Functional Role of Protein Kinase C Alpha in Endometrial Carcinogenesis

University of Nebraska Medical Center DigitalCommons@UNMC

Theses & Dissertations Graduate Studies

Spring 5-5-2018

Alice Hsu University of Nebraska Medical Center

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FUNCTIONAL ROLE OF PROTEIN KINASE ALPHA IN ENDOMETRIAL CARCINOGENESIS

Alice H. Hsu

A DISSERTATION

Presented to the Faculty of the University of Nebraska Graduate College in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

Cancer Research Graduate Program

Under the Supervision of Professor Jennifer D. Black

University of Nebraska Medical Center Omaha, Nebraska

April 2018

Supervisory Committee:

Adrian R. Black, Ph.D. Robert E. Lewis, Ph.D.

Adam R. Karpf, Ph.D. Jing Wang, Ph.D.

Kay-Uwe Wagner, Ph.D

This work is dedicated

To Nik,

Who led me in to the door

To Scott,

Who showed me how to have fun

To Jenny and Adrian,

Who gave me guidance and freedom to grow

iii

Acknowledgements

This work would not have been possible without the support and guidance from Jenny and Adrian, who have been and always will be my inspiration and role models. Under their, I have learned how to be critical with respect, be assertive but considerate, hold oneself to the highest standard but treat others with absolute grace. They have shown me the value of hard work and persistence, and tirelessly invested their time and energy to ensure my success in every experiment, every seminar, all the bits and pieces of writing and posters I have presented over the years, and most of all, as a scientist and a person. I truly could not have hoped for a better environment for my graduate training and am honored to be a member of the Black lab.

I have had the pleasure of working with wonderful colleagues from the Black lab, both past and present – Katie, Fang, Michelle, Misty, Xinyue, Navneet, Lisa, and Shane. Despite our different personalities, we have managed to create a cohesive, calming, productive, and accepting working environment that made it easy to hop and skip back to work the next morning following a long, frustrating day.

I would also like to express my gratitude to my committee members, Adrian Black,

Adam Karpf, Rob Lewis, Jenny Wang, and Kay Wagner for the insightful comments regarding our project and monitoring the development of my graduate career. They have generously provided their perspective to better our research, as well as imparted wisdom and encouragement to make me a better scientist.

Additionally, I want to thank all the support I have received from UNMC especially

Lijun Sun and Jiang Jiang from the Tissue Science Facility for their technical support for the extensive tissue work this project encompassed, our neighboring labs – the Dong lab, Lewis lab, and Rizzino lab for welcoming us into DRC and creating a harmonious research environment. I would also like to acknowledge those who have been my companions during my graduate studies, especially Erin, Beth, and Carter for challenging me both intellectually and personally. iv

In this long, testing journal of my graduate career, I have been blessed to be accompanied by an army of incredible friends. We have shared the important milestones of our lives with each other, celebrated many happy memories, but most important, when the going got hard, they have pulled me through the tough moments. I once read “Friends were family one chose”, and I am proud to say they are one of my best choices. Thank you Kelly L., Joanne, Simon, Daniel,

Patrick, Tzu-an, Jay, Danny, Jackal, Moashih, Enkang, Wa, Fang, Dianmu, Varun, Wanchi, and many more.

Finally, I would like to thank my parents, brother, and husband for their unconditional trust and support, who kept their distance to allow me the space and freedom to be unapologetically me but always know when to step in to assure me they are still behind me and

will always be. Hope I have done them proud.

Functional Role of PKCα in Endometrial Carcinogenesis Abstract Alice H. Hsu, Ph.D.

University of Nebraska Medical Center, 2018

Supervisor: Jennifer D. Black, Ph.D.

Protein kinase Cα (PKCα) is a member of the PKC family of serine/threonine kinases that regulate many fundamental cellular processes, including cell proliferation, differentiation, survival and transformation. The impact of PKCα on tumorigenesis, and whether it acts as an oncogene or tumor suppressor, has been debated over the years. The overall goal of this study was to establish the functional role of PKCα in endometrial carcinogenesis. Results from this study broaden our knowledge of PKCα signaling and add to our understanding of its role in tumor development.

As understanding of the role of PKCα signaling in the uterus is limited, we first examined the expression and activation of PKCα in the normal endometrium. Our analysis revealed cyclical activation of PKCα, linked to hormonal fluctuations and negatively correlated with epithelial proliferation, suggesting that PKCα is a growth inhibitory kinase in the endometrium.

Historically, PKC isozymes have been recognized as oncoproteins as they are activated by the tumor promoting phorbol esters. However, accumulating evidence indicates that PKCs can be inhibitory in some cancers, with recent findings propelling a shift in focus to understanding tumor suppressive functions of these enzymes. Here, we report that PKCα acts as a tumor suppressor in

PI3K/AKT-driven endometrial cancer. PKCα deficiency, due to transcriptional repression, is associated with aggressive disease and poor prognosis in endometrial cancer patients. Activation of PKCα induces PP2A activity, leading to hypophosphorylation of AKT and potently inhibiting the growth of endometrial cancer cells under anchorage-independent conditions. In murine models,

PKCα loss is a rate-limiting event for initiation of endometrial tumorigenesis, as demonstrated by vi

increased tumor burden in mice lacking PKCα.

More than 90% of endometrial cancer cases are characterized by AKT hyperactivation as the result of mutations in multiple PI3K/AKT pathway components, including PTEN, PIK3CA,

PIK3R1, and AKT. Mutations in these molecules often coexist in EC, indicating that EC development requires full, unopposed activation of PI3K/AKT signaling. Results from our study show that PKCα is a negative regulator of AKT activation in the endometrium, which can override hyperactivation of the PI3K/AKT pathway in endometrial cancer cells. Loss of this kinase in the context of other molecular aberrations, such as mutations in PTEN or PI3K, leads to further hyperactivation of AKT, enhancing tumor cell growth. Together, our data point to PKCα as a crucial tumor suppressor in the endometrium, with deregulation of a PKCα→PP2A signaling axis leading to robust AKT activation and enhanced endometrial tumorigenesis.

vii

Table of Contents Acknowledgements ...... iii

Abstract ...... v

List of Figures ...... x

List of Tables ...... xiii

List of Abbreviations ...... xiv

Chapter 1: Introduction ...... 1

1.1 The protein kinase C (PKC) family ...... 2 1.1.1 Structure, maturation, and activation of PKC isozymes ...... 2 1.1.2 PKC modulating agents ...... 7 1.1.3 Functions of PKC isozymes ...... 10 1.2 PKC in cancer ...... 17 1.2.1 PKC as a tumor promoter ...... 17 1.2.2 PKC as a tumor suppressor ...... 22 1.3 Protein kinase C alpha (PKCα) ...... 23 1.3.1 PKCα in intestinal homeostasis and carcinogenesis ...... 27 1.3.2 Impact of PKCα signaling in cancer ...... 33 1.4 AKT signaling network ...... 34 1.4.1 Regulation of AKT activation ...... 34 1.4.2 Downstream targets of AKT ...... 38 1.4.3 Hyperactivation of AKT in cancer ...... 45 1.5 Dissertation objectives ...... 47 Chapter 2: Materials and Methods ...... 49

2.1 Cell lines and reagents ...... 50 2.2 Transgenic mice ...... 52 2.3 Determination of estrous phases ...... 54 2.4 Human tissue microarray ...... 55 2.5 Immunohistochemistry ...... 56 2.6 Immunofluorescence ...... 57 2.7 Western blot analysis ...... 57 2.8 Laser-capture microdissection (LCM) ...... 59 2.9 Quantitative RT-PCR analysis ...... 60 viii

2.10 RNA interference ...... 61 2.11 Nascent RNA capture ...... 62 2.12 Subcellular fractionation ...... 62 2.13 PP2A activity assay ...... 63 2.14 Establishing EC cell lines stably expressing mutant AKT ...... 63 2.15 Adenoviral-mediated overexpression and soft agarose colony formation assay .. 64 2.16 Correlation between PKCα mRNA and protein levels ...... 65 2.17 Quantification of tumor burden and pAKT intensity relative to PKCα levels ...... 65 2.18 Statistical analysis ...... 66 Chapter 3: The Role of PKCα in the Endometrium ...... 67

3.1 Introduction ...... 68 3.1.1 Biology of the endometrium ...... 68 3.1.2 Human estrous cycle ...... 68 3.1.3 Rodent estrous cycle ...... 69 3.1.4 Effects of steroid hormone signaling: estrogenic response ...... 70 3.1.5 Effects of steroid hormone signaling: progesterone ...... 73 3.1.6 PKC signaling in the endometrium ...... 76 3.2 Results ...... 78 3.2.1 Survey of PKCα expression in the endometrium ...... 78 3.2.2 PKCα expression in mouse estrous cycle phases ...... 80 3.2.3 Membrane-associated PKCα at estrous phase ...... 82 3.2.4 Inverse correlation between PKCα activation and cell proliferation ...... 84 3.3 Discussion ...... 86 Chapter 4: Crosstalk Between PKCα and PI3K/AKT Signaling is Tumor Suppressive in the Endometrium ...... 89

4.1 Endometrial cancer ...... 90 4.1.1 Clinical and molecular classification ...... 90 4.2 Results ...... 98 4.2.1 Transcriptional suppression of PKCα is associated with high-grade EC ...... 98 4.2.2 PKCα expression is lost in mouse models of endometrial hyperplasia ...... 107 4.2.3 PKCα deficiency in EC is not a direct result of PTEN loss/ AKT activation and does not appear to drive loss of PTEN ...... 113 4.2.4 PKCα inhibits endometrial tumorigenesis in vivo and in vitro ...... 116 4.2.5 PKCα activity inhibits PI3K/AKT signaling in EC cells via a PHLPP 1/2- ix

independent, okadaic acid/ calyculin A-sensitive mechanism ...... 121 4.2.6 PKCα loss is associated with increased AKT activity in endometrial lesions in vivo ...... 136 4.2.7 Inhibition of AKT activity is an important component of the PKCα tumor- suppressive axis in EC cells ...... 137 4.3 Discussion ...... 140 Chapter 5: Conclusion ...... 146

5.1 PKCα as a potential anti-proliferative regulator in the endometrium ...... 147 5.2 PKCα suppresses endometrial carcinogenesis ...... 149 Appendix A: Estrogenic stimulation of PKCα expression ...... 154

Rationale: ...... 154 Results/Discussion: ...... 154 Appendix B: Regulation of PKCα Expression ...... 156

Rationale: ...... 156 Results/Discussion: ...... 156 Literature Cited: ...... 161

List of Figures

Fig. 1-1: Domain structure of PKC family members ...... 3

Fig. 1-2: Sequential maturation and activation of PKCs ...... 5

Fig. 1-3: PKC-mediated regulation of G1/ S transition ...... 13

Fig. 1-4: PKC-mediated regulation of G2/ M transition ...... 14

Fig. 1-5: Mechanism of PKCδ-mediated apoptosis ...... 16

Fig. 1-6: Phorbol ester as a tumor promoter ...... 19

Fig. 1-7: PKCα expression in human cancers ...... 25

Fig. 1-8: PKCα as a tumor suppressor ...... 27

Fig. 1-9: PKCα activation in the intestinal systems ...... 30

Fig. 1-10: PKCα expression in lost in intestinal tumors ...... 32

Fig. 1-11: AKT signaling network ...... 39

Fig. 3-1: PKCα expression in normal human uterus ...... 79

...... 79

Fig. 3-2: Correlation between PKCα and Ki67 expression during the estrous cycle ...... 81

Fig. 3-3: Membrane-associated PKCα at estrous ...... 83

Fig. 3-4: PKCα expression during the estrous cycle ...... 85

Fig. 4-1: PKCα expression in EC ...... 99

Fig. 4-2: Alterations in PKC isozyme mRNA expression in ECs s ...... 100

Fig. 4-3: Correlation between PKCα mRNA and protein levels in EC cell lines ...... 103

Fig. 4-4: Alterations in PKCα transcription in EC cells ...... 104

Fig. 4-5: Loss of PKCα is associated with higher-grade EC ...... 105

Fig. 4-6: Survival probability of EC patients with high and low PKC expression ...... 106

Fig. 4-7: Concurrent loss of PKCα and PTEN correlates with pAKT ...... 108

Fig. 4-8: Laser Microdissection of pAKT- and PKCα-labeled uterine epithelium for qRT- xi

PCR analysis of PKCα mRNA levels...... 110

Fig. 4-9: Loss of PKCα in uterine tumors of mice with cre-mediated conditional endometrial deletion of Pten...... 111

Fig. 4-10: Enhanced Id1 expression in endometrial hyperplasia correlates with loss of

PKCα ...... 112

Fig. 4-11: PKCα deficiency in EC is not a direct result of PTEN loss or AKT activation

and does not affect PTEN levels ...... 114

Fig. 4-12: PTEN knockdown did not alter the expression of other members of the PKC

family ...... 115

Fig. 4-13: PKCα inhibits endometrial tumorigenesis in vivo ...... 117

Fig. 4-14: PKCα inhibits endometrial tumorigenesis in vitro ...... 118

Fig. 4-15: PKCα kinase activity is required for its tumor suppressive function ...... 119

Fig. 4-16: siRNA-mediated knockdown of PKCα promotes tumorigenesis ...... 120

Fig. 4-17: PMA-induced activation of PKCα reduces AKT phosphorylation ...... 122

Fig. 4-18: The cPKC inhibitor ...... 123

Fig. 4-19: siRNA-mediated knockdown of PKCα prevents PMA-induced

hypophosphorylation of AKT ...... 124

Fig. 4-20: Knockdown of PKCα does not affect the expression of other members of the

PKC family in EC cells ...... 125

Fig. 4-21: The effects of PMA on PKCα and AKT activity are sustained in EC cells ... 126

Fig. 4-22: Knockdown of PKCα enhances steady-state AKT activation ...... 127

Fig. 4-23: PKCα signaling can inhibit AKT activity in EC cells with hyperactivation of

the PI3K/AKT pathway due to coexisting alterations in more than one pathway component .... 130

Fig. 4-24: PKCα downregulates Id1 in EC cells ...... 131

Fig. 4-25: Expression of PHLPPs and PP2A in EC cells ...... 132

Fig. 4-26: PP2A but not PHLPP inhibitors blocked PMA-mediated hypophosphorylation xii

of AKT ...... 133

Fig. 4-27: PMA induces PP2A activity in EC cells ...... 134

Fig. 4-28: PP2A knockdown reduces the effects of PKCα signaling on AKT phosphorylation/activity in EC cells ...... 135

Fig. 4-29: PKCα loss is associated with increased pAKT in endometrial lesions in vivo

...... 136

Fig. 4-30: Inhibition of AKT activity is a component of the PKCα tumor-suppressive axis

in EC cells ...... 138

Fig. 4-31: Mutant AKT partially rescues PKCα-mediated growth suppression ...... 139

Fig. 4-32: Methylation of PKCα CpG shores in cancer cells ...... 142

Fig. 5-1: Schematic representation of PI3K/PTEN and PKCα as regulators of AKT

activation endometrial cancer...... 153

Fig. A.1: PKCα expression following E2 stimulation of EC cells...... 155

Fig. B.1: Figure PKCα promoter activity in human EC cells...... 158

Fig. B.2: PKCα mRNA levels under RNA polymerase inhibition...... 159

Fig. B.3: PKCα levels under inhibition of protein synthesis...... 160

xiii

List of Tables

Table 4-1: Molecular alterations in endometrial cancer...... 93

Table 4-2: Top three most significantly mutated genes in 4 molecular subtypes of EC... 95

Table 4-3: Mutations in EC cell lines...... 102

xiv

List of Abbreviations

Abbreviation Definition

17-AAG 17-N-allylamino-17-demethoxygeldanamycin

4E-BP1 eIF4E binding protein

ACF Aberrant crypt foci

ANX8 Annexin VIII locus

AOM Azoxymethane

AP-1 Activator protein-1

APC Adenomatous polyposis coli

APCMin/+ Adenomatous polyposis coli gene with min mutation aPKC Atypical protein kinase C

ATF4 Cyclic AMP-dependent transcription factor

ATP Adenosine triphosphate

BASCs Bronchoalveolar stem cells

BCL-XL B-cell lymphoma-extra large

BH3 Bcl-2 homology 3

BIM-1/BisΙ Bisindolylmaleimides

Ca2+ Calcium

CDK Cyclin-dependent kinase

CML Chronic myeloid leukemia

COX2 Cyclooxygenase 2 cPKC Classical or conventional protein kinase C xv

DAG sn-1,2-diacylglycerol

DGK Diacylglycerol kinase

DiC8 1,2-dioctanoyl-sn-glycerol

DMBA 7,12-Dimethylbenz[a]anthracene

DNA Deoxyribonucleic acid

E2 Estrogen

E2F E2 factor

EC Endometrial cancer

EDTA Ethylenediaminetetraacetic acid

EGFR Epidermal growth factor receptor

EGTA Ethylene glycol-bis(β-aminoethyl ether)-N,N,N',N'-tetraacetic acid

EMT Epithelial-mesenchymal transition

ER Endoplasmic reticulum

ER Estrogen receptor

ERE Estrogen responsive elements

ERK Extracellular-signal-regulated kinase

ERαKO ERα knockout

ERβKO ERβ knockout

Ets2 Protein C-ets-2

EU Ethylene uridine

FBS Fetal bovine serum

FKBP12 12-kDa FK506-binding protein xvi

FOXO Forkhead box O

FSH Follicle-stimulating hormone

GADD45 Growth arrest and DNA damage 45

GLI1 Glioma-associated oncogene homolog 1

GPCR G-protein coupled receptor

GSK3 Glycogen synthase kinase 3

HEPES N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid

HER2 Human epidermal growth factor receptor 2

Hh Hedgehog

HIF-1α Hypoxia inducible factor-1 α

HSP90 Heat shock protein 90

Id Inhibitor of DNA binding

IEC-18 Intestinal epithelial cell-18

IGF-1 Insulin-like growth factor-1

IHC Immunohistochemistry

INPP4B Inositol polyphosphate-4-phosphatase

IP3 Triphosphoate

JNK c-Jun N-terminal kinase

KD-PKCα Kinase dead PKCα

KO Knockout

LCM Lasercapture microdissection

LH Luteinizing hormone xvii

LKB1 Liver kinase B1

LOF Loss of function

LSCC Lung squamous cell carcinoma

MARCK Myristoylated alanine-rich protein kinase C substrate

MCM Minichromosome maintenance protein miRNAs Micro RNAs

MMMT Malignant mixed Mullerian tumors mRNA Messenger ribonucleic acid

MSI Microsatellite instability

MSS Microsatellite stability

Mst1 Macrophage-stimulating 1 mTOR Mammalian target of rapamycin mTORC Mammalian target of rapamycin complex

Muc2 Mucin 2 myr-AKT Myristoylated AKT

NaF Sodium fluoride

NEAA Non-essential amino acids

NF-κB Nuclear factor kappa-light-chain-enhancer of activated B cells nPKC Novel protein kinase C

NRF2 Nuclear factor erythroid 2-related factor 2

NSCLC Non-small cell lung cancer

OAG 1-Oleoyl-2-acetyl-glycerol xviii

OCT Optimum cutting temperature

P4 Progesterone

PA Phosphatidic acid

Pax Paired box

PB1 Phox Bem 1

PBS Phosphate-buffered saline

PCR Polymerase chain reaction

PDAC Pancreatic ductal adenocarcinoma

PDBu Phorbol 12,13-dibutyrate

PDK1 Phosphoinositide-dependent kinase-1

PHLPP Pleckstrin homology domain and Leucine rich repeat Protein Phosphatases

PIK3CA Catalytic subunit of PI3 kinase or p110

PIk3R1 Regulatory subunit of PI3 kinase or p85

PIP2 Phosphatidylinositol 4,5-bisphosphate

PIP2 Phosphatidylinositol 3,4-bisphosphate

PIP3 Phosphatidylinositol 3,4,5-bisphosphate

PKB Protein kinase B

PKC Protein kinase C

PKD Protein kinase D

PLC Phospholipase C

PMA Phorbol 12-myristate 13-acetate

POLE DNA polymerase ε xix

PP2A Protein phosphatase 2A

PR Progesterone receptor

PRAKO PR-A knockout

PRAS40 Proline-rich Akt substrate of 40 kDa

PRBKO PR-B knockout

PRPS6 Plastid ribosomal protein S6

PTEN Phosphatase and tensin homolog qRT-PCR Quantitative reverse transcription polymerase chain reaction

RAC1 Ras-related C3 botulinum toxin substrate 1

RasGRP Ras guanyl-releasing protein 1

RB Retinoblastoma protein

RNA Ribonucleic Acid

RT-PCR Reverse transcription polymerase chain reaction

RUNX2 Runt-related transcription factor 2

S6K1 Ribosomal protein S6 kinase beta-1

SEM Standard error of the mean

SHIP Src homology 2 domain-containing Inositol 5′-Phosphatase siRNA Small interfering RNA

SIRT1 Sirtuin 1

SOX2 Sex determining region Y-box 2

SREBP1c Sterol regulatory element binding protein 1c

T-ALL T cell-acute lymphoblastic leukemia xx

TCA Trichloroacetic acid

TCGA The Cancer Genome Atlas

TCPA The Cancer Protein Atlas

TGFβ Transforming growth factor β

TMA Tissue microarray

TPA 12-O-tetra-decanoylphorbol-13-acetate

TRAIL TNF-related apoptosis-inducing ligand

TRAMP Transgenic adenocarcinoma of the mouse prostate

TSC2 Tuberous Sclerosis Complex 2

USDA United States department of agriculture

WT Wild type 1

Chapter 1: Introduction

1.1 The protein kinase C (PKC) family

Protein kinase C (PKC) is a member of the family of AGC kinases (including protein kinase

A, G, and C) that phosphorylates serine and threonine residues. It was discovered in the brain by

Nishizuka and colleagues in 1977 as a kinase whose activation is dependent on the presence of phosphatidylserine (PS) and calcium (Ca2+) (Inoue et al., 1977; Kishimoto et al., 1977). In 1979,

the same group showed that PKC can be activated by sn-1,2-diacylglycerol (DAG), a membrane-

associated lipid generated by phosphoinositide hydrolysis, in the presence of low Ca2+ (Kishimoto et al., 1980; Takai et al., 1979). Later, PKC was identified as a cellular target for tumor promoting phorbol esters, such as 12-O-tetra-decanoylphorbol-13-acetate (TPA, also referred to as phorbol

12-myristate 13-acetate (PMA)) (Castagna et al., 1982; Kikkawa et al., 1983), that bind to PKC at the same site as DAG. This study demonstrated the first link of PKC to cell proliferation and pointed to its potential contribution to carcinogenesis. After years of intensive study of these isozymes, PKC has been shown to be involved in the regulation of many fundamental cellular processes such as cell cycle progression, differentiation, cell survival and apoptosis. Subsequent studies have established the pathogenic association of PKC to important health concerns such as cardiovascular disease and diabetes (Ferreira et al., 2011; Geraldes and King, 2010; Inagaki et al.,

2006). In the late 1980s, multiple isoforms of PKCs were identified by molecular cloning

(Coussens et al., 1986; Parker et al., 1986). More than 30 years since its debut, current understanding defines PKC as a family of serine/threonine kinase consisting of ten isozymes, categorized into three classes based on their structure and coactivator requirements and has isozyme-specific, context-dependent functions.

1.1.1 Structure, maturation, and activation of PKC isozymes

There are three classes of PKC isozymes: the classical or conventional PKCs (cPKCs), the novel PKCs (nPKCs), and the atypical PKCs (aPKCs) (Figure 1-1). All PKC isoforms share

Fig. 1-1: Domain structure of PKC family members

(The three classes of PKCs – cPKCs, nPKCs, and aPKCs – share structural similarities in the catalytic domain (C3 and C4) and the hinge region, which connects the regulatory and kinase domains. Differences in the regulatory domains define the activation requirements for each class of PKCs: cPKC activation requires DAG, Ca2+, and phosphatidylserine (PS); nPKC activation requires DAG and phosphatidylserine; aPKC activation is mediated by protein-protein interactions. Adapted from (Mackay and Twelves, 2007)

structural similarities, with an N-terminal regulatory domain linked by a flexible hinge region to

the C-terminal carboxyl kinase domain. The catalytic C3 and C4 domains are conserved in all the

PKCs, which is similar to the catalytic domain of protein kinase A. However, each class of PKCs

has a unique regulatory domain that determines cofactor requirements for activation. The

pseudosubstrate domain is common to the regulatory domains of all PKCs, and plays a central role

in regulating PKC activity (House and Kemp, 1987; House et al., 1987). It contains a sequence

that binds to the substrate binding pocket of the catalytic domain and blocks the substrate binding,

keeping the kinase in its inactive state (House et al., 1987; Orr and Newton, 1994). The regulatory

domain of cPKCs consists of two cysteine-rich motifs (C1A and C1B) in the C1 domain and a Ca2+-

sensitive C2 domain. The C1 domain binds to DAG and phosphatidylserine, while the C2 domain

binds to Ca2+. The presence of Ca2+ in the C2 domain increases the affinity of cPKCs for DAG.

Thus, activation of cPKCs (including PKCα, the splice variants PKCβI and βII, and PKCγ) requires

DAG, Ca2+, and phosphatidylserine. While the nPKCs also have two cysteine-rich motifs in their

C1 domain, they have an “atypical” C2 domain that lacks Ca2+ binding residues. Differing from cPKCs, activation of the nPKCs (PKCδ, ε, η, θ) requires DAG and phosphatidylserine but not Ca2+,

as DAG exhibits higher binding affinity for the C1B component of nPKCs than for the C1A domain

of cPKCs before Ca2+ binding. The activating phorbol ester, PMA, binds to the C1B motif of the

regulatory domain of both cPKCs and nPKCs to enhance their membrane association and activation. Unlike other members of the family, activation of the aPKCs (PKCζ, λ/ι) is independent of both DAG and Ca2+. Atypical PKCs have a variant of the C1 domain that has only one cysteine-

rich motif and does not bind to DAG or phorbol esters. They also lack the C2 domain in their

regulatory region. Instead, aPKCs have a Phox Bem 1 (PB1) domain that is involved in protein-

protein interactions, providing the regulatory forces in their activation.

Fig. 1-2: Sequential maturation and activation of PKCs

The newly synthesized, immature PKC associates with the membrane where it is sequentially phosphorylated at the activation loop, turn motif, and hydrophobic motif mediated by PDK-1, and facilitated by mTORC2 and HSP90. The mature, phosphorylated PKC is then released into the cytosol.

Subsequently, PLC-mediated lipid hydrolysis generates membrane associated DAG and increased intracellular Ca2+ levels. Interaction with DAG creates a strong bond between the phospholipid in the membrane and the open, activated PKC, allowing it to act on appropriate downstream substrates. Adapted from (Wu-Zhang and Newton, 2013)

Maturation of PKC requires a series of highly regulated, ordered phosphorylation events

at three distinct sites of the kinase domain (Figure 1-2). While these events do not directly activate

PKC, each phosphorylation induces a conformational change that stabilizes the kinase in its

inactive form and protects it from phosphatase activity. The first of the three priming

phosphorylations occurs at the activation loop mediated by phosphoinositide-dependent kinase-1

(PDK-1) (Chou et al., 1998; Edwards et al., 1999; Le Good et al., 1998). The newly synthesized

PKC associates with the membrane in its open conformation, in which the substrate binding cavity is free of the pseudosubstrate domain, allowing PDK-1 to access the threonine residue at the activation loop (Dutil and Newton, 2000). It has been shown that cells lacking PDK-1 have markedly reduced PKC levels (Balendran et al., 2000). This phosphorylation initiates the maturation process of PKC and is required for subsequent phosphorylation (Dutil and Newton,

2000; Dutil et al., 1998). The second phosphorylation at the turn motif requires the mammalian target of rapamycin complex 2 (mTORC2), as cells lacking mTORC2 have diminishing levels of

PKC (Facchinetti et al., 2008; Ikenoue et al., 2008; Jacinto and Lorberg, 2008). Despite its essential role in the phosphorylation of the turn motif, mTORC2 was unable to phosphorylate PKC in vitro

(Ikenoue et al., 2008), raising the question whether mTORC2 mediates direct phosphorylation of

PKC. It has been proposed that mTORC2 might act as a chaperone to position the newly synthesized PKC for priming phosphorylation or activates another upstream kinase that phosphorylates the turn motif. The last step of this priming process is the autophosphorylation at the hydrophobic motif. Both heat shock protein 90 (HSP90) and mTORC2 have been shown to modulate the phosphorylation at this motif (Gould et al., 2009; Ikenoue et al., 2008; Ziegler et al.,

1999). However, this might be because mTORC2 is required for mediating the prerequisite phosphorylating event that is necessary for hydrophobic motif autophosphorylation. Completion of these sequential phosphorylation results in a mature, catalytically competent form of PKC. The fully phosphorylated PKC is then released from the membrane into the cytosol with its pseudosubstrate domain occupying the active site, preventing full activation of the kinase (Newton,

2010).

