TGF3 treatment, AMOT localization at the tight junction appeared intermittent and disrupted compared to vehicle controls in which AMOT localization at tight junction was continuous. This was confirmed by a reduction in Pearson’s correlation of AMOT and ZO-1 following treatment with

TGF1 and TGF3 (PCC=0.29 and PCC=0.28, respectively). Interestingly, TGF1/3 treatment resulted in redistribution of AMOT to the cytoplasm in a fiber-like orientation (Figure 3.6). To determine whether these AMOT fibers coincided with the F-actin scaffold, IF analysis of AMOT

and F-actin marker phalloidin following TGF ligand treatment was conducted. Following

TGF1/3 treatment, AMOT redistributed from the tight junction to predominantly the cytoplasm, where it co-localized with F-actin cytoskeleton fibers (Phalloidin) (Figure 3.7). Quantification confirmed that AMOT and phalloidin colocalization is increased following TGF1 and TGF3 treatment (PCC=0.64 and PCC=0.58, respectively) compared to vehicle control (PCC=0.19).

Overall, despite the reduction in AMOT 130 and 80 protein levels (Figure 3.5) following TGF1/3 treatment, TGF1/3 promoted striking changes in AMOT’s subcellular redistribution, in particular its colocalization with the cytoskeleton.

AMOT ZO-1 Merge IgG

40X 40X 40X 40X

LE AMOT ZO-1 DAPI

Figure 3.4 Co-localization of AMOT with tight junction protein ZO-1 in JEG3 choriocarcinoma cells

Representative images of immunofluorescence (IF) analysis of AMOT (green) and tight junction protein ZO-1 (red) in JEG3 cells cultured in standard conditions (n=3). Nuclei were stained with DAPI (blue). Normal Goat and Rabbit IgG were used as negative control. AMOT co-localization with ZO-1, as indicated by the white arrows, suggests AMOT localizes to primarily to the tight junctions in JEG3 cells. AMOT localization to the leading edge (LE) is indicated by the line of multiple arrows.

**** 1.5

***

Jeg3 Cells 1.0 Vehicle TGFβ1 T GFβ3 0.5

130 kDa Change Fold AMOT 0.0

Protein 130 AMOT 80 kDa Vehicle TGFβ1 TGFβ3 **

pSMAD2 58 kDa 1.5 * 50 kDa SMAD2 1.0

β actin 42 kDa 0.5 Fold Change Fold

AMOT 80 Protein Protein 80 AMOT 0.0 Vehicle TGFβ1 TGFβ3

Figure 3.5 Effect of TGFβ1/3 on AMOT protein levels in JEG3 cells

Representative Western blot (WB) showing AMOT 130 and AMOT 80 protein levels following treatment with 10 ng/mL TGFβ1 or TGFβ3 ligand for 24 hours. Densitometry analysis of AMOT 130 and AMOT 80 protein levels normalized β-actin and expressed as a fold-change relative to vehicle control (EtOH) (n=5). Statistical significance was assessed using pairwise non-parametric Mann-Whitney test (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001). AMOT 130 and 80 protein levels both significantly decreased following TGFβ1 and TGFβ3 treatment.

+ Vehicle + TGFβ1 + TGFβ3 IgG 63X 63X 63X

40X

AMOT

63X 63X 63X

n e

i f

f 0.8 **** e o

C ****

n 0.6 o 63X i 63X 63X t

* a l

e 0.4

* C

0.2

* s ’ erge

* * n

o M

s 0.0 r

a le 1 3 e ic β β

P F F eh G G AMOT ZO-1 DAPI V T T

Figure 3.6 Effect of TGFβ1/3 on AMOT and ZO-1 co-localization in JEG3 cells

Representative images of immunofluorescence (IF) analysis of AMOT (green) and tight junction protein ZO-1 (red), following treatment with 10 ng/mL TGFβ1 and TGFβ3 ligand for 24 hours (n=3). Nuclei were stained with DAPI (blue). Normal Goat and Rabbit IgG were used as negative controls. Colocalization analysis of AMOT and ZO-1 using Volocity Imaging Software and expressed as Pearson’s correlation coefficient (PCC) (Vehicle: PCC=0.56; TGFβ1: PCC=0.29; TGFβ3: PCC=0.28). Statistical significance was assessed using pairwise non-parametric Mann- Whitney test (****p<0.0001). Treatment of JEG3 cells with TGFβ reduces AMOT co-localization with ZO-1 at tight junction, as indicated by the white arrows pointing to intermittent AMOT/ZO- 1 colocalization. Blue asterisks indicate fiber like structures in the cytoplasm following TGFβ1/3 treatment.

+ Vehicle + TGFβ1 + TGFβ3

63X 63X 63X

AMOT

e i

c ****

f 63X f 0.8 63X 63X e ****

n 0.6

i t

e 0.4

C Phalloidin

0.2

’

s 0.0 63X 63X 63X r a le 1 3 e ic β β P F F eh G G

V T T

Merge

AMOT Phalloidin DAPI

Figure 3.7 Effect of TGFβ1/3 on AMOT and F-actin co-localization in JEG3 cells

Representative images of Immunofluorescence (IF) analysis of AMOT (green) and F-actin cytoskeleton probe phalloidin (red) levels following treatment with 10 ng/mL TGFβ1 and TGFβ3 ligand for 24 hours (n=3). Nuclei were stained with DAPI (blue). Colocalization analysis of AMOT and phalloidin using Volocity Imaging Software and expressed as Pearson’s correlation coefficient (PCC) (Vehicle: PCC=0.19; TGFβ1: PCC=0.64; TGFβ3: PCC=0.58). Statistical significance was assessed using pairwise non-parametric Mann-Whitney test (****p<0.0001). Treatment of JEG3 cells with TGFβ1/3 upregulates AMOT co-localization with F-actin cytoskeleton, as indicated by the white arrows pointing to AMOT/F-actin co-localized fibers.

3.3 AMOT 130 is regulated by Smad-dependent TGF pathway

As previously described, TGF ligands induce several signalling cascades, one of which includes the Smad-dependent pathway (also commonly known as the canonical pathway) (Budi EH 2018).

To assess whether TGF regulation on AMOT re-localization is mediated by the Smad-dependent pathway, I treated JEG3 cells with TGF1/3 in the presence of a small molecular inhibitor of the alk5 receptor (TGFR1), namely SB-431542, and analyzed changes in AMOT localization compared to treatment with TGF1/3 alone. IF analysis of AMOT and ZO-1 revealed that the redistribution of AMOT from tight junction to cytoplasm found following TGF treatment is abolished when alk5 inhibition by SB-431542 is introduced (Figure 3.8). This is evident by the retention of AMOT and ZO-1 at cell boundaries and the absence of AMOT cytoplasmic fibers in

TGF1/3+ SB-431542 conditions, versus the reduced AMOT and ZO-1 association and presence of AMOT cytoplasmic fibers following TGF1/3 exposure. No changes in AMOT/ZO-1 localization was observed following SB-431542 + vehicle treatment compared to vehicle control.

Despite being unable to distinguish the two AMOT isoforms by IF, AMOT 130 is the isoform known to associate with the F-actin cytoskeleton (Ernkvist et al., 2006). Considering this, it is reasonable to propose that inhibition of alk5 prevents AMOT 130 redistribution to the cytoplasm and association to the F-actin cytoskeleton, and as such TGF-mediated localization of AMOT to the cytoskeleton is regulated by the Smad-dependent pathway.

Next, AMOT protein levels following treatment with TGF1/3 +/- SB-431542 were assessed to determine if SMAD-dependent TGF signalling regulated quantities of AMOT protein.

Considering that inhibition of the alk5 receptor prevents phosphorylation of SMAD2 and further downstream signalling, I confirmed effective inhibition of Smad-dependent TGF pathway by

observing a reduction in pSMAD2:SMAD2 protein level ratio (Figure 3.9). By WB analysis,

AMOT 130 protein levels were significantly increased following TGF1/3 + SB-431542 treatment, compared to TGF1/3 treatment alone (p<<0.05) (Figure 3.9), whereas AMOT 80 protein levels remained unchanged. Overall, it appears that Smad-dependent TGF signalling preferentially regulates the localization and the quantity of AMOT 130 in trophoblast derived

JEG3 cells.

+ Vehicle + TGFβ1 + TGFβ3

+SB-431542 +SB-431542 +SB-431542 40X 40X 40X 40X 40X 40X

AMOT

40X 40X 40X 40X 40X 40X

1 -

40X 40X40X 40X 40X 40X 40X40X 40X *

* * Merge *

AMOT ZO-1 DAPI

Figure 3.8 Contribution of Smad-dependent TGFβ signalling on AMOT localization

Representative images of immunofluorescence (IF) analysis of; AMOT (green) and tight junction protein ZO-1 (red) following treatment with 10 ng/mL TGFβ1 and TGFβ3 ligand for 24 hours in the presence/absence of 5 M alk5 receptor inhibitor SB-431542 (n=3). Nuclei were stained with DAPI (blue). Treatment of JEG3 cells with TGFβ reduced AMOT co-localization with ZO-1 at tight junction and promoted AMOT localization in the cytoplasm. However, this effect was abrogated in the presence of SB-431542. Inhibition of alk5 promoted AMOT retention at the tight junction and less AMOT present in fiber like structures within the cytoplasm. White arrows indicate AMOT presence at tight junction, and blue asterisks indicate AMOT presence in cytoplasm.

+5uM 77 (-) SB-341542

V β1 β3 V β1 β3

AMOT 130 kDa

80 kDa

pSMAD2 58 kDa

SMAD2 58 kDa

B-actin 42 kDa

1.5 1.5 a

1.0 1.0

0.5 0.5

Fold Change Fold

Fold Change Fold AMOT 80 Protein Protein 80 AMOT 0.0 0.0 Protein 130 AMOT e 1 i i i le 1 3 i i i cl β 3 5 5 5 c B 5 5 5 i F β k k k i F B lk lk lk h F l l l eh F a a a e G G a a a 3 G G + + + V T T + + + c V T T le 1 3 d le 1 3 c β β ic B B i F F h F F h e G G e G G V V T T 2 T T f 1 g e

0 e 1 3 i i i l β β 5 5 5 ic F F lk k lk pSMAD2/SMAD2 pSMAD2/SMAD2 h a l a e G G + a + Protein Change Fold Protein V T T + le 1 3 ic β β h F F e G G V T T Figure 3.9 Contribution of Smad-dependent TGFβ signalling on AMOT protein levels

Representative Western blot (WB) showing AMOT 130 and 80 protein levels following treatment with 10 ng/mL TGFβ1 and TGFβ3 ligand for 24 hours in the presence/absence of 5 M alk5 receptor inhibitor SB-431542. Densitometry analysis of AMOT 130 and 80 protein levels normalized to β-actin and expressed as a fold-change relative to vehicle control (EtOH) (n=3). Statistical significance was assessed using pairwise non-parametric Mann Whitney test. Densitometry analysis of pSMAD2 protein levels normalized to total SMAD2 and expressed as a fold-change relative to vehicle control. Statistical significance was assessed using pairwise Mann- Whitney test [(a-TGFβ1 vs TGFβ1 +alk5i, b-TGFβ3 vs TGFβ3 +alk5i, p<0.05) (c-vehicle vs vehicle + TGFβ1; d-vehicle vs TGFβ3; e-vehicle vs vehicle +alk5i; f- TGFβ1 vs TGFβ1+alk5i; g- TGFβ3 vs TGFβ3+alk5i, p<0.01)

3.4 TGFβ promotes AMOT redistribution in migrating cells

Considering the literature surrounding the role of AMOT in endothelial cell migration (Aase et al.,

2007), as well as our finding of placental AMOT localization in a subset of EVT’s known to be characteristically migratory (Figure 3.3), I first assessed how AMOT distribution changes during cell migration. JEG3 cells are a suitable model to study trophoblast cell migration in vitro (Xu et al., 2016), therefore I performed a wound healing assay and IF staining in JEG3 cells to investigate

AMOT localization during migration in trophoblast derived cells (Figure 3.10A). Immediately following wound initiation, AMOT is primarily localized to the tight junctions, consistent with previous observations of endogenous AMOT localization in resting cells (Figure 3.4). By 3 hours following wound initiation, AMOT is still found at the tight junctions; however, it is also distinctly observed at the leading edge of migrating cells, and in the cytoplasm of cells directly underlying the migratory front. Of note, AMOT presence at the leading edge and in the cytoplasm are both in a fiber orientation (Figure 3.10A). Hence, these findings indicate that AMOT localization is altered during cell migration.

Various reports, in the placenta field as well as in cancer and immunology studies, have implicated a role of TGFβ as a regulator of cell migration (Kim et al., 2006; Wendt et al., 2012; Xu et al.,

2015). This, along with the prominent effect TGFβ has on AMOT subcellular localization, prompted me to investigate the effect of TGFβ stimulation on AMOT localization during migration

(Figure 3.10B). Analysis of AMOT localization 3 hours following wound initiation revealed that

TGFβ1 and TGFβ3 exposure promoted a dissolution of AMOT at the tight junction of cells distinctly beneath the migrating front, in addition to AMOT fibers localizing at the leading edge of the wound. Loss of AMOT at the tight junction in these cells following TGFβ treatment has potential to disrupt apical/basolateral cell polarity, which in turn, may promote cell migration.

