Signaling Effectors Required for G Protein-Coupled Estrogen Receptor-, GPER, Induced Events Associated with Breast Cancer Progression

Hilary Thompson Magruder

B.A. Wheaton College, 2009

A dissertation submitted in partial fulfillment of the requirements for the Degree of Doctor of Philosophy in the Division of Biology and Medicine at Brown University.

May, 2014

© Copyright 2014 by Hilary Thompson Magruder This presentation by Hilary Thompson Magruder is accepted in its present form by the Division of Biology and Medicine as satisfying the dissertation requirements for the degree of Doctor of Philosophy

Date______

Edward J. Filardo, Ph.D., Director

Recommended to the Graduate Council

Date______

Richard Freiman, Ph.D., Reader

Date______

Jonathan Reichner, Ph.D., Reader

Date______

Wentian Yang, Ph.D., Reader

Date______

Eric Prossnitz, Ph.D., Outside Reader

Approved by the Graduate Council

Date______

Peter M. Weber, Ph.D.

Dean of the Graduate School

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Hilary T. Magruder

29 High Hawk Road Portsmouth, RI 02871

(401) 662-9588 [email protected]

Education

Brown University (Providence, Rhode Island) Anticipated 2014 Ph.D. Candidate in Pathobiology under the Division of Biology and Medicine  Thesis: “Signaling Effectors Required for G Protein-Coupled Estrogen Receptor-, GPER, Induced Events Associated with Breast Cancer Progression” in Dr. Edward Filardo’s laboratory  Mentored Brown University undergraduate students in Dr. Jonathan Reichner’s laboratory  Teaching Assistant for a virology course  Nominated for the June Rockwell Levy Predoctoral Fellowship in the Fall of 2009

Brown University (Providence, Rhode Island) Anticipated 2014 M.A. in Pathobiology under the Division of Biology and Medicine

Wheaton College (Norton, Massachusetts) 2005 – 2009 B.A. in Biochemistry & Psychology (Double Major)  Cumulative GPA- 3.75; Biochemistry GPA- 3.77  Honors: The Villar’s Prize in Science, Julia R. Lange Fellowship, Phi Beta Kappa Honors Society, Psi Chi Honors Society, Tri Beta Biological Honors Society, Biochemistry Departmental Honors, Dean’s List, National Scholars Honors Society  Biochemistry Honors Thesis on GPR30 in ovarian cancer under the supervision of Dr. Edward Filardo from Rhode Island Hospital/Brown University and Dr. Elita Pastra-Landis from Wheaton College

Publications, Citations, and Abstracts

Magruder H, Quinn JA, Schwartzbauer JE, Reichner J, Huang A, Filardo EJ. The G-protein- coupled estrogen receptor, GPER-1, promotes fibrillogenesis via a Shc-dependent pathway resulting in anchorage-independent growth. Hormones and Cancer (in preparation).

Magruder H, Reichner J, Huang A, Filardo EJ. PTPN12 inhibits estrogen action via the G- protein-coupled receptor, GPR30/GPER-1. (in preparation).

Magruder H, Filardo EJ. Epidermal growth factor receptor transactivation and fibronectin matrix assembly by the G-protein coupled receptor, GPER, requires a transmembrane signaling complex consisting of PTPN12, integrin α5β1, and MMP-3. Abstract presented at the Experimental Biology Annual Meeting; April 2012, San Diego, California.

Dewan S, Magruder H. Re-treatment of vestibular schwannomas with gamma knife radiosurgery. Abstract Presented at The 14th International Meeting of the Leksell Gamma Knife Society; May 2008, Quebec, Canada.

McCormack E, Magruder H, Steinhoff MM, Gass J, Legare ED, Wiggins DL, Tejada-Berges T, Sikov W, Strenger R, Dizon DS. Clinicopathic analysis of tubular carcinoma of the breast: the experience from Women & Infants’ Hospital of Rhode Island. American Journal of Clinical Oncology, 2007; 30: 454-455.

Bonzagni A, Magruder H, Benoit JM. Mercury uptake from fish fertilizer by spinach plants. Eighth Annual Northeast Student Chemistry Research Conference (NSCRC) 2006; Cambridge, MA.

Work Experience and Internships

Wheaton College 2013 - Present  Laboratory Instructor of a general chemistry course and an organic chemistry course 2008 – 2009  Instructed a general and inorganic chemistry weekly review and graded papers under the supervision of Professor Matthew Evans 2007  Attempted to create an environmentally friendly stereoisomer of an amino acid through hydrogenation reactions under the supervision of Professor Christopher Kalberg 2006 – 2007  Teaching fellow in general and inorganic chemistry under the supervision of Professor Jani Benoit that included tutoring students and grading examinations 2006 – 2007  Kollett Center tutor for general and inorganic chemistry students

Memorial Hospital of Rhode Island 2008-2009

 Conducted a clinical research project on patient satisfaction in labor and delivery, and disparities in access to healthcare

American Cancer Society Fuller Fellowship 2007  Studied GPER at Alpert Medical School of Brown University/Rhode Island Hospital under the supervision of Dr. Edward Filardo

New England Gamma Knife Center 2007  Investigated vestibular schwannomas at Alpert Medical School of Brown University/Rhode Island Hospital under the supervision of Dr. Georg Noren  Research published in the Journal of Neurosurgery

NeuroHealth 2007  Explored the correlation between lack of smell and apathy in Parkinson’s patients under the supervision of Dr. Joseph Friedman and Dr. Megan Spencer

Merck Scholar Internship 2006  Investigated mercury levels in human hair and in leaves around the Wheaton College Vernal Pool under the supervision of Professor Jani Benoit

Women & Infants Hospital- Program in Women’s Oncology 2006 – 2007  Chaired and managed the Patient Advocate Fundraiser, which raised over $60,000  Structured and conducted a clinical research project on tubular carcinomas of the breast  Published an abstract in the American Journal of Clinical Oncology

Research Presentations

Experimental Biology Annual Meeting 2012 in San Diego 2012  Presented Epidermal growth factor receptor transactivation and fibronectin matrix assembly by the G-protein coupled receptor, GPER, requires a transmembrane signaling complex consisting of PTPN12, integrin α5β1, and MMP-3

Pathobiology Program Retreat 2012  Presented Epidermal growth factor receptor transactivation and fibronectin matrix assembly by the G-protein coupled receptor, GPER, requires a transmembrane signaling complex consisting of PTPN12, integrin α5β1, and MMP-3

19th Annual Hospital Research Celebration at Rhode Island Hospital 2011

 Presented Transmembrane signaling effectors that regulate GPER-mediated fibrillogeness and EGFR transactivation

Department of Medicine’s 17th Annual Research Forum- Brown University 2011  Presented Transmembrane signaling effectors that regulate GPER-mediated EGFR transactivation

Brown University Pathology II Course 2011  Presented Extracellular Matrix: Integrins and Matrix Metalloproteinases in Cancer

Brown University Virology Course 2011  Presented Herpesviridae

Pathobiology Program Retreat 2011  Presented Transmembrane signaling effectors that regulate GPER-mediated EGFR transactivation

18th Annual Hospital Research Celebration at Rhode Island Hospital 2010  Presented Role of GPR30 in breast tumor cell survival and its influence on mammary stromal fibroblast transformation

Pathobiology Program Retreat 2010  Presented Role of GPR30 in breast tumor cell survival and its influence on mammary stromal fibroblast transformation

Fuller and Stone Fellowship Reception 2007  Presented Expression of GPR30, a novel membrane estrogen receptor, in human breast and ovarian cancer

Wheaton College Academic Festival XVI 2007  Nominated and accepted to present Mercury Deposition to the Wheaton College Vernal Pool via Leaf Litter Fall

Wheaton College Research Symposium 2007  Presented Mercury Deposition to the Wheaton College Vernal Pool via Leaf Litter Fall

Achievements and Honors

June Rockwell Levy Predoctoral Fellowship Nominee 2009

The Villar’s Prize in Science 2009

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Julia R. Lange Fellowship 2009

Academic Festival XVIII Nominee and Participant at Wheaton College 2009

Outstanding Service as a Partner in Philanthropy Awarded by the 2007 Association of Fundraising Professionals

American Cancer Society Fuller Fellow 2007

Academic Festival XVI Nominee and Participant at Wheaton College 2007

Merck Scholar 2006

Wheaton Fellows Scholarship 2006 – 2007

Dean’s List Fall 2005 – 2009

Memberships

American Society for Pharmacology and Experimental Therapeutics 2011 - Present

Phi Beta Kappa Honors Society 2008 - Present

Psi Chi Honors Society in Psychology 2008 - Present

Pre-Health Society - President 2008 – 2009

Tri-Beta Biological Honors Society, Chi Nu – Vice President 2008 – 2009

American Cancer Society’s Fuller Fellow 2007 – 2008

National Scholars Honors Society 2007 - Present

Wheaton Fellow 2006 – 2007

Patient Advocate Fundraising Committee - Chair, Project Manager 2006 – 2007

Public Vaccination Council 2006

Pre-Health Society - Secretary and Events Coordinator 2006 – 2008

Biomedical Science Careers Program at Harvard 2006 - Present

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Program in Women’s Oncology Chemotherapy Committee 2006

Society of Surgical Oncology Application Committee 2006

Breast Fellowship Committee 2006

Certifications

NIH Clinical Research Training Course Certificate 2013  NIH Office of Clinical Research Training and Medical Education

Sheridan Teaching Certificate I 2010  Sheridan Center at Brown University

Training in Responsible Conduct in Research Certificate 2009  Brown University

Initiative to Maximize Student Development Certificate of Completion 2009  Brown University

Preface

The work presented in this Ph.D. thesis was conducted in the laboratories of Dr. Edward

Filardo and Dr. Jonathan Reichner. I have executed the work for all of the experiments presented herein with the following exceptions:

In Chapter 2, SKBR3 Shc Y315F, SKBR3Δ154, and 4T1Δ154 cells were generated previously in the Filardo lab. Experiments for Figures 1, 6, and 7 were performed by Dr.

Jeffrey Quinn in the Filardo lab.

In Chapter 3, phosphatase-dead PTPN12 cDNA was generated by the Westbrook lab at

Baylor College of Medicine.

In Chapter 4, experiments for Figures 3 and 5 were performed by Dr. Jeffrey Quinn in the Filardo lab. The proposal for Aims 3 and 4 were written with the assistance of Dr.

Edward Filardo for a NIH F31.

Acknowledgements

I would first like to thank my thesis adviser, Dr. Edward Filardo, for his support during my time at Wheaton College and Brown University. I have had the unique opportunity to work with Ed through multiple stages of my life. He has always been very attentive to me, but at the same time, he has helped me develop independence and prepare for my future. I am forever grateful for his personal and professional guidance.

I would also like to thank Dr. Jonathan Reichner for taking me into his laboratory following Dr. Edward Filardo’s departure from Brown University. Jonathan has been very supportive of my work. I cannot thank him enough for his guidance during the final two years of my graduate education. In addition, the members of Jonathan’s lab have been very helpful in guiding me through the thesis process. I would like to specifically thank

Dr. Xian O’Brien and Dr. Craig Lefort. Dr. Xian O’Brien answered my numerous troubleshooting questions and supported me through my thesis preparation and job search, and Dr. Craig Lefort provided me with valuable advice on the extracellular matrix.

I would also like to thank my committee members, Dr. Richard Freiman, Dr. Wentian

Yang, and Dr. Eric Prossnitz. They have all been a great support to me and have provided constructive guidance for my work.

I would also like to acknowledge the past members of the Filardo lab, Dr. Carl Graeber,

Dr. Shi-Bin Cheng, and Dr. Jeffrey Quinn. Carl, Shi-Bin, and Jeffrey taught me valuable laboratory techniques. It was a pleasure to work with them.

On a personal note, I would like to thank my family, Lauren Magruder (Mom), Andy

Magruder (Dad), Abigale Magruder (Sister), and Diane Thompson (Aunt). I cannot thank my parents enough for giving me the support needed to reach my goals. They have always stressed the importance of education, and I wouldn’t be where I am today without this. There aren’t enough words in the world to express my appreciation. My sister has always been by my side during difficult times, and I cannot thank her enough for this. She is the best sister one could ask for. My aunt has always been one of my biggest supporters. She has always believed in me and was the one to introduce me to the field of oncology. I love you all very much.

I would like to thank my fiancé, Chad Gaudet, for supporting me through the last few months of thesis writing and for listening to me incessantly talk about my thesis. His positive attitude has been a huge factor in surviving the last few months of graduate school. I love you, and I can’t wait to marry you this summer.

I would also like to thank all of my friends. They have been the biggest supporters in every aspect of my life. I would like to specifically thank Jennifer Ribeiro, a Pathobiology classmate. She has been there for me through every step of graduate school.

Finally, I am grateful to the Pathobiology Program, the Department of Surgery at Rhode

Island Hospital, and the Department of Medicine at Rhode Island Hospital for their financial support. I would like to specifically thank Patty and Tracey in the Department of Surgery, and Michelle Curry and Tami Mildner in the Pathobiology Program.

