The Role of Polarity Protein Angiomotin (AMOT) in the

Human Placenta

by

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

© Copyright by Abby Farrell 2018

ii

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.

TGF1/3 treatment decreased AMOT protein levels and redistributed AMOT from the tight-

junction to cytoplasmic F-actin in JEG3 cells. TGF1/3 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.

iii

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

iv

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 TGF1/3 ligand treatment reduces AMOT 130 and 80 protein levels ..................68

3.2.3 TGF1/3 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

v

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

vi

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.

vii

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

viii

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

ix

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

x

List of Tables

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

xi

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

xii

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

xiii

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

1

Chapter 1

Introduction

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.

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

3

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

4

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

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).

Angiomotin (AMOT)

130 kDa

80 kDa

1084 a.a

675 a.a

1 20 280 312 342 672 675

AMOT 130

AMOT 80

Angiostatin binding

PDZ binding

Coiled coil binding

106 LPTY

109

1 429 689 721 751 1081 1084239PPEY

242

284 PPEY287

867 1005

458 596

AMOT Like-1 (AMOTL1)

188 LPTY

191

1 438 694 729 953 956310PPEY

313

367 PPEY370

762 106 kDa956 a.a

AMOT Like-2 (AMOTL2)

86 kDa780 a.a

104 LPTY

107

1 308 581 777 780210 PPQY213

252 PPVF255

6

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.

7

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).

8

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.

9

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

10

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

11

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

12

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

13

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.

14

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

A

B

15

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).

16

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.

17

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

18

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

19

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 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 (TGF3) to maintain trophoblast cells in a proliferative, non-

invasive phenotype. Increased exposure to oxygen reduces HIF-1 and levels of TGF3, allowing

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

al., 2000; Ietta et al., 2006).

TGFs 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

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

A

B

Oxygen Gradient ~20 mmHg ~2-3% O2

~55 mmHg ~6-8 % O2

5-9 weeks 10-12 weeks

Fetal Side Maternal Side Decidua Myometrium

Anchoring villi column

Floating villi

ST CT

Proximal Intermediate Distal

pMSC

Spiral artery

Migratory EVT

Invasive EVT

Fetal Blood Vessel Stroma

Umbilical arteries

Umbilical veins

Chorionic Villi

Maternal spiral artery

Intervillous space

CT ST

EVT

21

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).

22

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 (TGF1, TGF2

and TGF3), 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, TGF3 is the only isoform demonstrated to exhibit a temporal expression

pattern, peaking at 7/8 weeks of gestation, and dropping thereafter. This is unlike TGF1 and

TGF2, which remained consistently expressed across gestation (Caniggia et al., 1999). As

mentioned, this temporal peak in TGF3 is attributed to the hypoxic environment during early

placenta development, where HIF-1 levels are abundant and positively regulate levels of TGF3

(Caniggia et al., 2000; Ietta et al., 2006). The drop in TGF3 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, TGF3 has proven to be an inhibitor of cytotrophoblast outgrowth

and invasion. Specifically, in vivo studies of placental explants showed that inhibition of TGF3,

but not TGF1 or TGF2, restored the invasive capabilities of trophoblast cells, and increased

both MMP production and increase fibronectin deposition (Caniggia et al., 1999). Moreover, only

the TGF3 isoform was found elevated in placentae complicated with preeclampsia. Altogether

these data suggest a role for TGF3 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 TGF1 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 TGF1

to supress trophoblast invasion by: (1) upregulating tissue inhibitors of metalloproteases (TIMPs),

23

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 TGF2 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

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

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

2002; Wrana, 2013). While seven types of TGFRI, 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 TGFRII, and promotes trans-phosphorylation and

24

activation of TGFRI (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 TGFRI (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 TGFRII, 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.

25

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

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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

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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.

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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).

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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

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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 51 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).

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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 el