3.5 AMOT 80 promotes JEG3 cell migration

AMOT is a scaffolding protein, and consequently is subject to dynamic movement throughout the cell at a given time. Mapping of AMOT through IF in fixed cells as shown in Figure 3.10, undoubtedly provided a clear picture of the differing AMOT localization at 0 versus 3 hours following wound formation but did not permit inference on AMOT movement in the cell. Thus, to fully appreciate the extent of AMOT redistribution and role during cell migration in real time, time lapse imaging in live cells was utilized.

Visualization of AMOT in live cells was permitted through the use of a YFP-labelled AMOT 80 construct transiently transfected in JEG3 cells. AMOT 80 is the dominant isoform involved in cell migration (Ernkvist et al., 2009), and therefore was the construct I used to assess AMOT in migrating JEG3 cells. Following the workflow in fixed cells, I assessed the subcellular movement of AMOT protein in JEG3 cells in a wound healing assay. In migrating cells, AMOT localization is in constant flux between the intracellular tight junction and the cytoplasm, as elucidated by the live cell videos in Figure 3.11. In the cytoplasm, AMOT is expressed as multiple, small punctate structures that are seen to dynamically redistribute around the cell during migration and division.

Following the visible redistribution of AMOT punctate structures, the cells overexpressing AMOT appear to accelerate their movement toward the wound. Quantification of migration rate revealed that JEG3 cells overexpressing AMOT 80 YFP migrated 2x faster than EV controls (p<0.01)

(Figure 3.12). These findings suggest that AMOT 80 has a role in promoting migration of JEG3 cells.

A B 0 hrs 3 hrs 3 hrs

40X 40X 40X LE LE + TGFβ1

TJ * *

TJ *

63X 63X 40X

LE LE + TGFβ3

TJ * TJ

AMOT/DAPI AMOT/DAPI IgG 40X

Figure 3.10 Effect of TGFβ on AMOT localization in migrating edge of JEG3 cells

(A) Representative images of immunofluorescence (IF) analysis of AMOT (green) at 0 and 3 hours following wound initiation in confluent monolayer of JEG3 cells (n=2). (B) Representative images of IF analysis of AMOT (green) 3 hours after wound initiation in the presence of 10ng/mL TGFβ1 or TGFβ3 ligand (n=3). Nuclei were stained with DAPI (blue). Normal Goat IgG was used for negative control. Solid white arrows indicate AMOT presence at tight junction (TJ) or leading edge (LE), blue asterisks indicate AMOT presence in cytoplasm, and dotted white arrows indicate lack of AMOT at tight junction.

Live Cell Videos 81

Click

Snapshots

Click

* *

ii) Click

Click

Snapshots

Click TJ * *

TJ TJ *

Figure 3.11 AMOT localization in live cell imaging of migrating JEG3 cells

Representative videos and corresponding still-pictures depicting cell migration in JEG3 cells overexpressing AMOT 80 YFP construct. The white-dotted line illustrates the location of the original wound. Videos were acquired on a spinning disc confocal microscope using Volocity Imaging software. Magnification of both videos is 20X. AMOT distribution is observed to dynamically change from tight junction (TJ) (indicated by white arrows), to punctate structures, in the cytoplasm (indicated by blue asterisks) during cell migration.

3 **

1 (Fold Change) (Fold

JEG3 Cell Migration Rate Migration Cell Rate JEG3 0 0 V 8 E T O M A E O

Figure 3.12 Effect of AMOT 80 overexpression on migration rate of JEG3 cells during wound healing.

Linear migration rates for JEG3 cells overexpressing AMOT 80 YFP expressed as a fold-change relative to pcDNA empty vector (EV) control (n=3). Statistical significance was assessed using non-parametric Mann Whitney test (**p <0.01). Migration rate of JEG3 cells overexpressing AMOT 80 was 2.3x faster than EV control.

3.6 Novel AMOT/Par6 interaction and its regulation by TGFβ

Interactions amongst cellular scaffolding proteins can facilitate or repress necessary cellular events including cell migration, protein trafficking and targeted recycling (Moleirinho et al., 2014).

Considering that scaffolding protein AMOT promotes reorganization of polarity proteins (Wells et al., 2006), I sought to establish if AMOT directly interacts with polarity protein Partitioning defective protein 6 (Par6). Par6 is under the regulation of Smad-independent TGFB signalling, and functions to disrupt apical basolateral cell polarity upon TGFβ ligand binding (Bose and Wrana,

2006; Ozdamar et al., 2005). Work from our lab has demonstrated the importance of Par6 in controlling trophoblast cell fusion and migration, as well as shown TGFβ signalling via Par6 is present in JEG3 cells (Sivasubramaniyam et al., 2013; Xu et al., 2015). However, the interplay of

Par6 and AMOT, and further the role of TGFβ signalling on this potential interaction, has never been investigated.

First, I assessed whether AMOT and Par6 colocalize in JEG3 cells by IF analysis (Figure 3.13).

In vehicle controls, AMOT and Par6 exhibit co-localization, particularly at the tight junction. Upon treatment with 10 ng/mL TGFβ1 or TGFβ3, AMOT and Par6 co-localization is still evident at the level of the tight junction, but more distinctly in the cytoplasm. Observation of a dissolution of

AMOT at tight junctions following TGFβ treatment is consistent with previous findings (Figure

3.6), and observation of increased Par6 fluorescence intensity is consistent with the literature (Xu et al., 2016).

After establishing that TGFβ treatment promoted an increased co-localization of AMOT and Par6,

I investigated if an association of the two proteins existed. Following immunoprecipitation (IP) of

Par6 followed by WB analysis of AMOT, an association of AMOT and Par6 was observed (Figure

3.14A). Further, this association was increased primarily following TGFβ3 treatment.

Interestingly, AMOT/Par6 association was only observed with the AMOT 80 isoform, as no bands at 130 kDa were present following IP of Par6 and WB of AMOT. To conclusively establish a novel interaction between AMOT and Par6 and its regulation by TGFβ, JEG3 cells were subjected to

Proximity Ligation Assay (PLA) following treatment with 10 ng/mL of TGFβ1 and TGFβ3, and vehicle control (Figure 3.14B). PLA analysis revealed an AMOT and Par6 interaction exists in

JEG3 cells, as identified by the presence of individual red PLA fluorophores (dots). Quantification of PLA signals per cell revealed that AMOT/Par6 interaction was significantly increased in both

TGFβ1 and TGFβ3 treated cells compared to vehicle control (p<0.05).

To confirm in vitro findings, I assessed co-localization of AMOT and Par6 in early placenta development by IF in human placenta tissue sections. Co-localization of AMOT and Par6 is distinctly observed in cytotrophoblasts of the floating placenta villi (Figure 3.15A). Unlike syncytiotrophoblasts, cytotrophoblasts are polarized cells and can function as precursors to EVT cells (DaSilva-Arnold et al., 2015). Further, AMOT and Par6 co-localization is also found in the

EVT’s comprising the placenta anchoring placenta villi (Figure 3.15A). IP analysis in placentae tissue lysates from 7-14 weeks of gestation revealed AMOT and Par6 association is present in tissue as early as 7 weeks of gestation, and peaks around 10-14 weeks of gestation (Figure 3.15B).

Vehicle TGFβ1 TGFβ3

AMOT 63X 63X 63X

Par6

63X 63X 63X

* * *

40X 40X * Merge 63X 63X 63X

AMOT Par6 DAPI

Figure 3.13 Effect of TGFβ1/3 on AMOT and Par6 co-localization in JEG3 cells

Representative images of Immunofluorescence (IF) analysis of; AMOT (green) and Par6 (red) following treatment with 10 ng/mL TGFβ1 and TGFβ3 ligand for 24 hours (n=3). Nuclei were stained with DAPI (blue). Treatment of JEG3 cells with TGFβ increases AMOT co-localization with Par6 at tight junction (TJ), as indicated by the white arrows, and cytoplasm, as indicated by blue asterisks.

IP Par6

AMOT 80 kDa

Par6 37 kDa B

Vehicle TGFβ1 TGFβ3

63X 63X 63X **

2.5 *

2.0

1.5 63X 63X 1.0

0.5 (foldchange) 0.0 Negative Controls signals/cell PLA minus probe absent plus & minus probes absent Par6 antibody absent Vehicle TGFβ1 TGFβ3 63X 63X 63X

PLA DAPI

Figure 3.14 Effect of TGFβ1/3 on AMOT and Par6 interaction in JEG3 cells

(A) Immunoprecipitation (IP) of Par6 followed by Western blot analysis of AMOT in JEG3 cells after treatment with 10 ng/mL TGFβ1/3 for 24 hours (n=3). (B) Representative fluorescence images depicting In situ Proximity Ligation (PLA) of AMOT with Par6 in JEG3 cells following treatment with 10 ng/mL TGFβ1/3, or vehicle for 24 hours (n=3 ). Nuclei were stained with DAPI (blue). Negative control was PLA reaction: missing minus (-) PLA probe, missing plus (+) and minus (-) probes and missing Par6 primary antibody. PLA interactions were counted blindly by two independent assessors, averaged and normalized to the number of whole nuclei. Data is expressed as a fold change relative to vehicle control. Statistical significance was assessed using non-parametric Mann-Whitney test (*p<0.05). An interaction between AMOT and Par6 proteins is observed, and TGFβ treatment upregulated this interaction.

ST ST CT CT EVT EVT

20x 20x

10x 10x

AMOT Par6 DAPI IgG

B 10x

IP Par6 IgG 7 8 9 10 11 13 14

AMOT 80 kDa

Par6 37 kDa Weeks of gestation

Figure 3.15 AMOT 80 and Par6 interaction in human placenta tissue

(A) Representative images illustrating AMOT (green) and Par6 (red) co-localization in placenta tissue sections from 5-6 weeks of gestation (n=3). Normal Goat and Mouse IgG were used as negative controls. Nuclei were stained with DAPI (blue). (B) Immunoprecipitation (IP) of Par6 and Western blot analysis of AMOT in human placenta tissue lysates from 7-14 weeks of gestation (n=14). ST(syncytiotrophoblast) CT(cytotrophoblast).

3.7 PDZ and coiled-coil binding domains are important for AMOT and Par6 interaction

AMOT contains multiple binding domains that promote its interaction with other proteins and are essential to its scaffolding function. The C-terminal PDZ binding domain found on both AMOT

130 and 80, is a known protein motif mediating protein-protein interactions within apical- basolateral polarity networks. Specifically, the PDZ-binding domain on AMOT has proven important for recruitment of polarity complex proteins and localization to tight junction (Wells et al., 2006). Additionally, the coiled-coil binding domain on AMOT 130 and AMOT 80 is important for AMOT binding to GTPase activating protein (GAP) Rich1 at the tight junction (Wells et al.,

2006). Considering this, the importance of the PDZ and coiled coil binding domains on AMOT

130 and 80 in the interaction with Par6 was investigated.

Plasmid DNA constructs of wild type AMOT 130, wild type AMOT 80, AMOT 130 with deletion in PDZ domain (AMOT 130 PDZ), AMOT 80 with deletion in PDZ binding domain (AMOT

80PDZ), and AMOT 130 with deletion in coiled coil domain (AMOT 130 cc) were obtained and validated (Figure 3.16). JEG3 cells were transfected with the aforementioned AMOT constructs along with wild type Par6, and association between AMOT and Par6 was assessed

(Figure 3.17A). Compared to EV controls, overexpression (OE) of wild type AMOT 80 and Par6 upregulated AMOT 80/Par6 association as assessed by IP. Examining the binding domains; overexpression of AMOT 80 PDZ decreased the association of AMOT 80 and Par6 compared to overexpression of wild type AMOT 80. Interestingly, overexpression of AMOT130 cc decreased the association of AMOT and Par6.

To compare with IP findings, I performed PLA analysis following over expression AMOT 130 and AMOT 80 constructs (Figure 3.17B). In EV control, AMOT and Par6 interaction is observed.

AMOT and Par6 interaction is increased following overexpression of both AMOT 130, and to a larger degree following over expression of AMOT 80. However, overexpression of all mutant constructs (AMOT 80 PDZ, AMOT 130 PDZ and AMOT 130 cc) resulted in a decreased

AMOT and Par6 interaction. Taken together, these data suggest that Par6 primarily interacts with

AMOT 80 and that the PDZ binding motif contained in AMOT 80 is critical for this interaction.