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Table of Contents

Title…………………………………………………………………………………………………………………………………i Copyright………………………………………………………………………………………………………………………..ii Signature Page……………………………………………………………………………………………………………….iii Curriculum Vitae……………………………………………………………………………………………………..…iv-ix Preface……………………………………………………………………………………………………………………………x Acknowledgements…………………………………………………………………………………………………..xi-xii Table of Contents…………………………………………………………………………………………………………xiii List of Figures……………………………………………………………………………………………………… …xiv-xvi Abbreviations……………………………………………………………………………………………………..…xvii-xix Abstract………………………………………………………………………………………………………………………1-2

Chapter 1- Significance, Specific Aims, and Background…………………………………………………3 Significance………………………………………………………………………………………………………………….…4 Specific Aims………………………………………………………………………………………………………………….5 Background……………………………………………………………………………………………………………….6-69 References………………………………………………………………………………………………………………70-79

Chapter 2- Aim 1………………………………………………………………………………………………………..…80 Abstract…………………………………………………………………………………………………………………….…81 Introduction………………………………………………………………………………………………………….…82-87 Materials and Methods……………………………………………………………………………………………88-93 Results…………………………………………………………………………………………………………………..94-108 Discussion……………………………………………………………………………………………………………109-116 References………………………………………………………………………………………………………..…117-127

Chapter 3- Aim 2…………………………………………………………………………………………………………128 Abstract……………………………………………………………………………………………………………………..129 Introduction………………………………………………………………………………………………………..130-134 Materials and Methods……………………………………………………………………………………….135-140 Results…………………………………………………………………………………………………………………141-156 Discussion……………………………………………………………………………………………………………157-163 References………………………………………………………………………………………………………..…164-169

Chapter 4- Overview, Future Directions, Clinical Implications….…………………………………170 Overview………………………………………………………………………………………………………….…171-173 Future Directions…………………………………………………………………………………………………174-198 Clinical Implications………………………………………………………………………………………….…199-200 References………………………………………………………………………………………………………..…201-207

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List of Figures

Chapter 1

Figure 1- Breast cancer subtypes.

Figure 2- Integrin signaling.

Figure 3- Focal adhesions.

Figure 4- PTPN12 structure.

Chapter 2

Figure 1- E2β-induced recruitment of integrin α5β1 to focal adhesions and the formation of actin stress fibers and FN fibrils are GPER- and Shc-dependent.

Figure 2- E2β stimulation of human ER-negative breast cancer cells induces the formation of focal adhesions and FN fibrils.

Figure 3- GPER stimulation promotes SKBR3 cell adhesion onto FN-coated, but not collagen-coated, substrata in a Shc-dependent manner.

Figure 4- GPER stimulation enhances haptotaxis of human SKBR3 breast cancer cells on

FN-coated, but not collagen-coated, substrata in a Shc-dependent manner.

Figure 5- GPER enhance FN-dependent, anchorage-independent growth of SKBR3 cells in a Shc-dependent manner.

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Figure 6- E2β stimulation alters colony morphology of ER-negative mouse breast cancer cell growth in soft agar.

Figure 7- GPER promotes FN fibril formation of mouse 4T1 breast cancer cells cultured in hanging drops.

Chapter 3

Figure 1- PTPN12 expression in human SKBR3 breast cancer cells stably transfected with

PTPN12 shRNA, WT PTPN12, PD PTPN12, or mutant NPLH PTPN12 as shown by western blot.

Figure 2- PTPN12 expression in human SKBR3 breast cancer cells stably transfected with

PTPN12 shRNA, WT PTPN12, or PD PTPN12 as shown by immunofluorescence.

Figure 3- PTPN12 expression in GE11β1 cells transfected with PTPN12 shRNA.

Figure 4- Overexpression of PTPN12 inhibits E2β-induced formation of focal adhesions.

Figure 5- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated formation of

FN fibrils in SKBR3 cells.

Figure 6- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated formation of

FN fibrils in GE11β1 cells.

Figure 7- Inhibition of PTPN12 results in constitutive activation of the EGFR.

Figure 8- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated cell adhesion onto FN-coated, but not collage-coated, substrata.

Figure 9- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated haptotaxis on

FN-coated, but not collagen-coated, substrata.

Figure 10- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated, anchorage- independent growth.

Chapter 4

Figure 1- Proposed mechanism of GPER action.

Figure 2- Stably transfected HEK-293 cells expressing human MMP-3.

Figure 3- Requirement of MMP-3 for GPER-mediated fibrillogenesis and EGFR transactivation.

Figure 4- Experimental pulmonary metastasis of 4T1 breast cancer cells.

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Abbreviations

5-FU- 5-flurouracil

BPA- Bisphenol A

CAMP- Cyclic adenosine monophosphate

Cdc42- Cell division control protein 42

CSK- C-terminal Src kinase

DOC- Deoxycholate

E2- Estrogen

E2α- 17α-Estradiol

E2β- 17β-Estradiol

ECM- Extracellular matrix

EGF- Epidermal growth factor

EGFR- Epidermal growth factor receptor

ER- Estrogen Receptor erbB1, erbB2, erbB3- Epidermal growth factor receptor

Erk- Extracellular regulated kinase

Erk-1- Extracellular regulated kinase 1

Erk-2- Extracellular regulated kinase 2

FAK- Focal adhesion kinase mABs- Monoclonal antibodies

MAPKs- Mitogen-activated protein kinases

MMPs- Matrix metalloproteinases

FN- Fibronectin

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G-1- 1-[4-(6-bromobenzo [1,3] dioxol-5yl)-3a,4,5,9b-tetrahydro-3H-cyclopenta- [c]quinolin-8-yl]-ethanone

G15- (3aS,4R,9bR)-4-(6-bromo-1,3-benzodioxol-5-yl)-3a,4,5,9b-tetrahydro-3H- cyclopenta[c]quinoline

G proteins- Guanine proteins

GDP- Guanosine diphosphate

GPCR- G protein-coupled receptor

GPER- G protein- estrogen receptor

GTP- Guanosine triphosphate

HEK 293- Human embryonic kidney 293

HER2- Human epidermal growth factor receptor 2

PARP1- poly [ADP-ribose] polymerase 1

PDGFR- Platelet-derived growth factor receptor

PI3K- Phosphoinositide 3-kinase

PKA- Protein kinase A

PKC- Protein kinase C

PR- Progesterone Receptor proHB-EGF- Pro heparin-bound epidermal growth factor

PTB- Phosphotyrosine binding domain

PTEN- Phosphatase and tensin homolog

PTP- Protein tyrosine phosphatase

SERMs- Selective estrogen receptor modulators

SH2- Src homology 2

SH3- Src homology 3 siRNA- small interfering ribonucleic acid

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TNBC- Triple negative breast cancer

VEGFR- Vascular endothelial growth factor receptor

WASP- Wiskott-Aldrich syndrome protein

WT- Wild-type

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Abstract

Stimulation of estrogen receptor (ER)-negative human breast cancer cells with 17β- estradiol (E2β) results in fibronectin (FN) matrix assembly and transactivation of the epidermal growth factor receptor (EGFR) via the G protein-coupled estrogen receptor

(GPER). This mechanism of action results in the recruitment of FN-engaged integrin

α5β1 to fibrillar adhesions and the formation of integrin α5β1-Shc adaptor protein complexes. Here, we show that GPER stimulation of murine 4T1 or human SKBR3 breast cancer cells promotes the formation of focal adhesions, actin stress fibers, and results in increased cellular adhesion and haptotaxis on FN, but not collagen. These actions are also induced by the xenoestrogen, bisphenol A, and the ER antagonist, ICI 182, 780, but not the inactive stereoisomer, 17α-estradiol (E2α). In addition, we show that GPER stimulation of breast cancer cells allows for FN-dependent, anchorage-independent growth and FN fibril formation in hanging drop assays, indicating that these GPER- mediated actions occur independently of adhesion to solid substrata. Furthermore, stable expression of Shc mutant Y317F lacking its primary tyrosyl phosphorylation site disrupts E2β-induced focal adhesion and actin stress fiber formation, and abolishes E2β- enhanced haptotaxis on FN and anchorage-dependent growth. We also show that overexpression of a phosphatase, PTPN12, known to interact with Shc inhibits focal adhesion and FN fibril formation, adhesion and haptotaxis on FN, and anchorage independent growth. Moreover, PTPN12 knockdown positively influences these events associated with cellular survival. Collectively, these data demonstrate that E2β action via GPER enhances cellular adhesivity and FN matrix assembly and allows for anchorage-

1 independent growth, cellular events that may allow for cellular survival and tumor progression.

Chapter 1

Significance, Specific Aims, and Background

Significance

Breast cancer is the most common malignancy among women [Sun, 2011], accounting for approximately 30% of all new cancer cases in women in 2013 [den Hollander, 2013].

Approximately one million new cases are diagnosed each year [Xunyi, 2013], and

232,340 new cases were diagnosed in the US alone in 2013 [den Hollander, 2013]. It is the most common cause of cancer-related death in women [Bozza, 2013], and it is estimated that approximately 40,000 women will die of breast cancer in 2013 [den

Hollander, 2013].

Patients with systemic breast cancer require aggressive therapeutic intervention, and despite treatment, these patients often have a poor prognosis. Estrogen (E2) is known to play an important role in promoting breast progression from primary cancer to metastatic disease; yet, the well known receptors, ERα and ERβ, that promote estrogen action are negatively associated with tumor progression variables. G protein- estrogen receptor (GPER) represents a newly appreciated estrogen receptor that is positively associated with tumor progression, and promotes cellular activities that may facilitate tumor implantation and invasion. Efforts to understand the specific biological mechanism of GPER action will serve as a foundation for the development of future therapeutic strategies that target this receptor and perhaps prevent metastatic disease

Specific Aims

1. To evaluate the role of Shc in GPER-mediated events associated with tumor cell

survival.

2. To evaluate the role of PTPN12 in GPER-mediated events associated with tumor

cell survival.

Background

Estrogen

E2 is a steroid hormone that promotes many biochemical actions [Filardo, 2006]. In women, it plays a role in the development of secondary sexual characteristics, regulation of gonadotropin secretion for ovulation, preparation of tissues for progesterone response, maintenance of bone mass, regulation of lipoprotein synthesis, prevention of urogenital atrophy, regulation of insulin responsiveness, and maintenance of cognitive function [Nelson, 2001]. It also prevents or delays age-associated changes in the skin, such as epidermal thinning, and has been shown to stimulate the proliferation and DNA synthesis of human keratinocytes [Kanda, 2004]. Most notably, E2 is important in the development and homeostasis of the mammary gland, and the growth of mammary tumors [Filardo, 2006].

E2 is produced from the granulose cells of the ovary and is synthesized from androgens by the enzyme aromatase [Nelson, 2001]. Aromatase is primarily expressed in ovarian granulose cells in premenopausal women, the placental syncytiotrophoblast in pregnant women, and the adipose and skin fibroblasts in postmenopausal women [Nelson, 2001].

E2 circulates in the blood bound to sex hormone-binding globulin, and is able to diffuse across the plasma and nuclear membranes of all cells, but is used only by cells that have estrogen receptors [Nelson, 2001]. Once bound to its receptor, there are two major signaling events that E2 promotes- rapid or pregenomic events, which occur within

6 minutes of E2 exposure, and genomic transcriptional responses, which occur over several hours after exposure [Filardo, 2006].

The estrogen receptor (ER) is a member of the nuclear steroid receptor superfamily

[Thomas, 2005]. There are two different forms of the ER, ERα and ERβ, and both function as hormone-inducible transcription factors and induce E2-dependent gene transactivation [Edwards, 2005]. The ER contains two highly conserved regions that are observed in other nuclear receptors [Nelson, 2001]. The first conserved region, domain

C, is in the middle of the protein and is involved in the interaction with DNA [Nelson,

2001]. The second conserved region, domain E/F, is in the carboxy-terminal region and binds hormones [Nelson, 2001]. Other functions of domain E/F include heat-shock protein association, dimerization, nuclear localization, and hormone-dependent transactivatin [Nelson, 2001]. The amino terminal region, domain A/B, has a ligand- independent activation function [Nelson, 2001].

Ligand-free ER forms a complex with heat shock proteins in the nucleus, but upon binding of E2, ER becomes active and heat shock proteins disassociate allowing dimerization of the receptor proteins [Nelson, 2001]. Active ER then binds as a homodimer to estrogen-response element DNA that is located in the 5’-flanking region of estrogen-responsive genes [Nelson, 2001]. These DNA sequences confer E2- inducibility on the genes [Nelson, 2001].

Normal Breast Tissue and Breast Cancer Overview

Overview-

Normal breast tissue is a lobulo-alveolar structure consisting of stratified myoepothelial cells that form the basal layer of ducts and alveoli, ductal epithelial cells that line the lumen of ducts, and alveolar epithelial cells that synthesize milk proteins [Dontu, 2003].

The stratified epithelium is located on a basement membrane [Chiorean, 2013]. All three cell types proliferate and differentiate in response to signals from the extracellular matrix (ECM) [Wozniak, 2005]. Upon differentiation, cells become polarized, growth- arrested, and highly organized into functional tissue [Wozniak, 2005]. However, with cancer development and progression, cells lose their normal interactions with the ECM and the cells de-differentiate, proliferate, and migrate [Wozniak, 2005].

Breast cancer is the most common malignancy among women [Sun, 2011], accounting for approximately 30% of all new cancer cases in women in 2013 [den Hollander, 2013].

Approximately one million new cases are diagnosed each year [Xunyi, 2013], and

Hollander, 2013].