Extracellular signals trigger signaling events leading to the generation of cofactors that are

required for PKC activation. Binding of ligands to their cognate tyrosine kinase receptors or G-

coupled protein receptors stimulates signal transduction that elicits intracellular molecular changes

such as activation of phospholipase C (PLC). PLC then catalyzes the hydrolysis of the lipid

phosphatidylinositol 4,5-bisphosphate (PIP2) to generate inositol 1, 4, 5-trisphosphoate (IP3) and

DAG, the physiological agonist of both cPKCs and nPKCs. DAG binds to the C1 domain of PKC

and anchors it to the plasma membrane. Thus, membrane translocation serves as a readout for PKC

activation (Kraft et al., 1982). IP3 binds to receptors in the smooth endoplasmic reticulum (ER)

and promotes the release of Ca2+ into the cytoplasm, resulting in elevated intracellular Ca2+

concentrations. To compensate for the weaker DAG binding affinity to cPKCs, Ca2+ enhances the binding between DAG and the C1 domain and is required for cPKC but not nPKC activation.

Because of their rapid metabolism at the cell membrane, DAG and its analogs are only transient

PKC-inducing signals. Works from our laboratory and others has shown that the activation of

DAG-stimulated PKC is acutely terminated by ‘reverse translocation’ - i.e., redistribution of the kinase to the cytoplasm (Akita, 2002; Di Mari et al., 2005; Leitges et al., 1996; Lum et al., 2016).

Additionally, prolonged activation of PKC such as that mediated by phorbol esters or chronic DAG stimulation, is terminated by enzyme degradation.

1.1.2 PKC modulating agents

As PKC signaling is associated with diseases such as cancer (Griner and Kazanietz, 2007),

hypertension, and diabetes (Ferreira et al., 2011; Geraldes and King, 2010; Inagaki et al., 2006),

deciphering the roles of individual PKCs in signal transduction and the dysregulation of these

processes is pertinent to further our understanding of human disorders. Pharmacological agents

that modulate PKC activity are valuable tools for the study of effects of this kinase family. Many

PKC modulating agents have been widely used experimentally, allowing for easy manipulation of

PKC activity. Yet, because of their promiscuous nature, these agents are also the culprits of

exacerbating the inherent complexity of PKC signaling, adding confusion to the interpretation of

results from their use. This following section provides an overview of the commonly used PKC

modulating agents, their mechanism of action and concerns that should be taken into account when

utilizing these compounds in experimental settings. Other pharmacological agents that also affect

PKC activity are reviewed in detail in (Wu-Zhang and Newton, 2013).

There are two major types of PKC agonists: DAG and its analogs and phorbol esters. Both are capable of activating cPKC and nPKC through their interaction with the C1 domain, but do not activate aPKC. As activation of PKC occurs at the plasma membrane (or possibly in other membrane compartments), all of these compounds promote membrane translocation of the enzyme,

where it can interact with additional activating cofactors such as phosphatidylserine. In addition to binding to PKC, these compounds can also interact with other families of signaling molecules that also have the C1 domain (e.g. PKDs, RasGRPs, and DGKs) (Blumberg et al., 2010). DAG, a product of PLC-mediated lipid hydrolysis, is the physiological agonist for typical C1-containing

PKC isozymes. Of note, elevated intracellular Ca2+ concentrations can also increase DAG levels

(Kunkel and Newton, 2010). The two short-chain synthetic analogs of DAG, DiC8 (1,2-dioctanoyl-

sn-glycerol) and OAG (oleoacetyle glycerol), have improved aqeous solubility compared with

DAG and are more commonly used in laboratories. DAG and its analogs are rapidly metabolized

in the cell, therefore, only inducing transient PKC activation. Although these compounds can bind

and activate both classical and novel isozymes, they exhibit preferential binding to the C1B motif

in nPKCs and interact with the C1A motif of cPKCs at reduced affinity. Interestingly, rather than

serving as an activating signal, interaction with the lipid kinase, DGK, converts DAG to

phosphatidic acid (PA) to abrogate its signaling ability (Baldanzi et al., 2016; Cai et al., 2009).

Tumor promoting phorbol esters are the active component derived from croton oil (Van

Duuren and Sivak, 1968). The most commonly used phorbol ester is PMA (phorbol 12-myristate

13-acetate), also known as TPA (12-O-tetradecanoylphorbol 13-acetate). The identification of

PKC as a receptor for PMA establishes its first association to carcinogenesis (Castagna et al., 1982;

Kikkawa et al., 1983). The phorbol ester derivative, PDBu (phorbol 12,13-dibutyrate) has

optimized potency and reduced lipophilicity comparing to PMA, making it more water soluble for

therapeutic use. These phorbol esters activate PKC by binding to the C1B motif. They are much

more potent PKC activators than DAG, effective at the nanomolar range. Unlike DAG and its analogs, phorbol esters are not readily metabolized and are capable of generating prolonged activation of PKC. Despite their shared high potency as PKC activators, the phorbol ester derivatives can induce different cellular responses (Blumberg, 1980, 1981).

The C1 ligand, bryostatin-1 (Kunkel and Newton, 2010), provides an example of a PKC agonist that can produce different effects when compared with PMA. Bryostatins are macrocyclic lactones that also bind to the C1 domain. Although these large complex molecules elicit the same effects as phorbol esters, albeit transiently, they actually antagonizes tumor formation (Blumberg et al., 2010). It is unclear how bryostatin-1 acts like a phorbol ester (binding to the C1 domain) to activate PKC on the one hand, while failing to contribute to carcinogenesis on the other. However, this unique property has brought bryostatin into clinical trials for cancer and Alzheimer’s disease

(Kortmansky and Schwartz, 2003; Mayer et al., 2010; Roffey et al., 2009).

The vast majority of effective PKC inhibitors are small molecules that target the ATP- binding site of PKC. These ATP-competitive PKC inhibitors have been the focus of the development of PKC-targeting drugs. There are two distinct chemical classes of these prototypical

ATP-binding inhibitors, bisindolylmaleimides and indolocarbazoles. All of the currently known

PKC inhibitors show some degree of promiscuity toward other kinases, despite some of them being highly selective for PKC. While these active site inhibitors target all three classical or novel PKCs, the inhibition effect is selective, rather than specific for certain isozymes (Wu-Zhang and Newton,

2013). For example, the non-isozyme specific inhibitors Gӧ6983 and bisindolylmaleimide 1 (BIM-

1 or BisI), when used at 500 nM, have been shown to be more effective at inhibiting PKC activity than other kinases (Wu-Zhang and Newton, 2013). Other bisindolylmaleimides, including Ro-31-

8220 and Ro-32-0432 developed by Roche, have been marketed as PKC inhibitors but also have the non-selective nature of other similar compounds. Among the ATP-competitive PKC antagonists, Gӧ6976 and staurosporine are the more widely used indolocarbazole-based inhibitors.

The pharmacological properties of Gӧ6976 have been characterized in multiple kinase inhibitor

studies (Anastassiadis et al., 2011; Bain et al., 2007; Davies et al., 2000), demonstrating that it is highly selective for cPKCs at the nanomolar range while having limited effects on other isozymes or non-PKC kinases. Thus, Gӧ6976 is commonly regarded as a cPKC selective inhibitor. On the other hand, staurosporine is one of the most promiscuous commercially available kinase inhibitors.

It binds to and potently inhibits all cPKC, nPKC, and even aPKC, as well as other non-PKC kinases

(Anastassiadis et al., 2011; Davis et al., 2011; Karaman et al., 2008).

Structure-based PKC inhibitors targeting less conserved domains have been designed with the hope of inhibiting these kinases with improved specificity. One example of these allosteric inhibitors is BisIV. Despite its chemical similarity to BIM-1 and its specificity for PKC, it is a much less potent inhibitor. Other non-active-site inhibitors targeting DAG-binding sites (e.g. calphostin C), the PKC priming kinases PDK-1 and mTORC2, or the maturation process by inhibiting HSP90 (e.g. 17-AAG), have also been employed to inhibit PKC activity. In addition to weak PKC inhibition, these agents also affect other molecular processes and present their own sets of caveats as PKC inhibitors.

Incorporation of PKC modulating agents in understanding the signaling roles of these kinases is a double-edged sword for the PKC field. While these agents allow in vitro manipulation of PKC activities and alteration of its interaction with other signaling molecules, their selective but non-specific nature makes it difficult to link the induced effects to PKC function. Therefore, interpretation of results generated from experiments using PKC modulating agents must be critically reviewed and confirmed with genetic approaches to exclude the involvement of other molecules that might also be targeted by these compounds.

1.1.3 Functions of PKC isozymes

PKC signaling is involved in a wide variety of molecular pathways, among which impact on cell cycle control and apoptosis are the most well-studied cellular processes influenced by this kinase family. Progression through the cell cycle is driven by the enzymatic activity of cyclin-

dependent kinases (Cdks) regulated by their association with cyclins, whose levels are determined by the cellular environment (e.g., mitogenic signals or DNA damage) (Otto and Sicinski, 2017).

PKC signaling can exert either positive or negative regulation of cell cycle progression primarily

at the G1/S and G2/M transitions. At early G1, pocket proteins such as the retinoblastoma protein

(RB), p107, and p130 bind to and inhibit the transcriptional activity of growth-related E2F transcription factors. Under appropriate mitogenic induction signals, Cdk4 and Cdk6 are activated by associating with one of the three D-type cyclins (D1, D2, and D3), leading to phosphorylation of pocket proteins, which relieves E2F repression and upregulates cyclin E (Cobrinik, 2005). The elevated cyclin E in late G1 binds and activates Cdk2 to initiate a program of transcriptional events necessary for S phase entry (Malumbres and Barbacid, 2005). Alternatively, when there are insufficient mitogenic signals for Cdk activation or pocket protein phosphorylation, the cell halts cell cycle progression and enters G0, the quiescent phase of the cell cycle. Continued inactivation of pocket proteins at the end of G1 phase allows accumulation of cyclin A. Once the cell enters S

phase, cyclin E is degraded through a proteasome-mediated mechanism to avoid re-initiation of

DNA replication. Instead, Cdk2 binds to cyclin A to facilitate completion and exit from S phase.

Cyclin A also binds to Cdk1 (Cdc2) at the end of S phase to sequester its activation. As the cell

progresses through G2, cyclin A is degraded while there is active synthesis and accumulation of

cyclin B. This results in the association of Cdk1 and cyclin B, triggering the cell to enter mitosis.

The effect of PKC signaling on cell cycle progression is highly dependent on the

timing and duration of PKC activation, the isozymes present in the cell, and the cellular

environment. Of the crucial molecules involved in cell cycle progression, PKC has been shown to

mediate its effect through regulation of cyclin D1 and p21 at the G1/S transition (Figure 1-3) and lamin B at G2/M (Figure 1-4). At early G1, PKC signaling can lead to increased transcription of cyclin D1 (Li et al., 2006; Soh and Weinstein, 2003) and the destabilization of p21 (Walker et al.,

2006), resulting in hyperphosphorylation of pocket proteins and progression into S phase.

However, activation of PKC signaling at mid-to-late G1 can decrease the transcription and

translation of cyclin D1 (Hizli et al., 2006; Pysz et al., 2009; Soh and Weinstein, 2003) and induce p21 expression (Black, 2000; Griner and Kazanietz, 2007), leading to inhibition of Cdk4/6 and

Cdk2 and cell cycle arrest at G0/G1. Lamin B is one of the essential proteins for nuclear structure

and function. During G2, PKCβII activation has been shown to result in lamin B phosphorylation and the disassembly of the nuclear envelope, allowing the cell to enter M phase and proceed to chromosome segregation (Thompson and Fields 1996). Alternatively, PKC can negatively regulate the G2/M transition by downregulation of Cdc25, preventing the dephosphorylation and activation of Cdk1 (Arita et al., 1998; Barth and Kinzel, 1994; Kosaka et al., 1993). PKC signaling can also induce p21 levels, likely through an ERK-dependent mechanism (Arita et al., 1998; Barboule et al., 1999; Oliva et al., 2008) causing a delay in G2/M transition

Fig. 1-3: PKC-mediated regulation of G1/ S transition

PKC-mediated regulation of the G1-S transition can have either positive (upper portion) or negative (lower portion) effects. In early G1, PKC activation increases cyclin D1 levels but leads to destabilization of p21, resulting in hyperphosphorylation of pocket proteins (e.g., Rb and p107) and progression into S phase.

However, activation of PKC in mid-late G1 reduces cyclin D1 levels while elevating p21. This results in hypophosphorylation of pocket proteins and G0 arrest.

Adapted from (Black, 2010).

Fig. 1-4: PKC-mediated regulation of G2/ M transition

Similar to its impact on the G1-S transition, PKC signaling can have opposing effects on G2-M progression. PKC-mediated phosphorylation of nuclear lamins promotes nuclear lamina disassembly and G2-M transition as depicted in the upper portion. Alternatively, PKC activation can block Cdc25 phosphorylation and result in accumulation of p21, in order to inhibit progression into M phase (bottom portion). Adapted from (Black, 2010).

Apoptosis is another fundamental cellular processes that is regulated by PKC

signaling. Appropriate induction of apoptosis is important for the maintenance of tissue

homeostasis and prevents diseases relating to abnormal cellular growth such as cancer. Apoptosis

pathways are defined by their upstream initiation signals; the extrinsic pathway is related to the activation of death receptors (e.g., TRAILR or Fas) and the intrinsic pathway is initiated by stimuli that trigger cytochrome c release from the mitochondria (Figure 1-5) (Ichim and Tait, 2016). The opposing regulatory roles of PKCε and δ in apoptosis are a clear example of the differential response of PKC isozymes as described in (Griner and Kazanietz, 2007) and (Reyland and

Bradford, 2010). The pro-survival effect of PKCε has been attributed to its ability to activate ERK and AKT, which are two well-known pro-mitogenic, pro-survival effector molecules (Lu et al.,

2006). This signaling crosstalk leads to the upregulation of pro-survival factors BCL-XL and XIAP

and protects the cell against TRAIL-induced apoptosis (Pardo et al., 2006). On the contrary,

activation of PKCδ can promote both intrinsic and extrinsic apoptosis. Upon DNA-damaging insult,

PKCδ undergoes caspase-3 mediated proteolysis, freeing the catalytic domain of the kinase to

translocate to the nucleus where it induces the phosphorylation of apoptotic related substrates

(Bharti et al., 1998; Cross et al., 2000; Frasch et al., 2000; Yuan et al., 1998). Additionally, PKCδ

translocates to the mitochondria and leads to cytochrome c release, which further amplifies caspase-

3 activation (Denning et al., 2002; Li et al., 1999a; Majumder et al., 2000). Therefore, PKCδ also

serves a positive feedback signal to trigger intrinsic apoptosis.

Much of our understanding of PKC functions is based on studies using non-specific phorbol

esters and pharmacological agents, which makes parsing the functional role of each of the ten PKC

isozymes a constant challenge of the field. Other factors that have contributed to the discrepancies

in the field include lack of antibody specificity, inclusion of validated controls in the analysis,

discordant levels of mRNA and protein expression, and the activation status of the enzyme (Garg

et al., 2014; Leitges, 2007). Studies using genetic approaches can provide important functional

insights of PKC enzymes. Despite some phenotypic abnormalities, mice lacking PKC isozyme do

not exhibit developmental defects with the exception of PKCι, in which deletion of this enzyme

was embryonic lethal (Suzuki et al., 2003), making it possible to examine the effect of PKC deletion

in combination with other genetic backgrounds (e.g. mutant Kras or p53 deletion). Therefore,

utilizing both pharmacological and molecular approaches with the incorporation of transgenic

models is critical for the elucidation of the role of PKC isozymes, especially in disease such as

cancer.

Fig. 1-5: Mechanism of PKCδ-mediated apoptosis

Upon DNA damage, either by exposure to UV or DNA-damaging agents, PKCδ is cleaved by caspase-3, resulting in the release of its catalytic domain, which is constitutively active without its regulatory domain. The catalytic fragment can then translocate to the mitochondria to stimulate cytochrome C release and initiate the intrinsic apoptotic pathway as shown in part a. Alternatively, the cleaved catalytic fragment of PKCδ can translocate into the nucleus to phosphorylate substrates involved in cell cycle or apoptosis. The full-length PKCδ can initiate extrinsic apoptosis by modulating the response of death receptors as shown in part b. Adapted from (Griner and Kazanietz, 2007).

1.2 PKC in cancer

The observation that exposure to PMA, which activates PKCs, promotes carcinogenesis in the presence of an additional initiating event (Barrett, 1993; Blumberg et al., 1983) sparked decades of research to develop PKC-targeted therapeutics for cancer. However, these earlier efforts have been futile as most of the inhibitors that were developed showed limited benefit in clinical trials

(Mochly-Rosen et al., 2012). As shown in later studies, this strategic failure was inevitable, due to the complex, context-dependent signaling roles of PKCs, their effects on carcinogenesis, and the promiscuous nature of the pharmacological agents. The reported enzyme depletion under prolonged activation raises the question whether tumor promotion is the result of PKC activity or its loss under prolonged stimulation, further challenging the role of PKC as a tumor promoting kinase. Using gene targeting approaches, PKC isozyme-specific knock-out transgenic mouse models were developed to identify and analyze PKC functions in vivo (Leitges, 2007). From the observations made in genetically modified PKC models, it is evident that the effect of PKC isozymes on carcinogenesis is isozyme-specific and cancer type-dependent (Garg et al., 2014). In human tumors, alterations in PKC expression can result in either increased or decreased levels of the enzyme (Garg et al., 2014). The following section summarizes the evidence supporting oncogenic as well as tumor suppressive properties of PKC from in vivo models and discusses the recent shift in the paradigm to focus on its tumor suppressive effects.

1.2.1 PKC as a tumor promoter

The concept of PKC as tumor promoter stemmed from its activation by phorbol esters, a class of compounds well-known for their ability to promote skin papilloma formation. In the two- step mouse model of skin carcinogenesis, the mutagen 7,12-dimethylbenz[a]anthracene (DMBA) was first applied to the skin. Subsequently, repeated exposure to phorbol esters led to the development of papillomas at the sites previously treated with DMBA (Figure 1-6) (Griner and

Kazanietz, 2007). Of note, no tumor was observed in mice treated with either DMBA or phorbol esters alone, suggesting that both agents have unique contributions to skin carcinogenesis.

Interestingly, reversing the sequence of exposure to these agents (repeated application of phorbol esters followed by DMBA), or lengthening the time between doses of phorbol ester, was insufficient to form papillomas. These observations indicated that, in this model, DMBA was an

“initiator” that acted to induce cell transformation, but was insufficient for tumor development.

Additional insults to the initiated cells, such as exposure to phorbol esters at the appropriate strength and timing, function as “promoters” and are necessary to complete the transformation and result in tumor formation. As signal transduction molecules, several PKC isozymes have been implicated in important pathways that promote tumor growth and proliferation, including crosstalk with

RAS/RAF/ERK and AKT signaling axes.

Fig. 1-6: Phorbol ester as a tumor promoter

In a mouse model, skin papillomas were only observed when treatment with a mutagenic initiator (e.g.

DMBA) was followed by repeated, frequent exposure to a tumor promoter (e.g., phorbol ester). No tumors were observed when only the initiator or promoter were applied, the order of initiator and promoter application was reversed, or the time between administrations of promoter was extended. This model defines the multi-stage nature of carcinogenesis and demonstrates the tumor promoting properties of phorbol ester. Adapted from (Griner and Kazanietz, 2007).

Elevated levels of PKCβII, one of the cPKCs, has been reported in colon cancer patients

(Davidson 1998, Spindler 2009) (although a recent study reported the opposite trend (Dowling et

al., 2016)). Transgenic mice overexpressing PKCβII in the intestinal epithelium led to hyper-

proliferation of the colonic epithelium and heightened sensitivity to carcinogens. These mice

showed increased formation of aberrant crypt foci (ACF), preneoplastic lesions of the colon, after

5-week administration of azoxymethane (AOM) compared with the wild-type (Murray et al., 1999).

In this context, PKCβII functions as an oncoprotein through 1) activation of the Wnt/adenomatous

polyposis coli (APC)/β-catenin pathway leading to decreased glycogen synthase kinase 3β activity

(GSK3β), a known inhibitor of AKT signaling (Murray et al., 1999); and 2) induction of ERK

activation through a RAS-PKCι/RAC1-MEK signaling axis (Zhang et al., 2004). Interestingly, the

colonic epithelial cells from PKCβII-overexpressing mice showed reduced levels of other cPKCs,

PKCα and PKCβI, suggesting possible opposing functions of these isozymes (Gokmen-Polar et al.,

2001).

PKCε, a member of nPKC subclass, is another oncogenic PKC isozyme that acts through

activation of the RAF/MEK/ERK pathway (Cacace et al., 1996). Overexpression of PKCε has

been reported in many human tumor types including breast cancer, non-small cell lung cancer

(NSCLC), and prostate cancer, and is associated with more invasive disease (Bae et al., 2007;

Cornford et al., 1999; Schleifenbaum et al., 2004). In the TRAMP prostate cancer model, deletion

of PKCε inhibited cancer development and metastasis (Hafeez et al., 2013), whereas its deletion in

the epidermis protected the skin from UV-induced transformation (Hafeez et al., 2016). Consistent

with its tumor promoting role in the prostate, tissue-specific overexpression of PKCε regulated by

the probasin promoter led to elevated phosphorylation of AKT and activation of its downstream

effectors S6 kinase and mTOR, as well as an increase in preneoplastic lesions (Benavides et al.,

2011). In the epidermis, PKCε overexpression resulted in induced inflammation, hyperkeratosis,

and hyperplasia (Jansen et al., 2001; Wheeler et al., 2004).

In recent years, PKCι has gained tremendous momentum as an oncogene and a prognostic marker in cancer. Accumulating studies have consistently demonstrated PKCι as an oncogene in

many human cancers (Murray et al., 2011; Regala et al., 2005b). Because of its distinct features as

an aPKC, PKCι has attracted much attention as a therapeutic target as inhibitors of this kinase might

not affect other PKC isozymes. Overexpression of PKCι has been reported in many cancer types

as summarized in (Murray et al., 2011). In most of these cancers, high PKCι expression is

predictive of poor patient survival, recurrence, and metastatic disease (Eder et al., 2005; Ishiguro

et al., 2009; Li et al., 2008; Murray et al., 2011; Regala et al., 2005b; Scotti et al., 2010; Weichert

et al., 2003). Elevated PKCι mRNA and protein can be attributed to gene amplification as reported

in NSCLC (Regala et al., 2005b), serous ovarian tumors (Eder et al., 2005; Zhang et al., 2006),

lung squamous cell carcinoma (LSCC) (Regala et al., 2005b), and esophageal squamous cell cancer

(Yang et al., 2008), or the result of aberrant signaling leading to transcriptional activation of PKCι

(Gustafson et al., 2004). While overexpression due to gene mutation is possible, it is an unlikely

culprit in this case as the most frequent mutation of PKCι affects the substrate binding dibasic motif

rather than increasing its expression (Linch et al., 2013; Murray et al., 2011). Functional analysis

of PKCι in vivo has focused on its oncogenic role in NSCLC, pancreatic ductal adenocarcinoma

(PDAC), and ovarian cancer. Disruption of PKCι expression in cells harboring KRAS mutation

inhibited the tumorigenicity of NSCLC and PDAC cells in a subcutaneous or orthotopic model,

respectively (Regala et al., 2005a; Scotti et al., 2010). In these tumors, PKCι acts as a critical

effector dowmstream of RAS, activating the RAC1/MEK/ERK signaling axis to drive cellular

transformation (Murray et al., 2004; Regala et al., 2005a; Scotti et al., 2010). In ovarian cancer,

PKCι is required for the maintenance of the tumor-initiating phenotype (Wang et al., 2013).

Additionally, PKCι has been found to co-amplify with SOX2 in LSCC, and is required for hedgehog (Hh) signaling and activation of the related transcription factor, GLI1 (Justilien et al.,

2014). These studies provided strong evidence for the ability of PKCι to functions as an oncogene.

Unlike other isozymes which could have opposing roles in carcinogenesis depending on the cellular

context, currently, no studies have reported PKCι as having tumor suppressive properties, further

cementing PKCι as a bona fide oncogene.

1.2.2 PKC as a tumor suppressor

Although PKC is well-known for being activated by the tumor promoting phorbol esters, several of these isozymes have been shown to inhibit cellular transformation and suppress tumor growth. One example is PKCα; mice lacking this kinase exhibited enhanced intestinal and lung tumorigenesis (Hill et al., 2014; Oster and Leitges, 2006). Using the DMBA/PMA skin tumor

model, PKCδ, one of the nPKCs, was similarly identified to function as a tumor suppressor. Mice

with elevated PKCδ expression were resistant to chemically-induced transformation (Reddig et al.,

1999), but PKCδ deficiency did not have the same protective effect against UV radiation-induced squamous cell carcinoma (Aziz et al., 2006). PKCζ has received little attention compared with the other aPKC, PKCι. While both up- and down-regulation of PKCζ has been reported in human cancers (Garg et al., 2014), observation from the Par4-deleted model suggested a tumor suppressive role of PKCζ. As mentioned in the previous section (chapter 1.1.1.), aPKCs are activated by protein-protein interaction; interaction with Par4 leads to PKCζ activation. Without this activating protein-protein interaction, as seen in Par4 knockout transgenic mice, reduced PKCζ activation

enhanced Ras-induced lung carcinoma (Joshi 2008), supporting the tumor suppressive function of

this PKC isozyme.

A recent survey of cancer-associated PKC mutations has propelled the paradigm shift to

focusing on the tumor suppressive function of PKCs. In Antal et al., despite the previously reported

low mutation frequency in genes encoding PKCs, analysis of 50 of the now more than 1000

mutations identified in these isozymes revealed that the majority lead to loss of function (LOF) of

the kinase (Antal et al., 2015). Two-thirds of identified mutations were found to be inactivating

while the rest were inert. These mutations can render the kinase inactive by impeding the binding

of second messengers or preventing priming phosphorylation. Most of these mutations are

heterozygous, yet mice completely devoid of individual PKC isozymes were not embryonic lethal

(with the exception of PKCι knockout mice) and exhibited limited phenotypic aberrations. Taken together, these observations raise the possibility that mutant PKC might have dominant negative functions, or can act as a scaffold to modulate signal transduction (Newton and Brognard, 2017).

It is also likely that maintenance of a threshold of PKC signaling is necessary for its oncogenic

effect – hyperactivation or complete loss of a signaling pathway can be detrimental to

tumorigenesis (Chen et al., 2015). In many of these tumors, the tumor suppression exerted by PKC

is dominant even in context of potent oncogenes such as KRAS (Antal et al., 2015; Hill et al.,

2014), suggesting a regulatory role of PKC in hindering the effect of oncogenes. Taken together, these paradigm-shifting, yet conflicting views on the tumor-related properties of PKC isozymes indicate an incomplete understanding of PKC functions in tumorigenesis, and warrant a thorough examination of each isozyme in a tumor-specific manner to better understand the signaling role of

PKC and its impact on tumor biology.

1.3 Protein kinase C alpha (PKCα)

PKCα is a member of the cPKC subclass of PKC isozymes that are activated by DAG (in the presence of Ca2+ and phosphotidylserine) and phorbol esters. Experimentally, PKCα can be

stimulated by pharmacological activators such as PMA, DiC8, and bryostatin-1 and inhibited by bisindolylmaleimide I (Bim-1; a pan-PKC inhibitor) and Gӧ6976 (a classical PKC inhibitor). PKCα activity is involved in the crosstalk of many signaling pathways that regulate a plethora of cellular processes including cell cycle progression, differentiation, and apoptosis (Nakashima, 2002).

Because this kinase is ubiquitously expressed in many tissue, the role of PKCα in normal tissue development as well as tumorigenesis has been the focus of many studies. However, these efforts have generated conflicting results and further mystified the signaling impact of PKCα. The current consensus in regard to the function of PKCα (and most other PKC isozymes) is that whether it

positively or negatively impacts cell growth or acts to promote or suppress tumor formation is dependent on the cellular context (Garg et al., 2014). As summarized in Griner 2007, overexpression of PKCα has been reported in glioma, breast and bladder cancers, while its downregulation was observed in cancers such as in the colon and renal cell carcinoma (Griner and

Kazanietz, 2007) (Figure 1-7). Specifically, 60% of colorectal tumors reported loss of or reduced

PKCα expression (Suga et al., 1998). In T cell acute lymphoblastic leukemia (T-ALL), low PKCα levels were predictive of poor clinical outcome (Milani et al., 2014).

Ex vivo manipulation of PKCα expression has generated evidence for the pro-tumorigenic function of this kinase in some cancer types. In glioma cells, PKCα enhances proliferation and confers resistance to radiation and chemotherapy agents (Baltuch et al., 1995; Blackburn et al.,

1998; Cameron et al., 2008; Mandil et al., 2001). Interestingly, depletion of PKCα but not the inhibition of its activity impairs growth of glioma cell lines, indicating PKCα functions independent of its activity to promote the growth of glioma cells (Cameron et al., 2008). Similarly, ectopic expression of PKCα in MCF-7 breast cancer cells led to phenotypic changes consistent with increased tumorigenic potential, including increased proliferation, enhanced anchorage- independent growth, and change in cell morphology that was indicative of EMT (Ways et al., 1995).