A AMOT 130 Constructs

N-terminal Coiled-coil PDZ binding extension domain ABD domain

AMOT 130 N - 1084 a.a - C

AMOT 130 ∆cc N - - C 1082 a.a

AMOT 130 ∆PDZ N - - C 1081 a.a

AMOT 80 Constructs

Coiled-coil PDZ binding domain ABD domain AMOT 80 675 a.a N - - C

AMOT 80 ∆PDZ N - - C 671 a.a

AMOT 130 kDa AMOT 80 kDa

-Tubulin -Tubulin 50 kDa 50 kDa

Figure 3.16 Description and validation of AMOT 130 and AMOT 80 plasmid constructs

(A) Protein structures for wild-type and mutated AMOT 130 and AMOT 80 plasmid DNA constructs in in vitro experiments. Pertinent binding domains are distinguished among constructs. a.a= amino acid (B) Validation of AMOT constructs by overexpression in JEG3 cells.

92  A AMOT 80 - - + - - PDZ AMOT 130 - + -  -  PDZ CC Par6 - + + + + + IP Par6 AMOT 80 kDa Par6 37 kDa

B OE OE Negative Controls EV -plus & minus AMOT 130 AMOT 80 -minus probe -Par6 antibody probes

40x 40x 40x

20x 40x 40x 3 b a OE AMOT OE AMOT OE AMOT 2 130  cc 130  PDZ 80  PDZ 1 c d e

(foldchange) PLA signals/cell signals/cell PLA 0 V 0 0 il z Z E 3 8 o d D 1 T c p T d a P O le lt ta O M i e l M o d e A c 0 d A E ta 3 0 E O el 1 8 O d T T 0 O O 3 1 M M T A A 40x 40x 40x O E E M O O A E PLA DAPI O Figure 3.17 Importance of PDZ and coiled-coil binding domains to AMOT-Par6 interaction

(A) Immunoprecipitation (IP) of Par6 and Western blot analysis of AMOT in JEG3 cells following overexpression with EV, AMOT 130, AMOT 80, AMOT 130 PDZ, AMOT 80PDZ, AMOT 130 cc (n=3). (B) Representative fluorescence images depicting In situ Proximity Ligation (PLA) of AMOT with Par6 in JEG3 cells following overexpression of the DNA constructs in panel A. Nuclei were stained with DAPI (blue). Negative control was PLA reaction: missing minus (-) PLA probe, missing plus (+) and minus (-) probes and missing Par6 primary antibody. PLA interactions were counted blindly by two independent assessors, averaged and normalized to the number of whole nuclei. Data are expressed as fold change relative to EV. Statistical significance was assessed using non-parametric Mann-Whitney test (a-EV vs OE AMOT130, p<0.05; b-EV vs OE AMOT 80, p<0.001; c-OE AMOT 130 vs OE AMOT 130 cc, p<0.01; d- OE AMOT 130 vs OE AMOT 130 PDZ, p<0.01; e-AMOT 80 vs AMOT 80 PDZ, p<0.001).

3.8 AMOT promotes dissolution of RhoA at the tight junction

Activation of the non-canonical TGFβ pathway via Par6/Smurf1 leads to targeted ubiquitination and degradation of tight junction residing GTPase RhoA, ultimately leading to the dissolution of tight junctions (Ozdamar et al., 2005; Wang et al., 2006). Having established an interaction between AMOT and Par6 proteins, the downstream consequence of this interaction was next to assess; particularly the interplay between AMOT and GTPase RhoA, a tight junction stabilizing protein. To investigate this, I overexpressed AMOT 130 and AMOT 80 and analyzed RhoA protein levels. Compared to EV controls, overexpression of both AMOT 130 and AMOT 80 resulted in a significant decrease of RhoA protein levels (p<0.01 and p<0.001, respectively) (Figure 3.18).

Having established the effect of AMOT on protein levels of RhoA, I validated and used a YFP tagged AMOT 80 construct, the isoform implicated in cell migration, and examined the effect of overexpressing AMOT 80 on the localization of RhoA (Figure 3.19). In EV controls, AMOT and

RhoA are primarily present at tight junctions as assessed by IF. RhoA is also present in the cytoplasm. Following overexpression of AMOT 80 YFP, RhoA fluorescence specifically at the tight junctions of JEG3 cells is reduced. This finding corroborates protein analysis (Figure 3.18), and further shows that AMOT 80 mediated reduction of RhoA occurs at the tight junction, potentially suggesting that overexpression of AMOT 80 results in tight junction destabilization.

OE OE EV AMOT AMOT 130 80

1.5

AMOT 130 130 kDa 1.0 * ** AMOT 80 80 kDa 0.5

Fold Change Fold RhoA Protein RhoA 22 kDa 0.0 OE OE EV AMOT AMOT B-actin 42 kDa 130 80

Figure 3.18 Effect of AMOT overexpression on protein levels of RhoA

Representative Western blot showing RhoA, AMOT 130 and AMOT 80 protein levels following overexpression of AMOT 130 and AMOT 80 in JEG3 cells. Densitometry analysis of AMOT 130 and AMOT 80 protein levels were normalized to β-actin and expressed as a fold-change relative to EV control (n=3). Statistical significance was assessed using unpaired Mann Whitney test (*p <0.01, **p<0.001). Overexpression of AMOT 130 and 80 decreased RhoA protein levels.

A OE AMOT 95 EV 80 YFP

40X 40X

AMOT 80 YFP Plasmid Construct

N - - C 40X 40X

B AMOT DAPI EV OE AMOT 80 YFP

20X 20X 40X

AMOT

20X 20X 40X IgG

RhoA 20X

20X 20X 40X

Merge Merge

AMOT RhoA DAPI

Figure 3.19 Effect of AMOT overexpression on RhoA localization

(A) Transfection of JEG3 cells with AMOT 80-YFP plasmid construct in JEG3 cells, compared to transfection of empty vector control and immunofluorescence staining with AMOT. (B) Representative images of Immunofluorescence analysis of RhoA (red) and YFP staining of AMOT 80-YFP in JEG3 cells (n=3). Nuclei were stained with DAPI (blue). Normal mouse IgG was used as negative control for RhoA. RhoA localization to tight junction is decreased in cells overexpressing AMOT 80 YFP.

3.9 AMOT protein levels and distribution is disrupted in preeclampsia

Reports from our lab have demonstrated that aberrant TGFβ signalling as a result of chronic hypoxia contributes to the impaired trophoblast differentiation and uteroplacental vascularization seen in preeclampsia (PE) (Caniggia et al., 1999; Caniggia et al., 2000). In this thesis, I have demonstrated that TGFβ regulates AMOT. Considering the established contribution of TGFβ signalling to the pathogenesis of preeclampsia, I sought to examine AMOT in this serious pregnancy disorder.

AMOT levels in placentae from early-PE (E-PE) and normotensive pre-term control (PTC) pregnancies were analyzed by WB analysis. Compared to PTC placentae, AMOT 130 and AMOT

80 protein levels were markedly reduced in placentae complicated with E-PE (p<0.0001) (Figure

3.20). IF analysis of floating villi from PTC placenta revealed that AMOT is widely localized in the stroma of the villi where mesenchymal cells reside. Further, AMOT was observed in the cytotrophoblast and syncytiotrophoblast layers (Figure 3.21A). However, analysis of floating villi from E-PE placenta revealed substantial differences. An overall reduction in AMOT fluorescence in all trophoblast layers was observed in placentae complicated with E-PE, corroborating findings of a decrease in AMOT protein levels. Furthermore, the minimal AMOT localization in the villous stroma appears restricted to cells surrounding the villous vessels (Figure 3.21B).

Appreciating the deficiencies in trophoblast invasion that typify preeclampsia, I also assessed

AMOT presence in EVTs. Specifically, I located the maternal-fetal interface and assessed the interstitial EVT population which had invaded and resided to the maternal decidua. In PTC, AMOT localization to the interstitial EVTs within the decidua is apparent, and is found expressed throughout the cytoplasm (Figure 3.21B). In E-PE, however, AMOT fluorescence in interstitial

EVTs is reduced, and appears localized to the outer membrane edge of cells (Figure 3.21B). These

deficiencies in AMOT observed in the interstitial EVTs occupying the decidua from E-PE pregnancies further corroborate the conclusion that AMOT protein levels and tissue distribution is altered in this disorder of pregnancy.

3.9.1 AMOT and Par6 interaction is impaired in preeclampsia

Having identified the discernable reduction of AMOT in E-PE, the question remained if the newfound interaction of AMOT and Par6 would also be disrupted. Also, the knowledge of altered

TGFβ signalling in preeclampsia along with the finding that interaction of AMOT and Par6 is regulated by TGFβ signalling (Xu et al., 2016), further prompted this investigation. Analysis of

AMOT protein levels following IP of Par6 in PTC and E-PE placentae was performed. Compared to PTC, there was reduced AMOT and Par6 association in pregnancies complicated with E-PE

(Figure 3.22A). IF analysis of AMOT and Par6 in the previously shown sections of maternal-fetal interface from both PTC and E-PE was also performed to examine AMOT and Par6 colocalization.

Initial examination of Par6 showed increased fluorescence in E-PE versus pre-term control, confirming previous reports from our lab that demonstrated an increased level of Par6 in PE

(Sivasubramaniyam et al., 2013). Furthermore, the co-localization of AMOT/Par6 appears reduced in E-PE compared to PTC (Figure 3.22B). This confirms IP findings and suggests that the interaction between AMOT and Par6 is impaired in PE.

2.0 1.5

1.0

Fold Change Fold 0.5 MOT 130 Protein Protein 130 MOT

PTC E-PE A 0.0 PTC E-PE AMOT 130 kDa 80 kDa ** 2.0

Ponceau 1.5 1.0

0.5

Fold Change Fold 0.0

Protein 80 AMOT PTC E-PE

Figure 3.20 AMOT protein levels in preeclamptic and normotensive pre-term control placentae

(A) Representative Western blot showing AMOT 130 and AMOT 80 protein levels in preeclamptic (E-PE) and pre-term control (PTC) placenta (E-PE, n=15; PTC, n=10). Densitometry analysis of AMOT 130 and AMOT 80 protein levels were normalized to entire lane following Ponceau staining and expressed as fold change relative to PTC. Statistical significance was determined using non-parametric Mann-Whitney test (*p<0.001). AMOT 130 and AMOT 80 protein levels are significantly decreased in E-PE compared to PTC placentae.

99 A PTC E-PE

ST x

S V

20x 20x

V S CT

ST 40x 40x

B PTC E- PE

EVT 40x 40x EVT

10x 10x

IgG Control IgG Control

10x AMOT DAPI 10x 10X 10X Figure 3.21 AMOT localization in preeclamptic and pre-term control placenta tissue sections

Representative immunofluorescence (IF) images in preeclamptic (E-PE) and pre-term control (PTC) placenta depicting AMOT distribution in (A) Floating villi and (B) fetal-maternal interface (E-PE, n=3; PTC, n=3). Normal Goat was used as negative control. ST= syncytiotrophoblast; CT= Cytotrophoblast; S= stroma; V=vessel; EVT=extravillous trophoblast).

100

IP Par6 IgG PTC E-PE

AMOT 80 kDa

Par6 37 kDa

B PTC E-PE

IgG

40x 40x 20x AMOT Par6 DAPI

Figure 3.22 AMOT 80 and Par6 association in preeclamptic and pre-term control placenta

(A) Immunoprecipitation of Par6 and western blot analysis of AMOT in preeclamptic (E-PE) and pre-term control (PTC) tissue lysates (E-PE, n=5; PTC, n=6). Normal Goat and Mouse IgG were used as negative controls. (B) Representative immunofluorescence (IF) images in preeclamptic (E- PE) and pre-term control (PTC) placenta depicting AMOT distribution in fetal-maternal interface (E-PE, n=3; PTC, n=3).

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3.10 AMOT protein levels and localization is disrupted in low oxygen

A major feature characterizing the pathogenesis of PE is persistence of low oxygen within the placenta, as elucidated by elevated levels of HIF-1 (Caniggia and Winter, 2002; Rajakumar et al., 2004; Soleymanlou et al., 2005) and impaired oxygen sensing machinery (Alahari et al., 2018;

Rolfo et al., 2010). During placental development, critical events including opening of the intervillous space, increase in maternal blood flow and proper trophoblast differentiation all rely on tightly regulated changes in oxygen tension. Further, changes in oxygen tension have been shown to regulate TGFβ signalling in the developing placenta (Caniggia et al., 2000). Hence, having established AMOT is disrupted in PE, I next investigated whether exposure to low oxygen would impact AMOT.

Exposure of JEG3 cells to 3% oxygen for 24 hours decreased AMOT 130 and 80 protein levels compared to JEG3 cells kept at 20% oxygen (physiologic oxygen) (Figure 3.23). To examine if low oxygen affected AMOT subcellular distribution, I performed IF analysis of AMOT/ZO-1 and

AMOT/Phalloidin in JEG3 cells following exposure to 3% oxygen (Figure 3.24A). At 20% oxygen, AMOT is consistently present at the tight junction, where it co-localizes with ZO-1.