Risk Factors-

There are several genetic, hormonal, and environmental risk factors that increase the probability of developing breast cancer [Mohanm, 2013]. Some of these risk factors include increasing age, past medical history of uterine cancer, ovarian cancer, or breast

8 cancer, and a family history of breast cancer [Mohanm, 2013]. In addition, postmenopausal hormone replacement therapy has been shown to increase the risk of breast cancer [Mohanm, 2013]. Patients with a history of breast cancer have a 30-50% chance of developing cancer in the contralateral breast [Mohanm, 2013]. Furthermore, dense breast tissue accounts for about 30% of breast cancers and is responsible for a four- to six-fold increased risk of developing breast cancer [Wozniak, 2005]. Dense breast tissue is a result of increased deposition of ECM proteins and fibroblasts in the stroma surrounding the epithelia cells [Wozniak, 2005].

Subtypes-

There are five major breast cancer subtypes (Table 1) [Wu, 2013]. They are defined by the presence or absence of the ER, the progesterone receptor (PR), and the receptor tyrosine kinase, human epidermal growth factor receptor 2 (HER2) [Sun, 2011]. Patients are considered ER-/PR-negative when immunohistochemistry staining for ER and PR is observed in under 1% of cells [Chiorean, 2013]. The first two subtypes are luminal A and luminal B [Wu, 2013]. Luminal A cancers are ER-positive and/or PR-positive and HER2- negative, while luminal B tumors are ER-positive and/or PR-positive and HER2-positive

[Wu, 2013]. Luminal cancers express genes associated with breast luminal cells [den

Hollander, 2013]. Patients with luminal cancers demonstrate the longest overall survival

[den Hollander, 2013]. The third subtype is basal-like, which are ER-negative, PR- negative, HER2-negative, and epidermal growth factor receptor (EGFR)- positive tumors

[Wu, 2013]. Basal-like cancers express genes active in breast basal epithelial cells [den

Hollander, 2013]. The fourth subtype is HER2-overexpressing, which are ER-negative,

PR-negative, and HER2-positive cancers [Wu, 2013]. Basal-like tumors and HER2- overexpressing tumors are associated with decreased survival [den Hollander, 2013].

The final subtype is unclassified and these tumors are negative for all markers [Wu,

2013].

The basal-like group is also known as triple negative breast cancer (TNBC) [Wu, 2013].

TNBCs demonstrate molecular heterogeneity and are highly aggressive [Albeck, 2011; den Hollander, 2013]. The majority of TNBCs are invasive ductal carcinomas; however, other types include invasive lobular carcinoma, metaplastic carcinoma, adenoid cystic carcinoma, neuroendocrine carcinoma, and secretory breast carcinoma [Chiorean,

2013]. Approximately 20% of all breast cancers are TNBC [Sun, 2011; Steinman, 2013], but for breast cancer patients under 50 years of age, the percentage of TNBC increases to 25-30% [Steinman, 2013]. Patients with TNBC generally present at more advanced stages, and 66-69% of TNBCs are grade 3 [Chiorean, 2013]. They have a high rate of reoccurrence [Chiorean, 2013], a high mitotic index, high nuclear-cytoplasmic ratio, frequent apoptotic cells, glameruloid microvascular proliferation, and lack of tubule formation [Chiorean, 2013]. While brain and lung metastases occur more frequently in

TNBC than in non-TNBC (30% versus 10% to brain, and 40% versus 20% to lung), liver and bone metastases occur less frequently in TNBC (20% versus 30% to liver, and 10% versus 40% to bone)[Chiorean, 2013]. Patients under 50 years of age with TNBC have a higher rate of brain metastases, and the time before brain metastases appear is shorter in TNBC patients (22 months for TNBC patients versus 51 months for non-TNBC patients)

[Chiorean, 2013]. The survival rate for patients after recurrence with TNBC is lower than that of non-TNBC (4 months versus 8 months) [Chiorean, 2013]. The rate of death at 5 years post-diagnosis in patients with TNBC is twice that of patients with non-TNBC

[Steinman, 2013].

Breast Cancer Subtype ER PR HER2 Characteristics Luminal A +/- +/- - Must be ER and/or PR positive; longest overall survival Luminal B +/- +/- + Must be ER and/or PR positive; longest overall survival Basal-like - - - TNBC; EGFR positive; shorter overall survival; highly aggressive HER2 Overexpressing - - + Shorter overall survival

Table 1- Breast cancer subtypes. Major breast cancer subtypes based on ER, PR, and HER2 expression, and their characteristics.

Staging-

Information collected upon baseline staging of breast cancer includes the tumor size, clinical evidence of nodal involvement, HER2 expression, PR expression, and ER expression [Bozza, 2013]. There are four stages of breast cancer, with several substages within each stage. Stage 0 is defined as carcinoma in situ with no lymph node metastasis and no distant metastasis [Yalcin, 2013]. Stage IA is when the primary tumor is less than

20 mm with no lymph node metastasis and no distant metastasis [Yalcin, 2013]. Stage IB is when there is either no evidence of primary tumor with metastasis to movable ipsilateral level I and II axillary lymph nodes, or the tumor is less than 20 mm with metastasis to movable ipsilateral level I and II axillary lymph nodes [Yalcin, 2013]. Stage

IIA is when there is either no evidence of primary tumor with metastasis to movable

11 ipsilateral level I and II axillary lymph nodes, or when the primary tumor is less than 20 mm with metastasis to movable ipsilateral level I and II axillary lymph nodes, or when the primary tumor is between 20 mm and 50 mm in diameter with no lymph node metastasis [Yalcin, 2013]. Stage IIB is either when the primary tumor is between 20 mm and 50 mm in diameter with metastasis to movable ipsilateral level I and II axillary lymph nodes, or when the primary tumor is greater than 50 mm with no lymph node metastasis [Yalcin, 2013]. Stage IIIA is when there is either no evidence of primary tumor with metastasis to ipsilateral level I and II axillary lymph nodes that are clinically fixed or matted or in ipsilateral internal mammary nodes, when the primary tumor is less than

20 mm with clinically fixed or matted ipsilateral level I and II axillary lymph node metastasis or ipsilateral internal mammary nodes, when the primary tumor is between

20 mm and 50 mm with metastasis to clinically fixed or matted axillary lymph nodes or ispilateral internal mammary nodes, when the primary tumor is greater than 50 mm with metastasis to movable ipsilateral and axillary lymph nodes, or when the primary tumor is greater than 50 mm with metastasis to ipsilateral internal mammary nodes and axillary lymph nodes [Yalcin, 2013]. Stage IIIB is when the primary tumor is any size with direct extension to the chest wall and/or the skin with no lymph node metastasis, when the primary tumor is any size with direct extension to the chest wall and/or the skin with metastasis to movable ipsilateral and axillary lymph nodes, or when the tumor is any size with direct extension to the chest wall and/or the skin with metastasis to ipsilateral and axillary lymph nodes that are clinically fixed or matted and ipsilateral internal mammary nodes [Yalcin, 2013]. Stage IIIC is when the primary tumor is any size

12 with metastasis in ipsilateral infraclavicular level III axillary lymph nodes with or without level I or II axillary lymph node involvement or in ipsilateral internal mammary lymph nodes with clinically evident level I or II axillary lymph node metastasis or metastasis in ipsilateral supraclavicular lymph nodes with or without axillary or internal mammary lymph node involvement [Yalcin, 2013]. Stage IV is when the primary tumor is any size with no or any lymph node involvement and distant detectable metastasis larger than

0.2 mm [Yalcin, 2013].

Stage IV, or metastatic cancer, is usually incurable because it reaches so many distant sites in the body that surgery becomes impossible [Ruoslahti, 1999]. In order for a tumor cell to become metastatic, it must be able to breach tissue barriers and proliferate at distant sites in the body [Ruoslahti, 1999].

Treatment and Prognosis of Breast Cancer

Overview-

The most common treatment options for breast cancer include chemotherapy, hormonal therapy, anti-HER2 therapy, and anti-angiogenic therapies [Mohanm, 2013].

The use of post-operative (adjuvant) and preoperative (neoadjuvant) chemotherapy and/or hormonal therapy have led to improvement in overall survival rate and progression-free survival [Mohanm, 2013]. Treatment options are selected based on factors such as tolerability, efficacy, and mechanism of overcoming resistance

[Mohanm, 2013]. Some specific treatment options include microtubule inhibitors, monoclonal antibodies, Epothilone, Taxane, tyrosine kinase inhibitors, poly [ADP-ribose]

13 polymerase 1 (PARP1) inhibitors, antiestrogens, and antimetabolites [Mohanm, 2013].

Microtubule inhibitors, such as Eribulin and Mesylate, cause irreversible mitotic block

[Mohanm, 2013]. Monoclonal antibodies act against HER2, such as Trastuzumab, or vascular endothelial growth factor receptor (VEGFR), such as Bevacizumab [Mohanm,

2013]. Epothilones, such as Ixabepiline, bind tubulin and initiate cell cycle arrest

[Mohanm, 2013]. Taxanes, such as Cabazitaxel, inhibit mitosis [Mohanm, 2013].

Tyrosine kinase inhibitors, such as Motesanib, inhibit VEGFR and platelet-derived growth factor receptor (PDGFR) [Mohanm, 2013]. PARP1 inhibitors, such as Iniparib, inhibit PARP1 leading to cell death [Mohanm, 2013]. Antiestrogens, such as Fulvestrant, block and degrade the ER and PR [Mohanm, 2013]. Antimetabolies, such as

Capecitabine, are converted to 5-fluoruracil (5-FU), which impairs growth [Mohanm,

2013].

Chemotherapy-

Chemotherapy is the primary treatment for breast cancer in women with tumors larger than 1 cm and in women with positive lymph nodes [Alken, 2013]. It is the only treatment option for metastatic breast cancer [Mohanm, 2013]. Chemotherapy is also used for some patients with early stage breast cancer with the hopes of treating micrometastatic disease [Alken, 2013].

The most effective chemotherapies include taxanes or anthracyclines [Mohanm, 2013].

Taxanes work by disrupting the equilibrium between polymerized and depolymerized forms of microtubules, which are required for cell division, inhibiting the ability of

14 mitotic spindles to separate DNA into two daughter cells during cell division [Alken,

2013]. Two available taxanes include docetaxel and paclitaxel [Alken, 2013]. Recent data demonstrates that anthracycline-containing regimens are better than non- anthracycline-containing multi-agent regimens such as cyclophosphamide, methrotrexate, and fluorouracil [Alken, 2013]. One study showed that anthracycline- based treatments given for 6 months reduce breast cancer death rates by 38% for women under 50 years at diagnosis and by 20% for women between 50 and 69 years at diagnosis [Alken, 2013]. Another study showed that taxane treatments improve recurrence-free survival when compared with non-taxane treatments [Alken, 2013].

Chemotherapy treatments come with side effects and toxicities [Alken, 2013]. Most of the side effects are reversible such as peripheral neuropathy, neurosensory disturbances, fluid retention, rash, bronchospasm, diarrhea, nail disorder, and neutropenia, but there are some irreversible side effects including cardiomyophahy, acute myelogenous leukemia, and myelodysplastic syndrome [Alken, 2013].

ER Antagonists-

Patients with hormone-receptor positive, lymph-node negative breast cancers can sometimes avoid chemotherapy and opt for endocrine therapy [Alken, 2013]. Endocrine therapeutics targeting ER- and PR- positive tumors lead to significant increases in patient survival [Sun, 2011]. Selective estrogen receptor modulators (SERMs), or ER antagonists, work by blocking the E2 binding sites on the ER to suppress E2 signaling pathways [den Hollander, 2013]. They are effective in inhibiting the growth of breast

15 tumors expressing ER [Filardo, 2006]. In addition, coexpression of the PR with ER generally leads to a more favorable response to hormonal therapy, likely because PR gene transcription is regulated by ER gene transactivation [Filardo, 2006]. Adjuvant endocrine therapy is recommended in all patients with 1% or greater ER- or PR- positive cells [Chiorean, 2013].

Some commonly used ER antagonists include tamoxifen and raloxifen [Vivacqua, 2006].

Tamoxifen is used to treat all stages of breast cancer, and was the first ER antagonist to be used for treatment of metastatic breast cancer [den Hollander, 2013]. Tamoxifen reduces recurrence and contra-lateral breast cancer by 40-50% in women with early breast cancers [den Hollander, 2013]. One study showed that tamoxifen reduces recurrence in women with a history of loblualr carcinoma by 56% [den Hollander, 2013].

Results from four studies investigating the effectiveness of tamoxifen in reducing breast cancer in women at high risk showed that tamoxifen reduces the overall incidence of breast cancer by 16-49%, and the incidence of ER-positive breast cancer by 31-69% [den

Hollander, 2013]. While many tamoxifen trials are effective in preventing ER-positive breast cancers, there is generally no reduction in ER-negative tumors [den Hollander,

2013]. In addition, not all patients with ER-positive tumors respond favorably to hormonal therapy. Approximately 25% of patients do not respond to tamoxifen based on data collected at 5-year follow-up for postmenopausal patients with ER-positive tumors [Filardo, 2006]. There are several possible explanations for this lack of response to tamoxifen: ER expression heterogeneity within the tumor; the development of mutant ERs with reduced affinity for ER antagonists; drug resistance; partial receptor

16 antagonism; and/or the presence or absence of trans-acting factors that influence ER functionality [Filardo, 2006]. The existence of another ER, whereby E2 may promote its actions on the cell, is another explanation for tamoxifen nonresponsiveness [Filardo,

2006].