In addition, PKCα has been implicated in facilitating the loss of estrogen receptor (ER) expression in breast cancer cells (Tonetti et al., 2000; Ways et al., 1995), leading to more aggressive disease that is insensitive to tamoxifen (Chisamore et al., 2001). Assessment of human breast tumors further corroborated the correlation of elevated PKCα levels with reduced progesterone receptor

(PR) expression, poor response to endocrine treatment, and worse prognosis in patients (Assender et al., 2007; Lonne et al., 2010; Tonetti et al., 2003).

Fig. 1-7: PKCα expression in human cancers

PKCα expression levels in multiple human cancer types based on the RNA-seq data generated from The

Cancer Genome Altas. Graph obtained from The Human Protein Atlas.

Despite the evidence supporting an oncogenic role of PKCα, transgenic models demonstrated its tumor suppressive effect in both intestinal and lung carcinogenesis (Figure 1-8).

In an ApcMin/+ background, mice lacking PKCα developed more aggressive intestinal tumors, larger in size, and had poor survival compared with their PKCα wild type littermate controls (Oster and

Leitges, 2006). Additionally, deletion of PKCα alone increased the number of spontaneous intestinal lesions independently of the ApcMin/+ mutation. Another key evidence of PKCα as a tumor

suppressor was from Fields and colleagues (Hill et al., 2014). As reported in this study, PKCα is

frequently lost in the tumors of lung cancer patients, and is associated with more advanced disease.

In mutant Kras-driven lung tumorigenesis, PKCα deletion enhanced tumor development and progression by facilitating the bypass of oncogene-induced senescence and expansion of the tumor- initiating bronchoalveolar stem cells (BASCs). Mechanistically, PKCα acts as a tumor suppressor in BASCs through activation of p38MAPK-TGFβ signaling axis, in part through inhibition of the expression of inhibitor of DNA binding 1-3 (Id1-3), which promote cell proliferation and inhibit differentiation (Ling et al., 2006). PKCα has also been shown to negatively regulate Id family protein expression in colon cancer (Hao et al., 2011; Pysz et al., 2009). In the skin, PKCα confers protection against skin tumor formation in a two-stage carcinogenesis system using DMBA as an initiator and PMA as a tumor promoter (as previously described in Chapter 1.2.1) (Hara et al.,

2005). However, there was no difference in tumor growth or progression between PKCα-/- or wild type mice, suggesting PKCα is involved in the early events of carcinogenesis but not progression

of chemically induced skin tumors (Hara et al., 2012)

Fig. 1-8: PKCα as a tumor suppressor

In the intestine, knockout of PKCα alone increased the number of spontaneous lesions formed. When paired with an initiating event such as in APCMin/+ and mutant Kras mouse models, loss of PKCα significantly enhances intestinal and lung tumorigenesis. Adapted from (Garg et al., 2014)

1.3.1 PKCα in intestinal homeostasis and carcinogenesis

The mammalian intestinal epithelium comprises many crypt-villus units that undergo

constant renewal every 4-5 days (for the small intestine). Within the crypt reside the proliferating

cells whose self-renewal capability repopulates the mucosal compartment. Starting at the crypt

bottom, the self-renewing stem cells give rise to proliferating progenitor cells, establishing the

proliferation zone. As these cells travel upwards along the crypt onto the villus, they undergo cell

cycle arrest and become committed differentiated cells. Near the tip of the villus, the cells undergo

apoptosis and eventually bud off into the lumen of the intestine (Figure 1-11 Picture of the small

intestine). Emerging from the extensive studies of the intestinal system are the crypt base columnar stem cell and the +4 stem cell models, whose namesake defined the cell population responsible for intestinal regeneration as described in (Clevers, 2013; van der Flier and Clevers, 2009). The uniquely, well-defined progression from proliferation to differentiation as the cells migrate along the crypt-villus axis makes the intestinal epithelium a valuable system for studying how the intestine maintains its homeostasis and regeneration, as well as the aberrant molecular changes associated with its malignant transformation.

One of the pivotal role of PKC isozymes in cellular growth is to regulate components of the cell cycle machinery, primarily at the G1S and the G2-M transitions (Black, 2000). Research from our laboratory has focused on understanding the role of PKCs (and PKCα in particular) in the tightly regulated intestinal regenerative process and its link to intestinal tumor development (Figure

1-9). In the non-transformed, immortalized rat intestinal epithelial cell (IEC-18) model system, our biochemical analyses detected multiple PKC isozymes, including α, βII, δ, ε, η, ζ and ι (Saxon et al., 1994). Among these isozymes, several PKCs (α, βII, δ, and ζ) are activated at the crypt-villus junction where the cells cease to proliferate and start to differentiate, as marked by their translocation from the cytoplasm to membrane compartments (Frey et al., 2000). This observation suggested that several PKC isozymes may be involved in negative regulation of intestinal cell proliferation. Data from our laboratory have further shown that activation of PKCs, and PKCα in particular, triggers a cascade of growth regulatory events, partly through an ERK-dependent mechanism (Clark et al., 2004). Contrary to the more well-known growth promoting property of

RAS/RAF/ERK signaling, sustained ERK activation as caused by prolonged PKCα activation resulted in growth inhibitory effects (Clark et al., 2004). Additionally, selective inhibition of PKCα or inhibition of ERK signaling abolished the ability of PKC agonists to induce cell cycle withdrawal

(Clark et al., 2004). Treatment of unsynchronized IEC-18 cells with non-selective pharmacological

PKC agonists led to cell cycle arrest in late G1 phase, as well as delayed transition through G2/M.

PKC activation resulted in induction of the known growth inhibitory cyclin-dependent kinase

inhibitors p21 and p27, decrease expression of p107 but increase expression of p130, as well as alterations in the expression and alteration in the phosphorylation of pocket proteins (e.g. p107, pRb, and p130) (Frey et al., 2000; Frey et al., 1997). These effects could be recapitulated by selective activation of PKCα alone (Frey et al., 2000). Furthermore, PKCα-mediated ERK activation was shown to downregulate growth promoting effectors (e.g., cyclin D1 and Id-1) through translational and/or transcriptional repression. Cyclin D1 senses extracellular mitogenic signals and promotes G1 transition through association with cdk4 and cdk6. In intestinal epithelial cells, PKCα signaling induces the binding of the translational repressor 4E-BP1 with the cap- dependent translational machinery, sequestering cyclin D1 mRNA and preventing its translation

(Guan et al., 2007; Hizli et al., 2006). In the case of the inhibitor of DNA binding (Id1-4) family

proteins, their decreased expression is attributed to the negative impact of PKCα activation on their

transcription (Hao et al., 2011). Members of Id family proteins are dominant negative antagonists

of antiproliferative, differentiation-inducing transcription factors such as basic helix-loop-helix

factors, Ets2, and Pax family members. Binding of Id1 negates the transcription activating function

of these factors by blocking their interaction with DNA. Interestingly, Id1 expression is generally

restricted to the proliferating crypt, opposite of the pattern observed for PKCα activity in the

intestine. Our work thus far demonstrated that PKCα helps to maintain intestinal homeostasis by

acting as a negative regulatory signal of epithelial proliferation.

Fig. 1-9: PKCα activation in the intestinal systems

PKCα activation, as indicated by its membrane association, correlates with intestinal epithelial cell growth arrest and differentiation. V, villus; C, crypt; P, progenitor cells; L, lumen. Adapted from (Saxon et al.,

1994).

The importance of PKCα as a gatekeeper for intestinal epithelial growth is further supported by the loss of its expression in intestinal tumors (Figure 1-10). Reduced PKCα expression is observed in multiple genetic models that develop intestinal neoplasms due to mutation or loss of Apc, β-catenin, KRas, and Muc2, (Pysz et al., 2009) and PKCα deficiency sensitizes the

animals to azoxymethane-induced carcinogenesis (unpublished data). Loss or reduction of PKCα expression is also observed in human colon tumors (Chen et al., 2016; Gwak et al., 2009; Hao et al., 2011). These findings strongly suggest that loss of PKCα is a general characteristic of intestinal tumors and is a critical step in colon tumor development. While PKCα signaling reduces intestinal tumorigenicity by blocking anchorage-independent growth of colon cancer cells, the disruption of

PKCα signaling results in a growth promoting cellular milieu, through modulation of the transcription and/or translation of cyclin D1 and Id1 (Hao et al., 2011; Pysz et al., 2009). While both PKCα and δ expression are reduced in intestinal tumors, restoration of PKCα confers stronger antitumor effects in colon cancer cells. Consistent with previous results in IEC-18 cells, the ability of PKCα to regulate cyclin D1 and Id1 in colon cancer cells requires ERK activation (likely via crosstalk at the level of Raf-1), and is independent of effects on PI3K/AKT signaling (Hao et al.,

2011). Our findings, together with the enhanced tumor formation in PKCα KO animals (Oster and

Leitges, 2006), provide strong in vivo and in vitro evidence for the growth and tumor suppressive role of PKCα in the intestine.

Fig. 1-10: PKCα expression in lost in intestinal tumors

Expression of PKCα is lost in A) multiple in vivo models of intestinal tumors; and B) human colon cancer cell lines; and C) human colorectal tumors. Adapted from (Pysz et al., 2009; Verstovsek et al., 1998)

1.3.2 Impact of PKCα signaling in cancer

As mentioned in previous sections, the active form of PKCα resides at the plasma membrane, allowing it to transduce signals from upstream receptors to many signaling pathways that are important in cancer, regulating major cellular processes like proliferation, apoptosis, differentiation, and motility. However, the biological outcome of PKCα signaling can be influenced by the duration of its activation, the interacting substrates available, and the molecular context of the cell. In breast cancer, PKCα may confer estrogen-independent, tamoxifen-resistant growth through Notch and ERK signaling (Pham and Tonetti, 2016; Tonetti et al., 2003; Yun et al., 2013), whereas PKCα-mediated ERK activation led to growth inhibition in hepatoma and intestinal epithelial cells (Clark et al., 2004; Wen-Sheng and Jun-Ming, 2005). Additionally, PKCα has also been shown to promote cell invasion and migration through NFκB activation (Cheng et al., 2009;

Mahanivong et al., 2008; Mut et al., 2010), while crosstalk with Wnt (Gwak et al., 2009) and

Hedgehog (Neill et al., 2003) signaling pathways inhibits tumor growth. PKCα activity has been shown to have opposing effects on AKT phosphorylation. Overexpression of PKCα has been shown to induce AKT activation in immune cells (Li et al., 1999b; Yang et al., 2010), keratinocytes

(Li et al., 2006), and endothelial cells (Partovian and Simons, 2004). In other cellular systems,

PKCα activity was found to suppress AKT phosphorylation (Motley et al., 2002) and dampen the activity of other components of the PI3K signaling pathway (Motley et al., 2003; Sipeki et al.,

2006). Despite the overwhelming evidence showing that PKCα indeed intercepts a wide network of signaling axes, understanding the biological impact of PKCα involvement requires careful examination in the cell type of interest. Our study presented here focuses on the role of PKCα on

PI3K/AKT signaling, specifically its impact on endometrial carcinogenesis driven by hyperactivation of AKT.

1.4 AKT signaling network

The discovery and cloning of AKT, which is also named protein kinase B (PKB) because of its similarity with PKA and PKC, identified it as a serine/threonine specific kinase activated downstream of PI3K (Bellacosa et al., 1991; Coffer and Woodgett, 1991; Jones et al., 1991; Staal,

1987). The mammalian genome encodes three isoforms of AKT - AKT1 (PKBα), AKT2 (PKBβ), and AKT3 (PKBγ). Many cell lines express all three isoforms of AKTs; however, AKT1 and

AKT2 are more ubiquitously expressed. Defining the isoform-specific functions of AKTs has been an exciting new frontier for the AKT field. Thus far, data from isoform-specific knockout models and RNA interference experiments have shown that AKT1 has significant developmental roles,

AKT2 is involved in glucose homeostasis, and AKT3 is important for brain development (Dummler and Hemmings, 2007). The importance of AKT as a signaling molecule is underscored by its numerous downstream targets that regulate key biological processes (e.g. metabolism, growth and survival) to establish a pro-proliferative cellular milieu. To promote cell survival, AKT blocks the function and expression of pro-apoptotic BH3-only proteins. AKT activity leads to activation of mTORC1 signaling, which in turn promotes protein translation and cell growth. Additionally, AKT is involved in responses to insulin and other metabolic events such as nutrient uptake and lipid and glucose metabolism. The following sections will discuss the regulation of AKT activation, the functions of its downstream targets GSK3β, PRAS40, and FOXO proteins, and hyperactivation of

PI3K/AKT signaling in cancer.

1.4.1 Regulation of AKT activation

When growth factors interact with their cognate receptor tyrosine kinases (e.g. EGFR,

HER2, and KIT) or G-protein coupled receptors (GPCRs) (Katso et al., 2001; Vanhaesebroeck and

Waterfield, 1999), PI3K is recruited to the cytoplasmic portion of the receptors either by direct interaction with p85 (the regulatory subunit of PI3K) or through an adaptor protein. The catalytic

subunit of PI3K, p110, then phosphorylates the lipid second messenger phosphoatidylinositol-3,4-

P2 (PIP2) to generate phosphoatidylinositol -3,4,5-P3 (PIP3) at the plasma membrane (Fruman et al.,

1998). Due to the high affinity of PIP3 for the N-terminal PH domain of AKT, the accumulation of PIP3 relocates inactive AKT to the plasma membrane. While PIP2 also binds to the PH domain,

it only induces partial AKT activation (Franke et al., 1997; Frech et al., 1997; Klippel et al., 1997).

Binding of PIP3 induces a conformational change in AKT, exposing two key residues – Thr308 in

AKT1 and 2/ Thr309 in AKT3 in the kinase domain and Ser473 in AKT1/ Ser474 in AKT2/ Ser472

in AKT3 in the C-terminal regulatory domain – giving kinases access to carry out phosphorylation

at these two sites for maximal AKT activation (Alessi et al., 1996). PIP3 also recruits the

phosphoinositide-dependent protein kinase 1 (PDK-1) to the membrane and puts it in proximity with AKT, leading to the phosphorylation of Thr308 (Alessi et al., 1997; Andjelkovic et al., 1997).

The full activation of AKT requires the mechanistic target of rapamycin complex (mTORC2) to phosphorylate AKT at Ser473 (Sarbassov et al., 2005). A phosphomimetic mutant of AKT

(T308D/S473D) generates constitutively active kinase that is PI3K independent (Alessi et al.,

1996). Both PDK-1 and mTORC2 also phosphorylate other AGC kinases such as PKC and S6K1.

However, unlike other kinases that are constitutively phosphorylated by PDK-1 and mTORC2, phosphorylation of AKT is PIP3-obligatory (Collins et al., 2003; Huang et al., 2008; McManus et al., 2005; Sarbassov et al., 2005). Recent evidence has shown that the pool of PIP3 and PIP2 found

on the endosomal membrane can also lead to AKT redistribution and activation, suggesting there

is an additional layer of spatial control of AKT phosphorylation and function yet to be explored

(Jethwa et al., 2015).

Termination of AKT signaling is achieved by a number of phosphatases that either counteract upstream lipid synthesis or directly dephosphorylate AKT. In the late 1990s, the phosphatase and tensin homolog (PTEN) was cloned and identified as a key regulator of PI3K signaling (Frech et al., 1997; Li et al., 1997; Maehama and Dixon, 1998). PTEN possesses dual

phosphatase function and can, therefore, act on both phosphoinositide and polypeptide substrates.

The lipid phosphatase function of PTEN dephosphorylates PIP3 at the 3-position and converts it to

P(4,5)P2, thus decreasing membrane PIP3 concentration and negating the effects of PI3K. Because

of this intricate balance of PTEN and PI3K functions, the lipid synthesis in response to growth

factor stimulation is often rapid and transient (Auger et al., 1989). Alternatively, the Src homology

2 domain-containing inositol 5’-phosphatase (SHIP) can dephosphorylate PIP3 at the 5-position and convert it to P(3,4)P2. The phospholipid is further hydrolyzed by inositol polyphosphate 4-

phosphatase (INPP4B) to P(3)P. This step-wise dephosphorylation of P(3,4,5)P3  P(3,4)P2 

P(3)P preferentially acts on phospholipids at endosomes and loss of INPP4B leads to AKT2

activation (Braccini et al., 2015; Fedele et al., 2010; Gewinner et al., 2009; Li Chew et al., 2015).

These observations suggest there is spatial restriction on the species of PI3K lipids present on membranes that might be involved in isoform-specific regulation of AKT activation. Together,

PTEN, SHIP, and INPP4B negatively regulate PI3K-mediated lipid synthesis and reduce the membrane concentration of P(3,4,5)P3. This reduction in membrane P(3,4,5)P3 not only affects the recruitment and activation of AKT but also other PH-domain containing proteins including PDK-

1, thus further exerting negative pressure on PI3K/AKT signaling (Manning and Toker, 2017;

Salmena et al., 2008).

Activation of AKT can also be terminated through direct modulation of its phosphorylation. Specific phosphatases act directly on AKT by removing the phosphate group from

Ser473 and Thr308, whose phosphorylation is critical for AKT activation. Protein phosphatase 2A

(PP2A) represents a family of holoenzymes that effectively dephosphorylate AKT (Andjelkovic et al., 1996)). This serine and threonine specific phosphatase is ubiquitously expressed in eukaryotic cells. Its involvement in major signaling pathways that regulate processes such as cell cycle, metabolism, migration, and survival demonstrates that reversible phosphorylation is a crucial means to ensure proper firing and fine tuning of signaling events in cells (Janssens and Goris, 2001;

Perrotti and Neviani, 2013; Schonthal, 2001; Seshacharyulu et al., 2013). The PP2A holoezyme consists of a core dimer of the structural subunit (A) and the catalytic subunit (C). Its substrate

specificity is dictated by the regulatory subunit (B), which binds to the subunit A/C dimer. In humans, 26 different B regulatory subunits have been identified and are categorized into 4 subfamilies: B/B55/PR55, B’/B56/PR61, B’’/B72/PR72, and B’’’/PR93 (SG2NA)/PR110

(Striatin). While the isoforms of the A and C subunits share high sequence homology, the B subunit subfamilies are distinct in their sequences and structures, which confers their substrate selectivity

and contributes to the diverse functions and location of PP2A holoenzymes (Chen et al., 2013;

Eichhorn et al., 2009; Sents et al., 2013). The PP2A-B55alpha holoenzyme has been shown to

mediate AKT dephosphorylation at Thr308 (Kuo et al., 2008), while the PP2A-B56 holoenzyme

acts to reverse phosphorylation of Ser473 (Rocher et al., 2007; Van Kanegan et al., 2005), making

this phosphatase a prominent negative regulator of PI3K signaling. In 2005, the Pleckstrin

homology domain leucine-rich repeat protein phosphatase (PHLPP) was identified as another

family of serine/threonine specific phosphatases that act on Ser473 of AKT (Gao et al., 2005).

There are three members in the PHLPP family: the splice variants PHLPP1alpha and PHLPP1beta

and PHLPP2. The PHLPP isoforms display preferential dephosphorylation of AKT isoforms.

PHLPP1 mainly acts on AKT2 and AKT3, while PHLPP2 mediates inactivation of AKT 1 and

AKT3 (Brognard et al., 2007).

In addition, other mechanisms triggered by active PI3K signaling can lead to negative

feedback inhibition. AKT activates mTOR signaling to enhance protein translation and cell growth.

As a result, mTORC1 promotes IRS1 and IRS2 degradation and phosphorylates GRB10 to

attenuate insulin-induced signaling (Harrington et al., 2004; Hsu et al., 2011; Shah and Hunter,

2006; Tzatsos and Kandror, 2006; Yu et al., 2011). Another kinase activated by mTOR signaling,

S6K, has been shown to phosphorylate mTORC2 component Rictor and decrease phosphorylation

of Ser473 of AKT by mTORC2 (Dibble et al., 2009; Julien et al., 2010; Liu et al., 2013a). Being

one of the most potent growth promoting molecules, a multitude of regulatory pathways

coordinated by phosphatases and intrinsic feedback responses ensure tight control of the timing and

duration of AKT activation. As a result, activation of AKT often occurs rapidly after mitogen

stimulation and is usually short-lived. Defects in these regulatory mechanisms can result in detrimental developmental abnormalities and are associated with overgrowth syndromes and cancer in humans.

1.4.2 Downstream targets of AKT

As the key effector of the PI3K signaling axis, AKT acts on over 100 downstream substrates that contribute to the expanding repertoire of AKT functions. The enzymatic property of AKT kinases is fully activated by phosphorylation at Thr308 and Ser473, and the active enzymes then phosphorylate its downstream targets at the serine and threonine residues within the minimal consensus motif R-X-R-X-X-S/T-Φ (X is any amino acid, Φ is a large hydrophobic residue). This phosphorylation leads to activation or inhibition of downstream targets. As summarized in the review by Manning and Toker (Manning and Toker, 2017), the most recognized signaling role of

AKT is mediated through three functional nodes: glycogen synthase kinase-3, TSC2/mTORC1, and Forkhead Box O transcription factors (Figure 1-11).

Fig. 1-11: AKT signaling network

When receptor tyrosine kinases are stimulated by extracellular signaling (e.g., insulin), PI3K is activated and converts PIP2 to PIP3, resulting in membrane recruitment of AKT to the proximity of its activating kinases (e.g., mTORC2 and PDK1). The phospholipid conversion can be reversed by PTEN to oppose

AKT activation. The fully activated AKT phosphorylates its downstream targets such a GSK3β,

PRAS40, and FOXO to regulate apoptosis, metabolism, translation, and protein and lipid synthesis.

Adapted from (Manning and Toker, 2017)

Glycogen synthase kinase-3 (GSK-3) was the first reported AKT substrate (Cross et al.,

1995). This serine/threonine protein kinase is ubiquitously expressed in all tissues and highly conserved across species. In 1990, cloning of cDNA from rat brain identified two distinct genes that encoded two different GSK-3 isozymes – GSK-3α and GSK-3β. Overall, the two GSK-3 proteins share 85% sequence homology, with an almost identical kinase domain, making specific targeting of these two isoenzymes a challenge. GSK-3α and GSK-3β are mostly functionally redundant; however, some studies have identified isozyme-specific functions in certain tissues

(Kaidanovich-Beilin and Woodgett, 2011). The multitude of GSK-3 functions is demonstrated by

its extensive number of downstream targets, which are involved in biological processes ranging

from development to circadian rhythm, affecting diseases such as diabetes and cancer

(Kaidanovich-Beilin and Woodgett, 2011). Amongst the vast number of targets is glycogen

synthase, the first known protein targeted by GSK-3. Initial discovery of GSK-3 identified it as the

regulatory kinase for glycogen synthase, a rate limiting enzyme in glycogen formation whose

activity is regulated by phosphorylation (Embi et al., 1980; Vandenheede et al., 1980). Unlike most

kinases, GSK-3 is active under steady state and becomes inactivated by extracellular signaling such

as insulin stimulation. Phosphorylation at tyrosine residue 270 for GSK3α and 216 for GSK3β,

mediated by kinases such as FYN2 and PYK2 or by autophosphorylation (Cole et al., 2004;

Lochhead et al., 2006), positively regulates GSK-3 activity. On the other hand, phosphorylation at

a serine residue at the N-terminal (Ser 21 for GSK3α and Ser9 for GSK3β) mediated by AKT

results in GSK-3 inhibition (Stambolic and Woodgett, 1994; Sutherland and Cohen, 1994;

Sutherland et al., 1993). It has been shown that the phosphorylated serine generates an intramolecular pseudosubstrate that binds to the substrate binding pocket of the kinase and obstructs actual substrate binding (Dajani et al., 2001; Frame and Cohen, 2001). Without stimulation, active

GSK-3 has inhibitory effects on its targets either by blocking their activity or destabilizing the

protein. Under the stimulation of extracellular factors, serine phosphorylation inhibits GSK-3

activity and releases the inhibitory regulation of GSK-3 targeted proteins. For example, GSK-3-

mediated phosphorylation hinders the activity of glycogen synthase and glycogen production.

When AKT is activated upon insulin stimulation, AKT phosphorylates GSK-3 at its serine residue, causing it to dissociate from glycogen synthase. Following this relief, there is an induction in glycogen synthase activity and an increase in glycogen formation and deposition in multiple tissues

(e.g. muscle and liver). This crucial involvement of GSK-3 in glycogen synthesis demonstrated its direct influence on cell metabolism. Later studies have identified additional GSK-3 targets such as sterol regulatory element binding protein 1 c (SREBP1c), hypoxia inducible factor 1 α (HIF-1α), and nuclear factor erythroid 2-related factor 2 (NRF2), showing that GSK-3 can also regulate metabolism in an indirect manner. For other targets, GSK-3-mediated phosphorylation results in their degradation. Proteins such as the pro-apoptotic protein Mcl-1 and pro-proliferative transcription factor c-Myc are first primed by kinases such as JNK and ERK respectively, which allows their phosphorylation by GSK-3 (Ding et al., 2007; Maurer et al., 2006; Morel et al., 2009;

Sears et al., 2000; Welcker et al., 2003). GSK-3-mediated phosphorylation of Mcl-1 and c-Myc marks them for recognition by specific E3 ubiquitin ligases, directing them to proteosomal degradation. Therefore, activation of AKT and subsequent GSK-3 inhibition stabilize both Mcl-1 and c-Myc to promote cell survival and proliferation. Additionally, GSK-3 activity can be regulated by its subcellular localization. Although GSK-3 is mostly observed in the cytoplasm, there is evidence that its nuclear localization occurs dynamically depending on cell cycle phase

(Bechard and Dalton, 2009; Liu et al., 2013b; Meares and Jope, 2007). It has also been reported that AKT can decrease nuclear levels of GSK-3 (Bijur and Jope, 2001, 2003). However, the significance of nuclear localization on regulation of GSK-3 activity and function remains unclear.

The second key signaling node regulated by AKT involves TSC2/mTORC1. Cellular growth requires increased production of proteins, lipids, and nucleotides. The mTOR signaling network, whose activation is controlled by AKT, is crucial in regulating the production of these biomolecules and the maintenance of metabolic balance in response to the cellular environment with the goal of promoting cell growth and division. Mechanistic (formerly known as mammalian)

target of rapamycin, or mTOR was first identified in 1994 as the direct binding target of the

FKBP12-rapamycin complex (Brown et al., 1994; Sabatini et al., 1994; Sabers et al., 1995). This interaction inhibited mTOR kinase activity, thus hindering cell growth and proliferation. This inhibitory effect on growth attracted significant attention to the clinical potential of rapamycin as

well as the pro-growth function of mTOR signaling. More than 20 years since its discovery, mTOR

is now identified as the catalytic component of two distinct complexes, mTORC1 and mTORC2

differing in their interacting partners. The core complex of mTORC1 consists of mTOR, mLST8,

and Raptor (Nojima et al., 2003; Schalm et al., 2003), which is substituted by Rictor in mTORC2

(Jacinto et al., 2006; Sarbassov et al., 2004). Functionally, the two mTORC complexes act at

different points along the PI3K/AKT signaling axis. As previously mentioned, mTORC2 activates

AKT by phosphorylating it at the Ser473 residue (Sarbassov et al., 2005). It is worth noting that

PKCα and other members of the PKC family are also phosphorylated by mTORC2 (Gan et al.,

2012; Jacinto et al., 2004; Li and Gao, 2014; Sarbassov et al., 2004; Thomanetz et al., 2013). On

the other hand, AKT acts upstream of mTORC1 to regulate its activation. Activated mTORC

signaling leads to increased production of biomolecules essential for cellular growth (e.g. protein,

nucleotides, and lipid) through phosphorylation of key effectors such as S6K1, 4EBP1, ATF4,

Lipin1, and HIF-1α and heightened glucose metabolism to meet the needs of a proliferating cell.

Additionally, mTORC1 suppresses catabolic pathways such as autophagy, shifting the metabolic

balance in favor of cellular growth. Under nutrient or serum deprivation, binding of the proline-

rich AKT substrate of 40 kDa (PRAS40) keeps mTOR in an inactive state (Sancak et al., 2007;

Vander Haar et al., 2007; Wang et al., 2007). As a direct substrate of AKT, PRAS40 is

phosphorylated upon insulin stimulation. This modification significantly weakens the physical

interaction between PRAS40 and mTORC1, thus relieving the inhibitory regulation on mTORC1,

allowing it to activate the aforementioned downstream, pro-growth effectors (Sancak et al., 2007).

To ensure unperturbed mTORC1 activation, phosphorylated PRAS40 is further sequestered by

binding to 14-3-3.