However, in 3% oxygen, AMOT presence at the tight junction is reduced as elucidated by less

AMOT and ZO-1 colocalization. Exposure to 3% oxygen increased AMOT localization in the cytoplasm (Figure 3.24B); however, AMOT and F-actin co-localization following 3% oxygen exposure is not as discernible as AMOT and F-actin co-localization following TGFβ treatment done in normoxia ( 20% oxygen). Nevertheless, it is interesting that the effect of low oxygen on

AMOT 130 and 80 protein levels and localization at the tight junction is similar to the effect of

TGFβ on AMOT. Considering the established regulation of TGFβ by oxygen, it is plausible that changes in oxygen tension regulate AMOT upstream of the TGFβ pathway.

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1.5

1.0 *

0.5 FoldChange

Jeg3 cells AMOT 130 Protein 0.0

20% O2 3% O2 20% O2 3% O2

AMOT 130 kDa 1.5

80 kDa 1.0 * β-actin 42 kDa

0.5 FoldChange

AMOT 80 Protein 0.0

20% O2 3% O2

Figure 3.23 Effect of low oxygen on AMOT protein levels in JEG3 cells

Representative Western blot showing AMOT 130 and AMOT 80 protein levels in JEG3 cells following exposure to 3% oxygen for 24 hours. Densitometry analysis of AMOT 130 and AMOT 80 protein levels normalized to β-actin and expressed as a fold-change relative to cells maintained at 20% oxygen (n=3). Statistical significance was assessed using non-parametric Mann-Whitney test (*p<0.05).

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A B 20% Oxygen 3% Oxygen 20% Oxygen 3% Oxygen

40X 40X 40X 40X

20% Oxygen 3% Oxygen 40X

40X 40X 40X 40X

TJ *

TJ * * AMOT ZO-1 DAPI AMOT Phalloidin DAPI

Figure 3.24 Effect of low oxygen on AMOT localization in JEG3 cells

Representative images of Immunofluorescence analysis of (A) AMOT (green) and tight junction marker ZO-1 (red) and (B) AMOT (green) and F-actin cytoskeleton probe phalloidin (red) levels following exposure to 3% oxygen for 24 hours, or maintenance at physiologic oxygen of 20% (n=3). Nuclei were stained with DAPI (blue). Exposure of JEG3 cells to 3% oxygen decreases AMOT and ZO-1 co-localization, and increases AMOT colocalization to F-actin cytoskeleton marker phalloidin. Solid white arrows indicate AMOT and ZO-1 colocalization at the tight junction (TJ), whereas dotted white arrows indicate loss of AMOT at TJ. Blue asterisks indicate AMOT co-localization with the F-actin cytoskeleton.

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3.11 Oxygen sensor JMJD6 positively regulates AMOT

Precise oxygen regulation is critical for proper placenta development and consequently it’s function. Recent work from our lab has revealed a role for novel oxygen sensor JMJD6 during human placenta development and in PE (Alahari et al., 2015; Alahari et al., 2018). JMJD6 requires oxygen to elicit its enzymatic functions; therefore, in hypoxic conditions, such as those seen in early placenta development and in PE, JMJD6 is rendered inactive. I first examined whether oxygen sensor JMJD6 elicits regulation over AMOT protein levels by loss- and gain- of function studies in JEG3 cells. Overexpression of JMJD6 resulted in an increase in AMOT 130 and AMOT

80 protein levels compared to EV controls (p<0.01) (Figure 3.25A). Following silencing of

JMJD6, AMOT 130 and AMOT 80 protein levels exhibited a trend towards decrease when compared to scrambled siRNA controls (Figure 3.25B). Considering the importance of subcellular localization when investigating scaffolding proteins, I assessed whether JMJD6 had affected

AMOT localization by IF analysis. Overexpression of JMJD6 promoted AMOT localization to the cytoplasm in fiber-like structures (Figure 3.26A). Subsequent IF analysis of phalloidin revealed

OE JMJD6 promoted co-localization of AMOT with the F-actin cytoskeleton (Figure 3.26B).

AMOT localization at the tight junction is retained following overexpression of JMJD6. These findings suggest for the first time that JMJD6 oxygen sensor is a positive regulator of AMOT polarity protein.

3.11.1 AMOT is subject to lysyl hydroxylation by JMJD6

Having established an interplay between oxygen sensor JMJD6 and AMOT, I next investigated the mechanism of this regulation. JMJD6 has dual enzymatic function as both an arginine demethylase and a lysyl hydroxylase (Chang et al., 2007; Mantri et al., 2011). Using the RF- hydroxysite in silico analysis tool that predicts probability of lysyl hydroxylation at various lysine

105

residues (Ismail et al., 2016), I assessed both AMOT 130 and AMOT 80 protein sequences.

Analysis revealed one shared lysine residue found on AMOT 130 (position 758) and AMOT 80

(position 349) with a score of 100% probability of lysyl hydroxylation (Figure 3.27). It is recognized that several other lysine residues were provided in the probability analysis report; however, none of these other lysine residues scored 100% probability of hydroxylation. To assess whether JMJD6 lysyl hydroxylates AMOT, a short peptide encompassing this high probability lysine residue was generated. Using this peptide, along with other necessary cofactors for enzymatic reactions, I performed an in vitro reaction in the presence and absence of JMJD6 enzyme. Following this in vitro reaction, analysis of the peptide mass profile in the presence and absence of JMJD6 was assessed by MALDI-TOF mass spectrometry (Figure 3.27). In absence of

JMJD6 enzyme, the primary peak is at 1130 mass units, corresponding to the mass of the peptide.

However, in the presence of the JMJD6 enzyme, the primary peak appears at 1146 mass units.

This shift in mass corresponds to the molecular mass of a hydroxyl group (16 mass units), and is not observed in the control reaction performed in the absence of JMJD6 enzyme. This finding demonstrates that JMJD6 executes lysyl hydroxylation at this residue found within AMOTs peptide sequence. Further, this may suggest that the JMJD6 regulation on AMOT we observed previously could be through lysyl hydroxylation.

To assess the contribution of lysyl hydroxylation on AMOT protein and cellular localization, I performed a preliminary experiment using the pharmacological inhibitor minoxidil, a widespread inhibitor of lysyl hydroxylation. Mechanistically, minoxidil inhibits activity of lysyl hydroxylase by drastically decreasing mRNA levels of lysyl hydroxlase 1, and to a lesser degree lysyl hydrolase

2 and 3 (Zuurmond et al., 2005). Treatment of JEG3 cells with 20 M minoxidil decreased both

AMOT 130 and AMOT 80 protein levels, compared to vehicle controls, as assessed by WB

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analysis (Figure 3.28). By IF analysis, AMOT localization in JEG3 following 20 M minoxidil treatment is reduced at the tight junctions compared to vehicle controls. (Figure 3.29). Overall, inhibition of lysyl hydroxylation elicited a reduction in AMOT protein levels and hindered AMOT localization to the tight junction. This may suggest that lysyl hydroxylation is a post translational modification that is important for AMOT’s cellular redistribution and function.

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2.5 A *

2.0

1.5 OE JMJD6 EV 1.0 0.5

AMOT 130 kDa FoldChange 0.0

AMOT 130 Protein 80 kDa EV OE JMJD6 2.0 JMJD6 50-55 kDa ** 1.5

1.0 β-actin 42 kDa 0.5

FoldChange

AMOT 80 Protein 0.0 EV OE JMJD6

B 1.5

1.0 ssRNA siJMJD6

0.5

AMOT 130 kDa FoldChange 0.0

AMOT 130 Protein ss siJMJD6 80 kDa 1.5

1.0 JMJD6 50-53 kDa

0.5 ld Change ld

β-actin 42 kDa Fo 0.0 AMOT 80 Protein ss siJMJD6

Figure 3.25 Effect of silencing and overexpressing JMJD6 on AMOT protein in JEG3 cells

(A) Representative western blot showing AMOT 130 and AMOT 80 protein levels in JEG3 cells following overexpression of JMJD6 (n=3). Densitometry analysis of AMOT 130 and AMOT 80 protein levels normalized to β-actin and expressed as a fold-change relative to scrambled siRNA (ss) control or empty vector (EV) control. Statistical significance was assessed using non- parametric Mann-Whitney test (*p<0.05 and **p<0.01). (B) Representative Western blot showing AMOT 130 and AMOT 80 protein levels in JEG3 cells following silencing of JMJD6 (n=2).

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A B

EV OE JMJD6 EV OE JMJD6

63X 63X 63X 63X

63X 63X 63X 63X TJ TJ *

* *

AMOT DAPI 63X 63X IgG TJ *

63X TJ *

AMOT F-actin DAPI

Figure 3.26 Effect of overexpressing JMJD6 on AMOT localization in JEG3 cells

Representative images of immunofluorescence analysis of (A) AMOT (green) and (B) AMOT (green) and F-actin cytoskeleton probe phalloidin (red) levels following overexpression of JMJD6 (n=3). Nuclei were stained with DAPI (blue). Normal Goat IgG was used as negative control. Overexpression of JMJD6 promoted AMOT distribution in the cytoplasm and promoted AMOT and F-actin colocalization. White arrows indicate AMOT presence at the tight junction (TJ) and blue asterisks indicate presence of AMOT fibers in cytoplasm.

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TOF/TOF™ Reflector Spec #1=>NF0.7[BP = 1130.7, 339]

100 1130.6956

B 338.9 1146

90 1130

60 No 50

Intensity % 40

30 1152.6837

20 1168.7002

1154.7073

1129.1895

1157.8268

1097.1244

1127.0179

1171.6606

1164.6920

1159.5782

1147.4136

1167.1805

1176.6860

1112.7667

1178.7712

1107.4961

1161.7686

1121.5780

1124.6862

1156.2012

1172.8276

1105.9299

1140.7609

1174.7313

1134.6899

1109.6676

1123.4436

1098.6656

1095.1165

1145.5021

1149.7592

1142.6097

10 1133.1875

1120.0658

1115.3167

1093.0830

1100.1812 enzyme 1118.4824 0 1091.0 1108.8 1126.6 1144.4 1162.2 1180.0 Mass (m /z) <> TOF/TOF™ Reflector Spec #1=>NF0.7[BP = 509.1, 613]

1146.8109 100 +16 239.9 90 + JMJD6 80 70 60

1168.8044 50

1130.7965

% Intensity % 40

1153.7958

1122.7545

1100.7706 enzyme 30 1132.4781

1124.7775

1149.8368

1145.7871

1105.7662

1093.5956

1109.7179

1113.8417

1103.7496

1126.7874

1128.8463

1136.7855

1143.3397

1173.8859

1116.8336

1111.3654

1164.8046

1098.7430

1158.8345

1141.8218

1167.8815

1108.5531

1134.7859

1091.9510

1161.8558

1148.4714

1102.4337

1175.4224

1138.7668

1097.3356

1157.5134

20 1118.1277

1176.8175

1120.5123

1178.3744 10 0 1091.0 1108.8 1126.6 1144.4 1162.2 1180.0 Mass (m /z)

TOF/TOF™ Reflector Spec #1=>NF0.7=>NR(2.00)[BP = 1130.7, 325] Magnified spectra

1130.6956 100 324.7

90 1130 80 70

1146 60 No 50

% Intensity % 40 30 20

1129.1903

enzyme 1127.0184

1147.4176

1141.1902

10 1133.1902 1140.7538

0 1127 1132 1137 1142 1147 1152 Mass (m /z) <> TOF/TOF™ Reflector Spec #1=>NF0.7=>NR(2.00)[BP = 509.1, 595]

+16 1146.8101 100 229.9 + JMJD6 90 80 1147.8119 70 60 enzyme 50

1130.7944

% Intensity % 40

30 1131.8282

1132.4794

1136.7836

1145.7876

1128.8479

1143.3438

1148.4948

1141.8148

1129.7677

1138.7533 1150.1907 20 1136.3610

1131.1309 10 0 1127 1132 1137 1142 1147 1152 Mass (m /z)

Figure 3.27 MALDI-Mass Spectrometry analysis of AMOT peptide mass profile following in vitro JMJD6 enzyme reaction

(A) Results from RF-Hydroxysite in silico analysis showing peptide sequence window on AMOT 130 and AMOT 80 with 100% probability of lysyl hydroxylation. (B) Mass spectrometry profiles of AMOT peptide following in vitro reaction in the presence and absence of JMJD6 enzyme (n=3).

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1.5

1.0 **

0.5

Fold Change Fold V minoxidil V minoxidil 0.0 130 Protein AMOT AMOT 130 kDa Vehicle Minoxidil

1.5 80 kDa

** Β-actin 42 kDa 1.0

0.5 Fold Change Fold 0.0

Protein 80 AMOT Vehicle Minoxidil

Figure 3.28 Effect of minoxidil induced inhibition of lysyl hydroxylation on AMOT protein levels in JEG3 cells

Representative Western blot showing AMOT 130 and AMOT 80 protein levels in JEG3 cells following treatment 20 M minoxidil for 48 hours (n=3). Densitometry analysis of AMOT 130 and AMOT 80 protein levels normalized to β-actin and expressed as a fold-change relative to vehicle (V) control (ETOH). Statistical significance was assessed using non-parametric Mann- Whitney test (**p<0.01)

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Vehicle + minoxidil

40X 63X 40X 63X

AMOT

40X TJ 63X 40X 63X

TJ Merge

AMOT DAPI IgG 63X

Figure 3.29 Effect of minoxidil induced inhibition of lysyl hydroxylation on AMOT localization in JEG3 cells

B) Representative images of immunofluorescence analysis of AMOT (green) following treatment with 20 M minoxidil for 28 hours (n=3). Nuclei were stained with DAPI (blue). Normal Goat IgG was used as negative control. Inhibition of lysyl hydroxylation by minoxidil reduces AMOT localization at the tight junction. Solid white arrows indicate AMOT presence at the tight junction (TJ), and dotted white arrows indicate loss of AMOT at TJ.