There are several side effects to tamoxifen treatment including increased risk of thromboembolisms, endometrial cancer, hot flushes, vaginal symptoms, and cataracts

[den Hollander, 2013]. It is also important to note that some studies have shown that tamoxifen displays agonistic activity in the uterus, bone, and cardiovascular system

[Vivacqua, 2006]. More specifically, tamoxifen induces uterine growth and stimulates the proliferation of human endometrial cells [Vivacqua, 2006].

Raloxifene works by blocking coactivator binding to the activating function 2 domain of

ER, inhibiting transcription of the ER-regulated gene [Gizzo, 2013]. In addition, raloxifen may reduce the human telomerase reverse transcriptase expression and vascular endothelial growth factor expression [Gizzo, 2013]. Raloxifene treatment for 4 or 8 years results in a 31% and 59% reduction in invasive breast cancer, respectively [den

Hollander, 2013]. Several studies show that long-term treatment reduces the rate of ER- positive breast cancer, but has no effect on ER-negative breast cancers [den Hollander,

2013].

Lasofoxifene is another ER antagonist that shows promise in the treatment of breast cancer, but it has not been FDA-approved. According to one study, patients treated with lasofoxifen for 5 years had an 81% decrease in ER-positive breast cancer [den Hollander,

2013]. Lasofoxifen also has lower toxicity when compared to tamoxifen and raloxifen

[den Hollander, 2013].

Aromatase Inhibitors-

Aromatase inhibitors work by blocking E2 synthesis [Albeck, 2011]. Specifically, they block the synthesis of E2 from androgens through the inhibition of the aromatase enzyme [den Hollander, 2013]. Aromatase inhibitors are effective in treatment of women with ER-positive breast cancer [den Hollander, 2013]. Some studies have demonstrated that aromatase inhibitors are more effective in the treatment of breast cancer, with an increased time to recurrence following treatment with aromatase inhibitors when compared to tamoxifen [den Hollander, 2013]. Commonly used aromatase inhibitors include letrozole, anastrozole, and exemestane [den Hollander,

2013]. A major side effect observed with the treatment of aromatase inhibitors is enhanced bone loss leading to an increased risk of bone fractures [den Hollander, 2013].

Other side effects of aromatase inhibitors include an increased risk of cardiovascular disease, and joint pain and stiffness [Lonning, 2013].

HER2 Inhibitors-

HER2-positive tumors are treated with a HER2 blocking antibody, Tratuzumab, or a HER2 kinase inhibitor, Trikerb [Albeck, 2011]. These drugs inhibit tumor growth and induce apoptosis [den Hollander, 2013]. Trastuzumab targets the extracellular domain of HER2.

Another dual EGFR/HER2 inhibitor, Lapatinib, inhibits kinase activity of HER2 and EGFR

[den Hollander, 2013].

Treatment for TNBC-

The prognosis for patients diagnosed with TNBC is poor because this subtype often reoccurs and becomes metastatic [Sun, 2011]. Furthermore, there is a lack of effective targeted treatments [Sun, 2011; Chiorean, 2013]. EGFR and HER2 inhibitors are ineffective in TNBC even though these tumors express high levels of EGFR [Albeck,

2011]. Cytotoxic chemotherapy is the frontline treatment for TNBC patients because specific targeted therapy has not been successful [Steinman, 2013]. TNBCs have higher response rate to chemotherapy initially, but they have a lower overall survival rate and greater risk or recurrence after treatment, which could be attributed to a faster adaptive response to the cytotoxic effects of chemotherapy in these tumors [Chiorean,

2013]. The first 3-5 years after initial treatment comes with the greatest risk of recurrence [Chiorean, 2013]. After 5 years, reoccurrence risk decreases, and after 10 years, patients have a good prognosis [Chiorean, 2013]. In order to create an effective pharmaceutical agent for TNBC, a target that is present in the majority of TNBC tumors must be discovered [Steinman, 2013].

Neoadjuvant Chemotherapy-

Neoadjuvant, or preoperative, chemotherapy was originally an option for patients with locally advanced breast cancer (stage III) to shrink the tumor before surgery, but it is now used for patients with early stage breast cancer when surgery is not an option, as well [Bozza, 2013]. Patients with HER2-positive tumors or TNBC respond more favorably

19 to neoadjuvant chemotherapy when compared to patients with HER2- negative tumors or ER-positive tumors [Bozza, 2013].

Neoadjuvant Endocrine Therapy-

Neoadjuvant endocrine therapy is generally used for postmenopausal women who are not good candidates for chemotherapy [Bozza, 2013]. These patients are given primary endocrine therapy for a minimum of 3-4 months [Bozza, 2013]. The option of surgery is then revisited [Bozza, 2013]. Fifty percent of patients become surgery eligible within 4 months of neoadjuvant letrozole treatment, and longer treatment of up to 8 months results in even greater tumor reduction in some cases [Bozza, 2013]. If the tumor does not respond significantly after 3-4 months of treatment, then 6 months or longer with clinical monitoring is an option [Bozza, 2013]. Aromatase inhibitors are generally used over tamoxifen in these patients [Bozza, 2013]. In one study, a higher response rate was observed in hormonal receptor positive tumors treated with neoadjuvant letrozole

(55%) versus tamoxifen (36%) [Bozza, 2013]. Another study showed that in postmenopausal women, aromatase inhibitors are more effective in downstaging tumors and reducing the need for mastectomy [Bozza, 2013]. In another study, anastrozone had a greater response rate than tamoxifen (35.4% versus 12.2%) [Bozza,

2013]. There is no difference in outcome of patients treated with letrozole, anastrozole, or exemestane [Bozza, 2013].

GPER

New Estrogen Receptor-

Studies in animal and cell models demonstrate that E2 promotes rapid biochemical effects that are not related to its known actions via the ER [Filardo, 2006]. One study showed that intrauterine administration of E2 in rats results in a rapid increase in cyclic adenosine monophosphate (cAMP) [Szego, 1967]. In addition, E2 promotes EGFR activation and second messenger signaling including calcium and inositol triphosphate activation [Filardo, 2006; Thomas, 2005]. This second messenger signaling is generally linked to seven transmembrane-spanning receptors [Filardo, 2006]; therefore, a new estrogen receptor may be responsible for the rapid E2 signaling. In support of this idea, the nonclassical E2 action is associated with G proteins [Thomas, 2005]. In addition, 25% of patients with ER-positive tumors do not respond to tamoxifen [Filardo, 2006]. In these patients, E2 may be exerting effects through this alternative ER. A candidate alternative ER is G protein-estrogen receptor (GPER), a G protein-coupled receptor

(GPCR).

GPCR Overview-

GPCRs are seven-transmembrane spanning receptors [Revankar, 2005]. With over 800 genes that encode GPCRs in the human genome, they are the largest class of signaling molecules [Filardo, 2005]. These receptors have a variety of extracellular stimulants such as light, odorants, neurotransmitters, vasoactive substances, chemokines, and peptide hormones [Filardo, 2005] that bind and initiate activation of heterotrimeric guanine proteins (G proteins) and downstream effectors [Filardo, 2005]. There are three

GPCR families. Family A consists of rhodopsin-type receptors, which have 20 highly

21 conserved residues [Rios, 2001]. Family B consists of secretin and glucagon receptors, which have a large N-terminal extracellular domain that contains well-conserved cysteine residues in addition to the 20 highly conserved residues in the transmembrane regions [Rios, 2001]. Family C consists of metabotropic neurotransmitter and Ca2+ sensing receptors with a very long N terminal extracellular domain that contains 20 cystine residues in addition to the 20 highly conserved residues of the transmembrane regions [Rios, 2001].

GPCRs signal via heterotrimeric G proteins, which consist of Gα, Gβ, and Gγ subunits

[Bondar, 2013]. They are responsible for transducing and amplifying signals from GPCRs to effectors within the cell [Bondar, 2013]. Upon activation of GPCRs by a ligand, an exchange of guanosine diphosphate (GDP)-bound to an inactive Gα subunit for guanosine triphosphate (GTP) occurs [Bondar, 2013]. The activated GTP-bound Gα subunit and the Gβγ dimer can now modulate activity of effectors [Bondar, 2013].

Common effectors include adenylate cyclase, phospholipase C, and G protein-regulated inward rectifying potassium channels [Bondar, 2013].

GPER Overview and Distribution-

GPER is widely expressed in numerous tissues throughout the body and is often highly expressed in cancer cell lines, particularly those from aggressive tumors [Smith, 2009].

GPER has been described in both normal and malignant neural, breast, placental, heart, ovarian, prostate, hepatic, vascular epithelial, and lymphoid tissues [Thomas, 2005]. It demonstrates mostly cytoplasmic staining in normal and malignant breast tissue

[Filardo, 2006]. It also demonstrates a subcellular distribution, which is likely due to receptor re-uptake [Filardo, 2006]. Revankar et al. report that GPER was observed on the endoplasmic reticulum and the golgi apparatus, but not in the mitochondria or the actin cytoskeleton [Revankar, 2005].

Structure of GPER-

GPER was discovered by molecular cloning, and demonstrates patterns of a plasma membrane ER [Filardo, 2002]. Typically, many cell surface receptors belong to the GPCR superfamily [Rios, 2001]. GPCRs have a seven-transmembrane spanning structure, and signaling molecules and drugs bind to the extracellular loops, while the intracellular regions are involved in signaling [Rios, 2001]. GPER shows structural sequence homology to angiotensin, inteleukin, and chemokine receptors [Thomas, 2005]. It has a 47 amino acid c-terminal domain, and a DRY sequence that is involved in signal transduction in the second intracellular loop [Thomas, 2005]. There are two conserved cysteines in the first two extracellular loops, which form disulfide bonds to help stabilize the receptor, and the second extracellular loop is 10-20 amino acids in length [Thomas, 2005]. The N terminal extracellular domain is 57-75 amino acids long, and contains glycosylation sites

[Thomas, 2005].

GPER Acts Independently of ER-

There is overwhelming evidence that GPER acts independently of the ER. First, expression of ERs and GPER is observed independently in both breast tumors and in breast cancer cells lines [Filardo, 2012], and cells expressing GPER, but lacking ER,

23 demonstrate specific E2 binding to the plasma membrane [Thomas, 2005]. Second, ER and GPER have different binding affinities for their ligands [Filardo, 205]. The binding affinity of the ER for E2β is typically greater than that of GPER [Filardo, 2005]. The dissociation constant of E2 binding to the ER is Kd=0.1-1.0 nM, while that of GPER is

Kd=3.3 nM [Filardo, 2005]. Specifically, membranes prepared from ER-negative, GPER- positive human breast cancer SKBR3 cells demonstrate a single, high affinity (Kd=2.7 nM), saturable, low capacity (Bmax=9pM) specific E2 binding site [Filardo, 2005]. In addition, the binding affinity of bisphenol A (BPA) to GPER is 8-50 times higher that

BPA’s affinity to the ER [Filardo, 2005], while other steroid hormones such as E2α, progesterone, cortisol, and testosterone demonstrate no affinity for GPER [Filardo,

2005]. The rates of association and dissociation of E2 binding to ER occur over several hours, while those of GPER occur over minutes, with a half-life of less than ten minutes

[Filardo, 2005]. Association (half-life=5.5min) and dissociation (half-life=8.1min) of E2 in

SKBR3 cells is rapid [Filardo, 2005]. This rapid binding time is characteristic of other membrane steroid receptors [Thomas, 2005]. Third, it has been shown the ER antagonists act as GPER agonists [Filardo, 2005]. Stimulation of breast cancer cells expressing GPER and lacking the ER with ICI 182,780 (Faslodex) and tamoxifen, both ER antagonists, results in E2-like effects in wild-type (WT) and recombinant GPER models

[Filardo, 2006]. Fourth, the activities associated with E2 signaling via ER and GPER are different. GPER is associated with activated G proteins and the upregulation of adenylyl cyclase activity, which is not characteristic of the ER [Thomas, 2005]. In a study by

Thomas et al., ER-negative and GPER-positive SKBR3 human breast cancer cells

24 demonstrated EGFR transactivation and adenylyl cyclase activity when stimulated with

E2, while these events were inhibited in SKBR3 cells treated with GPER small interfering ribonucleic acid (siRNA) and stimulated with E2 [Thomas, 2005]. In addition, in a study by Thomas et al., human embryonic kidney 293 (HEK-293) cells, lacking both ER and

GPER expression, demonstrated insignificant E2 binding or activation of G proteins.

However, in HEK-293 cells transfected with GPER, E2 stimulation resulted in E2 binding similar to that of SKBR3 cells [Thomas, 2005]. Activation of GPER by E2 may induce the dissociation of G-protein heterotrimers, Gsαβγ, into Gsα, which stimulates adenyl cyclase, and Gβγ, which promotes EGFR transactivation [Kanda, 2004]. Fifth, ER or GPER null mice each exhibit distinctly different phenotypes [Filardo, 2005]. Finally, ER and

GPER are associated with different clinicopathological markers of female reproductive cancers, such as disease free survival, metastases, primary tumor size, and lymph node invasion [Filardo, 2006].

A non-steroidal, GPER agonist, 1-[4-(6-bromobenzo [1,3] dioxol-5yl)-3a,4,5,9b- tetrahydro-3H-cyclopenta-[c]quinolin-8-yl]-ethanone (G-1), has been developed to further study GPER signaling independent of ER signaling, as G-1 binds GPER, but not ER

[Wang, 2012]. G-1 does not activate estrogen response elements, and does not bind 25 other GPCRs [Bologa, 2006]. In studies cited by Wang et al., activation of GPER by G-1 increased mobilization of intracellular calcium in COS7 cells overexpressing GPER, and activated phosphoinositide 3-kinase (PI3K) in SKBR3 and MCF7 cells [Wang, 2012].