The third important signaling node is through the Forkhead Box O (Fox) transcription factor family, specifically the “O” class proteins that are regulated by the insulin/PI3K/AKT signaling. The four members in this class – FOXO1, 3, 4, and 6 – share structural similarity and have four distinct functional motifs including forkhead, nuclear localization, nuclear export, and transactivation domains. Among the FOXO proteins, FOXO1, FOXO3, and FOXO4 are

ubiquitously expressed, although at different levels (Anderson et al., 1998; Furuyama et al., 2000),

whereas FOXO6 is primarily restricted to the central nervous system (Jacobs et al., 2003). The

FOXO genes encoding FOXO 1, 3, and 4 were first identified in chromosomal translocation events in rhabdomyosarcoma and acute myeloid leukemia (Dumont et al., 2012; Linardic, 2008; Schmitt-

Ney and Camussi, 2015; Wachtel and Schafer, 2015), suggesting their possible role in carcinogenesis. Soon after, studies of DAF-16, the homolog of FOXO proteins in C. elegans found the transcription factor to be crucial for the longevity of the organism and was negatively regulated by the insulin/PI3K/AKT signaling pathway (Hesp et al., 2015; Lin et al., 2001; Sun et al., 2017).

In C. elegans, the phenotype caused by deletion of AKT1 and 2 can be rescued if DAF-16 is also lost (Paradis and Ruvkun, 1998). This negative regulation of DAF-16/FOXO proteins is evolutionarily conserved. In mice, FOXO deletion (mainly FOXO1 in the liver) can reverse the severe hepatic insulin resistance and hyperglycemia caused by liver-specific deletion of AKT 1 and

2 (Lu et al., 2012). In vivo genetic manipulation reveals that the different FOXO proteins have non-redundant functions. Mice lacking FOXO1 succumb to defects at embryonic day 10.5 due to incomplete vascular development (Hosaka et al., 2004). On the other hand, FOXO3 null mice are viable, despite lymphocyte proliferation and autoinflammation associated with abnormal regulation of NFκB signaling (Lin et al., 2004), age-dependent infertility due to abnormal ovarian follicle development (Hosaka et al., 2004), and a decline in the neural stem cell pool (Renault et al., 2009).

Mice with FOXO4 deletion are also viable with exacerbated colitis in response to inflammatory stimuli (Zhou et al., 2009). As FOXO6 is mostly detected in the neural system, deletion of this isoform is reported to result in defects in memory consolidation (Salih et al., 2012). As they are

transcription factors, FOXO proteins are subjected to regulation by phosphorylation that alters their subcellular localization. Inactive FOXO proteins are found in the cytoplasm. Under oxidative or nutrient stress, kinases such as JNK, Mst1, or AMPK have been shown to phosphorylate FOXOs and induce their translocation from the cytoplasm to the nucleus (Coomans de Brachene and

Demoulin, 2016; Wang et al., 2014). On the other hand, AKT-mediated phosphorylation at three conserved sites (T24, S256, S319 for FOXO1; T32, S253, S315 for FOXO3A; and T32, S197,

S262 for FOXO4) generates a recognition motif for 14-3-3. This interaction facilitates the nuclear export of FOXO proteins and sequesters them in the cytosol, thus inhibits FOXO-regulated gene expression. Other post-translational modifications such as acetylation by SIRT1 and monoubiquitination have been shown to enhance FOXO-mediated transcription under oxidative stress (Boccitto and Kalb, 2011; Daitoku et al., 2011; Hariharan et al., 2010; Huang and Tindall,

2011). Similar to other transcription factors, the activity of FOXO proteins is also influenced by their binding to either coactivator or corepressors. FOXO-mediated transcription governs multiple cellular processes including proliferation, apoptosis, oxidative stress, etc. (Wang et al., 2014). In regard to cancinogenesis, FOXO proteins act as tumor suppressors. In colorectal cancer, there is decreased expression of FOXO4 (Xiang-qiang et al., 2011), while FOXO3 contributes to the repression of Myc function (Delpuech et al., 2007). Studies have shown that FOXO1 upregulates the expression of TNFα and FasL in prostate cancer and glioma, respectively (Ciechomska et al.,

2003; Modur et al., 2002), to initiate apoptosis. In CML and breast cancer, FOXO3 increases Bim expression in response to the chemotherapeutic agents STI571 and paclitaxel and inhibits tumor

+ growth (Essafi et al., 2005; Sunters et al., 2003). In HER2 breast cancer, FOXO4 promotes G1 and

G2 cell cycle arrest by induction of p27 and GADD45 (Yang et al., 2005). Both FOXO1 and

FOXO4 have been shown to limit metastasis by inhibiting RUNX2 activity or increasing ANX8

expression, respectively (Lee et al., 2009; Zhang et al., 2011). On the other hand, FOXO1- and

FOXO3-mediated transcription leads to expression of genes such as PIK3CA and confers chemo resistance in ovarian cancer and leukemic cells (Goto et al., 2008).

1.4.3 Hyperactivation of AKT in cancer

As described above, AKT alters a wide range of downstream effectors through GSK3β, mTOR, and FOXO signaling nodes to exert its pro-growth and pro-survival functions, making it a powerful oncogenic force in carcinogenesis. Indeed, hyperactivation of AKT is one of the most frequently observed molecular features in human solid and hematological malignancies (Samuels and Ericson, 2006; Shi et al., 2005). In cancer, AKT hyperactivation is often the result of defects

in its upstream regulators, either through 1) amplification or oncogenic mutations in EGFR, HER2,

or other RTKs, or kinases such as PDK1 and PI3K, and/or 2) inactivating mutations or loss of

heterozygosity in genes that negatively regulate PI3K/AKT signaling such as PTEN, INPP4B,

LKB1, and PHLPP as summarized in Table 1 (Cheung and Testa, 2013; Mayer and Arteaga, 2016).

Interestingly, PI3KCA mutations alone do not always correlate with hyperactivation of AKT and mutations in multiple proteins involved in the PI3K/AKT signaling axis is often observed in tumors

(Lien et al., 2017; Vasudevan et al., 2009). This suggests that components of the PI3K/AKT signaling network function co-operatively to regulate tumorigenesis. Although AKT mutation occurs at a lower frequency than its upstream regulators, an activating somatic E17K mutant has been observed in breast, colorectal, lung, ovarian, and endometrial cancers, as well as melanoma

(Carpten et al., 2007; Davies et al., 2008; Rudolph et al., 2016). This mutation occurs in the PH domain of AKT and allows constitutive membrane localization of AKT (Carpten et al., 2007;

Landgraf et al., 2008). Interestingly, the E17K mutant is insufficient for tumor transformation in the absence of other driver mutations (Lauring et al., 2010; Mancini et al., 2016), likely because the full activation of this AKT mutant still requires phosphorylation at the serine 473 and threonine

308 residues. However, tumors carrying mutant AKT have been reported to be more aggressive and have poorer prognosis (Bellacosa et al., 1995; Cheng et al., 1996; Parsons et al., 2005). In

gastric cancer, AKT1 with E17K mutation correlated with resistance to cisplatin treatment (Liu et

al., 2007) whereas a less common Q79K AKT1 mutant confers resistance to BRAF inhibitors in

melanoma patients (Shi et al., 2014).

In accordance with hyperactive AKT being a prominent oncogenic driver, many pharmacological compounds have been developed to target AKT with the aim to suppress cancer growth. There are generally two classes of AKT inhibitors – the ATP-competitive inhibitors that target the ATP-binding pocket in its kinase domain and the allosteric AKT inhibitors that target the

PH and catalytic domains of the kinase (Manning and Toker, 2017). Most of these agents are in either phase I or phase II clinical trials; however, results so far have been less than promising. Such outcomes could possibly be due to the associated toxicity of the agents or the lack of a patient stratification system as the molecular profile of the disease is likely to influence therapeutic response. Currently there are four ATP-competitive inhibitors of AKT - GSK690693, AZD5364,

GDC0068 or Ipatasertib, and GSK2110183 or Afuresertib. All of these inhibitors target all three

AKT isoforms except GSK2110183, which showed slight selective for AKT1 at subnanomolar concentrations and inhibition of AKT2 and 3 at the nanomolar range. Despite being efficacious in preclinical models, GSK690693 and GDC0068 did not fare well in phase I trials due to dose- limiting, on-target toxicity including hyperglycemia (Crouthamel et al., 2009; Rhodes et al., 2008) or lack of antitumor effects (Lin et al., 2013; Saura et al., 2017) respectively. GSK2110183 is currently being tested in phase I trials in combination with MEK inhibitors and appears to be well- tolerated (Dumble et al., 2014), while the efficacy of AZD5364 is being examined in phase II clinical trials for advanced breast cancer (Martini et al., 2014). Two allosteric inhibitors of AKT have been developed – MK2206 and ARQ092. Mechanistically, MK2206 targets the closed, inactive conformation of AKT, hindering its phosphorylation and activation (Barnett et al., 2005;

Hirai et al., 2010). As a single agent, MK2206 failed to elicit clinical efficacy (Ma et al., 2016); however, it is being tested in phase I trials in combination with trastuzumab in HER2-positive solid tumors (Hudis et al., 2013). One caveat of targeting the inactive form of AKT is the reduced sensitivity of mutant AKTs such as the E17K mutant that are constitutively membrane-associated, bypassing the function of the PH domain (Parikh et al., 2012). The structural allosteric inhibitor

ARQ092 has shown strong anti-tumor activity in multiple in vivo tumor models including ones

carrying E17K mutant AKT. Phase I trials are underway to evaluate ARQ092 in patients with

AKT-E17K and PIK3CA mutations (Lapierre et al., 2016; Yu et al., 2015).

1.5 Dissertation objectives

Since the discovery of PKCs, the field has made tremendous advances in understanding the differential function of PKC isozymes in certain tissues. Previous work from our laboratory has established PKCα as a key regulator of intestinal cell growth whose loss contributes to the development of intestinal tumors (Frey et al., 2000; Frey et al., 1997; Pysz et al., 2009; Saxon et al., 1994). This dissertation describes our effort to profile PKCα in the endometrium as well as to establish its functional contribution to endometrial carcinogenesis.

The first study, reported in chapter three, is our examination of PKCα expression and activation in the cycling uterus. Within a cycle, the rise and fall of steroid hormones regulates a program of coordinated molecular and physiological changes in the endometrium, which are necessary for the constant turnover of the tissue. Although PKCα is detected in multiple tissues, little is known of its expression, activation, or function in the endometrium. Therefore, the first objective of this dissertation was to examine PKCα in the human menstrual cycle and the mouse estrous cycle, to provide a basis for our understanding of PKCα expression and activation in the endometrium.

In the second study, presented in chapter four, our goal was (a) to survey alterations in

PKCα expression in human endometrial tumors and in lesions arising in mice harboring mutant

Pten, which are commonly used as in vivo models for oncogenic PI3K/Akt driven endometrial cancer, and (b) to identify the signaling functions of PKCα and its impact on endometrial cancer development. In light of the recent renewed understanding of PKC function in cancer and the increasing health risk posed by rising endometrial cancer incidence, a thorough examination of the role of PKCα in endometrial cancer will add new insight into PKC signaling in hyperactive

PI3K/Akt driven cancer as well its potential as a molecular biomarker for stratifying endometrial tumors.

Taken together, findings in this dissertation expand current knowledge of PKCα in the uterus. Our profiling of PKCα in the endometrium identifies the kinase as a potential regulator of endometrial renewal and function, and its ability to modulate AKT activation points to tumor suppressive properties in this tissue. Such understanding warrants further studies to examine the relationship between steroid hormones and PKCα activation in reproductive biology, and to determine whether PKCα impacts the clinical outlook of endometrial cancer such as patient prognosis and response to AKT or other small molecule inhibitors.

Chapter 2: Materials and Methods

2.1 Cell lines and reagents

Human endometrial cancer cell lines with diverse genetic mutations were obtained from the following sources:

Cell line Catalog # Source

HEC-1-A HTB-112 ATCC

HEC-1-B HTB-113 ATCC

HEC-50Co Dr. Kimberly K. Leslie (University of Iowa)

KLE CRL-1622 ATCC

AN3CA HTB-111 ATCC

Ishikawa Dr. Tim Hui-Ming (Ohio State University)

SK-UT-1B HTB-115 ATCC

RL95-2 CRL-1671 ATCC

ECC-1 CRL-2923 ATCC (discontinued)

SNG-II IFO50312 Japanese Collection of Research Bioresources (JCRB) Cell Bank

SNG-M IFO50313 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEC-6 JCRB1118 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEC-59 JCRB1120 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEC-108 JCRB1123 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEC-116 JCRB1124 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEC-151 JCRB1122 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEC-251 JCRB1141 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEC-265 JCRB1142 Japanese Collection of Research Bioresources (JCRB) Cell Bank

HEK293 CRL-1573 ATCC

All cells were maintained in culture conditions recommended by the providing source as described below:

Cell line Culture Media

HEC-1-A McCoy’s 5a + 10 % FBS

HEC-1-B, SK-UT-1B MEM + 10% FBS, 1% NEAA, 1 mM sodium pyruvate

HEC-50Co DMEM + 10% FBS, 1 mM sodium pyruvate

KLE DMEM F12 + 10% FBS

AN3CA, Ishikawa DMEM + 10% FBS, 1% NEAA, 1 mM sodium pyruvate

RL95-2 DMEM F12 + 10% FBS, 10 mM HEPES, 0.005 mg/ml insulin

ECC-1 RPMI-1640 + 5% FBS, 10 mM HEPES, 10 mM glucose, 1 mM sodium pyruvate

SNG-II, SNG-M Ham’s 12 + 10% FBS

HEC-6, 59, 116, 151, MEM + 15% FBS

251, 265

HEC-108 MEM + 10% FBS

Mechanistic studies were performed in cultured cells with the following pharmacological agents dissolved in the solvent listed:

Compound Concentration Solvent Source

PKC agonist PMA 100 nM EtOH Biomol

DiC8 20 µg/ml acetonitrile Caymen

cPKC inhibitor Gӧ6976 2.5 µM DMSO Calbiochem

PI3K inhibitor LY294002 40 µM DMSO Cell Signaling

AKT inhibitor MK-2206 100 or 250 nM DMSO EMD Millipore

Phosphatase inhibitors Calyculin A 10 nM DMSO Santa Cruz

Okadaic acid 1 µM DMSO Santa Cruz

NSC 45586 50 µM DMSO NCI

NSC 117079 50 µM DMSO NCI

Because of its rapid metabolism in cells, DiC8 was replaced every 30 minutes during the treatment

period. All phosphatase inhibitors and cPKC inhibitor were added to cells 30 minutes prior to

addition of PKC agonists, except for calyculin A, which was added 15 minutes prior to PKC

activation.

2.2 Transgenic mice

The allele-specific Pten mutant mice (PtenΔ4-5/+, PtenG129E/+, and PtenC124R/+) were generated and characterized by Dr. Gustavo Leone (Ohio State University) (Wang et al., 2010). The Pten conditional knockout animals were generated by mating Ptenflf and progesterone receptor promoter

driven- or lactoferrin promoter driven-Cre animals (PR-Cre or Ltf-Cre respectively) by Dr. Takiko

Daikoku and Dr. Sudhansu K. Dey (Cincinnati Children’s Hospital Medical Center) (Daikoku et al., 2008; Daikoku et al., 2014). Tissues from the aforementioned Pten transgenic models were provided by the corresponding laboratories for this study.

To generate Pten mutant mice lacking PKCα, SV129J/C57 BL6 Prkca-/- mice obtained from Dr. J.

Molkentin (Braz et al., 2004) were crossed with mixed strains of PtenΔ4-5/+ mice provided by Dr.

G. Leone (Wang et al., 2010). The Prkca+/-; PtenΔ4-5/+ mice were then intercrossed to generate animals with the following genotypes: Prkca+/+; Pten+/+, Prkca+/-; Pten+/+, Prkca-/-; Pten+/+,

Prkca+/+; PtenΔ4-5/+, Prkca+/-; PtenΔ4-5/+, and Prkca-/-; PtenΔ4-5/+. The “Y” shaped uterine horn was

halved at the cervix and cryopreserved in OCT or fixed in formalin for paraffin embedding. Uterine

tissue was collected from female mice from all six genotypes at 3, 5, 7, and 9 months of age;

however, 1- and 2-month uterine tissue was collected from Prkca+/+; PtenΔ4-5/+ and Prkca-/-; PtenΔ4-

5/+ only. The functional effects of PKCα were only examined in Prkca+/+; PtenΔ4-5/+ and Prkca-/-;

PtenΔ4-5/+ as we did not observe any uterine phenotypes in Prkca knockout only animals. The

genotypes were confirmed by PCR using the following primers and conditions:

Gene 5’-3’

Prkca

Primer 1 CCAAGTGTGAAGTGTGTGAG

Primer 2 AGCTAGGTCCTGTTGGTAAC

Primer 3 GCGCATCGCCTTCTTCGC

Cycling conditions 95°C 3 min

95°C 30 sec 35 cycles

57°C 30 sec

72°C 1 min

72°C 3 min

4°C Hold

Pten

CoA GAATGCCATTACCTAGTAAAGCAAGG

CoB GGGTTACACTAACTAAACGAGTCC

CoC GAATGATAATAGTACCTACTTCAG

Cycling conditions 95°C 3 min

95°C 30 sec 35 cycles

58°C 30 sec

72°C 1 min

72°C 3 min

4°C Hold

2.3 Determination of estrous phases

The phases of the estrous cycle of the mouse can be determined from visual observation or cytology of vaginal discharge as described in (Byers et al., 2012). In this study, estrous cycle phases (diestrus, proestrus, estrus, metestrus) were determined based mainly on the cellular makeup of the vaginal smear, using visual observation as a supporting tool. To ensure accuracy of phase determination, vaginal smears were obtained from 5-6 week-old virgin, female mice (when they reach sexual maturity) twice a day (once in the light cycle, once in the dark cycle) for a week to track the progression of estrous cycling. The vaginal discharge was collected using Pasteur pipets whose tips had been rounded by flaming and cooled prior to use. The vaginal opening of the mice was rinsed with 20-50 µl of 1X PBS and the vaginal discharge was collected in an Eppendorf tube, stained with 100-200 µl of 10% toluidine blue, transferred onto microscope slides and overlaid

with a coverslip. The phase of the estrous cycle was then determined based on the presence or absence of leukocytes, nucleated or cornified epithelial cells.

2.4 Human tissue microarray

Human endometrial cancer TMAs were generated by Dr. Carl Morrison at Ohio State University

(OSU) and Roswell Park Cancer Institute (RPCI). Two TMAs, displaying a total of 384 de- identified endometrial cancer tumors (304 endometrioid and 80 non-endometrioid), were generated at OSU with approval from OSU Institutional Review Board. All patients in the TMA were diagnosed with uterine malignancy between January 1, 1980 and July 31, 2003 at the Arthur James

Cancer Hospital of the Ohio State University Medical School. Specimens for controls within the

TMA consisted of 37 secretory endometrium, 30 proliferative endometrium, as well as multiple cores of normal tissue from 10 different organs including heart, colon, kidney, adrenal, ovary, myometrium, brain, thyroid, lung, and prostate. A third endometrial cancer TMA, consisting of 52 de-identified cases (26 endometrioid and 26 non-endometrioid) was generated at RPCI

(RPCI_GYNCa09) with RPCI Institutional Review Board approval. Cases were diagnosed at

RPCI between 1992 and 2011. For any case with variation in grade or cytological atypia, TMA cores of the donor block were always taken from the areas of the highest-grade tumor. Matching frozen tissue was also available for all of the RPCI cases and was used for determination of PKCα mRNA levels. Scoring of IHC staining was performed by three independent examiners who were blind to sample information. Patient survival analysis in correlation with PKCα expression was performed using the MDACC-endometrial-L3-S40 data set, consisting of 244 endometrial tumors of which 80% were of the endometrioid subtype or using TCGA data set. To perform parallel analysis of PKCα mRNA and protein levels in endometrial tumors, total RNA was isolated from frozen samples of 50 of the tumors included in the TMA. The correlation between PKCα mRNA and protein levels was determined by Spearman Rank Correlation analysis.

2.5 Immunohistochemistry

Murine uterine tissue was formalin-fixed, paraffin-embedded, and cross-sectioned or longitudinally-sectioned at 4 µm. Human and murine tissue sections were deparaffinized in xylene and rehydrated by incubation in serially diluted alcohol. Antigen retrieval used DAKO Targeting

Retrieval Solution (DAKO S1699) for 30 minutes in a steamer. Sections were blocked with 0.03% casein for 30 minutes at room temperature before incubation with primary antibody conditions listed below:

Antibodies Concentration Conditions Source Catalog #

PKCα 1:3000 - 6000 4°C overnight Abcam 32376

PTEN 1:100 4°C overnight Cell signaling 9559

pAKT Ser473 1:300 - 600 Room temperature 1 hour Cell signaling 4060

Id-1 1:1200 4°C overnight Biocheck BCH-1/195-14

Ki67 Stained by the Tissue Science Facility at UNMC

Following washes, incubation with secondary antibody (Vector BA-1000) was performed at room

temperature for 30 minutes, followed by addition of Vectastain Elite ABC reagent (Vector

Laboratories) and DAB chromogen solution (DAB Quanto; Thermo Scientific TA-060-QHDX).

Sections were counterstained with hematoxylin. Specificity of these antibodies was validated in previous studies; PKCα and Id-1 were detected as described in (Hao et al., 2011), and pAKT Ser473 and PTEN IHC followed procedures detailed in (Wang et al., 2010). PKCα immunostaining

specificity was further confirmed using PKCα knockout uterine tissue.

2.6 Immunofluorescence

Uterine tissue in various phases of the estrous cycle, as determined by vaginal smears, was collected and frozen in OCT. Frozen sections (4-6 µm) were fixed in 2% freshly depolymerized paraformaldehyde/PBS for 15 minutes at room temperature and 100% methanol for 10 minutes at

-20°C, prior to blocking and antibody incubation. Antibodies were as follows: anti-PKCα (1:250;

Abcam 32376), or Ki67 (1:1000; eBiosciences 11-5678) for 1 hour at room temperature, and anti-

rabbit Alexa 594 (1:300; Invitrogen A-21207) for 30 minutes at room temperature. All images

were obtained at the same exposure. Nuclei were detected by Hoechst dye.

A guide slide consisted of 2 serial sections (10 µm) was prepared and stained with

immunofluorescence to use for identification normal vs. tumor regions for LCM. The tissue was stained with primary antibodies: anti-PKCα (1:250; Abcam 32376) or anti-pAKT (1:200; Cell

Signaling 4060) for 1 hour and anti-rabbit Alexa 488 (1:300; Invitrogen A-32723) for 30 minutes at room temperature and stained with Hoechst dye for 5 minutes.

2.7 Western blot analysis

To prepare lysate for immunoblot analysis, cells were rinsed twice with cold 1X PBS before addition of pre-warmed lysis buffer (1% SDS, 10 mM Tris-HCl, pH 7.4). Cell lysates were collected, passed through a 27 g needle 5-10 times to shear DNA, and centrifuged at 16,000 x g for

20 minutes. Protein concentration was determined using the BCA Assay Kit (Pierce). Equal amounts of protein were subjected to SDS-PAGE and transferred to nitrocellulose membrane.

Membranes were blocked with 5% milk and probed with primary antibodies at 4°C overnight as listed below. Secondary antibodies were horseradish peroxidase-conjugated anti-rabbit or anti-

mouse IgG (1:2000) and detection used SuperSignal West (Pierce). Signal intensity was quantified using ImageJ.

Antibodies Concentration Source Catalog #

PKCα 1:10,000 Abcam 32376

PKCαa 1:20,000 - 30,000 Santa Cruz SC-208

PKCβI 1:2000 Abcam 195039

PKCβII 1:1000 Santa Cruz SC-210

PKCγ 1:1000 Santa Cruz SC-211

PKCδ 1:1000 Santa Cruz SC-213

PKCε 1:1000 Santa Cruz Sc-214

PKCθ 1:1000 BD Transduction Laboratories 610084

PKCη 1:500 Santa Cruz SC-215

PKCζ 1:8000 Santa Cruz SC-216

PKCι 1:1000 BD trans 610175

PTEN 1:1000 Cell signaling 9559

pAKT Ser473 1:8000 - 10,000 Cell signaling 4060

pAKT Thr308 1:8000 - 20,000 Cell signaling 2965

AKT 1:1000 Cell signaling 9272

pPRAS40 1:15,000 – 20,000 Cell Signaling 13175

PRAS40 1:10,000 Cell Signaling 2691

pFOXO1 1:1000 Cell Signaling 9401

FOXO1 1:1000 Cell Signaling 2880

pGSK3β 1:500 Cell Signaling 9323

GSK3β 1:10,000 BD Transduction Laboratories 610202

PP2AC 1:3000 Upstate 05-421

Id-1 1:1500-3000 Biocheck BCH-1/195-14

β-actin 1:15000 Sigma A2066

a To assess PKCα levels in adenoviral vector mediated PKCα overexpression experiments.

2.8 Laser-capture microdissection (LCM)

Murine uterine tissue was placed in OCT and frozen immediately after dissection. For LCM, the tissue was longitudinally sectioned at 10 µm and mounted on PET-membrane FrameSlides (Leica).

Slides were air-dried briefly, fixed in 70% EtOH for 30 seconds, washed in dH2O to remove excess

OCT, stained with hematoxylin until tissue was visible, and washed in dH2O to remove excess dye, followed by incubation in 95% and 100% EtOH for 30 seconds. Stained PET-slides were air-dried and placed in a cold dessicator until LCM. Additional serial sections were prepared on charged glass slides for IF analysis for PKCα and pAKT to serve as a guide to distinguish normal vs. tumor regions. Laser capture was performed at optimal laser strength that ensures precise dissection while minimizing tissue damage. Dissected tissue was directly collected into 30 µl lysis buffer place on

the cap of the collection tube. RNA was isolated from dissected sections using the RNAqueous-

MicroKit (Ambion) as directed by manufacturer’s instructions. All reagents used were RNAse free.

Slides and working surface were cleaned with RNaseZap.

2.9 Quantitative RT-PCR analysis

RNA was isolated using the RNAspin mini RNA isolation kit (GE Health) or Trizol reagent

(Thermo Fisher) and 10 ng total cellular RNA was analyzed using Brilliant II SYBR® Green

QRT-PCR 1-Step Master Mix (Agilent). In the nascent RNA capture experiments, cDNA was generated using the iScript™ cDNA Synthesis Kit and quantified using iTaq Universal SYBR

Green Supermix (BioRad). The primers used for each target are listed below:

Target 5’-3’

hPKCα

F’ GGAAGGGGACGAGGAAGGA

R’ TGATGACTTTGTTGCCAGCAG

hPHLPP1

F’ TGATGACTTTGTTGCCAGCAG

R’ AGTTCATTAAGCCCCCTGGC

hPHLPP2

F’ TGTACGCAAGGGAAAGACCC

R’ AGCAAGGGAGTATTGCCGTC

18s rRNA

F’ CATTGGAGGGCAAGTCTGGTG

R’ CTCCCAAGCTCCAACTACGAG

GAPDH

F’ TGAAGGTCGGAGTCAACGGA

R’ CCATTGATGACAAGCTTCCCG

2.10 RNA interference

For siRNA-mediated knockdown of PKCα or PP2AC, cells were transfected with 10 nM siRNA using RNAiMAX transfection reagent (Invitrogen) as per manufacturer’s protocol. Cell lysates were collected for Western blot analysis 72 hours after transfection. Knockdown of the catalytic subunits of PP2A was achieved by pooling PPP2CA and PPP2CB siRNAs. Information on the siRNAs used is provided below: siRNAs Part/ Catalog number Source siPKCα #1 4390827 Ambion Life Technologies

sense - GGAUUGUUCUUUCUUCAUATT

antisense - UAUGAAGAAAGAACAAUCCGA siPKCα #2 4390771 Ambion Life Technologies

sense - GAAGGGUUCUCGUAUGUCATT

antisense - UGACAUACGAGAACCCUUCAA siPP2CA ON-TARGET plus Human siRNA SMARTpool L- GE Dharmacon

003598-01 siPP2CB ON-TARGET plus Human siRNA SMARTpool L- GE Dharmacon

003599-00

Non-targeting D-001810-01-05 GE Dharmacon

Lentiviral vectors expressing PTEN targeting shRNAs were used to generate stable knockdown endometrial cancer cell lines. Cells were transduced with either or both clones of shPTEN vectors or a non-targeting vector at 20 moi (at 1×106 CFU/ml) in serum-free medium containing 8 µg/ml of polybrene and incubated at 37°C for 8 hours before adding 1 ml of culture medium. Cells were

passaged 4 days after transduction, selected with puromycin (10 µg/ml) on day 6, and cultured for at least 10 passages before collecting cell lysate for analysis or subsequent experiments.

Information on shRNAs used are listed as below:

shRNA Clone/ Catalog number Source

shPTEN #1 V2LHS_92317 Open Biosystems

shPTEN #2 V2LHS_231477 Open Biosystems

Non-targeting shRNA RHS 4348 GE Dharmacon

2.11 Nascent RNA capture

Labeling and capture of nascent RNA was performed using the Click-iT Nascent RNA Capture Kit

(Life Technologies) according to the manufacturer’s protocol. In brief, EU was added to cells for incorporation into nascent RNA to allow biotinylation of the synthesized RNA. The biotinylated

RNA was then captured using Streptavidin magnetic beads and cDNA was generated using the iScript cDNA Synthesis Kit (Bio-Rad). cDNA for PKCα and GAPDH was quantified by qPCR using iTaq Universal SYBR Green Supermix.