Chapter 4 Discussion

Discussion and Conclusions

The present study reveals a spatial and temporal regulation of AMOT polarity protein during early human placenta development, and demonstrates that the AMOT 80 isoform functions as a positive regulator of cell migration in JEG3 cells. Furthermore, in vitro findings indicate AMOT protein levels and localization are regulated by changes in oxygen and TGFβ signalling. Specifically,

TGFβ dependent redistribution of AMOT 130 to the F-actin cytoskeleton is mediated by Smad- dependent TGFβ signalling. I have further established a novel interaction between AMOT and polarity protein Par6, a component of the Smad-independent TGFβ signalling pathway that regulates tight junction dissolution and loss of cell polarity. This investigation has also revealed for the first time that AMOT 130 and AMOT 80 are regulated by the placenta oxygen sensor

JMJD6, potentially via JMJD6-mediated lysyl hydroxylation. Finally, this study reveals AMOT is disrupted in PE, a devastating pregnancy related disorder characterized by chronic hypoxia and aberrant TGFβ signalling. This is identified by reduced AMOT 130 and 80 protein levels and absent AMOT localization, particularly in EVTs at the fetomaternal interface.

4.1 General Discussion

Protein levels of AMOT 130 and 80 isoforms are both significantly elevated in placentae from 10-

15 weeks of gestation, compared to 5-9 weeks of gestation. AMOT has an established role in angiogenesis, and considering that the placenta becomes increasingly vascularized as gestation progresses, this could account for the temporal increase in AMOT protein levels. However, 10-12 weeks of gestation is the time frame for opening of the intervillous space and accompanying

112 113

increase in partial pressure of oxygen, a critical physiological step during placental development known to dictate trophoblast differentiation events (Ietta et al., 2006; Rodesch et al., 1992). Thus, the apparent increase in AMOT 130 and AMOT 80 protein levels around 10-weeks of gestation could be attributed to changes in oxygen tension, in line with data presented here showing that low oxygen downregulates AMOT protein expression. Spatial examination of AMOT in the human placenta tissue reveal that all trophoblast layers serve as a cellular source of AMOT. In the floating villi at 6-weeks of gestation, AMOT is predominantly localized to the progenitor cytotrophoblast cells; at 8-weeks of gestation, AMOT transitions to both cytotrophoblast and multi-nucleated syncytium; and at 13-weeks of gestation, AMOT is predominantly at the syncytiotrophoblast apical brush border. This temporal localization pattern of AMOT in these two functional layers of the floating villi suggest a potential role for AMOT in the fusion of cytotrophoblast into the overlaying syncytium, a process that is prevalent during this gestational window. Considering the established role of AMOT 130 and AMOT 80 in regulating apical-basolateral cell polarity

(Sugihara-Mizuno et al., 2007; Wells et al., 2006), its localization to the apical brush border of the syncytium may indicate an asymmetric distribution of proteins, lipids or transporters that contributes to unpolarized nature of this multi-nucleated specialized epithelial layer. Work from our group established polarity protein Par6 to be a negative regulator of trophoblast cell fusion by preserving tight junction integrity and controlling cytoskeletal dynamics (Sivasubramaniyam et al., 2013), revealing for the first time that polarity proteins are important in regulating trophoblast differentiation events. At the level of the anchoring villi, proliferative cytotrophoblasts residing at the proximal base of the anchoring column differentiate into migratory and invasive extravillous trophoblasts (EVT) that localize to the intermediate and distal regions, respectively. Our observation of selective AMOT localization to the intermediate and distal portions of the anchoring column suggest a potential role for AMOT in the migration and/or invasion of these trophoblasts.

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However, besides trophoblast cells, another cellular source of AMOT are the mesenchymal cells localized to the stroma of the chorionic villi. Considering that placental mesenchymal cells have been described to be involved in a cross-talk with overlying trophoblast cells via the exchange of specific growth factors (Chen et al., 2013; Lacey et al., 2002), it is possible that pMSC-derived

AMOT can regulate trophoblast cell polarity in a paracrine manner also.

A PCR array for multiple genes known to be involved in epithelial-mesenchymal-transition (EMT) was used to compare gene expression in primary isolated cytotrophoblast and EVT cells, and confirmed that the differentiation of first trimester CT to EVT is analogous to EMT (DaSilva-

Arnold et al., 2015). The general EMT processes requires the loss of junctional contacts between cells in order to remodel into a mesenchymal phenotype that is able to migrate away from original tissue (Kalluri and Weinberg, 2009). The localization of AMOT to the migratory and invasive population of EVTs, and the disruption of RhoA at the tight junctions following AMOT 130 and

80 overexpression further supports that AMOT may be involved in the EMT process of trophoblast differentiation. As anticipated, overexpression of AMOT 130 increased protein levels of this isoform; however, it is also surprisingly augmented protein levels of AMOT 80. Dissimilarly, overexpression of AMOT 80 did not affect protein levels of AMOT 130. As such, we can postulate that AMOT 130 may undergo replication slippage due to a downstream start site resulting in an

AMOT 80 product. Considering this, it is possible that the downregulated levels of RhoA observed following overexpression of AMOT 130 are not specific to this isoform, and AMOT 80 may be contributing to this negative regulation. Bearing in mind slippage events occur in regions of tandem repeats at the replication start site, a mutation event could be introduced in this region to decipher whether slippage could account for increased AMOT 80 protein levels following AMOT

130 overexpression, and to test if AMOT 130 has an independent role in regulating RhoA.

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The work described in this study establishes AMOT as a regulator of cell migration in the placenta, as trophoblast derived JEG3 cells overexpressing AMOT 80 migrated 2x faster than empty vector controls. These findings are in line with studies in endothelial cells which performed similar time- lapse imaging studies and determined overexpression of AMOT 80 significantly increased the migration rate (um/hr) compared to empty vector controls (Levchenko et al., 2004). Furthermore, studies in MDCK epithelial cell line revealed that AMOT 80 overexpression promoted a migratory phenotype characterized by extensions and lamellipodia whereas expression of AMOT 130 did not alter the cell phenotype (Ernkvist et al., 2008). An established mechanism by which AMOT

80 selectively promotes epithelial cell migration is via redistribution of components of the Par and

Crumbs complex that disrupts cell polarity and induces loss of tight junction integrity (Wells et al., 2006). The present study demonstrates the dynamic redistribution of AMOT 80 in migrating

JEG3 cells through the use of an AMOT 80-YFP construct in time-lapse imaging where AMOT is observed to redistribute between the tight junction and in punctate structures in the cytoplasm.

Further we show that OE AMOT 80, as well as AMOT 130, promoted decreased levels of RhoA

GTPase, a known G-protein that stabilizes intracellular tight junctions and polarity (Ozdamar et al., 2005; Wang et al., 2006), thereby implicating AMOT 80 in altering JEG3 cell polarity amidst its promotion of cell migration. The predominate tight junction localization of AMOT seen here is consistent with what is reported in the literature in polarized epithelial cells, which reveal that the tight junction localization of AMOT is contingent on its C-terminal PDZ motif (Wells et al.,

2006).

TGFβ is an integral regulator of EMT processes during development, as the addition of TGFβ to epithelial cells in culture is accepted as a convenient way to induce EMT in various epithelial cells

(Xu et al., 2009). Furthermore, TGFβ has been shown to promote trophoblast cell differentiation,

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namely migration, and thus possesses a fundamental role in the human placenta (Xu et al., 2016).

To my knowledge, the present study is the first to establish TGFβ as a regulator of AMOT protein levels and localization. TGFβ1/3 treatment promoted dissolution of tight junctions in JEG3 cells as indicated by the intermittent deposition of ZO-1 at cell boundaries, which is consistent with the effect of TGFβ on tight junctions in kidney epithelial cells, and confirms the presence of an EMT phenotype (Varelas et al., 2010). More notably, however, TGFβ1/3 treatment induced AMOT redistribution from tight junctions in JEG3 cells and association to the F-actin cytoskeleton. At the tight junction, it is established that AMOT regulates tight junction integrity via interactions with

GTPase activating protein Rich1, as well as with Crumbs polarity complex components Pals1 and

Patj and Par Complex protein Par3 (Wells et al., 2006). AMOT 130 is the preferential isoform bound to F-actin, and this interaction is implicated both in the negative and positive regulation of

YAP/TAZ, which mediate transcription genes involved in proliferation and EMT (Chan et al.,

2011; Hong, 2013; Hsu et al., 2015; Lv et al., 2016; Moleirinho et al., 2017). Aside from sequestration of YAP/TAZ that impacts Hippo pathway signalling, the interaction between AMOT

130 and F-actin also facilitates changes in cell shape, integrity of tight junctions and actin remodeling; all of which promote cell motility events (Ernkvist et al., 2006; Gagne et al., 2009;

Quiros and Nusrat, 2014). Considering this, it is plausible that in trophoblast cells, TGFβ promotes disruption of tight junctions and changes in cell shape via a direct regulation on AMOT subcellular redistribution, and this contributes to EMT or cell migration processes. Interestingly, TGFβ treatment resulted in an overall decrease in AMOT 130 and 80 protein levels. Taking into account the reduction in AMOT protein levels, it is possible that in addition to promoting binding to the F- actin cytoskeleton, TGFβ promotes AMOT localization to the cytoplasm for subsequent degradation. Studies have revealed a novel lipid binding domain on AMOT’s protein structure that targets it along with associated apical polarity proteins to the endosomal recycling compartment

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(Heller et al., 2010b; Wells et al., 2006). Thus, it is plausible that TGFβ -mediated re-localization of AMOT to the cytoplasm results in the degradation of AMOT and associated polarity components, further contributing to alterations in cell polarity. The current study demonstrates that

TGFβ1/3 mediated redistribution of AMOT in JEG3 cells is reliant on Smad-dependent signalling, as the inhibition of alk5 receptor prevented AMOT localization to the cytoplasm and promoted its retention at the tight junction. Notably, SB431542 is a selective inhibitor of alk5 mediated TGFβ signalling via Smad-proteins and has no effect on other signal transduction pathways whose activities are dependent on concerted activation of multiple kinases ERK, JNK or p38 MAP kinase pathways (Inman et al., 2002). Further, alk5 inhibition with SB-431542 restored AMOT 80 protein levels, and even increased AMOT 130 protein levels, suggesting that the TGFβ mediated reduction in AMOT protein is also via Smad-dependent TGFβ signalling. Previous studies have reported

JEG3 cells to be resistant to the effects of TGFβ as a mechanism to escape normal regulatory control of cell invasion (Graham et al., 1994). Specifically, JEG3 cells failed to show the normal

Smad3 signalling events following TGFβ treatment as a result of poor Smad3 expression (Xu et al., 2001). However, more recent studies have shown JEG3 cells are responsive to TGFβ exposure through Smad2 signalling(Xu et al., 2016), which are confirmed by the present investigation.

Altogether, these findings uncover a novel regulation of AMOT by Smad-dependent TGFβ signaling likely via a mechanism that involves Smad2.

As previously discussed, AMOT interaction with specific members of the Par and Crumbs complex, as well as other tight junction components, is implicated in cell polarity regulation. These protein-protein interactions are typically mediated by C-terminal PDZ binding domains and in some instances N-terminal coiled-coil domains, and are further regulated by Rho family GTPases and serine/threonine phosphorylation (Macara, 2004). For instance, the C-terminal PDZ binding

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domain on AMOT is critical to its interaction with Pals1/Patj polarity complex and its localization at the tight junction, whereas the coiled-coil/BAR domain promotes binding to GTPase activating protein Rich1, which promotes its apical localization and selective re-localization of Pals1 and

Par3 to early endosomal compartments (Sugihara-Mizuno et al., 2007; Wells et al., 2006). Despite these well-characterized studies on AMOT interaction with polarity complexes, the direct interaction between AMOT and Par6, an established cell polarity regulator in trophoblast cells

(Sivasubramaniyam et al., 2013; Xu et al., 2016), has never been investigated. Similar to AMOT,

Par6 polarity protein is also under the regulation of TGFβ signalling via a non-canonical signalling cascade that induces degradation of RhoA at the tight junction and leads to tight junction dissolution (Ozdamar et al., 2005; Wang et al., 2006). Furthermore, Par6, similar to other Par constituents, contains a PDZ binding domain and BAR/coiled coil domain within its protein structure (Assemat et al., 2008). Here, we reveal a novel interaction between AMOT and Par6, and show that this interaction is upregulated by TGFβ1/3 treatment. Immunofluorescence analyses revealed that TGFβ1/3 not only upregulated AMOT/Par6 co-localization, but shifted their association primarily to the cytoplasm. Considering Par6 presence at tight junction is required for tight junction maintenance and integrity, presence of AMOT/Par6 association within the cytoplasm following TGFβ1/3 treatment suggests an alteration in tight junction integrity. Similar to AMOT’s ability to recruit Pals1/Patj/Par3 away from the tight junction and into the endosome

(Heller et al., 2010a; Wells et al., 2006), it is plausible that AMOT effectively sequesters Par6 protein away from the tight junctions and contributes to TGFβ-mediated dissolution of tight junctions and loss of apical and basolateral cell polarity. In the developing placenta, TGFβ acts in a complex manner because it stimulates opposing functions in different trophoblast populations.