Albanito et al. showed that activation of GPER by G-1 results in proliferation of SKBR3 breast cancer cells [Albanito, 2007].

(3aS,4R,9bR)-4-(6-bromo-1,3-benzodioxol-5-yl)-3a,4,5,9b-tetrahydro-3H- cyclopenta[c]quinoline (G15), a selective GPER antagonist that does not bind classical

ERs, is able to block the ability G-1 induced mobilization of intracellular calcium and inhibit GPER-dependent PI3K activation in SKBR3 cells [Wang, 2012]. Other studies have also shown that G15 blocks the action of G-1 in various tissues [Wang, 2012].

GPER and Significance in Breast Cancer

GPER and Significance in Breast Cancer-

GPER has significance in breast cancer based on studies conducted in human breast tumors and breast cancer cell lines. In one study, while overexpression of GPER was not observed in primary invasive ductal tumors, about 50% of ER-negative tumors were

GPER-positive, suggesting that some ER-negative tumors may remain E2 responsive via

GPER [Filardo, 2006]. There were almost twice as many tumors that coexpressed ER and GPER than just ER-positive tumors [Filardo, 2006]. There was no correlation between GPER and PR expression; however, PR expression was twice as common in tumors that expressed both ER and GPER than in tumors that expressed only ER [Filardo,

2006]. As discussed above, coexpression of PR and ER generally leads to a more favorable response to hormonal therapy because the PR genes encode estrogen response elements that promote E2-dependent gene transcription [Filardo, 2006]. GPER expression may also predict the effectiveness of hormonal therapy because ER gene transactivation is increased by extracellular-regulated kinases 1 and 2 (Erk-1 and Erk-2), which are activated by GPER-dependent transactivation [Filardo, 2006]. Of most

26 importance, GPER expression in primary tumors is positively associated with tumor size,

HER2 expression, and the presence of extramammary metastases [Filardo, 2006], which will be discussed in greater detail later.

In another study conducted on breast tumors, the average age of patients with GPER- positive tumors was 45.8 years, while the average age of patients with GPER-negative tumors was 61.7 years [Steinman, 2013]. This difference is consistent with the finding that younger women have more aggressive tumors [Steinman, 2013]. Steiman et al. looked specifically at TNBC tumors, and found that 17 out of 18 tumors expressed GPER, further supporting the hypothesis that poor-prognosis tumors are more likely to express

GPER [Steinman, 2013]. Furthermore, 6 patients in the GPER-positive group (22.2%) had recurrences, while only 2 patients in the GPER-negative group (9.5%) had recurrences

[Steinman, 2013].

In ER-negative, GPER-positive SKBR3 human breast cancer cells, E2 and G-1 promote

GPER-dependent extracellular regulated kinase (Erk) activation and c-fos induction through EGFR transactivation, resulting in cell proliferation [Albanito, 2007]. This was inhibited when GPER antisense oligonucleotides were used [Albanito, 2007].

Furthermore, these proliferative responses were observed in breast cancer cells stimulated with genistein, a phytoestrogen [Filardo, 2005]. In addition to cell proliferation, stimulation of SKBR3 cells with E2 promotes pro heparin-bound epidermal growth factor (proHB-EGF) release, prolonged epidermal growth factor receptor (erbB1) tyrosyl phosphorylation, and Erk-1/-2 activation [Filardo, 2005]. In another cell model,

GPER-null MDA-MB-231 cells did not stimulate adenylyl cyclase. However, overexpression of GPER in MDA-MB-231 cells restored the activation of adenylyl cyclase by E2 [Filardo, 2005].

GPER and ER Antagonists-

GPER may have significance in the treatment of breast cancer patients. As discussed previously, patients with ER-positive breast tumors are often given tamoxifen or other

ER antagonists to prevent reoccurrence of the tumor; however, it has been shown that these ER antagonists are GPER agonists in cultured breast cancer cell lines [Thomas,

2005], which is consistent with the observation that prolonged tamoxifen use in women is correlated with endometrial hyperplasia [Filardo, 2005]. In addition, E2 effects have been observed in ER knockout mice treated with these ER antagonists [Filardo, 2005].

ER antagonists have been shown to stimulate adenylyl cyclase and intracellular cAMP in

MCF7 cells [Filardo, 2005]. Extended tamoxifen use has been associated with endometrial hyperplasia [Filardo, 2005].

GPER and Significance in Cell Survival and Metastasis-

There is no significant association between ER expression and breast cancer metastasis; however, GPER-positive breast tumors are almost twice as likely metastasize than GPER- negative tumors [Filardo, 2005]. One study showed an increase in lymph node invasion of GPER-positive tumors when compared to GPER-negative tumors [Filardo, 2005]. In breast cancer cell lines, E2 stimulation of GPER-positive cells results in EGFR transactivation, which has been linked to increased tumor survival, growth, and invasion

28 into distant tissues [Filardo, 2005]. In addition, GPER-dependent E2 action results in the activation of integrin α5β1, the primary FN receptor, and increased FN matrix assembly, which is significant in advanced disease because these activities allow for cell survival of metastatic cancer cells [Quinn, 2009]. Furthermore, FN matrix assembly is important in anchorage independent growth, which is a predictor of metastasis [Quinn, 2009].

GPER Significance in Other Cancers

Ovarian Cancer-

GPER has significance in ovarian cancer. In response to E2 or G-1 treatment, ovarian cancer cell lines expressing GPER proliferate at an increased rate [Smith, 2009]. In addition, Albanito et al. show that G-1 induces the proliferation of both ERα-positive and ERα-negative ovarian cancer cells in an EGFR-dependent fashion [Albanito, 2007].

Furthermore, E2 and G-1 induce the expression of c-fos, pS2, and cyclins A, D1, and E

[Albanito, 2007]. ERα-dependent transcription and PR expression were only altered in cells treated with E2, not G-1 [Albanito, 2007]. In support of these data, abrogation of

ERα and GPER expression by antisense oligonucleotides inhibited c-fos stimulation, Erk activation, and cell proliferation in cells treated with E2 or G-1 [Albanito, 2007]. In a study by Wang et al., knockdown of GPER inhibited G-1 induced cell proliferation in a granulosa tumor cell model [Wang, 2012].

GPER expression is correlated with 5 year survival rate, stage, and tumor grade of epithelial ovarian cancers [Smith, 2009]. In primary, grade 1 tumors with higher survival rates, GPER is expressed at a lower level, while GPER expression above the median was

29 associated with lower survival rates [Smith, 2009]. In addition, another study showed that GPER expression was observed in 23 of 24 endometriosis associated ovarian cancer specimens, while only 8 of the 32 benign specimens expressed GPER [Long, 2012].

Furthermore, GPER mRNA expression was increased in malignant tissue when compared to benign tissue [Long, 2012].

Estrogen replacement therapy action through GPER may be responsible for the malignant transformation of ovarian endometriosis [Long, 2012]. There are reports that breast cancer patients treated with tamoxifen undergo various endometriosis hyperplastic at a greater rate than those not treated with tamoxifen [Long, 2012]. Long et al. found that GPER expression of endometriosis associated ovarian carcinoma was correlated with late tumor stage, large tumor size, lymph node metastasis, and the development of distant metastatic disease [Long, 2012]. In addition, Long et al. found that GPER expression was correlated with matrix metalloproteinase-9 (MMP-9) expression in patients with endometriosis associated ovarian cancer [Long, 2012]. This is significant because MMP-9 is associated with matrix degradation, which is important in tumor invasion and metastasis, suggesting that GPER is important in tumor invasion and metastasis [Long, 2012].

Endometrial Cancer-

GPER also has significance in endometrial cancer. Endometrial cancer is divided into two types- type I and type II [Smith, 2007]. The 5 year survival rate of patients with type I tumors is 85.6%, whereas that of patients with type II tumors is 58.8% [Smith, 2007].

Type II tumors are generally classified as grade 3 nuclear and/or architectural, contain phenotypic features associated with lower survival rates (clear cell and mixed epithelial), invade deeply into the myometrium, lymphatic, and vascular spaces, are less likely to be

ER- or PR-positive, and are more likely to recur [Smith, 2007]. Only about 15-20% of recurrent tumors express ER or PR, so response to progestational therapy is poor

[Smith, 2007]. Smith et al., conducted a study characterizing 47 patients with endometrial carcinomas [Smith, 2007]. On the luminal or basal surfaces of normal, postmenopausal endometrial tissue, there was minimal GPER expression; however, there was intense GPER expression around the stroma without staining of the glands

[Smith, 2007]. GPER was inversely correlated with ER and PR expression, and positively correlated with EGFR expression [Smith, 2007]. EGFR expression had no correlation with

ER expression [Smith, 2007]. High levels of GPER expression were observed more frequently in tumors with lower survival rates, high grade, deep myometrial invasion, and advanced stage [Smith, 2007]. In addition, high levels of GPER expression were observed more frequently in high risk endometrial cancer subtypes including uterine papillary serous carcinomas, clear cell carcinomas, and carcinosarcomas [Smith, 2007].

Specifically, survival was 100% in patients with tumors that had low GPER expression and 62.5% in patients with tumors that overexpressed GPER, and GPER was overexpressed in all 8 cancer-related deaths [Smith, 2007].

The tamoxifen used to treat breast cancer has been shown to stimulate cell growth and proliferation in endometrial tissue [Ignatov, 2012]. Proliferation of endometrial tissue by tamoxifen increases the risk of endometrial abnormalities, such as polyps, endometrial

31 hyperplasia, and endometrial carcinoma [Ignatov, 2012]. The risk is increased with increased duration of tamoxifen use [Ignatov, 2012]. Ignatov et al. determined that tamoxifen-induced cell growth in HEC-1A cells occurs in a GPER-dependent manner

[Ignatov, 2012]. In addition, in a cohort of 95 patients, there was a significant correlation between GPER expression and tamoxifen-induced increased rate of endometrial polyps or hyperplasia [Ignatov, 2012]. Furthermore, there was an inverse correlation of GPER expression and the length of time between the start of tamoxifen therapy and the development of symptoms [Ignatov, 2012].

Uterine Cancer-

GPER has significance in uterine cancer. Hec50, an aggressive uterine cancer cell line, expresses elevated levels of GPER when compared to H, it associated normal uterine cell line [Revankar, 2005]. Furthermore, tamoxifen acts as an agonist in the uterus [Ignatov,

2012], and G15 has been shown to inhibit E2-induced proliferation of uterine epithelium in a mouse model [Smith, 2007].

Matrix and Survival

ECM Overview-

The ECM is important in normal and cancerous cell adhesion, migration, and tissue organization [Pankov, 2000; Geiger, 2011]. It also acts as a cellular cross-linker, connecting cells through integrin-ECM interactions [Caicedo-Carvajal, 2010]. It is made

32 up of signaling molecules that can activate intracellular complexes through recruitment to the adhesion site and stimulate growth factor receptors [Geiger, 2011].

Adhesion-

Adhesion is important in the maintenance of tissue, and many cell adhesion molecules and their ligands have been implicated in cancer [Ruoslahti, 1999]. The interaction between adhesion receptors and their ligands controls cell adhesion and can be regulated through reversible modulation of receptor function [Faull, 1993]. Integrins are a family of adhesion receptors [Faull, 1993]. Cellular adhesion begins with receptor- ligand interaction, followed by strengthening of adhesions through the organization of the cytoskeleton, the formation of focal adhesions, and cell spreading [Faull, 1993].

Adhesion is significant in migration. In order for a cancer cell to migrate, adhesion between cells must loosen [Ruoslahti, 1999], and upon contact with a distant site in the body, the cell must have the ability to adhere to this new site. Fibronectin (FN), a cell adhesion protein that mediates anchorage through integrins, plays a role in adhesion and migration [Ruoslahti, 1999].

Migration-

Cell migration is important in invasion of a wounded space, the translocation of lymphocytes and neutrophils to an inflammatory site [Angers-Loustau, 1999], embryogenesis, arthritis, atherosclerosis, osteoporosis, and cancer [Webb, 2012].

Specifically, in breast cancer, migration is important for metastasis [Ruoslahti, 1999]. To

33 be able to migrate, a cancer cell must loosen its adhesion at the primary site, invade a blood or lymphatic vessel, travel in the circulation, and anchor to a distant site

[Ruoslahti, 1999].

Cell migration is regulated by integrins; protein kinases, such as Src; focal adhesion kinase (FAK) and proline rich tyrosine kinase 2; small molecular weight G-proteins such as Rho, Rac, and cell division control protein 42 (Cdc42); guanine nucleotide exchange factors such at Vav; GTPase; adaptor molecules such as paxillin and p130Crk; and structural proteins such as tensin, talin, and actin [Zheng, 2009].

The process of migration begins with the extension of protusions at the cell front, which is driven by polymerization of the actin cytoskeleton [Webb, 2012]. Rac, a small molecular weight G-protein, is involved in the formation of new adhesions at the cell front, while Rho, another small molecular weight G-protein, is required for the maturation of these contacts [Webb, 2012]. The protusions are stabilized by attachment to the substratum [Webb, 2012]. The interaction of integrins with ECM proteins is responsible for the stabilization of these adhesions [Webb, 2012]. The adhesions transmit strong forces, and serve as traction points for the forces that move the cell forward [Webb, 2012]. At the rear of the cell, adhesions are released, partly by contractile forces, resulting in retraction of the cell tail and movement of the cell forward [Webb, 2012]. Rho promotes the release of adhesions [Webb, 2012].