2.12 Subcellular fractionation

To analyze PKCα subcellular partitioning after PMA-induced activation, cytosolic and particulate fractions were prepared from endometrial cancer cells after 6 hours of treatment with 100 nM phorbol 12-myristate 13-acetate (PMA; Biomol) dissolved in ethanol as previously described

(Saxon et al., 1994). Cells were rinsed twice with PBS, lysed with digitonin buffer (0.5 mg/ml digitonin, 20 mM Tris-HCL pH7.4, 2 mM EGTA, 2 mM EDTA, 10 mM NaF, and 1:100 protease inhibitor cocktail) and centrifuge at 100,000 x g at 4°C for 30 minutes to separate the cytosolic (in the supernatant) and the membrane/cytoskeleton (in the pellet) fractions. Cytosolic proteins were

precipitated by adding 100% trichloroacetic acid (TCA) to a final concentration of 10% TCA and

incubate briefly on ice. The pellet was washed with acetone four times and dissolved by incubation

with 100 mM NaOH at 4°C overnight. The cytosolic fraction was then neutralized with an equal

volume of 100 mM HCl. The pellet containing the particulate fraction was solubilized by addition

of 200-300 µl of 1% SDS-digitonin buffer with 1:100 protease inhibitor cocktail. The pellet was

fragmented by syringing and incubated on ice for 30 minutes at 4°C to further solubilize the

particulate fraction. An equal ratio of cytosolic and particulate fractions (relative to total protein)

was analyzed for the presence of PKCα.

2.13 PP2A activity assay

PP2A phosphatase activity was measured using a PP2A immunoprecipitation phosphatase assay

kit (Millipore) and as previously described in (Guan et al., 2007). Specifically, cells were transfected with 30 pmole of non-targeting or a combination of two siRNAs targeting PKCα for

three consecutive days to ensure sufficient knockdown. 24 hours after the last transfection, cells

were treated with PMA or vehicle control for 10 minutes, washed twice in cold TBS, lysed in lysis

buffer (20 mM imidazole-HCl, 2 mM EGTA, 2 mM EGTA, pH 7.0 with 1:100 protease inhibitor

cocktail) and subjected to immunoprecipitation and measurement of free phosphate with malachite

green following the manufacturer’s protocol.

2.14 Establishing EC cell lines stably expressing mutant AKT

To determine if the tumor suppressive effects of PKCα in the endometrium involves inhibition of

AKT hyperactivation, EC cell lines stably expressing constitutively active mutants, either the AKT

E17K mutant found in breast and endometrial cancers (Mancini et al., 2016; Rudolph et al., 2016;

Shoji et al., 2009) or the phosphomimetic AKT DD (T308D, S473D) mutant, were generated via

retroviral vector-mediated integration. Retroviral vector pBabe-puro-AKT1 was a gift from

Xinjiang Wang (RPCI, NY) and the E17K mutation was generated through site-directed mutagenesis. The plasmid encoding pEGFP-AKT-DD was a gift from Julian Downward (Addgene plasmid # 39536) (Watton and Downward, 1999) and was subcloned into retroviral vector pBMN-

I-GFP, which was gifted by Garry Nolan (Addgene plasmid # 1736). First, pEGFP-AKT-DD was digested with BamH1 and pBMN-I-GFP was digested with NotI. The ends of the digested fragments were filled in by Klenow-mediated DNA polymerization. The cloning vector and insert were digested by XhoI and SalI, respectively, and ligated to generate pBMN-AKT-DD for subsequent viral vector production. Retroviral vectors expressing AKT mutants were generated by transfecting the packaging cell line 293GP (stably expressing gag and pol viral components) with pVSVG and either control vector (pBabe-puro or pBMN-I-GFP) or the mutant AKT encoding plasmids (pBabe-AKT-E17K or pBMN- AKT-DD) in DMEM medium containing 10% heat- inactivated serum. The viral supernatant was collected 48 hours after transfection and centrifuged to pellet cell components. The target EC cells were incubated with the cleared viral supernatant for 6-8 hours before changing to culture media. The infection was repeated 3 times to achieve sufficient viral transduction and robust mutant AKT expression (approximately 4-fold increase of endogenous level). EC cells transduced with pBabe-puro or pBabe-AKT-E17K were subjected to puromycin selection (10 µg/ml) and cells expressing pBMN-I-GFP or pBMN-AKT-DD were selected with 600 µg/ml of G418 for at least four passage before subsequent analysis by soft agarose colony formation assays.

2.15 Adenoviral-mediated overexpression and soft agarose colony formation assay

Adenoviral vectors expressing either LacZ, PKCα, or kinase-dead PKCα were amplified in

HEK293 packaging cells and the titer of viral supernatants was determined using Adeno-X Rapid

Titer Kit (Clonetech). Adenovirus expressing myr-AKT1 was purchased from Vector Biolabs.

Cells (5×105) were transduced with adenovirus at indicated moi in reduced (2.5%) serum medium for 24 hours and then incubated in complete culture medium. 48 hours post transduction, cells (1-

5×103 cells per well) were plated in 0.6% low-melting agarose as described in (Pysz et al., 2009) in duplicate, and colonies were allowed to grow for 1-4 weeks prior to staining with crystal violet.

Whole plates were then scanned and visible colonies were counted.

For AKT mutant-expressing cells lines, Cells (3×105) were transduced with the lowest moi (at

which Ad-PKCα inhibits colony formation) in 2.5% serum medium for 24 hours. 2 days after

infection, 2×104 cells were plated in 0.6% low-melting agarose as previously described, and allow to grow for 7 days before analysis. For analysis of the effect of co-expression of AKT mutants and

PKCα, images from 4 independent fields per well were taken at 5X magnification at 4 different focal planes. Images of colonies in different focal planes were superimposed in Adobe Photoshop

CC software and the number of colonies per field was determined using the particle analysis tool in ImageJ software.

2.16 Correlation between PKCα mRNA and protein levels

To perform parallel analysis of PKCα mRNA and protein levels in endometrial tumors, total RNA was isolated from frozen samples of 50 of the tumors included in the TMA. PKCα mRNA levels were measured by qRT-PCR. A score was given to the tumors based on PKCα signal intensity from the IHC analysis. Results from the mRNA analysis were paired with the corresponding IHC score for the 50 tumors. The correlation between PKCα mRNA and protein levels was determined by

Spearman Rank Correlation analysis.

2.17 Quantification of tumor burden and pAKT intensity relative to PKCα levels

For quantification of uterine tumor burden in mice, two to four longitudinal sections per animal were stained for pAKT Ser473 to identify endometrial lesions (Wang et al., 2002) and positive signal was quantified using DEFINIENS software and expressed relative to total endometrial epithelium area (as %). For determination of pAKT intensity relative to PKCα expression, staining corresponding to negative-low, medium and high pAKT staining was determined for each slide without knowledge of corresponding PKCα staining. Analysis was performed using Adobe

Photoshop CC. pAKT staining was abstracted from the image using the Color Range tool and copied to a separate layer. Thresholds corresponding to the boundaries between staining levels were set and used to trace areas corresponding to each level. Areas of positivity on slides probed for PKCα were similarly identified and abstracted to a separate layer without reference to pAKT staining. Staining layers from consecutive slides stained for PKCα and pAKT were overlaid and the area (in pixels) corresponding to each level of pAKT staining in PKCα positive and PKCα negative areas was determined.

2.18 Statistical analysis

Data are presented as the mean ± SEM of three or more independent experiments unless otherwise stated. Statistical analysis was performed using Microsoft Excel, GraphPad Prism or SAS software version 9.3 (SAS Institute Inc., Cary, NC). p-values of less than 0.05 were considered statistically significant. Significance was assessed using Student's t-test for comparison of means, Chi-square or Fisher’s exact test for categorical analysis of two groups, Wilcoxon rank-sum test for nonparametic comparison of two samples, or two-way ANOVA test for multiple comparisons.

Pairwise comparisons were adjusted for multiple comparisons with Tukey’s method. Model assumptions were examined with residual plots.

Chapter 3: The Role of PKCα in the Endometrium

Portions of the content covered in this chapter are the subject of an article under

consideration at Cell Reports by Hsu A.H. et al.

3.1 Introduction

3.1.1 Biology of the endometrium

The main function of the uterus, which develops from the Müllerian tubes (also referred to as

the female ducts or paramesonephric ducts), is to house the developing embryo during its early life.

The uterus consists of two compartments - the myometrium, the smooth muscle that controls uterine

contraction, and the endometrium that lines the inner cavity of the uterus. Functionally, the

endometrium facilitates the transport of the sperm and provides the nurturing environment for

implantation and embryo development. The endometrium is one of the most dynamic tissues in the body, undergoing periodic remodeling throughout the reproductive period. In each cycle, both the stroma and epithelium of the endometrium are restructured in response to the rise and fall of luteinizing hormone (LH) and follicle-stimulating hormone (FSH) produced from the pituitary gland, and estrogen (E2) and progesterone (P4) secreted by the ovaries. This intricate hormonal

balance governs the proliferation, differentiation, and regression of the endometrium. Disruption

of this tightly regulated process triggers the onset of menopause or, in the event of molecular dysregulation, results in uncontrolled, hyperplastic growth of the endometrium and the eventual development of endometrial carcinoma. This next section describes the regeneration of the endometrium during the menstrual cycle (human) and estrous cycle (mouse), and actions of E2 and

P4 during these events.

3.1.2 Human estrous cycle

In humans, the endometrium regenerates every 28-30 days through a process that is defined

as the menstrual cycle. This breakdown and rebuilding of the endometrium was first observed in

other primates (Bartelmez, 1951; Bartelmez et al., 1946). Subsequent studies identified the basalis

layer of the endometrium situated immediately atop the myometrium to be quiescent at the

beginning of the cycle (the first day of spotting). The progenitor cells of the endometrium were

thought to reside in this layer as depletion of the basalis abolished the regenerative capability of the endometrium (Wyss et al., 1996). With the increase in E2 levels in the follicular phase, a follicle starts to develop and mature in the ovary. This process generally spans the first 14 days of a typical

28-day cycle. During this time, the stromal and epithelial cells, as well as endothelial vasculature in the basalis of the endometrium, proliferate in response to estrogenic stimulation, giving rise to the functionalis layer of the thickening endometrium. This phase is referred to as the “proliferative” phase of the endometrium. At the end of this phase, E2, LH, and FSH reach peak levels and rapidly decline following ovulation or the release of the oocyte. This marks the entry to the ovarian luteal phase (or the secretory phase of the endometrium). At this time, although there is moderate amount of E2, P4 is the predominant steroid, stimulating cell differentiation and secretion of growth factors in the endometrium and leading to decidualization. In the case of successful fertilization, P4 levels are maintained throughout pregnancy. Otherwise, the fall of P4 levels leads to shedding of the

functionalis, leaving only the basalis to regenerate the endometrium for the next cycle. Because of

advances in cell-based therapy, there has been effort to identify and isolate progenitor cells from

the endometrium as a source of stem/ progenitor cells (Emmerson and Gargett, 2016).

3.1.3 Rodent estrous cycle

The rodent uterus is a widely used model for reproductive studies because of the ease with

which it can be physically and genetically manipulated. While the rodent endometrium shares many

similarities with its primate counterpart, there are distinct features that maximize rodent fertility.

The rodent uterus is a ‘Y” shaped organ with horns that are joined at the cervix. This unique shape

allows the implantation and development of multiple embryos in a single pregnancy. Unlike the

human menstrual cycle, the rodent estrous cycle typically lasts 4-6 days and has four phases:

proestrus, estrus, metestrus, and diestrus. Each phase can be differentiated by the physical

appearance of the genital area and cellular makeup of vaginal smears (Byers et al., 2012). At the

beginning of proestrus, a new batch of eggs enters the maturation process. The accompanying surge of E2 at proestrus induces cell proliferation in the endometrium leading to the thickening and enlargement of the mouse uterus (Wood et al., 2007). In rodents, E2 peaks prior to ovulation and the surge of P4, LH, and FSH soon follows (Yoshinaga, 1973). Levels of these hormones sharply

fall after ovulation. The rise of P4 converts the endometrium from a growth to a receptive state.

Without implantation, the cycle proceeds to metestrus and diestrus at which time the endometrium

is resorbed and primed to start the next cycle. These last two phases are equivalent to the secretory phase of the human menstrual cycle. Many studies have profiled the proliferation activities of the endometrium during the estrous cycle. Maximal proliferation is observed in proestrus followed by metestrus and diestrus, while estrus is repeatedly associated with minimal proliferation (Marusak et al., 2007; Mendoza-Rodriguez et al., 2003; Wood et al., 2007). In addition, increased apoptosis

is also reported during estrus (Dery et al., 2003; Marusak et al., 2007). As part of our study to

understand the role of PKCα in endometrial cancer, we examined the expression and activation of

PKCα in the normal mouse endometrium and its correlation with proliferation as indicated by Ki67

immunohistochemistry.

3.1.4 Effects of steroid hormone signaling: estrogenic response

The cyclic endometrium remodeling is orchestrated by the fluctuation of follicular E2 and P4

and the activation of their cognate nuclear receptors. Rising E2, observed in proestrus of the estrous cycle or the proliferative phase of the menstrual cycle, initiates early events associated with the induction of uterine growth. Sustained estrogen receptor (ER) activation and E2 bioavailability

then leads to late E2 responses that result in uterine epithelial proliferation. Thus, E2 functions as a

pro-proliferative signal in the uterus. Increased P4 levels, following E2 induction, opposes ER

activation to mediate cell differentiation, switching the uterus from a proliferative to a receptive

state for possible implantation.

The surging E2 stimulates both physiological and molecular changes in the uterus (Barton et al., 1998). The early events, which occur between one to six hours after E2 induction, include

increases in RNA and protein synthesis and nuclear ER occupancy (Hewitt et al., 2016). These

changes prime the endometrial epithelial cells for subsequent proliferation. Physiologically, there

is also an increase in water intake of the uterus (or water imbibition) and influx of blood and immune infiltration. Sustained E2 elicits late uterine responses that peak 24 to 72 hours after the initial E2 surge. At this time, there is continuing increase in DNA synthesis and significant mitosis

in the uterus, which leads to an increase in uterine weight. There is also further accumulation of

immune cells. Together, these events ensure that the physical fitness of the uterus is permissive

for a subsequent P4-mediated switch from proliferating to differentiated endometrium.

Studies using transgenic models have provided strong evidence that these uterine changes

in response to E2 are mainly mediated by ERα. There are two isoforms of ER expressed in the uterus, - ERα and ERβ - encoded by two distinct genes. Although the ligand-binding domain is conserved and E2 indiscriminately binds to both receptor subtypes, ERα is the predominant isoform

expressed in the endometrium, while ERβ is expressed at a lower level (Lecce et al., 2001).

Furthermore, the uteri of ERα knockout mice (ERαKO) were shown to be hypoplastic and

unresponsive to E2 stimulation. In contrast, the uteri of ERβ knockout (ERβKO) mice were not different from those of wild type (WT) mice and were able to undergo cyclic remodeling when stimulated with E2 (Hewitt et al., 2016; Krege et al., 1998; Lubahn et al., 1993). Gene expression analysis has also shown that ERα is responsible for the E2-induced genomic responses. The ERβKO and WT mice had indistinguishable expression profiles in response to E2 stimulation, whereas the

ERαKO uterus showed no changes in gene expression. The genes induced by E2 also clustered into two profiles, mirroring the previously described biphasic physiological response to E2 stimulation

in the uterus (Hewitt et al., 2003). These observations indicate that E2 induces its major effects in

the endometrium through ERα activation and that ERα but not ERβ is essential for normal

endometrial development.

Ovariectomized mouse models are commonly used in studies of hormonal response in vivo, taking advantage of the ease of manipulation and the similarities between rodent and human reproductive biology. Studies have shown that E2 stimulates an ERα-dependent biphasic response

in the uterus (Hewitt et al., 2003). In early response to E2 stimulation, events such as ER nuclear

occupancy, transcription of early response genes (including c-fos and c-jun), increased protein and

RNA synthesis, and water imbibition occur within hours. Nuclear ER occupancy peaks again

approximately at 8 hours after E2 stimulation, followed by late responses such as transcription of late phase genes (e.g. lactoferrin), DNA synthesis, mitosis, accumulation of immune cells, and increase in uterine weight (Groothuis et al., 2007; Hewitt et al., 2003).

The binding of E2 triggers a conformational change in ER through which ER assumes its

transcriptionally active state capable of promoting gene expression via interaction with estrogen

responsive elements (ERE) and other transcriptional mediators (Hewitt et al., 2016). The activated

ER leads to the expression of genes involved in crucial biological processes such as cell cycle

progression, growth factor-mediated signaling, and apoptosis (Basu and Rowan, 2005; Murphy,

1991). In the case of ERα, one of the genes upregulated through direct interaction with ERE is insulin-like growth factor 1 (Igf1), an important signaling molecule in stimulating uterine proliferation (Adesanya et al., 1999; Hewitt et al., 2009; Zhu and Pollard, 2007). Although

elevation of IGF-1 production occurs in all compartments of the endometrium, it was most apparent

in the uterine stroma (Cunha et al., 2004; Zhu and Pollard, 2007). IGF-1 is essential for E2-

stimulated uterine proliferation, as administration of E2 fails to induce a proliferative response in mice lacking Igf1 genes (Adesanya et al., 1999). IGF-1 binds to IGF-1 receptor (IGF-1R) to activate downstream signaling mediators and inhibit negative regulators including IRS1 (Richards et al., 1996), AKT, and GSK3β (Zhu and Pollard, 2007), leading to growth promoting effects such as nuclear accumulation of cyclin D1 (Tong and Pollard, 1999).

In addition to the IGF-1-mediated activation of PI3K/AKT signaling, E2 also exerts mitogenic effects through another arm of signal transduction that involves the MAPK/ERK

pathway. In vitro studies have shown that estradiol activates MAPK in breast, bone, and neuronal cells (Endoh et al., 1997; Migliaccio et al., 1996; Watters et al., 1997). In the endometrium, administration of E2 in ovariectomized mice leads to phosphorylation of ERK in uterine epithelium

(Wang et al., 2015). The activation of ERK prominently upregulates the activity of AP-1 transcription factors such as c-Fos and c-Jun in response to E2 or tamoxifen stimulation (Babu et al., 2013; Karin et al., 1997; Treisman, 1996; Whitmarsh and Davis, 1996). AP-1 proteins then form active dimers (homodimers or heterodimers) to regulate genes that play important roles in cell proliferation and survival – for example, upregulation of cyclin D1 and cyclin D3 to promote growth or repression of growth inhibitory effectors such as p21, p27, and p53 (Shaulian, 2010;

Shaulian and Karin, 2001; Zhuang et al., 2001).

Continuing research of E2 actions in the endometrium has revealed other mechanisms by

which E2 promotes the proliferation of endometrial cells (e.g., induction of non-genomic effects

through the G-protein coupled receptor 30 (GPR30)) (Bamberger et al., 2007; Gielen et al., 2007).

Elevated exposure to E2 or tamoxifen is one of the main risk factors for endometrial cancer, increasing the risk of developing this malignancy (Morice et al., 2016) by supporting the pro- proliferative and -survival properties of E2 and ER signaling in the endometrium.

3.1.5 Effects of steroid hormone signaling: progesterone

The main role of progesterone (P4) in the uterus is to oppose the actions of E2 to maintain

uterine homeostasis by inhibiting cell growth and inducing differentiation, through binding to the

progesterone receptor (PR). Like ER, there are two isoforms of PR – PR-A and PR-B. Rather

than being encoded by two separate genes (like ERα and ERβ), the two PRs share a single PR gene

located on chromosome 11, regulated by different transcriptional start sites, thus generating two

mRNA transcripts encoding either PR-A or PR-B (Conneely et al., 1989). While both PR-A and

PR-B are detectable in endometrium, IHC analysis has found that PR-A is mainly in the stroma

and PR-B is the predominant PR in the epithelial compartment (Mote et al., 2006; Mote et al.,

1999). In the uterus, E2 stimulation decreases PR levels in the epithelium while stromal PR is elevated in response to E2 (Patel et al., 2015; Tibbetts et al., 1998). Yet, both epithelial and stromal

PR act to antagonize E2-responsive proliferation, while epithelial PR mediates expression of P4- targeted genes and inhibition of proliferation (Franco et al., 2012; Liu et al., 2009b; Smid-

Koopman et al., 2003; Zhang et al., 2013). With the understanding of differential PR isoform levels in the uterine compartments, this suggests that PR-A and PR-B have distinct functions in regulating uterine growth and homeostasis.

The predominance of each PR isoform in different endometrial compartments suggests there is differential response to estrogenic stimulation mediated by these molecules. Interestingly, elevated PR levels and activation, in turn, suppress ERα expression (Haluska et al., 1990; Mote et al., 2006), generating a feedback loop to maintain the intricate balance of ER and PR signaling and uterine homeostasis. The essential role of PR in the reproductive tract was further demonstrated by transgenic models. When lacking both PRs, the mice were infertile due to defects in ovulation and implantation. The animals also showed evidence of uterine hypertrophy and inflammation of the glandular epithelium of the endometrium (Mulac-Jericevic et al., 2000). Using isoform specific knockout models (PRAKO for PR-A knockout and PRBKO for PR-B knockout), it was shown that functional PR-A alone is sufficient and necessary for normal ovarian and uterine functions as

PRAKO mice were infertile (similar to PR-null animals), while normal implantation and decidualization were observed in PRBKO mice (Mulac-Jericevic et al., 2003; Mulac-Jericevic et al., 2000). PRAKO mice also showed increase endometrial hyperplasia and inflammation (Mulac-

Jericevic et al., 2000). Unexpectedly, P4 stimulated abnormal epithelial proliferation in the uterus in PRAKO mice that was not observed in PR null mice, suggesting P4 promotes uterine epithelial proliferation through PR-B activation, and that PR-A acts to limit PR-B-mediated proliferation

(Mulac-Jericevic et al., 2003; Mulac-Jericevic et al., 2000). Together these observations

demonstrated 1) PR-A is required in the anti-proliferative response of PR signaling and; 2) the P4- mediated uterine response is essential for maintenance of uterine homeostasis, blastocyst implantation and pregnancy.

P4 can exert its functions in the uterus through a genomic pathway by binding and activating the transcriptional activity of PR or by non-genomic activation of signal transduction. The ligand bound PR (both PR-A and PR-B) can form either homodimers or heterodimers, leading to receptor phosphorylation, and subsequent interaction with hormone responsive promoters to regulate targeted gene expression (Conneely et al., 2003). In addition, P4 also induces non-genomic responses through PR-dependent activation of Src/RAS/MAPK signal transduction and PR- independent activation of membrane receptors, whose physiological contribution is less clear

(Ballaré et al., 2006; Mulac-Jericevic and Conneely, 2004). In general, P4 antagonizes the action of E2 by negatively impacting the regulation of genes activated by ER signaling (Yang et al., 2011).

In contrast to the induction of AP-1 expression by E2, P4 downregulates growth promoting proteins such as the AP-1 family of transcription factors, particularly c-Jun, cyclin D1, and minichromosome maintenance proteins (MCMs) (Chen et al., 2005; Dai et al., 2003; Pan et al.,

2017; Yang et al., 2011). P4 also exerts its growth inhibitory effect through reprogramming of the

AP-1 mediated transcriptional network, leading to opposing effects on many of the E2-reponsive genes. In addition to shunting AP-1 from binding to growth promoting effectors such as cyclin D1,

P4 also upregulates the expression of p53 and CDKIs such as p21, p27 by enhancing the binding of AP-1 to their respective promoters (Dai et al., 2003; Hagan et al., 2011; Yang et al., 2011).

Other PR-responsive genes, such as FOXO-1 that mediates cell senescence, have been shown to be anti-proliferative (Kyo et al., 2011; Ward et al., 2008). Although P4 binds to both PR-A and PR-

B, the receptors have distinct, isoform-specific genomic functions, providing a likely explanation for the aforementioned differential physiological roles of PR isoforms. Studies have found that activated PR-A mainly acts as a dominant negative inhibitor of both ER and PR-B (Kraus et al.,

1997; McDonnell et al., 1994; Vegeto et al., 1993), while PR-B exhibits stronger transcriptional

activating capability than PR-A (Jacobsen et al., 2002; Kumar et al., 1998; Leslie et al., 1997;

Vegeto et al., 1993). However, only a subset of PR-responsive genes are regulated by PR-B.

Among the genes related to implantation (e.g., calcitonin, histidine decarboxylase, lactoferrin),

RNA analysis showed that PR-B alone was sufficient for P4-mediated transcriptional changes in

histidine decarboxylase and lactoferrin but not calcitonin (Mulac-Jericevic et al., 2000).

Interestingly, despite the hyper-proliferative effect of P4 in PRAKO mice, PR-B activation

upregulated IGFBP-1, which exerts P4-induced anti-proliferative effects in cooperation with

FOXO-1(Nakamura et al., 2013).

There is mounting evidence that steroid hormones can modulate gene expression profiles

through regulation of microRNAs (miRNAs) (Nothnick et al., 2010). Alteration in miRNAs is

associated with uterine abnormalities including endometriosis, uterine leiomyoma and endometrial

carcinoma (Burney et al., 2009; Kuokkanen et al., 2010), strengthening the important regulatory

role of miRNAs in the endometrium. In a high-throughput miRNA analysis in the endometrial epithelium of ovariectomized mice under hormone stimulation, P4 was found to upregulate many miRNAs with known biological importance, including inhibition of proliferation (Lim et al., 2005;

Yuan et al., 2014). For example, miR-133a has been shown to target cyclin D2 to exert its P4-

induced anti-proliferative response (Liu et al., 2008; Pan et al., 2017). Other P4-induced miRNAs, such as miR-145/143 and miR-152, are involved in P4 related growth inhibition by suppressing

levels of cyclin D2 and Wnt-1, respectively (Yuan et al., 2015). It remains elusive whether there

are PR isoform-selective effects on the regulation of miRNA and its downstream targets.

3.1.6 PKC signaling in the endometrium

Several PKC isozymes (e.g., PKCα, βII, δ, η, and ζ) have been detected in the rat uterus,

particularly in the myometrium, where they have been implicated in mediating uterine contraction

(Jofre et al., 2013; Kim et al., 1999). PKC activity has been shown to upregulate urocortin production through activation of the urcortin promoter (Bamberger et al., 2007). The fine-tuning of PKC activation has been implicated in the control of myometrial contractility and determining the duration of gestation (Karteris et al., 2004; Vitale et al., 2016). Activated PKC has also been found to induce COX2 levels, allowing the production of prostaglandin to mediate uterine contractility (Wouters et al., 2014). In contrast to the inductive effect of PKC activation in uterine myometrium contractility, PKCα activity, not its abundance, was reported to significantly increase in preterm women (not in labor) when the myometrium is quiescent (Jofre et al., 2013), suggesting that PKCα is important in regulating the duration of pregnancy. This is in accordance with the role of PKCα as an inhibitor of smooth muscle contractility in the heart (Braz et al., 2004; Hambleton et al., 2006; Liu et al., 2009a). Clearly, further investigation is needed to elucidate the isozyme specific effects of PKCs on uterine contractility.

There is limited knowledge of the role of PKCs in the endometrium. PKC signaling has been shown to be involved in generating a suitable environment for embryo implantation. Many reproduction-related signaling molecules, such as Lewis Y antigens and calcitonin, which are actively secreted by the endometrial epithelial cells during implantation, were found to activate

Ca2+-dependent PKCs to facilitate trophoblast-endometrial interaction (Kalbag et al., 1991; Li et al., 2008; Li et al., 2009). The work of Pollard and coworkers on hormone-regulated changes in the uterine epithelium suggested the involvement of PKC in the normal physiology of endometrial epithelium. They reported that E2 exerts mitogenic effects through 1) IGF-1 signaling that activates PI3K/AKT pathways, leading to cyclin D1 nuclear accumulation, cell cycle progression, and DNA synthesis (Zhu and Pollard, 2007); 2) retention and chromatin loading of minichromosome maintenance proteins (MCMs) to permit DNA replication licensing (Pan et al.,

2006); and 3) regulation of protein synthesis by a PKC-ERK-mTOR signaling pathway (Wang et al., 2015). In addition, this group found that, while P4 inhibits DNA synthesis mediated by an IGF-

1/PI3K/AKT signaling axis, ERK-mTOR dependent protein synthesis was undisturbed (Wang et

al., 2015). Interestingly, the induction of PKC activity, as indicated by the phosphorylation of

MARCKS (myristoylated alanine-rich protein kinase C substrate), coincided with the switch from

non-receptive to receptive state in the uterus, which is regulated predominantly by P4 in

cooperation with nidatory E2 (Chen et al., 2003). Activation of PKC was found to be involved in

P4-mediated gene expression such as induction of immune response gene-1 (Irg1) (Chen et al.,

2003), suggesting that the PKC signaling pathway may modulate steroid hormone responsiveness

of the uterine epithelium. Together, these studies suggest that PKC signaling may have important

roles in regulating the physiological changes of the uterus and female fertility, although little is

known about isoform-specific functions in these processes.