This can be attributed to different TGFβ signalling pathways being activated in a cell-specific manner throughout development. Establishing a role for AMOT in both the Smad-dependent and

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Par6-mediated TGFβ signalling pathways further underscores the complexity of TGFβ in trophoblast cell populations and suggests a role for AMOT in various trophoblast functions.

Interestingly, Par6 interacts preferentially with the 80 kDa isoform of AMOT, as co- immunoprecipitation studies revealed no association of Par6 with AMOT 130. This selective interaction is also seen in other studies which demonstrated AMOT 80 to be the predominant

AMOT isoform involved in interactions with Pals1 and Patj (Wells et al., 2006). Considering that the present study and others have emphasized the role of AMOT 80 in cell migration versus the role of AMOT 130 in cell shape alterations (Ernkvist et al., 2006; Ernkvist et al., 2008), it is possible that AMOT/Par6 interaction is primarily involved in promotion of cell migration via cell polarity changes. Analysis of the binding domains required for this newfound interaction revealed the PDZ domain on AMOT 80 to be important for Par6 binding, as elucidated by their reduced association following over expression of AMOT 80 with a deletion mutation in the last 3 amino acids in the PDZ domain. This is in line with studies that show mutations in C-terminal PDZ domain of AMOT 80 impairs its ability to localize to the tight junction proteins such as MUPP1

(Sugihara-Mizuno et al., 2007), and impairs the interaction of AMOT 80 with the Pals1/Patj/Par3 complex (Wells et al., 2006). Further, the PDZ binding domain on AMOT 80 has proven to be essential for its function in cell migration, as cells expressing PDZ mutant form of AMOT exhibited defective migration (Levchenko et al., 2003). Additional investigation is required to determine whether disruption of AMOT 80/Par6 interaction via deletion within the PDZ binding domain impedes cell polarity and downstream migration of these cells. Nonetheless, this study reveals a novel binding partner for AMOT 80 at the putative PDZ binding domain.

Changes in oxygen tension and TGFβ signalling are intricately linked during placenta development. Specifically, work from our lab has shown that in conditions of hypoxia such as that

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seen in early pregnancy, hypoxia-inducible factor 1-alpha (HIF-1) mediates trophoblast differentiation events via regulation of TGFβ3 (Caniggia et al., 2000). In line with the negative effect of TGFβ1/3 treatment on AMOT protein levels and localization in JEG3 cells, exposure of

JEG3 cells to hypoxia (3%) reduces protein levels of AMOT 130 and AMOT 80. These similar findings suggest that upstream or potentially independent of TGFβ signalling, AMOT is also regulated by oxygen changes. No previous studies have looked at the impact of hypoxia on AMOT

130 or AMOT 80 expression; however, another motin family member AMOTL2 has been reported to be upregulated by hypoxia. A study looking at the effect of low oxygen on the transcriptome in early human placental villi showed upregulation of several genes involved in angiogenesis following short exposure to hypoxia (2% O2), one of them being AMOTL2 (Mondon et al., 2005).

A more recent study revealed that exposure of HeLa cells to hypoxia (2-0.1% O2) for 8 hours increased protein and mRNA levels of a spliced form of AMOTL2 (Mojallal et al., 2014). Further, hypoxia induced expression of AMOTL2 was shown to be associated with disruption of apical- basal cell polarity and promotion of tumour cell invasion in colon and breast cancer (Mojallal et al., 2014). Firstly, these studies focused on AMOTL2. Thus, it is plausible that these effects are not transferable to other motin proteins, such as AMOT 130 or AMOT 80. Secondly, these studies examined the effect of hypoxia on AMOTL2 in the context of cancer, where hypoxia is typically correlated with hyper-invasion and metastases. In the placenta, however, chronic hypoxia is typically associated with hypo-invasion of trophoblast cells, as highlighted in the placenta pathology PE (Caniggia and Winter, 2002; Genbacev et al., 1996). Considering this, the upregulation of AMOTL2, or other motin proteins, observed in these other hypoxic studies, may not be seen in the hypoxic placentae where migration and invasion is known to be impaired.

Further investigation into the other motin proteins during human placenta development, and in hypoxic conditions could address these concerns. Nonetheless, the present findings show that

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exposure of JEG3 cells to 3% O2 promotes downregulation of AMOT 130 and AMOT 80 protein levels and redistribution from the tight junction to the cytoplasm. Considering this, the presence of low oxygen in the early developing placentae could contribute to the downregulation of AMOT

130 and 80 seen in early gestation.

In the human placenta, novel oxygen sensor JMJD6 responds to changes in oxygen, whereby it is paradoxically upregulated by hypoxia (Alahari et al., 2015). JMJD6 has been shown to affect diverse cellular processes by promoting alterations in gene transcription and/or protein stability via its two main enzymatic functions: lysine hydroxylation and histone demethylation (Alahari et al., 2015; Alahari et al., 2018; Chang et al., 2007; Mantri et al., 2011; Webby et al., 2009). For instance, in breast cancer cells, JMJD6 was found to promote transcription of genes involved in cell proliferation, as well as migration and invasion (Lee et al., 2012; Poulard et al., 2015). In this study, I show through loss and gain of function studies that AMOT 130 and 80 protein levels are positively regulated by JMJD6 protein. Further, JMJD6 overexpression results in both AMOT localization to the cytoplasm, and maintenance of AMOT at the tight junction. Considering that

JMJD6 requires molecular oxygen to remain enzymatically active, it is possible that the hypoxia- mediated downregulation of AMOT protein levels seen here is a result of non-functional JMJD6, and as such JMJD6 mediates the effects of oxygen on AMOT. In silico analysis of AMOT protein sequence revealed a lysine residue with 100% probability of being lysyl hydroxylated, suggesting

AMOT as a potential target for JMJD6-mediated lysyl hydroxylation. An identified limitation of this prediction software was changing the length of the amino acid sequence window parameter yielded differences in the probability values for the lysine residues. However, through MALDI-

TOF mass spectrometry, I revealed that this specific predicted lysine residue (located at amino acid position 758 on AMOT 130 and 349 on AMOT 80) was in fact a substrate for JMJD6-

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mediated lysyl hydroxylation. Analysis of the characterized domains on AMOT reveal that this particular lysine residue resides in a linker region between the coiled-coil/BAR domain and the angiostatin binding domain on both AMOT isoforms. Further investigation of other high probability lysyl residues within AMOT protein sequence for potential JMJD6-mediated lysyl hydroxylation is warranted to address the possibility that JMJD6 hydroxylates AMOT at multiple lysine residues. Nonetheless, these findings demonstrate a novel regulation of AMOT 130 and 80 by JMJD6, and suggest for the first time that this could occur post-translationally through lysyl hydroxylation.

Post translational modifications of AMOT have only recently begun to be uncovered, thus a lot remains unknown regarding the post translational regulation of AMOT protein. Omics-type approaches identified modifications of AMOT to include phosphorylation, ubiquitination, acetylation and glycosylation (Moleirinho et al., 2014). The ability of post translational modifications to impact AMOT localization and function is demonstrated by studies showing how phosphorylation of AMOT 130 at serine 175 via LATS1/2 kinases inhibits AMOT binding to the

F-actin cytoskeleton, impairs cell migration and angiogenesis, whilst freeing it to interact with

Hippo pathway activators such as YAP and TAZ (Dai et al., 2013; Hirate et al., 2013). Another post-translational modification that has been studied is ubiquitination of AMOT 130 at lysine 481 by Atrophin-1 interacting protein 4 (AIP4), an E3 ligase, which targets AMOT 130 for 26S proteasomal degradation (Adler et al., 2013; Wang et al., 2012). The present study deems JMJD6- mediated lysyl hydroxylation important for AMOT stability, as addition of the lysyl hydroxylase inhibitor minoxidil reduced total AMOT protein levels and altered deposition of AMOT fibers in the cytoplasm. It is important to note that minoxidil promotes wide-spread inhibition of lysyl hydroxylase activity; therefore, one limitation is that its effects are not specific to JMJD6 mediated

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lysyl hydroxylation. To properly assess the consequence of inhibiting lysyl hydroxylation solely by JMJD6, the JmjC catalytic domain on JMJD6 that permits its lysyl hydroxylase and arginine demethylase activities (Kwok et al., 2017; Tsukada et al., 2006) could be mutated, and AMOT protein levels and localization could be assessed. This would reveal whether regulation of AMOT by JMJD6 occurs by its canonical enzymatic functions. This is important to distinguish because recent studies have found a novel function for JMJD6 that is independent from its canonical enzymatic activities, where it is secreted as a soluble protein that can functionally interact with extra cellular matrix components and interfere with their assembly (Miotti et al., 2017).

Furthermore, to strengthen the finding that hydroxylation of AMOT occurs primarily at the lysine

758 (AMOT 130) and lysine 349 (AMOT 80) residues, an AMOT peptide with a point mutation at these outlined lysine residues could be generated and the same in vitro JMJD6 hydroxylation and MALDI-TOF mass spectrometry could be performed. Additionally, it could be of great value to digest endogenous immunoprecipitated AMOT protein into small peptide fragments, rather than using a synthesized peptide sequence, and the resultant peptide fragments could then be assessed by MALDI mass spectrometry. Nonetheless, these findings reveal AMOT as a novel potential substrate for JMJD6-mediated lysyl hydroxylation.

PE is a serious disorder of pregnancy, is characterized by inadequate trophoblast differentiation leading to insufficient EVT invasion of spiral arties and impaired trophoblast turnover (Arnholdt et al., 1991; Naicker et al., 2003). In PE, I show a marked reduction in AMOT 80 and AMOT 130 protein levels, as well as disrupted localization of AMOT in EVT cells residing at the fetomaternal interface compared to pre-term age-matched controls. Considering the established role for AMOT

80 in cell migration presented in this study and others, and changes in cell shape mediated through

AMOT 130, it is possible that the drastic reduction of total AMOT impacts EVT ability to migrate

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and invade the uterine tissue and transform the spiral arteries and establish adequate blood flow to the fetus. Furthermore, disruption of the newfound interaction between AMOT and Par6 in PE could also exacerbate the immature preeclamptic phenotype. I have proposed that the collective

AMOT and Par6 re-localization to the cytoplasm may be contributing to cell migration, as well as promoting the endosomal recycling of Par6 polarity protein in order to induce changes in apical- basolateral cell polarity. Thus, as a consequence of reduced AMOT and Par6 interaction, Par6 is free to elicit its regulatory function at the tight junction. Work from our lab has established Par6 as a negative regulator of cytotrophoblast cell fusion by stabilizing intracellular tight junctions, and further found that its elevated expression in PE could be contributing to the aberrant trophoblast fusion and cell turnover that is typical of this pathology (Sivasubramaniyam et al.,

2013). Thus, elevated amounts of Par6 in PE could be attributed to reduced interaction with the

AMOT scaffolding protein, leading reduced Par6 degradation.

Persistence of low oxygen, and aberrant TGFβ signalling are molecular characteristics of preeclampsia (Caniggia et al., 1999; Caniggia et al., 2000; Xu et al., 2016). Specifically, in vivo analysis of placental explants from PE revealed significantly elevated levels of TGFβ3 (Caniggia et al., 1999). The present data demonstrating reduced AMOT protein levels following treatment with TGFβ1/3 and exposure to 3% O2 are consistent with the reduction of AMOT seen in PE. This suggests that the impaired AMOT levels and localization in PE could be a result of prolonged hypoxia and aberrant downstream TGFβ signalling, both of which have been shown to contribute to the immature phenotype of trophoblast cells that is implicated in the pathogenesis of this disease.

Furthermore, JMJD6 enzymatic function is impaired in PE as a result of lowered oxygen availability and reduced intracellular iron availability (Alahari et al., 2015; Alahari et al., 2018).

Thus, impaired JMJD6 regulation of AMOT in PE could account for disrupted AMOT protein

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levels and altered distribution in EVTs. In the future, measurement of AMOT lysyl hydroxylation in PE versus age-matched controls could be conducted to investigate this mechanism further.