There are many other proteins involved in the process of migration. FAK activity has been correlated with migration rate [Angers-Loustau, 1999], which has also been

34 correlated with the hyperphosphorylation of p130cas by Src, a tyrosine kinase [Angers-

Loustau, 1999]. Fibroblasts from FAK-null mice demonstrate reduced migration and spreading rates, and an increase in the number and size of adhesions [Webb, 2012].

Overexpression of Src has been associated with increased motility and invasiveness of tumor cells [Chiorean, 2013], and increased metastases in breast carcinomas [Chiorean,

2013]. Fibroblasts from Src-null mice demonstrate a decrease in cell spreading and reduced motility [Webb, 2012].

Both formation and breakdown of focal adhesions can occur at the same time to increase turnover rate [Angers-Loustau, 1999]. Proteases are necessary to break down adhesion, and integrins often control the expression of proteases [Ruoslahti, 1999].

Without the necessary components for focal adhesion breakdown, the adhesions between cells and the ECM are too strong, and the cell cannot migrate [Angers-Loustau,

1999]. Fibroblasts lacking SHP-2, a phosphatase, have an increased number of adhesions and decreased rates of spreading and migration [Webb, 2012].

Fibronectin-

FN is a soluble plasma protein that is secreted as a disulfide-bonded FN dimer

[Takahashi, 2007]. Each subunit is 230 kDa to 270 kDa [Huynh, 2013] and contains three repeating modules- FN-I, FN-II, FN-III [Takahashi, 2007]. It is produced in the liver and secreted from hepatocytes into circulation [Huynh, 2013]. It is essential in processes such as embryogenesis, development, blood vessel formation, inflammation, atherosclerosis, and wound healing [Quinn, 2009; Takahashi, 2007]. It is also imperative

35 in tumor cell survival and migration. These functions are dependent on the conversion of soluble FN into an insoluble fiber during FN matrix assembly [Quinn, 2009; Zheng,

2009]. In studies cited by Quinn et al., FN enhances tumorigenicity and allows for resistance to apoptosis by chemotherapy [Quinn, 2009]. There are survival advantages for tumors that interact with FN [Quinn, 2009]. It has been shown that mammary adenocarcinomas can convert soluble FN into fibrils resulting in enhanced responsiveness to growth factors and improved anchorage-independent growth, which is a predictor of metastatic potential [Quinn, 2009]. In addition, lung carcinomas often express high levels of FN [Quinn, 2009]. In another study, implantation of mammary tumor xenografts in immunocompromised mice is enhanced with the introduction FN

[Quinn, 2009].

RGD is a soluble peptide fragment of the FN recognition sequence (arginine, glycine, aspartic acid) [Quinn, 2009]. The interaction of the RGD motif of FN with α5β1 or αv integrins is an essential step in FN fibril formation [Takahashi, 2007].

Matrix Assembly-

Matrix assembly involves the conversion of soluble FN into fibrillar structures containing insoluble FN [Pankov, 2000]. It occurs at the cell surface, and is triggered by the unfolding of the FN dimer [Pankov, 2000]. Integrins are major players in this event, as upon unfolding of the FN dimer, the interaction between FN and integrins promotes cellular adhesion, haptotaxis, and survival [Quinn, 2009]. FN assembly begins when FN is secreted in a compact or inactive conformation, followed by integrin binding, and the

36 conversion of FN into an active and expanded dimer [Takahashi, 2007]. Next, FN binding induces integrin clustering and the accumulation of integrin-bound FN, which allows for

FN-FN interactions and the formation of fibrils [Takahashi, 2007].

ECM and Cancer-

Matrix assembly is implicated in cancer. FN and integrins are expressed in the mammary gland and are affected by E2 [Quinn, 2009]. As cited by Quinn et al., the ability of mammary adenocarcinoma cells to convert soluble FN into fibrils results in increased responsiveness to growth factors and anchorage-independent growth [Quinn, 2009]. E2 treatment of breast cancer cells results in cytoarchitectural changes, and actin stress fiber and focal adhesion formation have been observed in breast cancer cells treated with tamoxifen [Quinn, 2009]. Integrins bind to the RGD cell attachment sequence on

FN, but treatment of breast cancer cells with alternative peptides containing the RGD sequence results in antimetastatic effects [Ruoslahti, 1999]. As cited by Ruoslahti et al., fewer colonies in the lungs appear when RGD peptides are injected with tumor cells

[Ruoslahti, 1999].

ECM and Anoikis-

Anoikis, or cell death, occurs as a result of forcing anchorage-dependent cells into suspension [Ma, 2007]. Cells require integrin-dependent attachment for survival; therefore, anoikis occurs following the withdrawl of integrin-related survival signals

[Ma, 2007]. Constitutave activation of FAK, Src, Akt, and/or Ras aid in the ability of cells to bypass detachment-induced cell death [Ma, 2007]. In addition, integrins control

37 anoikis by sensing mechanical properties of the ECM [Ma, 2007]. It is thought that cells initiate a process to gauge substrate stiffness in an integrin-mediated inside-outside-in feedback loop, but there is little known about the components of this loop [Ma, 2007].

Integrin α5β1

Integrin Overview-

Integrins make up a family of protein receptors [Geiger, 2011]. They are membrane- spanning heterodimers consisting of one alpha subunit and one beta subunit [Pankov,

2000; Geiger, 2011], each with a molecular weight of 100-200 kDa [Ruoslahti, 1999].

There are 15 alpha subunits and 8 beta subunits, which combine to make 25 integrins

[Ruoslahti, 1999]. The majority of the peptide is outside the cell, and the cytoplasmic domains are about 30-50 amino acids long [Ruoslahti, 1999].

Integrins are bidirectional transmembrane signaling effectors [Quinn, 2009] with distinct functions [Geiger, 2011]. Integrins are important in the formation, maturation, and function of cell adhesions [Geiger, 2011], and they mediate cell migration [Pankov,

2000]. They signal via both outside-in and inside-out mechanisms. Outside-in signaling occurs when a matrix protein binds to the integrin on the outside of the cell, promoting cellular signaling inside the cell [Quinn, 2009]. Inside-out signaling occurs when proteins inside the cell bind to the cytoplasmic domain of the integrin, promoting cellular signaling outside the cell. Seven-transmembrane spanning receptor ligands have been shown to promote integrin clustering and the formation of focal adhesions [Quinn,

2009].

Inactive Activation Active Conformation Conformation

Ligand

Out

Inside Outside

Figure 1- Integrin signaling. Integrins signal via outside-in and inside-out mechanisms. Outside-in signaling occurs when matrix proteins binds to the integrin outside of the cell, promoting signaling inside the cell. Inside-out signaling occurs when proteins inside the cell bind to the cytoplasmic domain of the integrin, promoting events outside the cell.

Common ligands for integrins include collagens, laminins, FN, vitronectin, and fibrin

[Geiger, 2011]. There are several integrins that recognize ligands containing the RGD peptide including five αV integrins, α5β1, α8β1, and αIIbβ3 [Humphries, 2006]. These

RGD-binding integrins bind to a large number of ECM ligands [Humphries, 2006]. LDV binding integrins include α4β1, α4β7, α9β1, four β2 integrins, and αEβ7 [Humphries,

2006]. Laminin/collagen-binding integrins include α1, α2, α10, and α11 combined with

β1 [Humphries, 2006]. Other laminin-binding integrins include α3, α6, and α7 combined with β1, and α6β4 [Humphries, 2006].

Adhesion Structures-

Integrins form many types of adhesion structures, including focal complexes, focal adhesions, elongated fibrillar adhesions, and podosomes or invadopedia [Geiger, 2011].

Focal complexes are short-lived adhesions that transform into focal adhesions. They are

39 formed along lamellipodial protusions [Geiger, 2011]. Focal and fibrillar adhesions will be discussed in detail in the next sections, but in short, fibrillar adhesions are formed along matrix fibrils and are abundant under the central areas of the cell [Geiger, 2011].

Podosomes are small adhesions formed around active bundles, and invadopedia are thin protusins from the actin-cortactin core [Geiger, 2011]. Inactive integrins are distributed throughout the plasma membrane, but upon activation they localize in focal contacts and focal adhesions [Pankov, 2000]. After detachment of cells from their substrate, both contacts disappear [Pankov, 2000]. There have been over 50 proteins found in adhesions [Webb, 2012].

Functions of Adhesion Structures-

Adhesion structures have many functions. They are important in tissue and organ morphogenesis and in cascades of signaling events stemming from interactions with the

ECM [Geiger, 2011]. These signaling events effect cell proliferation, transcriptional activity, migration, and survival [Geiger, 2011].

Integrin α5β1 Overview-

Integrin α5β1 is the primary FN receptor. It is responsible for FN matrix assembly

[Pankov, 2000], and it is involved in the formation of focal and fibrillar adhesions

[Geiger, 2011]. It exists in at least four states, differing by ligand occupancy, degree of activation, and cell surface distribution [Pankov, 2000]. Integrin α5β1 is associated with migration and proliferation in cancer [Ruoslahti, 1999].

Role of Integrin α5β1 in FN Matrix Assembly-

Many integrins bind soluble FN, but cannot convert this FN into fibrils to the same extent as integrin α5β1 [Pankov, 2000]. Integrins such as αvβ3, αIIbβ2, and α5β1 have been shown to mediate matrix assembly when activated, and αvβ1 has been shown to temporarily replace α5β1 during embryogenesis of α5 null mice [Mao, 2005]. In addition, αvβ3 and αIIbβ3 both bind FN, but these integrins are less able to form a dense and delicate fibrillar network, which is thought to be due to their subcellular localization [Takahashi, 2007]. Furthermore, small GTPases such as RhoA are activated upon integrin α5β1 binding to FN, but not when αv integrins bind FN [Takahashi, 2007].

Integrin α5β1 is the major receptor in FN matrix assembly that is able to convert soluble

FN to its insoluble form [Pankov, 2000]. Integrin α5β1 is present in both focal and fibrillar adhesions, and FN matrix assembly is dependent on the translocation of activated integrin α5β1 to these adhesions [Pankov, 2005]. FN matrix assembly occurs when integrin α5β1 binds soluble FN and converts it into fibrils that make up the matrix

[Ruoslahti, 1999]. Specifically, integrin α5β1 binds the RGD motif of FN [Takahashi,

2007] and FN-FN interactions occur resulting in the formation of deocycholate (DOC)- soluble fibrils, which are then converted into a DOC-insoluble fibrillar network [Mao,

2005].

In an inactive state, integrin α5β1 has a low affinity for its ligand. Upon activation, integrin α5β1 is converted to a high affinity state for its ligand. Cells preparing for migration express integrin α5β1 in a low affinity state, while cells preparing for adhesion

41 express integrin α5β1 in a high affinity state. Leukocytes in circulation express low affinity integrin α5β1, but integrin α5β1 coverts to its high affinity conformation upon adherence of the cell to a site of injury during wound healing [Faull, 1993]. Furthermore, an erythroleukemic cell line binds FN with a low affinity, but the anti-β1 mAB 8A2 induces a 20 fold higher affinity conformation of integrin α5β1 for FN [Faull, 1993]. This affinity modulation is dependent on soluble FN concentrations [Faull, 1993]. This increase in affinity results in an increase in cell adhesion [Faull, 1993].

Cells transfected with integrin α5β1 survive longer when cultured without serum on FN

[Ruoslahti, 1999]. In addition, truncation of the integrin β1 subunit results in the abolishment of FN matrix assembly [Pankov, 2000]. Of most interest, E2 stimulation of human breast cancer cells promotes GPER-dependent recruitment of integrin α5β1 to focal and fibrillar adhesions [Filardo, 2002]. FN matrix assembly is increased when CHO cells are transfected with integrin α5, while FN matrix assembly is inhibited when cells are treated with monoclonal antibodies (mABs) against α5 or β1 subunits or an RGD peptide [Pankov, 2000]. When α5-null CHO cells are transfected with integrin α5, they gain the ability to form spherical aggregates when treated with exogenous FN [Caicedo-

Carvajal, 2010]. In addition, surface tension increases with moderate levels of integrin

α5 expression [Caicedo-Carvajal, 2010]. Interestingly, when integrin α5 expression was increased in CHO cells from a moderate level to a high level, there was a drop in surface tension, suggesting that there is an optimal level of integrin expression for FN matrix assembly and migration [Caicedo-Carvajal, 2010]. CHO aggregates expressing low or high levels of integrin α5 exhibit properties of motile cells, while CHO cells expressing

42 moderate levels of integrin α5 appear to be locked in place [Caicedo-Carvajal, 2010].

Furthermore, CHO cells expressing intermediate levels of integrin α5β1 generate a rich

FN matrix where fibers extend from cell-to-cell, while CHO cells expressing high levels of integrin α5β1 generate a reduced matrix with short fibers [Caicedo-Carvajal, 2010].

Integrin α5β1 and Migration-

Integrin α5β1 is essential in each step of migration [Ayoub, 2013]. Integrins regulate cell migration through coupling of cytoskeletal and signaling molecules in focal adhesions

[Ayoub, 2013]. FAK is an important signaling molecule that interacts with integrin α5β1 during migration [Ayoub, 2013], and FAK has the ability to influence protein tyrosine phosphorylation to initiate the reorganization of the cytoskeleton [Ayoub, 2013].