While the main focus of our study is to decipher the functional role of PKCα in endometrial

cancer, it is important to survey its expression in the normal endometrium. The following section

reports our findings on PKCα levels and activation in the cycling human and mouse endometrium.

3.2 Results

3.2.1 Survey of PKCα expression in the endometrium

As a first step in characterizing PKCα expression in the endometrium, we analyzed PKCα levels in human uterine tissue using immunohistochemistry. While PKCα was detected in all samples, there was disparity in its levels among the different endometrial compartments (Figure.

3-1A). In both the secretory and proliferative uterine tissue, there were high levels of PKCα in the stroma, as indicated by strong IHC signal. In contrast, PKCα was expressed at a much lower level in the epithelium when compared to adjacent stroma. It has been shown previously that the activation of PKCs is associated with membrane translocation of the enzyme (Black and Black,

2012). Therefore, a membrane-associated pattern of PKC staining is indicative of its activation.

Interestingly, multiple regions of membrane-associated PKCα were observed in the secretory, but

not in the proliferative epithelium (Figure. 3-1B), suggesting that PKCα is activated when the

endometrium switches from a proliferative to a secretory state and that activity of this enzyme

might have a signaling role relating to the secretory uterine phenotype.

Fig. 3-1: PKCα expression in normal human uterus

(A) IHC analysis of PKCα expression in secretory and proliferative endometrium, S indicates the stromal compartment of the tissue. Arrow points to the endometrial epithelium. (B) Membrane associated PKCα in secretory endometrium. Arrow indicates epithelial regions with membrane- associated PKCα. Scale bar = 50 µm.

3.2.2 PKCα expression in mouse estrous cycle phases

The uterus undergoes significant physiological changes and remodeling throughout the estrous cycle. These changes are governed by an intricate network of signal transduction. As part

of the effort to understand if PKCα plays a role in regulating uterine physiology, we performed

IHC analysis on uterine tissues from all four estrous phases: proestrus, estrus, metestrus, and

diestrus, aiming to assess PKCα expression in relation to the murine estrous cycle. The estrous phases of the animals were determined from the cytological makeup of the vaginal smear. Vaginal

secretion was collected from sexually mature (5-6 weeks), virgin female mice and examined using microscopy. In our previous studies of the intestinal epithelium, we have shown that PKCα is

activated at the crypt-villus junction where there is a program switch from proliferation to

differentiation (Frey et al., 2000). As the cycling of the endometrium also involves a physiological

switch between these states as it progresses through the estrous cycle, we hypothesized that PKCα

may also be growth inhibitory in the endometrium and play a role in mediating this proliferation-

to-differentiation switch. To test this hypothesis, we investigated the correlation between PKCα

expression and proliferation, marked by positive Ki67 staining, in formalin-fixed, paraffin-

embedded uterine tissue collected from mice at each phase. PKCα was detectable in all four phases

of the estrous cycle (Figure 3-2). Levels of PKCα were relatively lower in metestrus, diestrus, and

proestrus. The enzyme was predominantly cytoplasmic during these phases. However, in estrus,

PKCα showed robust expression. Interestingly, there was reduced staining for the proliferation

marker Ki67 in the same phase. These observations suggest a negative correlation between PKCα

expression and proliferative activity in the epithelium of the uterus.

PKCα Ki67 Proestrus

100 μm 100 μm Estrus

100 μm 100 μm Metestrus

100 μm 100 μm Diestrus

100 μm 100 μm

Fig. 3-2: Correlation between PKCα and Ki67 expression during the estrous cycle

IHC analysis of PKCα expression and proliferative activity as indicated by Ki67 staining in.uterine tissues from mice at different phases of the estrous cycle: proestrus, estrus, metestrus, and diestrus, as determined by the cytology of vaginal smears.

3.2.3 Membrane-associated PKCα at estrous phase

Upon closer examination, we found that PKCα exhibited a distinct membrane-associated

staining pattern in both late proestrus and estrous phases (Figure 3-3A). In late proestrus, PKCα was predominantly inactive, as most of the detected PKCα resided in the cytoplasm. However, the glandular epithelium not only had higher PKCα levels, but staining for PKCα displayed a membrane-associated pattern, indicating that the enzyme was in its active form. This membrane-

associated PKCα staining intensified in estrous tissue, extending to both the glandular and luminal

epithelium and providing evidence for continuous strengthening of PKCα activating signals as the

cycle progresses from proestrus to estrus. However, PKCα maintained a cytoplasmic localization

in both metestrus and diestrus indicating a cessation of activating signals in the later phases of the

estrous cycle as shown in Figure 3-2. The membrane-associated staining in proestrous and estrous

endometrium was further confirmed in frozen sections using immunofluorescence (Figure 3-3B).

Fig. 3-3: Membrane-associated PKCα at estrous

(A) IHC analysis of PKCα in the luminal (LE) and glandular (G) compartments of uterine tissue during late proestrus and estrus. Membrane-associated localization of PKCα indicates activation of the kinase.

In proestrus, PKCα expression is cytoplasmic in the luminal compartment but membrane-associated in the glands (arrows). In estrus, membrane association of PKCα is detected in both the luminal and glandular epithelium (arrows). (B) Immunofluorescence staining for PKCα in frozen sections of luminal and glandular epithelium in estrus phase, confirming the localization seen in paraffin sections.

3.2.4 Inverse correlation between PKCα activation and cell proliferation

Thus far, our data have shown that membrane association of PKCα was most evident in the estrous phase, where the least proliferation was detected. In order to further confirm if there is a correlation between PKCα activation and cell proliferation, we examined PKCα expression/subcellular distribution and Ki67-marked proliferation by performing IHC analysis on serially sectioned uterine tissue at proestrus and estrus phases. As shown by images in Figure 3-4, proliferative activity was restricted to the luminal epithelium (indicated by arrowhead) and the glandular epithelium was completely devoid of Ki67 signal (shown by arrow). Interestingly, this compartmental delineation of staining correlated with PKCα levels and activation. Furthermore, in the lower panel, there was a striking correspondence between a Ki-67 negative region and strong membrane association of PKCα as indicated with arrows. Taken together, these observations demonstrated a negative correlation between PKCα activation and epithelial proliferation in the endometrium, suggesting that PKCα might have an inhibitory effect in the cycling uterus.

Fig. 3-4: PKCα expression during the estrous cycle

Serial sections from late proestrus and estrus phase uterine epithelium were analyzed for PKCα and the proliferation marker, Ki67. Top Panels: Survey sections show markedly reduced proliferation in the luminal epithelium (LE) during estrus phase; glands (G) are negative for Ki67 staining in both late proestrus and estrus phases. Block arrows in the lower panels indicate areas of PKCα membrane association/activation; arrowheads indicate areas that lack membrane-associated PKCα. Note the strong nuclear Ki67 staining in areas which lack membrane-associated PKCα. In contrast, areas with low levels of Ki67 exhibit strong membrane association/activation of the enzyme.

3.3 Discussion

The constant remodeling of the endometrium makes it one of the most dynamic tissues in the body. The cyclic breakdown and rebuilding of this tissue requires a complex signaling network

that is intricately regulated to ensure appropriate proliferation, differentiation, and apoptosis of the

epithelium. Molecular aberrations that tip this balance lead to outgrowth of the endometrium,

resulting in tumor development in this tissue. As part of our effort to decipher the role of PKCα in

endometrial cancer, we first profiled PKCα expression in the normal endometrium. Through IHC

analysis, we found that PKCα is ubiquitously expressed in the both human and mouse

endometrium, regardless of the stage of the menstrual cycle or estrous cycle, respectively.

However, PKCα levels are significantly lower in the endometrium when compared to the adjacent

stroma in the human uterus and in the mouse uterus in estrous phase, while there was little

difference between these two compartments in the mouse uterus in other phases. In the human

tissue, membrane-associated PKCα was only observed in the secretory phase, while this staining

pattern was detected in both late proestrus and estrus phases in the murine endometrium. This

observation, which was consistent in our analysis of paraffin-embedded and frozen murine uterine

tissues, suggests that activation of PKCα is initiated in proestrus and continues in estrus of the

mouse estrous cycle. Interestingly, there was minimal epithelial proliferation, as marked by

reduced Ki67-positive staining, when active PKCα was detected. This led us to hypothesize that

PKCα signaling may be anti-proliferative and tumor suppressive in the endometrium. The data presented in this section provide the basis for future studies to further investigate the hormonal regulation of PKCα activation and the signaling effects of PKCα in the endometrium.

A limited number of studies have demonstrated that E2- and P2- mediated responses in the uterus require PKC activation (Chen et al., 2003; Wang et al., 2015). Chen et al. showed that PKC activity is involved in implantation-related uterine responses induced by the synergistic actions of

P4 and E2 and the upregulation of P4-regulated genes. However, a subsequent study from the same group identified a PKC-ERK-mTOR signaling axis that activates E2-induced protein synthesis

(Wang et al., 2015). Because of the opposing actions of E2 and P4 in the endometrium, these findings raise questions on how these hormones regulate PKC activation in this tissue. Both of these studies assessed PKC activation using whole cell lysates prepared from uterine epithelial cells

isolated from ovariectomized mice after hormone treatment in vivo. PKC activity was then

determined by the phosphorylation of MARCKS, shown by Western blotting. Although the

hormone administration was performed in vivo, assessment of PKC activity was indirect and used

a non-selective PKC substrate. Additionally, previous studies have also shown uneven distribution

of ERα/β and PR-A/B expression in the different compartments of the endometrium; thus, the

purity of the epithelial isolate might be a confounding factor. Because of the high PKC levels in

the endometrial stroma (in the human), it is not clear whether epithelial or stromal PKC is

responsible for the hormone-induced responses in the uterus, or if there is isoform selectivity in

PKC-mediated signaling programs in the endometrium. In our study, active PKCα, demonstrated

by its membrane-associated staining, was detected at late proestrus and estrus, suggesting that

PKCα activation might be a late response to E2 stimulation or the result of rising P4. Staining of the uterine tissue directly assesses the presence of active enzyme in the uterine tissue and allows us to visualize its compartmental localization in vivo. However, the current data are observational and can only demonstrate a correlative relationship between PKCα activation and the cycling endometrium. To establish a mechanistic link between hormonal stimulation and PKCα activation, we propose to examine PKCα localization in uterine tissue from ovariectomized mice after treatment with E2, P4 or E2P4.

In a previous study, estradiol was shown to stimulate endometrial proliferation and induce

sustained PKC activation for at least 8 hours (Wang et al., 2015). Another study has demonstrated

elevated PKCα mRNA in response to tamoxifen and E2 in isolated human endometrial cells (Wu et al., 2005). However, it was unclear whether the enzyme was in its active state. Based on our examination of uterine sections from proestrus and estrus, PKCα activation was associated with reduced proliferation, which is thought to be a P4-induced phenotype. Furthermore, PKCα activity

is increased in pregnant women when sustained P4 is necessary to maintain pregnancy, which

supports the notion that PKCα activation is more closely associated with PR signaling. Thus,

although E2 might induce activation of other PKC isozymes, PKCα activation is likely to be regulated by P4. We expect to detect strongest membrane-associated PKCα staining in the uterus

when the animals are given E2P4 synergistically as the presence of low E2 levels is necessary for the proper activation of PR signaling (Chen et al., 2003). It is also possible that P4 alone can induce

PKCα activation; however, we expect it would be at a much lower level than in the E2P4 treated group. Meanwhile, the tissue will also be assessed for proliferation by Ki67 staining and for

downstream signaling effects on cyclin D1, p-mTOR, and pRPS6, whose expression or

phosphorylation have been shown to be hormonally regulated in the endometrium (Wang et al.,

2015). If our data demonstrate PR-mediated activation of PKCα, this will support a role for this

kinase in exerting anti-proliferative signaling in the endometrium and highlight its involvement in

the transition from a proliferative to differentiated state. Alternatively, we might observe

membrane-associated PKCα when mice are given E2 alone if PKCα is activated by ER.

Together with the current data, results from the proposed studies will provide a more comprehensive view of the role of PKCα signaling in the biology of normal, cycling endometrium, as well as a basis for its effects during endometrial tumorigenesis. Further investigation in ER or

PR isoform specific knockout models will be necessary to decipher isoform selective responses of

PKCα.

Chapter 4: Crosstalk Between PKCα and PI3K/AKT Signaling

is Tumor Suppressive in the Endometrium

Portions of the content covered in this chapter are the subject of an article under

consideration at Cell Reports by Hsu A.H. et. al

4.1 Endometrial cancer

Endometrial cancer (EC) is the most common gynecological malignancy in the United States,

with an estimated 60,380 new cases and 10,920 deaths in 2017 (Siegel et al., 2017). With an overall

5-year survival rate of >80%, EC has attracted less public attention than other cancers. Most cases

are diagnosed at an early stage when the patients experience unusual spotting, heavy bleeding or severe menstrual pain. When diagnosed early, patients are treated with surgery. For more advanced cases, chemotherapy or radiation may follow. However, advanced (which has spread beyond the uterine cavity) and recurrent disease is refractory to treatment and the prognosis for these patients is dismal, with survival estimates of less than one year (Engelsen et al., 2009). Because of its high incidence, EC is the sixth leading cause of cancer death in women, accounting for more deaths than melanoma, cervical cancer, glioblastoma, all lymphomas, or all leukemias (Siegel et al., 2017).

Alarmingly, the incidence and mortality for EC are on the rise, with a >40% increase since 2005.

Since obesity is a major risk factor for the disease (Fader et al., 2009), EC will become an even greater health concern as the effects of increased societal obesity become evident in coming years.

4.1.1 Clinical and molecular classification

EC has historically been classified into two histopathological subtypes. Type I tumors (85-90% of cases) are of endometrioid histology, while Type II non-endometrioid tumors can be further classified by histological appearance as serous, clear cell, mixed cell, or malignant mixed Mullerian tumors (MMMT), with serous being the predominant subtype in this category (Suarez et al., 2017).

The original landmark paper published in 1983 described the clinical features observed in each of the subtypes (Bokhman, 1983). A majority of EC patients are diagnosed with Type I or endometrioid subtype. Many of these patients are post-menopausal women who are also likely to have elevated estrogen exposure (related to hormone replacement therapy, obesity, late onset of menopause, nulliparity), diabetes, hypertension, obesity, and hyperlipidemia. Formation of Type I

ECs follows a sequential course of progression initiated by hyperplasia, atypical hyperplasia, and benign hyperplasia, followed by premalignant neoplasia (Dedes et al., 2011). Disease in these patients often presents as low grade hyperplasia, with histology resembling the proliferative epithelium, anovulatory bleeding, and sometimes infertility (in women at reproductive age). Type

I tumors have lower invasive potential, often only penetrating the superficial surface of the myometrium. These tumors are more sensitive to progesterone-mediated growth inhibition. Most of type I patients are successfully treated with surgery, progestin therapy, and radiation for more advanced cases. Patients with Type I disease have more favorable prognosis and have an overall

5-year survival rate of 85% (Morice et al., 2016).

As discussed above, Type II non-endometrioid tumors encompass several histological types including serous, clear cell, malignant mixed Mullerian tumors, and mixed epithelial types. These cancers generally have an accelerated course of disease progression, and tumors are more invasive with a higher likelihood of recurrence. Although Type II EC only accounts for ~20% of all endometrial cancer cases, it is responsible for 50% of EC-related mortality (Lobo and Thomas,

2016). At the time of diagnosis, these tumors have usually already invaded the myometrium and lymphovasculature and are often are metastatic. Despite the difference in disease type, patients with Type II EC also receive surgery as first line therapy. Due to the more aggressive nature of the disease, patients with Type II EC also receive adjuvant radiation and combinational chemotherapy to limit recurrence. Progestin therapy is rarely administered to these patients as Type II tumors are often insensitive to P4. However, if the disease recurs, the tumors are often resistant to existing

therapies and the prognosis for these patients is dismal (median survival of 8-16 months) (Dellinger

and Monk, 2009). The simplistic classification of EC established in 1983 has guided clinical

prognosis and treatment for this disease in the following decades. As the EC field gains more

understanding of the biology of these tumors, the classification of ECs continues to refine this

dualist model.

The first evolution of this two-category classification of EC disease was fueled by advances in the molecular understanding of ECs (Table 4-1) (Dedes et al., 2011). Molecular analysis of these tumors has identified genetic alterations that are more prevalent in each EC type. Because of the high prevalence of mutations in PTEN (83%), loss of PTEN function is considered a primary feature of Type I tumors (Mutter et al., 2000; Risinger et al., 1998). Additional oncogenic mutations are found in other molecules in the PI3K/AKT pathway, including activating mutations in PIK3CA

(~ 30%) and AKT (3%) (Catasus et al., 2008; Miyake et al., 2008; Shoji et al., 2009). The frequent co-existence of mutations in PI3K subunits and PTEN highlights the tumorigenic advantage of hyperactive AKT signaling in endometrial carcinogenesis (Oda et al., 2005). The microsatellite instability phenotype of Type I ECs has been attributed to inactivation of DNA repair genes (e.g.

MLH1, MSH2, MSH6, and PMS2) (MacDonald et al., 2000; Mackay et al., 2010; Miturski et al.,

2002) Other molecular aberrations commonly found in Type I ECs include KRAS mutations,

FGFR2 mutations, and nuclear accumulation of β-catenin (Koul et al., 2002; Lax et al., 2000;

MacDonald et al., 2000; Pollock et al., 2007; Schlosshauer et al., 2000; Stefansson et al., 2004). In contrast, KRAS mutations and FGFR2 mutations are mutually exclusive (O'Hara and Bell, 2012).

Prevalence in Prevalence in Alteration Type I (%) Type II (%)

PIK3CA mutation ~ 30 ~ 20

PIK3CA amplification 2-14 46

KRAS mutation 11-26 2

AKT mutation 3 0

PTEN loss of function 83 5

Microsatellite instability 20-45 0-5

Nuclear accumulation of β- 18-47 0

catenin

E-Cadherin loss 5-50 62-87

TP53 mutation ~ 20 ~ 90

p16 loss of function 8 45

HER2 overexpression 3-10 32

HER2 amplification 1 17

FGFR2 mutations 12-16 1

Table 4-1: Molecular alterations in endometrial cancer. Adapted from (Dedes et al., 2011)

The most commonly observed aberration in Type II EC disease is mutation in p53, which is found in 90% of Type II tumors, particularly in the serous subtype (Jia et al., 2008; Khalifa et

al., 1994; Lax et al., 2000). In serous EC, somatic mutations of p53 are believed to be an initiating event (O'Hara and Bell, 2012). Interestingly, serous ECs share clinical and pathological similarities

with serous ovarian tumors in which p53 mutations have significant pathogenic and therapeutic

impact (Di Cristofano and Ellenson, 2007). Alterations in p16 levels are more commonly detected

Ink4 in Type II than in Type I ECs. p16 negatively regulates progression into the G1/S phase by

binding to cyclin dependent kinases Cdk4 and Cdk6 and is, thus, considered a tumor suppressor

(Liggett and Sidransky, 1998). Loss of p16 in EC is associated with aggressive disease with poor prognosis (Ignatov et al., 2008; Salvesen et al., 2000). However, other studies have reported overexpression of p16 in ECs (O'Hara and Bell, 2012). Elevated p16 is thought to be indicative of oncogene-induced senescence in benign or pre-malignant hyperplasia, or the result of the cell’s effort to arrest proliferation when the Rb pathway is defective (Romagosa et al., 2011). Other molecular alterations that are more commonly associated with Type II EC include loss of E-

Cadherin and elevated levels of HER2 due to overexpression or gene amplification. Although molecular profiling provided a more detailed distinction of EC types, some aberrations are found in both types. For example, mutations in PIK3CA are found at similar frequencies in both Type I and II ECs (although there appears to be a preference for mutations in exon 20 in Type II disease, whereas Type I ECs have mutations in both exon 9 and 20 of PIK3CA) (Catasus et al., 2008).

Interestingly, PIK3CA amplification is more prevalent in Type II than Type I ECs. The classical subtyping of EC disease based on tumor histology was particularly problematic for high-grade endometrioid tumors (Clarke and Gilks, 2010). Newly identified molecular alterations in ECs can serve as biomarkers to facilitate more accurate subtyping and better guidance in treatment selection

(Alkushi et al., 2010; Darvishian et al., 2004; Yemelyanova et al., 2009). Using phenotypic markers such as MLH1, MSH2, p16, cyclin D1, ERBB2, and p53, 37% of grade 3 tumors exhibited molecular profiles of endometrioid tumors while 63% of the cases shared molecular similarities with serous carcinomas (Alvarez et al., 2012). This suggests that a comprehensive molecular profiling is needed to maximize the accuracy of the classification of ECs. Together, these findings provided new insights into the molecular alterations in ECs and the prognostic potential of these molecules, and also revealed opportunities for targeted therapy for the disease.

In 2013, The Cancer Genome Atlas project (TCGA) completed their integrated genomic, transcriptomic, and proteomic analysis of 373 ECs including 307 low or high grade endometrioid,

53 serous, and 13 mixed histology cases (Kandoth et al., 2013). The top most significantly mutated

genes for each subgroup are listed in Table 4-2.

Gene Mutation Molecular subtype Gene name symbol frequency POLE ultramutated POLE Polymerase epsilon, catalytic subunits 100 % PTEN Phosphatase and tensin homolog 94% CSMD3* CUB and Sushi multiple domains 3 94% TAF1 TAF1 RNA polymerase II, TATA box binding protein 83% (TBP)-associated factor, 250KDa FBXW7 F-box and WD repeat domain containing, E3 83% ubiquitin protein ligase Hypermutated/ PTEN Phosphatase and tensin homolog 88% microsatellite- unstable PIK3CA Phosphatidylinositol-4,5-bisphosphate 3-kinase, 54% catalytic subunit alpha PIK3R1 Phosphoinositide-3-kinase, regulatory subunit 1 40% (alpha) Copy number low/ PTEN Phosphatase and tensin homolog 77% microsatellite-stable PIK3CA Phosphatidylinositol-4,5-bisphosphate 3-kinase, 53% catalytic subunit alpha CTNNB1 Catenin, beta 1, 88 KDa 52% ARID1A AT rich interactive domain 1A (SWI-like) 42% Copy number high/ TP53 Tumor protein 53 92% serous like PIK3CA Phosphatidylinositol-4,5-bisphosphate 3-kinase, 47% catalytic subunit alpha PPP2R1A Protein phosphatase 1, regulatory subunit A, alpha 22% FBXW7 F-box and WD repeat domain containing, E3 22% ubiquitin protein ligase Table 4-2: Top 3 most significantly mutated genes in 4 molecular subtypes of EC. *High mutation frequency of this gene is likely a statistical artifact. Adapted from (Le Gallo and Bell,

2014)

While mutations in known EC-associated genes were detected, this comprehensive analysis revealed many more significantly altered genes that were not identified to drive endometrial tumorigenesis (e.g. BCOR, MECOM, ARID5B, and NKAP etc.). Based on results from this analysis, four molecular subgroups of ECs have been recognized: POLE ultramutated, hypermutated/microsatellite-unstable (MSI), copy number low/microsatellite-stable (MSS), and copy number high/serous like, as reviewed in (Le Gallo and Bell, 2014). This marked the second evolution of EC classification. DNA polymerase ε (POLE) exonuclease, encoded by the POLE gene, catalyzes the synthesis of the leading strand during DNA replication (Pursell and Kunkel,

2008). Mutations in the POLE gene, such as those identified in ECs, disrupt the enzyme’s proofreading ability leading to an ultramutated phenotype (232 × 10-6 mutations/Mb; 867-9714

mutations/tumor), as well as high incidence of C>A transversions. Other studies have also reported

POLE mutations in EC (Church et al., 2013). Despite the extraordinarily high mutation numbers

in POLE ultramutated tumors, patients in this group have the most favorable prognosis among all

EC subgroups. All POLE ultramutated tumors are of endometrioid histology, accounting for 10%

of all endometrioid tumors examined (6.4% of low grade and 17.4% of high grade endometrioid

tumors). Another subgroup that exclusively consists of endometrioid tumors is the

hypermutated/MSI category. As its name indicates, the hypermutated phenotype in these tumors is

the result of reduced expression of the mismatch-repair protein MLH1 and frequent methylation of the MLH1 promoter. This is also consistent with previous reports of high MSI frequencies in ECs

(An et al., 2007; Konopka et al., 2007; Zighelboim et al., 2007). MSI tumors comprise 28.6% of low grade and 54.3% high grade endometrioid cases in the TCGA cohort. The next subgroup includes the copy number low/MSS tumors. The MSS tumors account for 60% of all low grade endometrioid tumors, making it the most common subgroup of endometrioid tumors. Unlike the previous two subgroups, the MSS subtype includes serous and mixed-histology carcinomas. In contrast to the higher mutation frequencies in KRAS and FGFR2 in MSI endometrioid tumors,

CTNNB1 mutations are more common in MSS endometrioid tumors (Byron et al., 2012). Despite the frequent occurrence of high grade endometrioid histology in MSI tumors and the inclusion of the more aggressive serous or mixed-histology MSS carcinomas, these two subtypes displayed similar trends in progression free survival. The most distinct subgroup of ECs is known as copy number high/serous-like. Almost all serous cases are categorized in this subgroup (97.7% of serous and 75% of mixed-histology). Phenotypically, these tumors are more similar to high-grade serous ovarian tumors than to other ECs (Merritt and Cramer, 2010). Genetic profiling of serous EC has also identified prominent mutations in TP53 in these tumors, mirroring serous ovarian tumors, which suggests that mutant p53 is the main driver for formation of serous EC. Mutation in other

genes such as PTEN, CTNNB1, and PIK3R1 that are highly prevalent in other subgroups are

significantly less frequent in serous EC, although they are still present. As expected, these tumors

generally also have decreased pAKT levels, differing from the AKT hyperactivation that commonly

drives carcinogenesis in other EC subtypes. In addition, TCGA analysis also identified focal

amplification of MYC, ERBB2, and CCNE1 in 23-25% in this subgroup. Because of distinct

molecular features and the close resemblance between serous endometrial and ovarian tumors,

targeted therapies developed against high grade serous ovarian tumors might also be effective in

copy number high/serous-like EC (Iijima et al., 2017).

The rapid advances in sequencing technology and high-throughput omics analysis sparked the

second evolution of EC classification. The concerted effort by the TCGA research network

provided a comprehensive view of EC tumor biology, and strongly urged re-definition of EC

classification, which will also impact clinical decisions on appropriate treatments based on the disease subgroups.

4.2 Results

4.2.1 Transcriptional suppression of PKCα is associated with high-grade EC

As a first step toward understanding the role of PKCα signaling in EC, we performed IHC analysis to profile its expression in a panel of 436 human endometrial tumors (330 endometrioid;

106 non-endometrioid) in tissue microarray (TMA) format. While PKCα was detected in normal secretory and proliferative endometrium (as shown in the Figure 3-1) and was retained in some lesions, the enzyme was absent or markedly reduced in >60% of endometrioid ECs and >50% of all non-endometrioid tumors, with the proportion of PKCα deficient tumors ranging from 31% of

MMMT to 70% of clear cell ECs (Figure 4-1, middle panel).

The availability of frozen tissue for a subset of these ECs (24 endometrioid; 26 non- endometrioid) allowed parallel quantification of PKCα mRNA by qRT-PCR. A significant correlation was noted between PKCα mRNA and protein levels in both tumor subtypes (Figure 4-

1, bottom table), indicating that downregulation of PKCα protein in human EC is controlled primarily at the level of mRNA expression. Analysis of TCGA data confirmed that PKCα mRNA is markedly downregulated in human ECs (p=2.77e-9), and pointed to increased or decreased expression of several additional PKC family members in these tumors (Figure 4-2). Like PKCα,

PKCβ, ε, η, and θ, showed reduced mRNA expression in tumors relative to normal endometrial tissue, while levels of other PKC isozymes were elevated (e.g. PKCδ and ξ) or unchanged (e.g.

PKCγ and ι) (Figure 4-2). Note that PKCα is one of the most profoundly downregulated PKCs in

this tumor type.

Grade 1 Grade 3

PKCα +ve 80 PKCα -ve

40 % of Cases

0 Endometrioid Mixed MMT Serous Clear cell epithelial (n= 330) (n = 26) (n = 16) (n = 54) (n = 10) Non-endometrioid

SpearmanRank Correlation of PKCα mRNA and Protein Levels

rs P-value EC tumors 0.46 0.001 Endometrioid 0.51 0.01 Non-endometrioid 0.41 0.02

Fig. 4-1: PKCα expression in EC

IHC analysis of a panel of ECs including endometrioid and non-endometrioid subtypes. Decreased

PKCα expression was found in over 60% of endometrioid tumors and 50% of non-endometrioid subtypes. PKCα signal in the stroma (S) serves as a positive control for IHC staining in Grade 3 tumors.