The functional consequences of reduced or abrogated AMOT expression in the placenta remain undetermined. Investigations using Amot knockout mice have been conducted, where mice display embryonic lethality at E11-E11.5, as well as produce smaller sized embryo and placentae when measured at E10.5 (Aase et al., 2007). Notably, embryonic days E9-12 during mouse pregnancy is critical for embryonic growth and viability because it is the timeframe for when placenta development is undergoing completion. Thus, these studies are consistent with a potential role for

AMOT in the placenta. It should be noted that the embryonic lethality in Amot knockout mice is sensitive to genetic background, as crosses of mice with specifically 129/SvEv background lead to death of Amot-deficient embryos at E7.5 (Shimono and Behringer, 2003). Initial morphological characterization of Amot-deficient mice placentae by H&E histology revealed no obvious structural differences between Amot deficient and wild-type mice placentae (Aase et al., 2007).

However, further in-depth investigation beyond H&E staining of whole mouse placentae, and investigations using in vivo models of human placenta such as placental explants, are necessary to adequately assess the contribution of reduced AMOT to the developing placenta.

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4.2 Conclusions

AMOT plays a fundamental role in regulation of cell polarity, changes in cytoskeletal rearrangements, and promotion of cell migration in both endothelial and epithelial cells. During human placenta development, differentiation of cytotrophoblast cells into migratory and invasive

EVT is critical for proper invasion of the maternal decidua and establishment of utero-placental blood flow via remodeling of the spiral arteries. However, the contribution of cell polarity regulators to this differentiation process in the placenta remains poorly understood. This study is the first to establish functional importance for AMOT in the human placenta and trophoblast derived JEG3 cells, specifically its role in cell migration and interaction with Par6 polarity protein, and its regulation by oxygen and TGFβ. Altogether this study adds to the current knowledge of the characterized regulatory processes that guide trophoblast cell differentiation in the human placenta, and contributes to our understanding of the disrupted biological processes that contribute to pathogenesis of preeclampsia.

My first objective was to establish a temporal and spatial expression pattern of AMOT in the early human placenta. I systematically addressed this by mapping both isoforms of AMOT, AMOT 130 and AMOT 80, in whole placenta tissue lysates as well as in sections of human placenta from 5-

15 weeks of gestation. I determined that AMOT 130 and 80 protein levels increase with advanced gestation, as well as AMOT localized to the polarized cytotrophoblast cells and migratory/invasive

EVTs in the intermediate and distal portions of the anchoring villi in the early placenta.

Furthermore, through the use of JEG3 choriocarcinoma cell line which mimic features of EVT cells, I established both AMOT 80 and AMOT 130 to be endogenously expressed and further revealed a role for AMOT 80 in promoting JEG3 cell migration.

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My second objective was to establish if TGFβ signalling and changes in oxygen, two fundamental regulators of early trophoblast differentiation, exerted regulatory control over AMOT. I determined that TGFβ1/3 act as regulators of AMOT abundance and subcellular localization in

JEG3 cells, and this occurs via the Smad-dependent TGFβ pathway. As a scaffolding protein, the localization of AMOT is of paramount importance and often indicative of its function. I demonstrated Par6 as a novel interacting partner for AMOT, and further determined the putative

PDZ binding domain to be important for this newfound interaction. Furthermore, levels of oxygen mediated through JMJD6 oxygen sensor positively regulated AMOT protein levels, and also affected its subcellular redistribution. A mechanism by which this occurs was demonstrated to be

JMJD6-mediated lysyl hydroxylation, revealing a new post translation modification on AMOT that could distinguish its function in the placenta.

My final objective was to investigate AMOT in the placenta pathology preeclampsia, a disease characterized by impaired oxygen sensing, persistence of hypoxia and aberrant TGFβ signalling.

I determined that protein levels of AMOT are reduced in PE, and its localization in invasive EVTs is disrupted. I propose that reduced AMOT seen in PE contributes to the impaired trophoblast migration seen in this disease, as this study highlights the importance of AMOT in the process of cell migration. A putative model summarizing the present findings on the regulation and role of

AMOT in the context of normal and pathological pregnancy is demonstrated in Figure 4.1.

128 Normal Placentation Preeclampsia

Smad AMOT ↓O TGFβ1/3 2

AMOT Par6 ↑TGFβ3 F-actin non- Smad O JMJD6 2 TGFβ1/3 JMJD6

(inactive)

↑ AMOT ↓ AMOT Par6

Trophoblast Cell Impaired Trophoblast Cell Migration Migration

Figure 4.1 Putative model depicting the role and regulation of AMOT in normal placentation and in preeclampsia.

In trophoblast cells during normal placentation, AMOT redistribution to the F-actin cytoskeleton is mediated by the Smad-dependent TGF pathway. Simultaneously, TGF stimulates a novel interaction between AMOT and polarity protein Par6, an established component of the Smad independent/Par6 mediated TGF pathway that regulates tight junction dissolution and cell migration. Most importantly, JMJD6 is enzymatically active in the presence of oxygen and positively regulates AMOT expression. Altogether, these events favor AMOT’s role in promoting migration of trophoblast cells. In PE, however, the prescience of chronic hypoxia upregulates levels of TGF3 to subsequently downregulate AMOT protein levels. Also, the interaction between AMOT and Par6 is disrupted. Most notably, the lack of molecular oxygen in PE renders JMJD6 inactive and unable to regulate AMOT. This further contributes to the downregulation of AMOT, favoring the impaired trophoblast migration phenotype seen in PE.

Chapter 5 Future Directions

Future Directions

The present study is the first to systematically investigate AMOT in the developing human placenta, particularly the role of AMOT in controlling trophoblast cell migration. The use of JEG3 choriocarcinoma cells provided insight into EVT cell behaviour, as they closely resemble EVT cellular phenotype and are an established model used to study events linked to proliferation, migration and invasion (Huang et al., 2017). However, investigation of AMOT using in vivo cell models of such as placental explants, could significantly strengthen the present findings.

Furthermore, to properly assess AMOTs contribution to human placenta development, investigations should not be limited to solely trophoblast cells. Another cell type in the human placenta that is fundamental to proper development, despite them being presently understudied, is the placental mesenchymal cells (pMSC). These pMSC cells are located in the stromal core of floating villi, and contribute to healthy placental development by aiding in vascularization of the villi, and more notably through their role in regulating trophoblast invasion into the maternal decidua (Chen, 2014). Additionally, this study primarily focused on AMOT interactions in the context of cell polarity regulation and its implication on cell migration, however it is clear AMOT plays critical role as a regulator of the Hippo pathway, particularly in the sequestration and binding of TAZ transcription factor (Chan et al., 2011; Hirate et al., 2013; Piccolo et al., 2014). Considering that the Hippo pathway also controls cell phenotype and differentiation events, it would be worthwhile to investigate the contribution of the AMOT/TAZ axis in trophoblast cells.

Additionally, we implicated JMJD6 as a novel regulator of AMOT through lysyl hydroxylation, however JMJD6 also functions as a histone arginine demethylase to regulate gene transcription

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(Chang et al., 2007). Investigation into whether JMJD6 elicits its histone demethylation capabilities on AMOT could further strengthen JMJD6-mediated regulation on AMOT. Lastly, this report proposes AMOT binding to the endosomal recycling compartment as a potential mechanism in which it regulates cell polarity in trophoblast cells. Interestingly, our lab has demonstrated autophagy and lysosomal degradation as cellular processes important for proper human placenta development, and upregulated in the placenta pathology preeclampsia (Ermini et al., 2018; Melland-Smith et al., 2015). Thus, investigation into AMOT’s degradation via these pathways in physiological and pathological placenta could further establish the importance of

AMOT in the placenta.

5.1 Using in vivo villous explants to assess AMOT’s role in trophoblast cell differentiation

In vivo placental explants reflect the uterine environment more faithfully than the current in vitro cell lines, and as such represent a useful model to investigate EVT differentiation events including migration and invasion (Aplin, 2000; Caniggia et al., 2000; Caniggia et al., 1997; Miller et al.,

2005). Use of placental explants from early gestation provides an opportunity to translate our in vitro findings and further establish a role for AMOT in EVT migration and invasion that occurs in anchoring villi during first trimester of gestation. I propose isolation of villous trees from first trimester human placentae, and culturing them in supplemented DMEM media containing Matrigel coated dish inserts, as previously described (Caniggia et al., 1997). Considering villous explants are amenable to treatment with small molecular inhibitors or antisense-induced inhibitors

(Caniggia et al., 1997; Chauvin et al., 2015), I propose to selectively inhibit both AMOT isoforms using antisense inhibitors against AMOT and assess its impact on EVT differentiation and outgrowth. This would include morphological analysis by assessing flattening of the distal ends of villous trips, adherence to Matrigel and appearance of EVT breakthrough from the villous tips

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through immunostaining, as described by Genbacev et al. (Genbacev et al., 1992). It could also entail quantification of specific epithelial and mesenchymal markers in explants considering how

EVT differentiation from proliferative to invasive has been characterized as an EMT process

(DaSilva-Arnold et al., 2015). E-cadherin, a transmembrane protein involved in cell-cell adhesion; epidermal growth factor receptor (EGFR); and occludin, tight junction stabilizing protein are markers of epithelial cells and thus could be measured to indicate proliferative cytotrophoblast cells. On the other hand, mesenchymal markers including fibronectin, an extracellular matrix protein implicated in cell adhesion and migration; matrix metalloproteinases, enzymes responsible for degradation of extracellular matrix proteins; and vimentin, a mesenchymal intermediate filament protein could be measured as an indices of trophoblast differentiation towards a migratory and invasive phenotype. If there is a differential effect on EVT differentiation following inhibition of AMOT, mechanistic studies could be performed to elucidate how AMOT is regulating this process. Similar to the present study, AMOT interaction with other polarity proteins and F-actin cytoskeleton could be assessed determine if any defect in EVT differentiation is a result of impaired cell polarity or cytoskeletal rearrangements.

Cultured primary isolated trophoblast cells from first trimester placenta are another favourable biological model; however, it has been established that these cells undergo spontaneous fusion in culture over time and thus are unable to be passaged and maintained in culture for extended periods

(Li and Schust, 2015). Consequently, the timeframe to study polarized, cytotrophoblast events, such as migration and invasion, are limited. The present study remained focused on extravillous differentiation events; however, fusion of cytotrophoblasts to form the functional syncytium is also vital to proper placenta development. Utilizing the fact that primary isolated trophoblast cells spontaneously syncytialize, we could also establish whether AMOT plays a role in the fusion of

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cytotrophoblast into syncytiotrophoblasts. For instance, AMOT expression could be silenced in primary isolated trophoblast cells grown in culture for up to 72 hours, and the effect on syncytialization could be assessed by measuring levels of syncytin-1, an established fusogenic marker, as well as examining of cell shape and morphology by immunostaining. Additionally, assessment of E-cadherin levels, which would act as a marker of epithelial, non-fused cells, could further indicate extent of syncytialization. Our lab has used primary isolated trophoblast cells as a model of cell-fusion, in which polarity protein Par6 was discovered to be a negative regulator of this cytotrophoblast fusion (Sivasubramaniyam et al., 2013). Cytotrophoblast proliferation and fusion occurs in a temporally regulated manner during placenta development, and interestingly, we have found a unique temporal and spatial distribution of AMOT in the developing placenta.

Thus, investigation in how AMOT may be implicated in this process will deepen our understanding of its role in trophoblast cells, and may also provide additional explanation for its temporal expression pattern.

5.2 Deciphering the role of AMOT in placental mesenchymal cells

As mentioned, placental derived mesenchymal cells (pMSC) are a relatively understudied placental cell type, despite their apparent role in placenta development (Chen, 2014). pMSC secrete a wide array of soluble cytokines and growth factors including insulin-like growth factor

1(IGF-1) and hepatocyte growth factors (HGF), which have been shown to trigger EVT migration and invasion (Lacey et al., 2002). Mechanistically, HGF is secreted by pMSC and binds to receptors on trophoblast cells which facilitates an increase MMP9 expression, and induction of cyclic AMP (cAMP) mediated signalling that leads to integrin 1 expression (Chen et al., 2013), both of which contribute to the migratory and invasive trophoblast phenotype. In addition, pMSC have also been demonstrated to participate in vascularization of the placental villous. One study

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revealed pMSC differentiate into endothelial cells and contribute to new blood vessel formation through integrin 51 signalling, both in vitro and in vivo (Lee et al., 2009). AMOT’s implicated role in epithelial cell migration and invasion, as demonstrated by this research and previous studies, as well as its widely acknowledged function in angiogenesis, make AMOT a relevant protein to investigate in this subpopulation of placental cells.

Currently, our lab is actively isolating and studying pMSC from first trimester, pre-term, term, and preeclamptic pregnancies to expand the current knowledge on the contribution of pMSC to development and pathology of the human placenta. Immunofluorescence analyses of AMOT in pMSC isolated from term placenta confirmed endogenous AMOT expression in these cells

(Figure 5.1A). Additionally, co-localization of AMOT with F-actin probe phalloidin was observed in pMSC (Figure 5.1A). Interestingly, AMOT and phalloidin localization remained predominantly in the leading edge of pMSC, suggesting a potential role for AMOT in the migration and/or polarity of these cells. To test this, investigations into AMOT movement and contribution to pMSC migration should be assessed. This could be done using YFP AMOT plasmid constructs and time- lapse imaging as demonstrated in this study. Our lab has determined pMSC to be a robust cell model as they are highly amendable to transfection and can be passed and maintained in culture.