Integrin α5β1 and EGFR Transactivation-

Integrin α5β1 is a required intermediary for GPER-dependent EGFR transactivation

[Filardo, 2012]. Addition of soluble RGD peptide to SKBR3 breast cancer cells treated with E2β results in the inhibition of GPER-mediated EGFR tyrosine phosphorylation

[Quinn, 2009]. Additional evidence that integrin is required for EGFR transactivation involves Src. Src is required for general GPCR-mediated EGFR transactivation, and because Src also plays an important role in integrin activation, integrin α5β1 is likely required for GPER-mediated EGFR transactivation [Quinn, 2009]. Furthermore, GPER uses proteases to cleave and release HB-EGF, and integrin α5β1 provides a mechanism for localizing proteases at the cell surface [Quinn, 2009].

Integrin α5β1 and Breast Cancer-

Integrin α5β1 has importance in the maintenance of breast tissue and in breast cancer.

Faraldo et al. generated a transgenic mouse model to express a form of integrin β1 that lacks the ability to bind ECM proteins in mammary epithelium [Faraldo, 2001]. This mutant integrin β1 consists of cytoplasmic and transmembrane domains, but the extracellular domain contains a T-cell differentiation antigen, CD4, which acts as a dominant-negative mutant [Faraldo, 2001]. In cultured cells, this mutant affects cell spreading, migration, and matrix assembly [Faraldo, 2001]. In the mutant female mice there are mammary gland growth defects [Faraldo, 2001]. Reduced epithelial cell proliferation and increased apoptosis rates were also observed [Faraldo, 2001].

Furthermore, there was a decrease in phosphorylation of Erk-1 and Erk-2, which are mitogen-activated protein kinases (MAPKs) that are involved in the control of cell cycle progression [Faraldo, 2001].

Focal Adhesions

Focal Adhesion Overview-

Focal adhesions, also known as focal contacts, are adhesion sites that anchor stress fibers and provide cells with substrate attachment [Pankov, 2000]. Integrins cluster at focal adhesions and link actin filaments to the ECM [Ruoslahti, 1999; Angers-Loustau,

1999]. Other signaling molecules concentrate in focal adhesions, and many of them are phosphorylated at tyrosine residues [Ruoslahti, 1999]. Major structural proteins found in focal adhesions include, talin, tensin, and alpha-actinin. Major tyrosine kinases

44 include FAK, Src, and Csk, and these tyrosine kinases are activated upon ECM binding

[Angers-Loustau, 1999]. Adaptor proteins localized in focal adhesions include p130CAS,

Shc, Grb2, Crk, and Nck [Angers-Loustau, 1999]. Focal adhesions inhibit cell migration

[Webb, 2012].

Shc

CSK CAS Paxillin

FAK Vinculin

Src

Focal Adhesion Complex

Figure 2- Focal adhesion. Tyrosyl phosphorylated proteins localize to focal adhesions upon integrin binding of its ligand.

Formation of Focal Adhesions-

During the formation of focal adhesions, while laemellapoium and lamella advance, adhesions stay attached to the substrate, increase their length and total area, and associate with the termini of the actin stress fibers [Geiger, 2011]. The physical tension caused by stress fiber formation in the cells under the control of Rho GTPase results in integrin cluster association with α-actinin and tensin [Angers-Loustau, 1999]. These

45 changes are associated with alterations in protein composition [Angers-Loustau, 1999], such as activation of protein kinases, including FAK [Ruoslahti, 1999]. FAK is a cytoplasmic nonreceptor protein tyrosine kinase that binds to the cytoplasmic domain of integrins and is important in integrin signaling [Ruoslahti, 1999; Zheng, 2009].

Activated FAK provides a docking site for other signal transduction proteins [Angers-

Loustau, 1999], and results in the activation of downstream signaling molecules such as

Erk and PI3K/AKT [Zheng, 2009]. Cell migration leads to the turnover of focal adhesions

[Zheng, 2009].

Focal Adhesions to Fibrillar Adhesions-

The transformation of focal adhesions into fibrillar adhesions involves the translocation of integrin α5β1 out of focal contacts [Pankov, 2000]. It also involves an increase in isometric tension resulting in the stretching and unfolding of bound FN [Pankov, 2000].

Fibrillar Adhesions

Fibrillar Adhesions Overview-

Fibrillar adhesions, also known as ECM contacts, are elongated matrix contacts that are associated with FN fibrillogenesis [Geiger, 2011]. They bind FN parallel to actin bundles and tensin, and are lacking in focal adhesion proteins such as paxillin and vinculin

[Pankov, 2000].

Formation of Fibrillar Adhesions-

Fibrillar adhesions form when integrin α5β1 and tensin translocate away from focal adhesions, resulting in forces that promote FN fibril formation [Geiger, 2011]. While integrin α5β1 translocates, αv integrins, vinculin, and paxillin remain in place [Pankov,

2000].

Tensin is the major cytoskeletal protein in fibrillar adhesions [Pankov, 2000]. It is involved in the interaction between integrins and the cytoskeleton, and has actin- capping, filament-binding, and cross-linking sites that are important in the formation of fibrillar adhesions and the translocation of integrin α5β1 [Pankov, 2000]. In a study by

Pankov et al., cells with a dominant-negative form of tensin were able to form focal adhesions, but were not able to form fibillar adhesions [Pankov, 2000].

FN Fibrils

FN Fibril Overview-

FN fibrillogenesis is the conversion of soluble FN into insoluble FN fibrils in the ECM

[Huynh, 2013]. This is a multi-step process that begins with the binding of soluble FN dimers to integrins, followed by conformational changes of the bound FN [Huynh, 2013].

New fibrils are thin and DOC detergent-soluble, but then grow in length and thickness and become a DOC detergent-insoluble matrix [Huynh, 2013].

Formation of FN Fibrils-

Upon binding with FN, integrin α5β1 undergoes conformational changes resulting in increased receptor affinity for FN [Quinn, 2009]. FN fibril formation occurs following the

47 translocation of clustered, FN-occupied integrin α5β1 out of focal contacts and fibrillar adhesions along actin microfilament bundles [Pankov, 2000]. The cytoskeleton- generated forces due to integrin α5β1 translocation that are required for the initiation of FN fibrillogenesis [Pankov, 2000]. This process is dependent on integrin α5β1, actin,

FN, and tensin [Pankov, 2000].

In a study conducted by Robinson et al., integrin α5β1-mediated cell aggregation, compaction, and cohesion were correlated with FN matrix assembly [Robinson, 2004].

Cells cultured in the absence of external mechanical support have the ability to induce

FN matrix assembly which promotes integrin-mediated aggregate compaction and cohesion, whereas inhibition of FN matrix assembly results in cell dispersal [Robinson,

2004]. In an anchorage-independent growth model, no surface exists for FN fibril formation other than neighboring cells, suggesting that cell-FN-cell interactions generate enough tension to support FN fibril assembly in the absence of FN-substrate and cell-substrate adhesion [Robinson, 2004].

FN Fibrils and Cancer-

FN fibril formation is important in cancer progression. Some studies have shown that organization of FN into a network is implicated in the transition of dormancy to metastatic growth [Barkan, 2008]. Barkan et al. found that a metastatic mammary cell line transitioned from an inactive state to a proliferative state upon FN signaling through integrins [Barkan, 2008]. Quinn et al. show that E2 action via GPER promotes the spreading of human breast cancer cells and FN fibril formation [Quinn, 2009].

EGFR Transactivation

Integrins and Growth Factor Receptors-

Integrins and growth factor receptors are both linked to cancer independently, are present in cell adhesions, and have been shown to interact with eachother [Geiger,

2011]. It is accepted that members from the β1, αv, β7, and β4 integrin subgroups crosstalk with many growth factor receptors including EGFR, MET, PDGFR, and VEGFR

[Brizzi, 2012]. Integrins are required for growth factor receptor signaling and growth factor receptors are important in the regulation of integrin activation [Brizzi, 2012].

Specifically, integrin and growth factor receptor communication is important in receptor transactivation, receptor coordination, receptor pathway modulation, and receptor compartmentalization [Brizzi, 2012]. These interactions are important in normal and malignant cell proliferation, survival, and differentiation [Brizzi, 2012]. For example, β1 integrins have been shown to induce ligand-independent EGFR transactivation, while ligand-dependent EGFR signaling from the plasma membrane to the cytoskeleton continues [Brizzi, 2012]. Integrin β1 is required for sustained EGFR signaling in cancer cells and breast cancer cell growth and invasion [Brizzi, 2012].

GPER and EGFR Transactivation-

Many ligands including endothelin, thrombin, carbachol, and lypophsphatidic acid use

GPCRs to transmit signals via EGFR transactivation [Filardo, 2002]. For example, angiotensin II transactivates the EGFR through GPCR signaling to activate Erk1 and Erk2

[Thomas, 2005].

E2 results in increased concentrations of EGF [Filardo, 2005], and E2 activates MAPK via

GPER-dependent, EGFR transactivation, and c-fos expression in ER-negative breast cancer cells [Thomas, 2005]. EGFR transactivation occurs following the release of HB-

EGF from the cell surface via a Gβγ-Src-Shc signaling pathway [Thomas, 2004]. Details of this Gβγ signaling have not been sorted out yet, but it has been linked to the activation of matrix metalloproteinases (MMPs), which may be responsible for the cleavage and release of HB-EGF from the cell surface [Filardo, 2005]. Intrauterine administration of E2 resulted in an increase in EGF and an increase in EGFR phosphorylation [Filardo, 2005].

In addition, neutralizing antibodies to EGF resulted in the inhibition of E2-induced proliferation in the uterus [Filardo, 2005]. Furthermore, E2 signaling to Erk1 and Erk2 was blocked by specific inhibitors of EGFR tyrosine kinase and downregulation of pro-

HB-EGF from the cell surface by diphtheria toxin mutant, CRM-197 [Filardo, 2005]. Cells lacking GPER function due to a carboxyl-terminal deletion mutant blocks E2-dependent

EGFR transactivation [Quinn, 2009]. In addition, the ER antagonist, Faslodex, activates the EGFR in SKBR3 breast cancer cells, and expression of the truncated GPER blocks

EGFR transactivation [Quinn, 2009].

Shc

Shc Overview-

The phosphorylation and dephosphorylation of proteins on tyrosine residues is important in normal and malignant cell regulation [Habib, 1994]. Shc, an adaptor protein, is a substrate for tyrosine kinases following treatment of cells with growth

50 factors and hormones [Habib, 1994]. It is responsible for transmitting activation signals from the receptor or tyrosine kinases to downstream signaling components [Faisal,

2002]. It has no catalytic activity [Habib, 1994]. Three genes (ShcA, ShcB, and ShcC) encode Shc proteins [Faisal, 2002]. All three genes have an amino-terminal phosphotyrosine binding domain, a central proline/glycine rich region, and a carboxyl terminal Src homology 2 (SH2) domain [Faisal, 2002]. The central region contains two sites of phosphorylation at Tyr239/Tyr240 and Tyr313 residues [Zheng, 2012]. ShcA encodes three isoforms (p46, p52, and p66), which differ in their amino-terminal sequence and are produced through alternative splicing [Faisal, 2002]. Proteins p46 and p52 differ in an additional stretch of 45 amino acids at the amino terminus of p52

[Habib, 1994]. Protein p66 is the long isoform [Ma, 2007]. Proteins p46 and p52 are known to facilitate survival and proliferative signals, while p66 is known as a proapoptotic protein [Ma, 2007].

Shc regulates signaling events through mediation of the assembly of tyrosine phosphorylated signaling complexes via the SH2 and phosphotyrosine binding/phosphotyrosine interaction domains and through tyrosine phosphorylation on residue Tyr317 [Charest, 1996]. All three isoforms bind activated EGFR [Habib, 1994], and serine/threonine residue phosphorylation of p66 has been shown to occur as a result of epidermal growth factor (EGF) stimulation [Faisal, 2002]. Additional EGF- induced phosphorylation sites on Shc include Ser29, Thr214, Tyr240, Tyr313, and Ser334

[Zheng, 2012]. Shc has been shown to mediate the signaling output of EGFR [Zheng,

2012]. EGFR phosphorylation of Shc stimulates the Ras-Erk-MAPK and pI3K/AKT

51 pathways, which may mediate Shc feedback phosphorylation at Ser/Thr sites according to Zheng et al [Zheng, 2012]. Upon EGF stimulation, tyrosine phosphorylated Shc binds

Grb2, which stimulates mitogenic pathways [Zheng, 2012]. However, Shc-/- cells antagonize EGFR mitogenic signaling [Zheng, 2012]. Furthermore, treatment of cells with the EGFR inhibitor, AG1478, inhibited the phosphorylation of all Shc Tyr and

Ser/Tyr sites [Zheng, 2012].

Shc in an Unfamiliar Position-

It is widely accepted that Shc binds EGFR [Habib, 1994]. In addition, Shc associates with activated EGFR, insulin receptors, ErbB2, ErbB3, and Trk [Trub, 2005]. However, a recent study by Quinn et al. shows that Shc acts in a novel position in EGFR signaling, upstream of the EGFR [Quinn, 2009]. A mutant form of Shc lacking its primary tyrosine phosphorylation site at residue 317 inhibits E2-induced EGFR tyrosine phosphorylation, but not EGF-induced activation [Quinn, 2009].