Scale bars, 50 µm. Spearman Rank correlation analysis of PKCα protein and mRNA in endometrial tumors Analysis done with assistance from Adrian Black.

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Fig. 4-2: Alterations in PKC isozyme mRNA expression in ECs s

PKC isozyme mRNA levels in normal vs. tumor tissue based on analysis of TCGA database. Expression of PKCα, β, ε, η, and θ mRNA is reduced in endometrial tumors, while ECs have increased levels of

PKCδ and ξ mRNA. Levels of PKCγ and ι mRNA are comparable in the normal endometrium and endometrial tumors.

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To further examine the regulation of PKCα in EC, we determined its expression in a panel of 16 well-characterized human EC cell lines harboring a variety of genetic mutations characteristic

of human endometrial tumors (Table 4-3) (Weigelt et al., 2013). Six of these cell lines were found

to express markedly reduced levels of PKCα (Figures 4-3, top panel) and, consistent with findings

in EC tissues, a significant correlation was observed between PKCα protein and mRNA levels

across the panel of EC cell lines analyzed (Figure 4-3). To test for changes in mRNA transcription,

RNA was pulse-labeled with 5-ethynyl uridine (EU) in two PKCαhigh (HEC-1-A; HEC-50Co) and two PKCαlow (RL-95; Ishikawa) EC cell lines. PKCαlow cells incorporated markedly lower amounts of EU into PKCα mRNA than PKCαhigh cells, with the extent of labeling closely paralleling the

differences in overall levels of PKCα mRNA in the cells (Figure 4-4). Together, these data confirm

that PKCα is downregulated in a significant fraction of human ECs and EC cell lines, and that

PKCα deficiency primarily reflects a reduction in the transcription rate of the PKCα gene.

Further analysis of the TMA data revealed a positive correlation between decreased levels

of PKCα and disease aggressiveness in the endometrioid EC subtype, with 56% of grade 1, 61% of

grade 2, and 76% of grade 3 lesions showing loss of the enzyme (Figure 4-5, top panel). A significant association was also observed between loss of PKCα and risk of high-grade endometrioid disease (OR = 3.4) (Figure 4-5, bottom panel). PKCα deficiency also correlated with

disease severity when only PTEN negative endometrioid tumors were considered (OR = 2.8),

indicating that PKCα loss is a risk factor for aggressiveness in human EC even in the presence of

activating mutations in the PI3K/AKT pathway. Consistent with this conclusion, reverse phase

protein array analysis of 244 EC cases (~80% endometrioid (Liang et al., 2012)) from The Cancer

Proteome Atlas (TCPA) showed that PKCα expression trends with enhanced survival of EC

patients (Figure 4-6). In contrast, lower expression of PKCε or ι was associated with higher

survival probability, consistent with the oncogenic properties of these isozymes in other systems

(Bae et al., 2007; Benavides et al., 2011; Cacace et al., 1996; Eder et al., 2005; Murray et al., 2004;

Regala et al., 2005a). Collectively, the data show that loss of PKCα can be an early event in EC

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and that PKCα deficiency is associated with more aggressive disease, pointing to a potential tumor suppressive function of PKCα signaling in the endometrium.

Cell line PTEN PIK3CA PIK3R1

HEC-1-A WT G1049R

HEC-1-B WT G1049R

HEC-50 WT E468InsGEYDRLYE

KLE WT

AN3CA R130fs R557_K561>Q

Ishikawa V317fs; V290fs L570P

RL95-2 M134I; R173H; N323fs R386G

SNG-II K6fs

SNG-M K164fs; V290fs R88Q

HEC-6 V85fs; V290fs R108H; C420fs

HEC-59 Y46H; R233X; P246L; L265fs R38C K567E; S460fs

HEC-108 K6fs; E288fs A331V

HEC-116 R55_L70>S; R173C R88Q

HEC-151 I33del; Y76fs C420R

HEC-251 S10N; E299X M1043I

HEC-265 L318fs H180fs; Q586fs

Table 4-3: Mutations in EC cell lines. Adapted from (Weigelt et al., 2013)

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Fig. 4-3: Correlation between PKCα mRNA and protein levels in EC cell lines

Survey of PKCα protein and mRNA levels in a panel of EC cell lines using WB analysis and qRT-PCR respectively. The mRNA quantification was normalized against 18s rRNA. Vertical lines in the upper panel indicate rearrangement of lanes from a single membrane for clarity. Black bars: PKCαhigh. Grey bars: PKCαlow. Spearman Rank correlation analysis of PKCα protein and mRNA in EC cell lines.

Together, these data demonstrated a correlation between PKCα mRNA and protein levels. Data represent results from 2-3 biological replicates ± SEM. Analysis done with assistance from Adrian

Black.

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Fig. 4-4: Alterations in PKCα transcription in EC cells

RNA was pulse-labeled with EU in PKCαhigh and PKCαlow EC cell lines. Levels of labeled PKCα mRNA were measured by qRT-PCR. PKCαhigh EC cells incorporated significantly higher amounts of EU than

PKCαlow cells, pointing to differential transcription rates in these cells. Data represent results from 2-3 biological replicates. Experiments performed by Adrian Black.

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Fig. 4-5: Loss of PKCα is associated with higher-grade EC

The bar graph depicts PKCα expression status in endometrioid tumors of different grades. Included in the table is the odds ratio of tumor grade based on PKCα expression. Grade 1 and 2 tumors are considered low grade and Grade 3 tumors are high grade. ** p < 0.01, *** p < 0.005 (2-sided Fisher’s exact test).

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Fig. 4-6: Survival probability of EC patients with high and low PKC expression

Patients with higher levels of PKCα have higher survival probability. Additionally, patients that have lower levels of PKCε or ι also have better survival prognosis. Results were obtained from analysis of

TCPA (for PKCα) or TCGA (for PKCε and ι) databases.

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4.2.2 PKCα expression is lost in mouse models of endometrial hyperplasia

The expression of PKCα was also explored in endometrial lesions arising in mice with single allele knockin of Pten mutations seen in Cowden syndrome patients (frameshift/truncation

PtenΔ4-5/+ mutation, and missense PtenG129E/+ or PtenC124R/+ mutations). These mice develop

endometrial hyperplasia with high penetrance upon loss of expression of the wild-type (WT) Pten

allele (Figure 4-7A) (Wang et al., 2010). Loss of the WT allele is reflected in absence of PTEN

staining in PtenΔ4-5/+ lesions and a reduction in PTEN staining in PtenG129E/+ or PtenC124R/+ lesions

(Figure 4-7B, C). As expected, loss of PTEN function was accompanied by a marked increase in

AKT activity, as indicated by enhanced pAKTSer473 signal. Remarkably, these lesions uniformly

lacked PKCα protein expression, with a perfect correspondence between PTEN deficiency, AKT

activation, and PKCα loss. The sharp delineation between PKCα-expressing normal endometrial

tissue and PKCα-deficient, pAKT positive lesions is clearly evident in these images (arrows and boxed areas, and Figure 4-7D). qRT-PCR analysis of normal and tumor tissue collected by laser capture microdissection demonstrated that loss of PKCα protein is associated with a reduction in

PKCα mRNA in these early lesions (Figure 4-8). PKCα loss was also observed in uterine tumors of mice with cre-mediated conditional endometrial deletion of Pten in the epithelium and stroma

(Ptenpr-/-) or the epithelium alone (Ptenltf-/-), regulated by the progesterone receptor promoter or the

lactoferrin promoter, respectively (Figure 4-9) (Daikoku et al., 2008; Daikoku et al., 2014).

Interestingly, these lesions expressed increased levels of Id1, a gene known to be suppressed by

PKCα signaling (Figure 4-10) (Hao et al., 2011). The loss of PKCα in multiple in vivo models of

PI3K/AKT-driven endometrial neoplasia further supports a role for PKCα deficiency in uterine

tumorigenesis.

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Fig. 4-7: Concurrent loss of PKCα and PTEN correlates with pAKT

See next page for figure legend

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Immunohistochemical analysis of PTEN, pAKT, and PKCα expression in consecutive uterine sections from three strains of Pten-mutant mice - PtenΔ4-5/+ (A), PtenC124R/+ (B), and PtenG129E/+ (C). Arrows indicate hyperplastic lesions lacking functional PTEN and PKCα expression, and strongly positive for pAKT. In (B), inserted are magnified images of the boxed regions. (D) Image depicting the loss in PKC in the endometrial hyperplasia and its retention in the adjacent normal endometrium (NE) and endometrial hyperplasia (EH). Scale bar, 50 µm. Images are representative of all hyperplastic lesions observed in these animals. Experiments performed with assistance from Kathryn Curry and Kang-Sup

Shim.

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Fig. 4-8: Laser Microdissection of pAKT- and PKCα-labeled uterine epithelium for qRT-PCR analysis of PKCα mRNA levels.

Immunoflourescence analysis of serial sections of frozen uterine tissue from 9 month PtenΔ4-5/+;Prkca+/+ mouse identifies normal epithelium (PKCαpos; pAKTneg, indicated by large arrows) and tumor lesions

(PKCαneg; pAKTpos, indicated by small arrows). The tissues were isolated by laser microdissection and qRT-PCR analysis was performed to quantify PKCα mRNA levels in the corresponding regions.

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Fig. 4-9: Loss of PKCα in uterine tumors of mice with cre-mediated conditional endometrial deletion of Pten.

Immunohistochemical staining of PKCα in uterine tumors of mice with cre-mediated conditional endometrial deletion of Pten in the epithelium and stroma (Ptenpr-/-) or the epithelium alone (Ptenltf-/-), regulated by the progesterone receptor promoter or the lactoferrin promoter, respectively. Note the

PKCα positive stroma and myometrium and PKCα negative endometrial tumor tissue. Scale bar, 50

µm.

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Fig. 4-10: Enhanced Id1 expression in endometrial hyperplasia correlates with loss of PKCα

IHC analysis of PKCα and Id1 expression in endometrium from PtenG129E/+ and PtenC124R/+ mice. Note the strong nuclear staining for Id-1 in hyperplastic regions that lack PKCα (block arrows). Arrowheads indicate normal endometrial glands with high PKCα and low Id1 expression. IHC staining performed by Kathryn Curry.

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4.2.3 PKCα deficiency in EC is not a direct result of PTEN loss/ AKT activation and does not appear to drive loss of PTEN

To determine if PKCα deficiency in EC is a direct consequence of PTEN loss of function/AKT activation, PTEN was stably knocked down in three PTEN WT, PKCαhigh human

EC cell lines (Figure 4-11A). While the functional consequences of PTEN knockdown were

confirmed by increased AKT phosphorylation, loss of PTEN did not affect the levels of PKCα (or

of any other PKC isozyme, Figure 4-12). Similarly, inhibition of aberrant AKT activity in PTEN

mutant/PKCαlow EC cell lines by restoration of PTEN expression (Figure 4-11B) or treatment with

AKT or PI3K inhibitors (MK-2206 or LY294002, respectively) (Figure 4-11C), failed to affect

PKCα levels. Taken together, these data indicate that PKCα deficiency is not a mere passenger event of PI3K/AKT hyperactivation, and suggest that loss of the enzyme may contribute directly to endometrial tumor development. To test the possibility that PKCα deficiency drives loss of

PTEN in endometrial cells, we explored the effects of PKCα knockdown in PTEN WT, PKCαhigh

EC cell lines. As shown in Figure 4-11D, PKCα knockdown failed to affect PTEN levels in these cells.

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Fig. 4-11: PKCα deficiency in EC is not a direct result of PTEN loss or AKT activation and does not affect PTEN levels

Western blot analysis of PKCα levels in EC cells with A) stable expression of PTEN shRNA, B) adenoviral restoration of PTEN, and C) treatment with AKT (MK-2206) or PI3K (LY294002) inhibitors. siRNA-mediated knockdown of PKCα, in turn, failed to affect PTEN levels (D). Data represent results from at least 3 biological replicates.

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Fig. 4-12: PTEN knockdown did not alter the expression of other members of the PKC family

Western blot analysis of the expression of indicated PKC isozymes in EC cells stably expressing shRNA targeting PTEN. Data represent results from at least 3 biological replicates.

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4.2.4 PKCα inhibits endometrial tumorigenesis in vivo and in vitro

To directly examine the effects of PKCα deficiency on endometrial tumor development,

+/+ Δ4-5/+ -/- Δ4-5/+ we generated Prkca ;Pten and Prkca ;Pten mice. Uterine tissue was collected at 1, 3, 5,

7, and 9 months and hyperplastic lesions were identified by pAKTSer473 staining and

histopathological analysis (Wang et al., 2002) (Figure 4-13, top and middle panels). Quantification

of the proportion of pAKT positive endometrium pointed to a statistically significant 3-fold

increase in tumor burden in PKCα-/- animals at both 1 and 3 months compared with PKCα+/+

littermate controls (Figure 4-13, bottom graph). However, this difference diminished at later times.

Blinded histological analysis of H&E stained sections by a pathologist (Dr. Yuri Sheinin)

+/+ Δ4-5/+ confirmed this conclusion (not shown). Since hyperplastic lesions arising in Prkca ;Pten mice uniformly lacked PKCα, the increased tumor burden in Prcka-/-;PtenΔ4-5/+ mice indicates that PKCα

loss is a rate limiting step in endometrial tumorigenesis in this model.

To test the effects of PKCα on the transformed phenotype of human EC cells, PKCα expression was restored in PKCαlow cell lines by adenoviral transduction or silenced in PKCαhigh

cells using siRNA, and anchorage-independent growth was assessed by colony formation in soft

agarose. Exogenous PKCα markedly inhibited colony formation of PKCαlow EC cells, and colonies that did form were substantially smaller than those of LacZ-transduced cells (Figure 4-14). One infection unit/cell (moi) of PKCα adenovirus was sufficient to suppress anchorage-independent growth of these cells (Figure 4-15), pointing to a strong tumor suppressive activity of PKCα in human EC cells. The requirement for PKCα kinase activity in these effects was tested using kinase dead enzyme (KD-PKCα). Since KD-PKCα is unstable in cells (Pysz et al., 2009), 25 or 50 moi of

KD-PKCα adenovirus were used to ensure levels of mutant protein expression comparable with

WT PKCα at 1 moi. KD-PKCα failed to affect colony formation, even at the highest level of expression (Figure 4-15). The effects of PKCα knockdown were examined in PKCαhigh HEC-6 cells, which grow less efficiently in soft agarose than other EC cells in our panel (Figure 4-16).

Notably, PKCα depletion led to an approximately two-fold increase in colony formation of these

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cells. Thus, PKCα kinase activity is a potent inhibitor of the transformed phenotype of human EC

cells.

Fig. 4-13: PKCα inhibits endometrial tumorigenesis in vivo

Uterine tissue from PtenΔ4-5/+;Prkca+/+ and PtenΔ4-5/+;Prkca-/- transgenic mice aged for 1, 3, 5, 7, or 9 months was analyzed for pAKT by immunohistochemistry, as shown in the representative images from

1- and 3-month cohorts. At least two sections were analyzed for each animal. Percent pAKT-positive uterine epithelium was measured as a proportion of the total epithelial compartment of the endometrium.

* indicates p < 0.05 as determined by Wilcoxon Rank Sum test. Grey circles, PtenΔ4-5/+;Prkca+/+. Black circles, PtenΔ4-5/+;Prkca-/-. Scale bar, 200 µM.

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Fig. 4-14: PKCα inhibits endometrial tumorigenesis in vitro

PKCαlow endometrial cancer cell lines (Ishikawa, RL95-2, HEC-59, HEC-116, HEC-251) were infected with adenoviral vectors expressing either PKCα (α) or LacZ (Z) at a moi of 20 and analyzed for PKCα expression by immunoblotting; β-actin serves as a loading control (upper panel). Infected cells were plated in soft agarose and quantification of colony formation is presented relative to the LacZ control

(bottom graph). Data are representative of at least 3 independent experiments.

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Fig. 4-15: PKCα kinase activity is required for its tumor suppressive function

Assessment of anchorage independent growth of EC cells transduced with WT-PKCα or KD-PKCα.

The higher mobility of KD-PKCα reflects the absence of priming phosphorylation of the enzyme

(Carpenter et al., 2004). * p < 0.05, ** < 0.01, or ns (not significant) as calculated by 2-sided student’s t test. Error bars indicate mean ± SEM from 3 biological replications. β-actin serves as a loading control.

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Fig. 4-16: siRNA-mediated knockdown of PKCα promotes tumorigenesis

Assessment of anchorage independent growth of EC cells with siRNA-mediated knockdown of PKCα.

* p < 0.05, ** < 0.01, or ns (not significant) as calculated by 2-sided student’s t test. Error bars indicate mean ± SEM from 3 biological replications. β-actin serves as a loading control.

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4.2.5 PKCα activity inhibits PI3K/AKT signaling in EC cells via a PHLPP 1/2-independent, okadaic acid/ calyculin A-sensitive mechanism

Since a majority of ECs are highly dependent on PI3K/AKT signaling (Weigelt et al.,

2013), we next examined the effects of PKCα activity on this pathway using a panel of human EC cell lines (Table 4-3). Treatment of PKCαhigh cells with the PKC agonist phorbol 12-myristate 13- actetate (PMA) markedly decreased the phosphorylation of AKT as well as its downstream substrate, PRAS40 (Figure 4-17A). Similarly, the short-chain diacylglycerol 1,2-

Ser473 dioctanoylglycerol (DiC8), a more physiological PKC activator, also led to decreased pAKT

levels (Figures 4-17B). Consistent with the involvement of PKCα, PMA treatment did not affect

AKT phosphorylation in PKCαlow cells (Figure 4-17C), an effect that was rescued by restoration of

PKCα by adenoviral transduction (Figure 4-17D). Furthermore, PKCα inhibition with Gö6976, which targets the classical PKCs, blocked the ability of PMA to reduce pAKT in PKCαhigh cells

(Figure 4-18). Finally, knockdown of PKCα in PKCαhigh cells with two different siRNAs prevented

PMA-induced AKT hypophosphorylation (Figure 4-19). Since PKCα siRNA had no notable effect on the expression of other PKC isozymes (Figure 4-20), these data confirm a requisite role for

PKCα in suppression of AKT activity in EC cells. Interestingly, PKCα knockdown also led to increased basal AKTSer473 phosphorylation, pointing to a tonic repressive effect of the enzyme on

PI3K/AKT signaling in these cells (Figure 4-19, compare lane 1 with lanes 3-6 in both panels). Full activation of AKT requires phosphorylation at both S473 and T308 residues. Both of these residues are targeted by PKCα in EC cells, as indicated by (a) the parallel decrease in their phosphorylation following PMA treatment, a sustained effect that is supported by the continued activation of PKCα

(Figure 4-21), and (b) the robust increase in phosphorylation at both sites induced by PKCα knockdown in PKCαhigh cells (Figure 4-22).

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Fig. 4-17: PMA-induced activation of PKCα reduces AKT phosphorylation

Western blot analysis of the phosphorylation status of AKT and its downstream target PRAS40 in

PKCαhigh and PKCαlow EC cell lines treated with vehicle (C) or 100 nM PMA (P) for various times as indicated (A and C) or with 20 µg DiC8 for 6 hr (B). DiC8 was added in fresh medium every 30 min to account for its rapid metabolism in cells. (D) Adenoviral restoration of PKCα rescued the effect of PKC agonists in PKCαlow cells. Data are representative of at least 3 biological replications. β-actin or total

AKT serve as a loading control. Western blot analysis performed with assistance from Michelle Lum.

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Fig. 4-18: The cPKC inhibitor Gӧ6976 blocks PMA-induced AKT hypophosphorylation

Treatment with Gӧ6976 (2.5 µM for 30 min prior to addition of PMA), which inhibits cPKCs, blocked the ability of PMA to reduce AKT phosphorylation. Data are representative of at least 3 biological replicates. β-actin serves as a loading control.

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Fig. 4-19: siRNA-mediated knockdown of PKCα prevents PMA-induced hypophosphorylation of AKT

Western blot analysis of AKT phosphorylation in EC cells transfected with non-targeting (nt) or PKCα siRNAs (siRNA 1 or 2). Note the increase in steady-state levels of pAKT on PKCα knockdown. Data are representative of 3 biological replicates. β-actin serves as a loading control.

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Fig. 4-20: Knockdown of PKCα does not affect the expression of other members of the PKC family in EC cells

Western blot analysis of PKC isozymes in EC cells transfected with non-targeting (nt) siRNA or siRNA against PKCα (#1 or 2). Data are representative of 3 biological replicates. β-actin serves as a loading control.

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Fig. 4-21: The effects of PMA on PKCα and AKT activity are sustained in EC cells

A) Western blot analysis of the effects of PMA on the phosphorylation of both Thr308 and Ser473 residues of AKT in a time course PMA treatment. Despite the progressive downregulation of PKCα,

AKT phosphorylation remains suppressed over the 6 h period. Immunofluorescence staining (B) and subcellular fractionation (C) confirmed the presence of active, membrane-associated PKCα after 6 h of

PMA treatment. β-actin and AKT serve as loading controls. Arrows in (B) indicate membrane- associated PKCαlpha. In (C), Particulate indicates the membrane fraction. Data are representative of 3 biological replicates.

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Fig. 4-22: Knockdown of PKCα enhances steady-state AKT activation

PKCα was silenced in KLE, HEC-6 and SNG-M EC cells and effects on phosphorylation/activity of

AKT were determined by Western blot analysis of pAKT473 and pAKT308 and of AKT substrates

FOXO1 or PRAS40. Data are representative of 3 biological replicates. Western blot analysis performed with assistance from Michelle Lum.

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Coincident mutations in PTEN, PIK3CA and PIK3R1/2 are commonly seen in EC (Weigelt

et al., 2013). To determine if PKCα signaling can inhibit AKT activity in EC cells with coexisting

PI3K/AKT pathway alterations, PTEN was knocked down in PKCαhigh, PTEN WT HEC-1-A and

HEC-50Co cells, which harbor activating mutations in the catalytic (PIK3CA) and regulatory

(PIK3R1) subunits of PI3K, respectively (Weigelt et al., 2013). As expected, PTEN knockdown further increased PI3K/AKT signaling, as indicated by enhanced basal levels of phosphorylated

AKT (at both Ser473 and Thr308) and its downstream effector PRAS40 (Figure 4-23). However, loss of PTEN did not prevent PMA-induced hypophosphorylation of these sites, indicating that

PKCα signaling can suppress AKT even in the face of multiple activating events in the pathway.

This finding is consistent with the ability of PKCα to inhibit AKT in SNG-M and HEC-6 cells, which naturally harbor coincident mutations in PTEN and PI3K subunits. These data, combined with evidence for downregulation of the PKCα target gene Id1 (Figure 4-24), a master regulator of tumor aggressiveness (Hao et al., 2011), further support a role for PKCα as a tumor suppressor in

EC.

Since PP2A and PHLPPs have been identified as the main phosphatases that reverse AKT phosphorylation (Gao et al., 2005; Van Kanegan et al., 2005), we investigated their involvement in

PKCα-mediated AKT hypophosphorylation in EC cells. These phosphatases are expressed and functional in EC cells (Figure 4-25). A role for PHLPP1/2 in regulation of AKT in EC cells was confirmed by the ability of the PHLPP1/2 inhibitors NCS45586 and NCS117079 (Sierecki et al.,

2010) to increase basal AKT phosphorylation (Figure 4-26A). Enhanced AKT phosphorylation was similarly observed following inhibition of PP1 and PP2A with calyculin A, or PP2A alone with okadaic acid (Figure 4-26B). PMA was still able to suppress pAKT in the presence of the

PHLPP1/2 inhibitors, indicating that the effects of PKCα are independent of PHLPP1/2 (Figure 4-

26A). In contrast, both calyculin A and okadaic acid blocked PMA-induced AKT hypophosphorylation, pointing to a PP2A-dependent mechanism (Figure 4-26B). More importantly, PMA led to an induction of PP2A activity, as determined by PP2A

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immunoprecipitation phosphatase assays that was abolished by siRNA-mediated PKCα knockdown (Figure 4-27). The involvement of PP2A was further explored by siRNA-mediated

knockdown of the catalytic subunit of the phosphatase (PP2AC). PP2AC knockdown enhanced

basal AKT phosphorylation and inhibited the ability of PMA/PKCα to reduce pAKT levels (Figure

4-28), although the effect was less pronounced than seen with pharmacological inhibition, likely

reflecting the incomplete knockdown of PP2AC achieved with siRNA. Together, these results

indicate that PKCα inhibits PI3K/AKT signaling through a PHLPP-independent, PP2A-dependent mechanism. It should be noted that, although PP2A may preferentially dephosphorylate AKT on the T308 site, it can also dephosphorylate AKT on the S473 site, as described in numerous reports using a variety of biological systems (Andrabi et al., 2007; Li et al., 2003).

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Fig. 4-23: PKCα signaling can inhibit AKT activity in EC cells with hyperactivation of the

PI3K/AKT pathway due to coexisting alterations in more than one pathway component

EC cells harboring mutations in the catalytic (PIK3CA, HEC-1-A) or regulatory (PIK3R1, HEC50Co) subunits of PI3K and stably expressing non-targeting (nt) or two different PTEN (PTEN1+2) shRNA were treated with EtOH (C) or PMA (P) and subjected to Western blot analysis for the indicated proteins.

Data are representative of 3 biological replicates. β-actin serves as a loading control. Western blot analysis performed with assistance from Michelle Lum.

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Fig. 4-24: PKCα downregulates Id1 in EC cells

Western blot analysis showing downregulation of Id1, a growth/ tumor promoter, following treatment with ethanol (C) or PMA (P) in PKCαhigh but not PKCαlow cells. Data are representative of 3 biological replicates. β-actin serves as a loading control

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Fig. 4-25: Expression of PHLPPs and PP2A in EC cells

Assessment of the expression of two families of phosphatases that are known to modulate AKT phosphorylation, PHLPPs and PP2AC (the catalytic subunit of PP2A), in EC cells by qTR-PCR or

Western blot analysis. The mRNA quantification was normalized against 18s rRNA. β-actin serves as a loading control for Western blots. Data are representative of 3 biological replicates and error bars are

± SEM.

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Fig. 4-26: PP2A but not PHLPP inhibitors blocked PMA-mediated hypophosphorylation of

AKT

EC cells were pretreated with A) 50 mM PHLPP inhibitor NSC 45586 or 117079 (15 min), or B) 10 nM calyculin A (CA) (15 min) or 2.5 µM okadaic acid (OA) (30 min) prior to treatment with vehicle (C) or

100 nM PMA (P) for 6 hr. Western blot analysis demonstrated the effects of phosphatase inhibition on

AKT phosphorylation. β-actin serves as a loading control and data are representative of 3 biological replicates.

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Fig. 4-27: PMA induces PP2A activity in EC cells

Measurement of PP2A activity using a malachite green based assay in PKCα knockdown EC cells

(achieved by a combination of 2 siRNAs targeting PKCα) treated with vehicle (C) or 100 nM PMA (P) for 10 min. Non-targeting siRNA was used as a control. Results are representative of multiple biological replicates and error bars indicated mean ± SEM. *** p < 0.005, n.s. not significant. Experiment performed with assistance from Adrian Black.

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Fig. 4-28: PP2A knockdown reduces the effects of PKCα signaling on AKT phosphorylation/activity in EC cells

Immunoblot analysis of AKT phosphorylation in EC cells transfected with non-targeting (nt) siRNA or a combination of two siRNAs targeting the catalytic subunit of PP2A (siPP2AC). β-actin serves as a loading control and data are representative of 3 biological replicates.

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4.2.6 PKCα loss is associated with increased AKT activity in endometrial lesions in vivo

While universal loss of PKCα was seen in uterine lesions from PtenΔ4-5/+, PtenG129E/+ and

PtenC124R/+ mouse models, PKCα-retaining areas were detected in lesions arising in the Ptenpr-/-

mouse. This allowed comparison of AKT activity in PKCα-retaining and PKCα-deficient cells in

situ. pAKT staining was stratified as none-low, medium or high and association between pAKT

staining intensity and the presence or absence of PKCα signal was quantified in serial sections.

PKCα expression was associated with significant differences in AKT staining intensity (p <

0.0001), with PKCα negative areas predominantly exhibiting high intensity pAKT staining, and

PKCα positive areas largely coinciding with low intensity pAKT staining (Figure 4-29, compare

areas indicated with block and open arrows). This correspondence supports a role for PKCα in

suppression of PI3K/AKT signaling in endometrial tumor cells in vivo as well as in vitro.

Fig. 4-29: PKCα loss is associated with increased pAKT in endometrial lesions in vivo

Serial sections of uterine tissue from Ptenpr-/- mice immunostained for PKCα or pAKT(Ser473).

Quantification of different intensities of pAKT staining (none-low, medium, or high) in PKCα positive

and negative areas. Data are representative of 3 independent experiments. Error bars indicate SEM.

Analysis performed with assistance from Adrian Black.