Thus, in addition to the overexpression and subsequent migration studies which would implicate a role for AMOT in mesenchymal cell motility, it is important to distinguish whether AMOT is involved in the differentiation into endothelial cells and subsequent vascularization of the chorionic villi. To test AMOT’s role in mesenchymal-to-endothelial differentiation, AMOT loss and gain of function studies in pMSC could be conducted, and resulting pMSC cells induced to differentiate, using supplemented culture medium with necessary endothelial growth factors as outlined by Lee et al (Lee et al., 2009). Here, we could effectively assess the impact of AMOT on

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mesenchymal-to-endothelial transition by analyzing differences in cell morphology, as well as measuring respective mesenchymal and endothelial markers. Alternatively, pMSC could be induced to differentiate into endothelial cells first, and then the resulting differentiated cells subjected to AMOT loss and gain of function studies. Following this, in vitro vessel formation assays could be performed to establish the impact of AMOT overexpression or silencing on pMSC- mediated angiogenesis.

In line with the presented findings in JEG3 cells, preliminary immunofluorescence staining revealed reduced AMOT in term pMSC cells exposed to low oxygen (3% O2) compared to cells cultured in normoxic conditions (8% O2) (Figure 5.1B). Furthermore, the leading edge localization of AMOT is absent in pMSC exposed to low oxygen (3% O2). I propose further quantification of

AMOT protein levels by WB analyses to confirm an oxygen mediated regulation of AMOT in pMSC cells. Nonetheless, these pilot experiments reveal AMOT is expressed in pMSC and appears to be differentially regulated by oxygen tension. Investigation into how low oxygen impacts AMOT potential functions in pMSC, such as migration and differentiation into endothelial cells, could provide insight into mechanisms which are implicated in preeclampsia. This study demonstrated significantly reduced levels of AMOT protein, and disrupted EVT localization of

AMOT in PE. Exciting preliminary immunofluorescence staining reveals a striking reduction in

AMOT fluorescence, and localization, in pMSC isolated from preeclamptic placentae compared to term control (Figure 5.2). Furthermore, through visual analyses it is clear that pMSC from PE are much smaller than those from term placentae. Reiterating the aforementioned experimental concept, pMSC from PE cells could be subjected to migration, differentiation and vessel forming assays and the role of AMOT could be ascertained.

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Term pMSC

A AMOT Phalloidin Merge

20X 20X 20X LE LE LE

AMOT Phalloidin DAPI

Term pMSC

8% O2 3% O2 B 20X 20X

AMOT DAPI

Figure 5.1 Investigating AMOT in placenta mesenchymal cells (pMSC) isolated from term placentae.

Representative immunofluorescence images of: (A) AMOT (green) and F-actin cytoskeleton probe phalloidin (red) in term pMSC (n=2), (B) AMOT (green) in term pMSC cultured in normoxia (8% O2) and hypoxic (3% O2) conditions for 24 hours (n=2). Nuclei were stained with DAPI (blue).

LE= Leading Edge

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pMSC

Term PE 20X 20X

AMOT

20X 20X

Phalloidin

20X 20X

Merge

AMOT Phalloidin DAPI

Figure 5.2 Investigating AMOT localization term and preeclamptic placental mesenchymal cells (pMSC)

Representative immunofluorescence images of AMOT (green) and F-actin cytoskeleton probe phalloidin (red) in pMSC isolated from term and preeclamptic (PE) placentae (n=2). Nuclei were stained with DAPI (blue).

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5.3 Establishing a role for the AMOT/TAZ axis in trophoblast cell differentiation

The functional consequence of AMOT interaction with Hippo pathway effector TAZ has been investigated in various cancers (Chan et al., 2011; Gomez et al., 2014; Hong, 2013; Lv et al., 2015;

Moroishi et al., 2015; Yi et al., 2013b), in endothelial cells during angiogenesis (Dai et al., 2013) and even in lineage differentiation in the early embryo development (Hirate et al., 2013; Hirate and Sasaki, 2014). Yet, the role of AMOT/TAZ axis in trophoblast cell differentiation in the first trimester human placenta remains unknown. A future direction of this project could entail investigation into AMOT and TAZ interaction in trophoblast cells, and determine how this interaction contributes to trophoblast proliferation, migration or invasion. Preliminary immunofluorescence analyses in 5 week placentae sections reveals an increase in AMOT/TAZ co- localization in the intermediate and distal portions of the anchoring column, further confirmed by fluorescence quantification (Figure 5.3). Here, EVT cells are in closer proximity to the maternal decidua and spiral arteries, and are exhibiting characteristics of migration and invasion (DaSilva-

Arnold et al., 2015). Notably, TAZ fluorescence was also present in the proximal region of the anchoring column. Considering the established role of TAZ in promoting cell proliferation, these findings inspired the following concept. In proximal EVTs, TAZ is free to induce cell proliferation; however, in the intermediate and distal EVTs, AMOT interacts with TAZ and sequesters it from the cell nucleus and into the cytoplasm where they collectively regulate cell migration and invasion. This could occur either through AMOT/TAZ binding to F-actin and participate in cytoskeletal rearrangements, or AMOT/TAZ localizing to the tight junction and coordinating cell contact- induced inactivation of TAZ or participate in regulation of epithelial cell polarity.

Association between AMOT and TAZ can be confirmed by co-IP or PLA, in placenta tissue lysates and in vitro JEG3 cells. It would also be of value to assess AMOT/TAZ interaction by PLA during

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wound healing (JEG3 cells) or during EVT outgrowth (villous explants) as this could provide insight into how these two proteins interact during trophoblast cell migration and invasion. It is known the AMOT/TAZ interaction occurs via the PPxY motif in the N-terminal extension of

AMOT 130, and the tryptophan binding domain (WW) of TAZ (Chan et al., 2011). Thus, it could also be of value to disrupt AMOT/TAZ association using AMOT 130 constructs with mutations in its PPxY domain, and assess the impact on migration/invasion of JEG3 cells or villous explants.

In the present study, I showed that TGFβ1/3 promotes redistribution of AMOT from the tight junction to the cytoplasm in JEG3 cells. Studies have outlined an interplay between TGFβ signalling and TAZ activity, where TAZ binds to Smad proteins, translocates into the nucleus and contributes to Smad-dependent TGFβ mediated gene transcription (Varelas et al., 2008; Varelas et al., 2010). Additionally, studies have demonstrated TGFβ signalling to upregulate TAZ expression

(Miranda et al., 2017). In preliminary studies, I found that AMOT and TAZ co-localize in the cytoplasm of JEG3 cells, and this co-localization is upregulated following TGFβ1 and TGFβ3 treatments (Figure 5.4). Additionally, I observed an increase in TAZ nuclear fluorescence following TGFβ1/3 treatment, corroborating what is seen in the literature (Figure 5.4). Follow-up co-IP and PLA assays could be performed to quantify the effect of TGFβ1/3 on AMOT/TAZ interaction. Furthermore, considering the implications of TGFβ signalling in early placenta development, and its disruption in preeclampsia, it is of value to investigate whether TGFβ mediated alterations in trophoblast cell migration or invasion occurs by modulating the

AMOT/TAZ interaction, or their collective localization. This could be achieved by treating JEG3 cells or villous explants with TGFβ1/3, impeding the interaction between AMOT/TAZ either by silencing or by the use of AMOT constructs with mutations in its PPxY domain, and finally assessing migration and invasion using the functional assays mentioned previously.

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PC DC

DC PC

20X 20X

AMOT TAZ DAPI 10X

35000 14000 e

e c

n 30000 n 12000

e e

c c

s s

e 25000 10000 e

r r

u y

u t

t 20000 8000 i

i o

s l

n n

15000 F

e 6000

t Z

n n I 10000

I 4000 A

M T

5000 2000 n

a a

e 0 e 0

M PC DC M PC DC Region of anchoring column Region of anchoring column

Figure 5.3 Investigating AMOT and TAZ spatial association in the human placenta

Representative immunofluorescence images illustrating AMOT (green) and TAZ (red) co- localization in placenta tissue sections from 5-6 weeks gestation (n=3). Nuclei were stained with DAPI (blue). Images were acquired with spinning disc confocal microscope and Volocity Imaging software. Fluorescence was quantified as the mean fluorescence intensities of green (AMOT) and red (TAZ) using Volocity quantification software. PC= Proximal Column DC= Distal Column.

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Vehicle TGFβ1 TGFβ3

TAZ AMOT DAPI

Figure 5.4 Effect of TGFβ1/3 on AMOT and TAZ localization in JEG3 cells

Representative images of immunofluorescence analysis of AMOT (green) and TAZ (red) following treatment with 10ng/mL TGFβ1 and TGFβ3 ligand for 24 hours (n=3). Nuclei were stained with DAPI (blue). Treatment of JEG3 cells with TGFβ promotes AMOT co-localization with TAZ in the cytoplasm.

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5.4 Investigating lysosomal degradation of AMOT

The present study postulates degradation of AMOT via the endocytic pathway as a mechanism to explain the downregulation of AMOT protein. This is in part due to the lipid binding domain on

AMOT that has been shown to promote its interaction with endosomes (Heller et al., 2010a), as well as the established role of AMOT in promoting the internalization and redistribution of apical polarity complex proteins to endosomes (Wells et al., 2006). During the endocytic recycling pathway, endocytosed proteins from the plasma membrane are assembled into endosomes, which ultimately fuse with lysosomes containing proteolytic enzymes to promote protein degradation

(Elkin et al., 2016). Thus, immunofluorescence analysis of AMOT and markers of lysosome, such as lysotracker or lysosomal associated membrane protein (LAMP-1), in vitro or in placentae tissue would be of value because it could determine whether AMOT is actually present at the lysosome.

Further, to determine if AMOT is actually subject to lysosomal degradation, one could treat JEG3 cells with ammonium chloride, an inhibitor of phagosome to lysosome fusion, and assess AMOT protein levels. If AMOT is subject to lysosomal degradation, inhibition of the proteolysis by ammonium chloride would intuitively result in an increase levels of AMOT protein, which could be assessed by western blot analyses. Considering that the internalization of proteins at the plasma membrane precedes their lysosomal degradation (Elkin et al., 2016), an alternative approach could be inhibiting endocytosis and assessing AMOT/lysosome dynamics. Cytochalasin D is a chemical inhibitor of endocytosis via preventing the polymerization of the F-actin cytoskeleton (Dutta and

Donaldson, 2012). Thus, if treatment of JEG3 cells with Cytochalasin D attenuates the association between AMOT and lysosomes, or ultimately increases AMOT protein levels, one could propose that AMOT is undergoing lysosomal degradation by the endocytic pathway. Interestingly, recent work from our lab has discovered that the number of lysosomes, as well as the levels of lysosomal markers including as LAMP-1 and TFEB, are significantly increased in PE placentae vs age-

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matched controls (Ermini et al., 2018). Considering this, it would be of value to also investigate

AMOT and lysosome dynamics using the same methodology in preeclamptic placentae, as this could potentially account for the significantly decreased levels of AMOT found in PE. The lipid binding domain within AMOT has been shown to selectively bind with high affinity to membranes containing monophosphorylated phosphatidylinositol (PIPs) and cholesterol (Heller et al., 2010a).

Thus, one final experiment could be to assess the membrane levels of these specific lipids in varying in vitro conditions, such as low oxygen exposure or following TGFβ treatment, and/or in preeclamptic and age-matched control placentae via liquid chromatography tandem-mass spectrometry (LC-MS/MS). Differential expression of these lipids could affect AMOTs level of involvement in the endocytic pathway, as well as altering its potential degradation by the lysosome.

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Appendix - Statement of Contributions

The following individuals have contributed to the collection of materials and generation of data:

Mount Sinai Hospital Biobank (Toronto, ON, CAN) supplied first-trimester, preeclamptic and normal (age-matched control) placentae for immunohistochemistry, immunofluorescence, and

RNA and protein analyses.

Dr. Sruthi Alahari for her help in performing the in vitro JMJD6 hydroxylation reaction.

Dr. Leonardo Ermini for performing MALDI-TOF mass spectrometry and analyzing AMOT peptide profiles.

Dr. Michael Litvak for his immeasurable help in performing time lapse imaging in JEG3 cells following wound healing assay, as well as his assistance in calculating linear migration rates of

JEG3 cells.

Dr. Lubna Nadeem for her ongoing help and advice while I was conducting the proximity ligation assay experiment.

The work presented in this thesis was supported by Canadian Institute of Health Research

(CIHR) grant held by Dr. Isabella Caniggia. In addition, I received the Ontario Student

Opportunity Trust Fund (OSOTF) Award from Mount Sinai Hospital, as well as the Institute of

Medical Science Entrance Scholarship and the Institute of Medical Science Open Fellowship

Award.