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4.2.7 Inhibition of AKT activity is an important component of the PKCα tumor-suppressive axis in EC cells

The role of inhibition of PI3K/AKT signaling in the tumor suppressive effects of PKCα in

EC cells was tested using constitutively active AKT: myristoylated AKT1 (myr-AKT), AKT-DD

(T308D, S473D), and AKT-E17K (an active mutant identified in EC (Mancini et al., 2016; Rudolph et al., 2016)). These constructs were introduced into PKCαlow HEC-59, RL95-2, HEC-116,

Ishikawa, or HEC-251 EC cells by adenoviral or retroviral transduction and the ability of

exogenous PKCα to inhibit anchorage-independent growth was assessed. The amount of

constitutively active AKT introduced into EC cells was titrated to produce a 2- to 5-fold increase

in basal pAKT levels, as determined by quantification of immunoblots. In the context of strong

endogenous AKT activation in EC cells, this increase in AKT activity had only slight effects on

anchorage-independent growth on its own, with myr-AKT, AKT-DD, and AKT-E17K causing a modest increase in overall colony size (Figures 4-30, 4-31) and a small, non-statistically significant increase in colony number in some cell lines. In contrast, constitutively active AKT had a marked effect in cells transduced with PKCα, restoring both colony number and overall colony size, albeit to different extents in different cell lines (Figures 4-30, 4-31). While the inability of constitutive to fully reverse the effects of PKCα suggests that other factors may be involved, these data point to inhibition of PI3K/AKT signaling as a key mechanism mediating the tumor suppressive actions of

PKCα in the endometrium.

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Fig. 4-30: Inhibition of AKT activity is a component of the PKCα tumor-suppressive axis in EC cells

Colony formation in soft agarose of EC cells transduced with adenovirus expressing LacZ (moi = 50), myr-AKT1 (moi = 50), and/or PKCα (moi = 1) as indicated. * p < 0.05; **p < 0.01 (2-sided student’s t test). Error bars indicate SEM from 3 independent experiments. Quantification was performed with assistance from Adrian Black.

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Fig. 4-31: Mutant AKT partially rescues PKCα-mediated growth suppression

Colony formation in soft agarose of EC cells stably expressing mutant AKT generated by transduction with AKT-DD or E17K or empty retroviral vectors. Mutant AKT-expressing or control cells were subsequently transduced with Ad-LacZ or Ad-PKCα as indicated. * p < 0.05; **p < 0.01; ***p < 0.005

(2-sided student’s t test). Error bars indicate SEM from 3 independent experiments. Quantification was performed with assistance from Adrian Black and Michelle Lum.

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4.3 Discussion

The current study provides the first evidence for a tumor suppressive role of PKCα signaling in PI3K/AKT-driven EC, via a mechanism that involves PP2A-mediated inactivation of

AKT. The role of PKCα in tumorigenesis appears to be multifaceted. While the enzyme has been linked to tumor promotion in some systems, our studies add to a growing list of tumor types (e.g., colon and lung) in which PKCα signaling has tumor suppressive effects.

Our IHC and qRT-PCR analysis of human ECs revealed loss of PKCα protein and mRNA in a majority of cases. Notably, more than half of grade 1 ECs showed deficiency of the enzyme,

indicating that PKCα suppression can occur early during human endometrial tumorigenesis. This

notion is supported by our analysis of mouse models of PI3K/AKT-driven endometrial neoplasia,

which demonstrated loss of PKCα expression in pre-malignant hyperplasia regardless of the nature

of PTEN dysregulation. Additional supportive evidence is provided by previous transcriptome

analysis of early Type I ECs, which demonstrated a 3-fold downregulation of PKCα mRNA in

tumors relative to normal endometrium (Risinger et al., 2013; Saghir et al., 2010), and by IHC

analysis of a limited set of grade 1 ECs, which revealed regions of low or undetectable PKCα

protein in a subset of lesions (Haughian and Bradford, 2009).

Although loss of PKCα mRNA and protein has been noted in other tumor types (e.g., colon cancer (Verstovsek et al., 1998), non-small cell lung cancer (Hill et al., 2014), basal cell carcinoma

(Neill et al., 2003), T-cell Acute Lymphoblastic Leukemia (T-ALL) (Milani et al., 2014)), the mechanisms underlying PKCα deficiency have yet to be defined. Our qRT-PCR and IHC analysis argues that decreased mRNA expression is the major mechanism involved in disabling PKCα functions in the endometrium, although TCGA data indicate that PKCα may be inactivated by mutation in up to 5 % of ECs (Antal et al., 2015). We also show, for the first time, that PKCα loss can be mediated by transcriptional repression of the PKCα gene. Initial mechanistic analysis excluded methylation of the PKCα promoter in EC (data not shown) or differential promoter activity in cells with high or low levels of PKCα mRNA/protein. Recent analysis of the regulatory

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region of PKCα found differential methylation of the CpG shores, defined as 2kb up- or downstream of the CpG island, with hypermethylation of the right CpG shores in cells with reduced

PKCα mRNA (Figure 4-32A). In line with this new finding, we have observed increased PKCα expression when EC cells are treated with decitabine, a DNA hypomethylating agent (Figure 4-

32B). Experiments are underway to confirm these results in additional cells lines.

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RKO Caco2 Lovo DLD-1 HCT-116 RL-95 Ishikawa HEC-6 SW480 HEC-116 HEC-251 KLE HEC-1A HCEC SNG-M SW620 HEC-1B SW48 HCT-15 CT26 HT29 HEC-50Co IEC-18 TENN HEC-59 Colo205 CBS FET B

Decitabine Decitabine µM 0 0.01 0.05 0.1 0.5 1 0 0.01 0.05 0.1 0.5 1

PKCα Ishikawa RL95-2

Fig. 4-32: Methylation of PKCα CpG shores in cancer cells

A) Methylation analysis showed hypermethylation in the right CpG shore in PKCαlow cell lines, as indicated in red, whereas the same region is generally hypomethylated in cell lines with high PKCα expression. B) Increased PKCα levels in PKCαlow cells (Ishikawa) treated with decitabine. Methylation analysis was performed with assistance from Xinyue Li, Chun-wei Chen, and Dr. Tim Huang.

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The precise correspondence between PKCα loss, PTEN deficiency, and AKT activation in murine endometrial lesions suggested that transcriptional suppression of PKCα could be a direct result of hyperactivation of the PI3K/AKT pathway, at least in some ECs. However, modulation of

PTEN expression or AKT activity in human EC cells excluded this possibility, indicating that

PKCα downregulation is not merely a passenger effect of aberrant PI3K/AKT signaling but, instead, is an alteration that is selected for because of specific contributions to endometrial tumor development. The possibility that PKCα deficiency drives the loss of PTEN was also tested and excluded in PKCα knockdown experiments. Anchorage-independent colony formation assays provided direct evidence for a tumor suppressive function of PKCα in human endometrial cells: restoration of PKCα expression/activity profoundly inhibited colony formation of human EC cells in soft agarose, while PKCα knockdown enhanced the transformed properties of EC cells that retained expression of the enzyme. The importance of early loss of PKCα was highlighted by the marked acceleration in uterine hyperplastic transformation observed in PKCα deficient PtenΔ4-5/+

mice, a finding that pointed to abrogation of PKCα signaling as a rate limiting step for tumor

initiation in the endometrium. In advanced cases of human EC, the proportion of PKCα negative

tumors increased to >75%, suggesting that PKCα deficiency defines a subset of ECs with high risk

of progression, a notion that is consistent with TCPA data pointing to an association between

reduced PKCα expression and decreased survival of EC patients. The finding that PKCα deficiency

is associated with a 3-fold increase in risk of high grade disease is consistent with previous reports

in other tumor types. Genetic deletion of PKCα led to increased tumor burden, adenoma-to-

carcinoma progression, and reduced survival in the APCMin/+ mouse model of intestinal neoplasia

(Oster and Leitges, 2006) and in KRas-mediated models of lung tumorigenesis (Hill et al., 2014).

Low PKCα expression identified a new subgroup of childhood T-ALL with an extremely poor outcome (Milani et al., 2014) and appears to enhance the tumorigenic potential of Gli1 in basal cell carcinoma (Neill et al., 2003). It will be interesting to determine if common mechanisms mediate

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PKCα loss in these tumor types.

A majority of ECs harbor coexisting alterations in two or more PI3K/AKT pathway components, with additive effects on AKT activation that are crucial for EC development (Chen et al., 2006; Oda et al., 2005; Weigelt et al., 2013). Remarkably, knockdown of PKCα in cells with genetic hyperactivation of the PI3K/AKT pathway further enhanced the activity of AKT, highlighting the ability of PKCα deficiency to contribute to the strength of PI3K/AKT signaling in

EC cells. The demonstration that PKCα negative uterine regions in Ptenpr-/- mice are associated with significantly higher levels of pAKT than PKCα-retaining regions indicates that PKCα loss also contributes to PI3K/AKT hyperactivation during endometrial tumorigenesis in vivo. PKCα activity may also restrain AKT in the normal endometrium, since increased expression and membrane association of the enzyme coincide with the reported downregulation of AKT activity during the proestrus-to-estrus transition in rats (Dery et al., 2003). These findings, together with the demonstration that agonist-induced PKCα activation suppresses AKT in EC cells, identify

PKCα as a potent inhibitor of PI3K/AKT signaling in the endometrium, capable of disabling the

AKT oncoprotein even in the context of multiple activating mutations in upstream signaling intermediates (e.g., PTEN and PIK3CA in HEC-251 cells; PTEN, PIK3CA and KRAS in SNG-M cells).

Inactivation of AKT appears to be a key component of the tumor suppressive actions of

PKCα in endometrial cells, since constitutively active AKT [myr-AKT, AKTDD (T308D,

S473D), and AKT(E17K)] markedly diminished the inhibitory effects of PKCα on EC cell colony formation in soft agar. Together, our findings support the idea that disruption of PKCα signaling may be critical for achieving the robust PI3K/AKT pathway activation that is required to drive EC development and progression (Figure 4-8D). To demonstrate the potential clinical impact of PKCα

loss in EC, future experiments will investigate whether PKCα deficiency will enhance the

sensitivity of EC tumors to AKT inhibitors or other targeted agents. The efficacy of agents that are

currently under development or in clinical trials is largely limited by their dose-dependent toxicity

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(Manning and Toker, 2017). As cells with PIK3CA mutations exhibited increased sensitivity to

agents targeting the PI3K/AKT pathway (e.g. PI3K inhibitors, AKT inhibitors, and mTOR

inhibitors), the loss of PKCα-mediated negative regulation of AKT activation may similarly

enhance the sensitivity of these cells to PI3K/AKT pathway inhibition. Thus, PKCα deficiency

may provide an additional layer of patient stratification, indicative of therapeutic responsiveness to

PI3K/AKT targeting agents in clinical trials.

Crosstalk between PKCα and the PI3K/AKT pathway has been noted in other systems.

PKCα can inactivate upstream modulators of PI3K such as growth factor receptors and scaffolding

proteins (Koese et al., 2013; Oriente et al., 2005) and can dampen PI3K activity directly (Hoshino et al., 2012; Sipeki et al., 2006). Here, we demonstrate that PKCα activates the serine/threonine phosphatase PP2A in EC cells and inactivates AKT via a PP2A-dependent mechanism. PKCα- induced AKT inactivation may involve direct effects of PP2A on activating phosphorylation at

T308 and S473 (Manning and Toker, 2017), although potential effects on upstream regulators of

AKT activity remain to be formally excluded. A role of PP2A in the antitumor effects of PKCα in

the endometrium is consistent with known functions of the phosphatase as a bona fide tumor

suppressor, with key activities in maintenance of cellular homeostasis (Perrotti and Neviani, 2013).

Future studies will explore the specific PP2A trimeric holoenzyme complex(es) responsible for

mediating PKCα-induced inactivation of AKT.

Together, our findings highlight disruption of a PKCα→PP2A AKT signaling module as

a key step in the development of EC, ensuring robust, unopposed AKT⟞ activation and promoting

transformation of endometrial cells (Figure 4-8D). These studies provide a foundation for exploring the prognostic value of PKCα expression in EC and the potential of harnessing the PKCα→PP2A

module for therapeutic benefit in a subset of ECs.

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Chapter 5: Conclusion

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PKCα is ubiquitously expressed in many tissues and regulates the physiology of a cell by relaying signals to a wide molecular network. Understanding the functional role of PKCα in maintenance of tissue homeostasis and its involvement in the carcinogenesis process in different systems has been a long-standing interest of our laboratory. Past work from our lab, focusing on the intestinal system, found PKCα signaling to be growth inhibitory and tumor suppressive. In this study, we examined the expression and activation of PKCα in the normal endometrium. Previous studies have shown that PKCα mediates growth, invasion and survival response to DNA damage in endometrial cancer cells (Haughian and Bradford, 2009; Haughian et al., 2006; Haughian et al.,

2009). However, its impact on oncogenic PI3K signaling, the driving force of the majority of endometrial tumors, remained elusive. Therefore, as part of our effort to understand the functional role of PKCα in cancer, we investigated how signaling mediated by this kinase impacts endometrial carcinogenesis driven by hyperactive PI3K/AKT signaling.

5.1 PKCα as a potential anti-proliferative regulator in the endometrium

The endometrium undergoes cyclic remodeling, in which the stroma and epithelium proliferate and regress/shed in response to hormonal fluctuation. Aberrations in this tightly regulated process disrupt the homeostasis of the endometrium and, eventually, results in uncontrolled growth and tumor formation. PKCα is among the several isozymes that have been reported to be expressed in the uterus (Jofre et al., 2013; Kim et al., 1999). However, little is known of its specific functions in this tissue. Unlike other isozymes, which have been shown to induce uterine contraction, activation of PKCα was observed in preterm uterus when the myometrium is quiescent and PKCα has been shown to inhibit myometrium contractility by regulating the expression of prostaglandin and urocortin, in order to maintain pregnancy (Bamberger et al., 2007;

Jofre et al., 2013; Wouters et al., 2014). In the epithelium, PKC has been implicated in promoting protein synthesis through ERK-mTOR in response to estrogenic stimulation (Wang et al., 2015).

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PKC activation was also shown to coincide with the switch to the secretory phase that is predominantly regulated by P4 (Chen et al., 2013). Based on these studies, it is not clear how PKCα

is involved is regulating E2/P4 responses and epithelial remodeling in the endometrium or which

PKC isozyme(s) is the major regulator of these events.

Current understanding indicates that PKC functions are dependent on cellular context and that individual isozymes can have overlapping or opposing actions (Garg et al., 2014). To decipher the role of PKCα in cancer, it is crucial to first understand its involvement in the normal physiology in the tissue of interest. In our study, we observed PKCα expression in the stromal and epithelial compartments of the uterus, though expression was higher in the stroma in both human and murine uterine tissues (Figure 3-1). In the human endometrium, PKCα activation, as indicated by its membrane association, was observed in the secretory phase. In mouse, there was regional membrane-associated PKCα in the epithelium in proestrus, which became more prevalent in the subsequent estrous phase (Figure 3-3). This increase in PKCα activation negatively correlated with proliferation, as shown by the reduced Ki67 signal in estrus compared with other phases (Figure 3-

2, 3-4). Interestingly, this timing coincided with a transition from massive proliferation to a state primed for embryo implantation. Together, our data linked the expression and activation of PKCα to hormone-driven cycling of the tissue, suggesting that PKCα signaling may trigger an anti- proliferative program that is involved in endometrial remodeling, implantation, and maintenance of pregnancy. Our IHC analysis also revealed a regional staining pattern for PKCα, suggesting that the endometrial epithelium might be heterogeneous, exhibiting differential responses or temporal discrepancies in their response to hormone fluctuation (Figure 3-3, 3-4). To further explore the role of PKCα in the endometrium, mechanistic studies focusing on how E2 and P4 influence PKCα

signaling (and other PKCs) may reveal the role of this kinase in regulating endometrium

regeneration, thus painting a more complete picture of how these enzymes function cooperatively

to regulate uterine biology.

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5.2 PKCα suppresses endometrial carcinogenesis

The question of whether PKCs promote or suppress cancer has been a subject of long- lasting debate in the field. Historically, PKCs are mostly recognized for their association with the carcinogenic phorbol esters, and their ability to promote tumor formation that encouraged the development of PKC inhibitors as cancer therapeutics of which had disappointing results in clinic trials (Garg et al., 2014; Mochly-Rosen et al., 2012). Subsequent studies revealed that these enzymes possess tumor suppressive ability, an observation that is bolstered by the recent survey of cancer-associated PKC mutations, which found these mutations to cause loss of kinase function

(Antal et al., 2015; Hill et al., 2014; Newton and Brognard, 2017; Oster and Leitges, 2006).

Previous work from our laboratory has shown that PKCα inhibits intestinal tumor formation through downregulation of cyclin D1 and Id1 and increased expression of p21 (Frey et al., 2000;

Hao et al., 2011; Pysz et al., 2009). In this study, we demonstrated that PKCα acts as a tumor suppressor in the endometrium by negatively regulating AKT activation. Together, our work further supports the current paradigm shift in the function of PKCs in cancer.

Our analysis and TCGA data both point to reduced PKCα expression in ECs due to

alterations in its gene transcription (Figure 4-1 – 4-3). Although loss of PKCα protein and mRNA

has been reported in other cancer types, we provided the first evidence that transcriptional

repression is responsible for its absence in cancer (Figure 4-4). Furthermore, loss of PKCα

correlated with poorer prognosis as it was associated with more aggressive disease and reduced

patient survival probability (Figure 4-5, 4-6). In multiple mouse EC models carrying mutant PTEN or generated by allelic PTEN deletion, PKCα loss was uniformly observed in lesions arising in the endometrium (Figure 4-7, 4-9). However, it is unlikely that loss of PTEN led directly to reduced

PKCα transcription as we did not observe changes in PKCα levels in EC cells with PTEN knockdown or overexpression, or with inhibition of PI3K/AKT signaling (Figure 4-11).

Conversely, manipulation of PKCα expression did not affect PTEN levels, indicating that PKCα deficiency itself is unlikely to drive PTEN LOH to give rise to endometrial lesions. However, the

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specific mechanisms that cause the reduction in PKCα transcription remain elusive. Although our initial analysis has excluded methylation at the PKCα promoter as a means of regulating its expression (data not shown), future experiments will explore other possible mechanisms such as differential binding of regulatory elements (e.g., transcription factors, co-activators, or corepressors) or methylation of other regulatory regions (e.g., CpG shores).

Our study finds that PKCα acts as a tumor suppressor and the reduced expression of this kinase diminishes protection against endometrial transformation. In mice carrying loss-of-function

Pten mutations, concurrent PKCα deficiency increased tumor burden by 3-fold at 1- and 3-months, although the differences diminished at later times of disease progression (Figure 4-13). This suggests that PKCα acts as a “gatekeeper” and loss of this kinase is likely a rate-limiting event in endometrial tumorigenesis. Indeed, re-introduction of PKCα markedly inhibits EC cell growth under anchorage-independent conditions (Figure 4-14 ), while PKCα knockdown has the opposite effect. Interestingly, kinase-dead PKCα had no effects, demonstrating that the tumor suppressive effects of PKCα in EC are indeed due to its kinase function (Figure 4-15). Data from this study further demonstrated that PKCα activation stimulates PP2A activity and leads to hypophosphorylation of AKT and its downstream targets PRAS40 and FOXO1 (Figure. 4-17, 4-

22). This negative regulation of AKT can be abolished by inhibitors of PKCα or PP2A or by

PKCα/PP2A RNA interference (Figure 4-18, 4-22, 4-26, 4-28). However, the co-expression of constitutively active AKT (myr-AKT or AKT-DD) or hyperactive AKT found in breast and endometrial tumors (AKT-E17K) only partially rescues the growth inhibitory/tumor suppressive effects of PKCα in EC cells (Figure 4-30, 4-31)), suggesting that PKC can suppress EC tumorigenesis through other mechanisms in addition to negative regulation of AKT activity. PKCα has been shown to downregulate the expression of Id1 family proteins (Figure 4-10, 4-24), which promote cell cycle progression, tumor cell growth and migration, through ERK-dependent signaling (Hao et al., 2011). Our analysis also demonstrated an inverse correlation between PKCα and Id1 in EC tumors, presenting a possible additional mechanism for PKCα-mediated suppression

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of EC. However, in-depth mechanistic studies are needed to confirm this hypothesis and also to explore other pathways by which PKCα functions as a tumor suppressor in EC.

The present study also showed that PP2A but not PHLPP is involved in PKCα-mediated hypophosphorylation of AKT (Figure 4-26). In contrast to previous studies showing a residue- specific dephosphorylation of AKT by PP2A and PHLPP on threonine 308 and serine 473, respectively, PP2A was responsible for PKCα-mediated dephosphorylation of AKT at both of these sites (Figure 4-26, 4-28). The specificity of the PP2A holoenzyme is determined by the B, or regulatory, subunits in the complex (Chen et al., 2013; Eichhorn et al., 2009; Sents et al., 2013).

Although our data demonstrated that PKCα activation induces PP2A activity (Figure 4-27), we do

not know the specific makeup of the holoenzyme involved in the effect. To determine how PKCα

mediates PP2A activation, future studies will examine the formation of PP2A holoenzymes

following PKCα activation, and how the individual holoenzyme(s) accounts for the

dephosphorylation of AKT.

In comparison with other more widely studied cancer types, EC patients have a relatively

optimistic prognosis as 1) many EC cases are symptomatic and detected at an early stage, and 2)

the disease is usually contained in the uterus at the time of diagnosis, thus curative through

hysterectomy (Engelsen et al., 2009). However, approximately 20% of EC patients experience

recurrent disease that is resistant to radiation or currently available chemotherapeutics. Recurrent

disease is usually aggressive and accounts for a majority of EC-related mortality (Engelsen et al.,

2009). Molecular analysis of EC tumors has found most ECs are driven by hyperactivation of

PI3K/AKT signaling, most commonly due to loss of PTEN, activating mutations in PI3K or AKT,

or the combination of multiple genetic aberrations in this pathway (Dedes et al., 2011). As PTEN

loss and AKT hyperactivation are prevalent in many cancers, small molecule inhibitors of AKT

have received much attention as an emerging class of targeted therapeutic against ECs (Manning

and Toker, 2017). However, results from clinical trials thus far have been disappointing as the

efficacy of these agents is limited by their accompanying dose-limiting toxicity (Ma et al., 2016;

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Saura et al., 2017). Apart from developing new agents that are less toxic, a system that can stratify patients who are more sensitive to AKT inhibition, and thus more likely to respond at a lower dosage, would also be beneficial to the clinical outcome of these trials (Manning and Toker, 2017).

From our study of the role of PKCα in endometrial tumorigenesis, we have shown that PKCα acts as a negative regulator of AKT activation (Figure 4-17). While PTEN loss and oncogenic PI3K mutation indeed result in elevated phosphorylated AKT, PKCα signaling can still exert negative pressure on the kinase (Figure 4-23). Therefore, concurrent loss of PKCα leads to robust, unopposed AKT activation and more aggressive EC phenotypes (as depicted in our model in Figure

5-1). Our data strongly supports a PKCα-PP2A-mediated mechanism that acts directly on AKT; however, the possibility that crosstalk exists between PKCα-PP2A and other components of PI3K signaling (e.g. upstream receptor tyrosine kinases or PTEN) should also be considered.

Additionally, future experiments examining whether PKCα status affects the sensitivity of EC cells to currently available AKT inhibitors could improve the clinical benefits of these inhibitors. Based on our findings, PKCα is able to suppress AKT phosphorylation regardless of the status of PTEN or PI3K, suggesting it is a potent inhibitor of AKT activation. It is likely that EC cells that have

PKCα and, by extension, retain part of the regulation of AKT activation, could be more sensitive to small molecule AKT inhibitors, allowing them to inhibit tumor growth with limited, better tolerated side effects.

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Fig. 5-1: Schematic representation of PI3K/PTEN and PKCα as regulators of AKT activation endometrial cancer.

The model depicts that in addition to the well-known PI3K/PTEN signaling pathway, PKCα-PP2A mediates another regulatory axis that maintains balanced activation of AKT in EC cells. Mutations of

PI3K and PTEN have been shown to result in hyperactivation of AKT. However, our study demonstrated that disruption of the PKCα-PP2A regulatory axis ensures robust, unopposed AKT activation and promotes the transformation of endometrial cells.

154

Appendix A: Estrogenic stimulation of PKCα expression

Rationale:

Based on our examination of normal mouse endometrium under estrous cycle, PKCα

expression increased during later proestrous and was robustly expressed during estrous,

coinciding with the peak of estrogen level.

Results/Discussion:

To yesy whether estrogenic stimulation induces PKCα expression, Ishikawa, an EC cell line that retained ER expression, was treated with estradiol (E2). Both PKCα mRNA and protein were examined at 3 and 24 hours following E2 stimulation. Cells were maintained in charcoal-

cleared culture medium and experiments were performed with assistance from the laboratory of

Dr. Vimla Band.

Results from our analysis showed that E2 robustly increased PKCα mRNA expression

after 24 hours but no change in protein level was observed. The induction of cyclin D1, a well-

known E2-responsive target, following E2 treatment demonstrated that the cells were indeed responsive to the stimulation. Our finding is consistent with previous report showing that estrogenic stimulation induced PKCα expression in primary endometrial cancer cells (Wu et al.,

2005). However, failure to translate PKCα mRNA to protein indicated possible translational

defect in Ishikawa, a human EC cell line, which might account for the decreased PKCα

expression in these cells.

155

Fig. A.1: PKCα expression following E2 stimulation of EC cells.

The graph showed PKCα mRNA levels at 3 and 24 hours after stimulation with E2 or vehicle control

(Ethanol) measure by qRT-PCR analysis relative to 18sRNA. Shown in bottom panel is Western Blot analysis of PKCα protein levels following E2 stimulation. Cyclin D1, a well-known E2 target gene, was assessed as demonstration of E2 effect.

156

Appendix B: Regulation of PKCα Expression

Rationale:

Parallel analysis of PKCα mRNA and protein from human endometrial tumors, mouse

endometrial lesions, and endometrial cancer cell lines revealed that reduced PKCα mRNA

correlated with lower protein levels. We explored possible mechanisms that might be involved in

regulating PKCα expression in EC cells including differential promoter activity, mRNA stability,

and protein stability.

Results/Discussion:

To explore mechanism that might be involved in regulating PKCα expression in EC cells, we assessed promoter activity, mRNA stability, and protein stability of PKCα using methods described below:

1) PKCα promoter activity assay

Plasmids encoding firefly luciferase regulated by PKCα promoters (-1571/+277, -

1571/+77, -331/+77, -279/+77, -260/+77) were transfected into EC cells using Fugene 6 or

XtremeGENE 9 as per manufacturer’s protocol. Plasmids encoding renilla luciferase regulated

by thymidine kinase promoter was transfected in all samples and served as normalization for

transfection efficiency PKCα promoters within a cell line. Promoter activity was expressed at

relative to the minimal PKCα promoter (-260/+77)

2) mRNA stability

To examine PKCα mRNA stability, EC cells were treated with α-amanitin, a selective

RNA polymerase II and III inhibitor, or vehicle control (ethanol) for 12 hours. PKCα mRNA

levels were assessed by qRT-PCR and normalized to the vehicle control.

3) Protein stability

Protein stability was examined by a time course (6 hour interval in 24 hours) treatment

157

with cycloheximide, which inhibits protein synthesis or the vehicle control (DMSO). PKCα protein levels were assessed by Western Blotting analysis.

Results from these experiments did not present a general mechanism that regulates the broad alteration of PKCα expression in these cell lines. The promoter analysis showed possible repressive element binding to the PKCα promoters in 4 out of the 7 EC cell lines (SK-UT-1B,

Ishikawa, AN3CA, KLE) examined, as demonstrated by the decreasing luciferase activity with increase PKCα promoter length (Figure B.1.). In three EC cell lines (SN3CA, Ishikawa, SK-UT-

1B), there was reduced PKCα mRNA when mRNA synthesis was inhibited by α-amanitin (Figure

B.2.). Interestingly, despite the long half-life of PKCα protein, it was more readily degraded in

AN3CA than other PKCαlow cells (e.g. SK-UT-1B) (Figure B.3.). Together, these results demonstrated that multiple mechanisms may be involved in the regulating PKCα expression in

EC cells. However, the mechanisms explored here did not explain the broad PKCα loss in EC cells or tumors.

158

Fig. B.1: Figure PKCα promoter activity in human EC cells.

Plasmids of PKCα promoter-regulated firefly luciferase plasmids were used to measure promoter activity. All expressions were represented as relative to the minimal promoter (-260/+77) of each cell lines and normalized to renilla luciferase expression to account for transfection efficiency. Error bars indicate mean ± SEM from 3 biological replications.

159

Fig. B.2: PKCα mRNA levels under RNA polymerase inhibition.

PKCα mRNA was measured using qRT-PCR after 12 hours of treatment with the a-amanitin, a RNA polymerase inhibitor or vehicle control (ethanol), and quantified relative to 18rRNA. Error bars indicate mean ± SEM from 3 biological replications

160

Fig. B.3: PKCα levels under inhibition of protein synthesis.

EC cells treated with cycloheximide to inhibit protein synthesis or vehicle control (DMSO) in a time course. PKCα protein levels were analyzed by Western Blotting.

161

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