Shc and GPER-

Shc and integrin β1 are both required for mammary gland homeostasis [Quinn, 2009].

Transgenic mice expressing an interfering mutant of integrin β1 have defects in mammary gland development and Shc signaling [Quinn, 2009]. In light of this fact, Quinn et al. show that integrin α5β1 and Shc are both necessary for GPER-mediated transactivation [Quinn, 2009]. Shc is required for GPER-dependent EGFR transactivation and FN fibril formation upon E2 stimulation in SKBR3 breast cancer cells [Quinn, 2009].

SKBR3 cells transfected with Shc317Y/F, a mutant form of Shc lacking its tyrosine

52 phosphorylation site, inhibits GPER-dependent FN matrix assembly and EGFR transactivation following treatment with E2 [Quinn, 2009]. Interestingly, after GPER stimulation of SKBR3 with E2, Shc 317 Y/F associated with integrin α5β1 more readily than WTShc or endogenous Shc isomers, suggesting that phosphorylation of Tyr317 is not required for the association of Shc with integrin α5β1 [Quinn, 2009]. A possible explaination for this observation is that phosphorylated 317 may result in Shc disengagement from integrin or dephosphorylation of this tyrosyl residue may cause

Shc-integrin disassembly [Quinn, 2009].

Shc and Integrin α5β1-

Shc is known to associate with integrins [Ma, 2007]. Specifically, association of Shc with

β1 integrins is known to occur indirectly through caveolin and Fyn through Fyn SH3 domain, through proline-rich motifs contained within the CH1 domain of Shc [Ma,

2007]. It is suggested that integrins activate the MAPK pathway via either a pathway involving FAK or a pathway involving Shc [Faraldo, 2001]. FAK and Shc signaling pathways have been shown to be activated independently, but function in parallel

[Faraldo, 2001]. Upon the binding of integrins to ECM proteins, FAK is autophosphorylated at Tyr397, resulting in the activation of Erk [Faraldo, 2001]. The autophosphorylation of FAK creates a binding site for Src kinase, which then phosphorylates FAK at Tyr925 [Faraldo, 2001]. The phosphorylation at Tyr925 results in the creation of a binding site for the Grb2 complex, which transmits signals to Ras, Raf, and Erk [Faraldo, 2001]. FAK activation is also followed by the phosphorylation of other

53 signaling molecules such as p130cas and paxillin [Faraldo, 2001]. There was a decrease in Erk activation in the mammary gland of transgenic mice expressing a β1 integrin mutant, but FAK activity is unaffected, suggesting that FAK cannot account for the decrease in Erk activation in these mice [Faraldo, 2001].

Shc may account for the decrease in Erk activation in transgenic mice expressing β1 integrin mutant [Faraldo, 2001]. Shc is an important intermediate of MAPK activation by integrins, as Shc-/- fibroblasts show a decrease in activation of Erk in response to ECM ligands [Faraldo, 2001]. Upon binding of integrins to ECM proteins, Shc is phosphorylated at Tyr317, resulting in the recruitment of the Grb2 complex [Faraldo,

2001]. Inhibition of β1 integrin function in the mammary gland of mice results in a decrease in Shc phosphorylation, failure to recruit Grb2, and failure to activate the Ras pathway [Faraldo, 2001].

Shc in Focal Adhesions-

Shc proteins are known to associate with integrins, but the role of Shc in integrin structures and binding mechanisms are not well understood [Ma, 2007]. Shc p52 and p66 have been shown to localize to focal adhesions. Cells lacking functional Shc have adhesion defects and changes in focal contact and actin fiber organization [Ma, 2007]. In one study, vector control 293 cells demonstrated small and peripheral focal contacts identified by DSRed-zyxin, and the cells were rounded with sparse actin stress fibers

[Ma, 2007]. When these same 293 cells were transfected with p66 Shc, the focal adhesions were more robust, and the cells were more spread with numerous stress

54 fibers [Ma, 2007]. Furthermore, human umbilical vein endothelial cells, which express high levels of endogenous p66 Shc, are polarized and have abundant focal adhesions; however, knockdown of p66 in these cells results in a loss of cell polarity and a decrease in stress fibers and focal adhesions [Ma, 2007].

Shc and Anoikis- p66 Shc is known as a proapoptotic protein and may target focal adhesions as a sensor for anoikis [Ma, 2007]. Vector control 293 cells form colonies in soft agar; however, when transfected with p66, colony formation is inhibited [Ma, 2007]. Furthermore, 293 cells transfected with a p66 S36E mutant are able to escape anoikis and grow in an anchorage-independent fashion [Ma, 2007].

Shc and PTPN12-

PTPN12 is a phosphatase that interacts with p52 and p66 Shc proteins, but not the p46 isoform [Habib, 1994]. Shc and PTPN12 interact at the NPLH sequence in the carboxyl terminus of PTPN12 [Faisal, 2002]. It appears that PTPN12 is a negative regulator of Shc because overexpression and antisense experiments show that PTPN12 down-regulates lymphocyte stimulation by dephosphorylation of Shc and downregulating the Ras pathway [Faisal, 2002]. The interaction of PTPN12 with Shc has been shown to be regulated by a GPCR agonist that activates kinase C [Habib, 1994]. In this case, PTPN12 would act as a suppressor of runaway activation of ligand-induced signaling [Habib,

1994].

Shc and PTPN12 are both implicated in EGFR signaling. Feedback phosphorylation of Shc requires PTPN12, which occupies the Shc phosphotyrosine-binding domain (PTB) domain through the NPLH motif [Zheng, 2012]. PTPN12 antagonizes pro-mitogenic EGFR signaling by displacing Shc from the EGFR and by dephosphorylating its Grb2 binding

XYN motifs [Zheng, 2012].

Protein Tyrosine Phosphorylation Signaling

Tyrosine signaling networks consist of an interconnected web of substrates that are regulated by tyrosine kinases and phosphatases [Albeck, 2011]. When one kinase or phosphatase is inhibited, the whole network can collapse [Albeck, 2011]. The web can also be flexible, depending on the cell’s profile of kinases and phosphatases [Albeck,

2011]. Tyrosyl phosphorylation of signaling molecules can alter their structure and function, enzyme activity, protein localization, and assembly of other signaling complexes [Zhang, 2009]. Furthermore, altered regulation of phosphorylation can result in altered protein-protein interactions, protein instability, and abnormal protein activity, which can lead to disease such as cancer [Zhang]. Protein tyrosine phosphorylation plays an important role in cancer [Sun, 2011]. Specifically, phosphorylation is important in cell growth, division, differentiation, adhesion, motility, and death [Zheng, 2013]. Abnormal activation of EGFR occurs commonly in breast cancer [Sun, 2011]. In addition, tyrosine kinase inhibitors that target three or more tyrosine kinases are used in the treatment of cancer [Albeck, 2011].

Protein Tyrosine Phosphatase

PTP Families-

There are over 100 protein tyrosine phosphatases (PTPs) encoded in the human genome

[Zheng, 2013]. There are three groups within the PTP superfamily including phosphor- tyrosine-specific PTPs, dual specificity phosphatases that dephosphorylate both phospho-serine/phospho-threonine and phospho-tyrosine residues in proteins, and low molecular weight PTPs [Zheng, 2013]. There are about 40 PTPs that make up the classical phospho-tyrosine specific PTP group within two subfamilies- transmembrane receptor-like proteins and non-receptor proteins [Zheng, 2013]. Receptor-like PTPs are found at the plasma membrane, while non-receptor PTPs are found in the cytosol, plasma membrane, and endomplasmic reticulum [Zheng, 2013].

PTP Structure-

All PTPs have a core structure composed of a central parallel beta-sheet with flanking alpha-helices containing a beta-loop-alpha-loop with the PTP signature motif, but they have differences in their active site pockets and their immediate surrounding environment for substrate specificity [Zheng, 2013]. Regulatory sequences near the catalytic domain control catalytic activity by interacting with residues at active sites

[Zheng, 2013]. The structural diversity of PTPs accounts for the differences in regulation and function [Garton, 1994].

PTP Locations-

PTPs regulate tyrosine phorphorylation as antagonists to tyrosine kinases [Sun, 2011].

PTPs have activites one to three times greater than protein tyrosine kinases [Garton,

1994]. PTPs have different subcellular locations, which limits the range of substrates

[Garton, 1994]. However, there are some PTPs that are cytosolic, which are less restricted in their available substrates and are subject to strict control [Garton, 1994].

PTP1B is targeted to the endomplasmic reticulum, and SH-PTP2 is targeted to membrane complexes by binding to specific phosphotyrosine-containing sequences in growth factor receptors via SH2 domains [Garton, 1994]. TCPTP can be nuclear or targeted to the endoplasmic reticulum, depending on the splice form expressed

[Garton, 1994]. PTPH1 is thought to be localized to the cytoskeleton because it has sequences similar to those of several cytoskeleton-associated proteins [Garton, 1994].

Some PTPs require activation [Garton, 1994]. SHP1 and SHP2 are relatively inactive in vitro, but can be activated by C-terminal truncation, addition of certain phospholipids, or SH2 domain-mediated binding of phosphotyrosine-containing peptides [Garton,

1994]. PTPN12, on the other hand, is highly active in vitro, and because it is soluble, may act on many substrates in the cytoplasm [Garton, 1994].

PTP Functions-

PTPs have many functions, and they often play a role in tumor suppression [Sun, 2011].

Many PTPs, including SHP1, SHP2 (PTPN11), PTPα, and PTPN12, are involved in focal adhesion disassembly and cell migration [Zhang, 2009]. Specifically, PTPN11 regulates focal adhesions, cell spreading, and migration [Zheng, 2013]. Dominant-negative SHP2

58 mutants have altered receptor tyrosine kinase signaling and integrin-dependent activation of the Erk pathway, gene activation, focal adhesion and stress fiber turnover, cell spreading and migration, and cell proliferation [Zheng, 2013]. Furthermore, EGF- induced Ras activation is inhibited in SHP2-null cells [Zheng, 2013]. The increased stress fiber and focal adhesion formation in SHP2-null cells may be due to an increase in Rho activity [Zheng, 2013].

Overexpression of a dominant-negative form of PTP1B resulted in a defect in integrin adhesion and signaling in fibroblasts [Angers-Loustau, 1999]. A transmembrane PTP,

LAR, has been shown to translocate to focal adhesions and promote their disassembly

[Angers-Loustau, 1999]. PTPα has been shown to regulate Src activity and cellular adhesion [Angers-Loustau, 1999]. Yersinia YopH, a bacterial PTP, has been shown to dephosphorylate p130Cas and FAK, resulting in disruption of focal adhesion formation

[Angers-Loustau, 1999].

PTPN12 Overview and Structure-

PTPN12 is a member of the non-receptor PTP subfamily and is located in the cytoplasm

[Zheng, 2013]. It is ubiquitously expressed in many tissues and cell types including the hematopoietic system, particularly in the thymus, spleen, and liver, but it is also found in the brain and heart [Zheng, 2013].

PTPN12 is located in 7q11.23 [Xunyi, 2002]. It contains 780 amino acids, with a molecular weight of 112 kDa [Zheng, 2013]. It has 6 protein-protein binding domains, including four proline rich domains numbered P1 to P4, an NPLH motif, and the CTH

59 domain [Cousin, 2004]. It also has a tyrosine phosphatase NH2-terminal catalytic domain (amino acid residues 1-300) with the sequence 231CSAGC, which is vital for its catalytic activity [Angers-Loustau, 1999]. The sequence 229-IHCSAGCGRTG-239, also in the N-terminal domain, conforms to the conserved phosphatase signature motif

(I/V)HCXAGXXR(S/T)G [Zheng, 2013]. The cysteine residue within this motif is required for its PTP activity [Zheng, 2013]. The protein-protein binding domains, also known as the COOH-terminal domain (amino acid residues 304-775), has PEST sequences [Zheng,

2013]. P1 (amino acid residues 332-348) binds the Src homology 3 (SH3) domain of p130Cas, the SH3 domain of Hefl, and the SH3 domain of Sin [Cousin, 2004]. P2 (amino acid residues 647-690) binds SH3 domain of Csk and the N-SH3 domain of Grb2 [Cousin,

2004]. P3 (amino acid residues 355-374) binds LIM3/LIM4 of paxillin and LIM3 of Hic5

[Cousin, 2004]. NPLH (amino acid residues 599-602) binds the PTB domain of Shc

[Cousin, 2004]. CTH binds coiled coil of PST-PIP/CD2BP1 and PST-PIP2 [Cousin, 2004].

Catalytic domain Polyproline rich region N PTP Cas Pax P3 Shc Csk PEST C

Figure 3- PTPN12 structure. PTPN12 has a tyrosine phosphatase NH2-terminal catalytic domain, four proline rich domains, an NPLH motif, and a PEST sequence.

Regulation by PTPN12-

PTPN12 is important in the regulation of proteins involved in focal adhesion formation such as FAK, PYK2, p130Cas, and paxillin [Zheng, 2013]. These proteins become tyrosine- phosphorylated in response to the activation of the integrin signaling pathways, and

60 dephosphorylated in response to the activation of PTPN12 [Zheng, 2013]. Thus, PTPN12 controls the rate of focal adhesion turnover and cell migration [Zheng, 2013].

Overexpression of PTPN12 reduces integrin and platelet-derived growth factor induced phosphorylation of p130Cas and inhibits motility [Zheng, 2013]. PTPN12-deficient fibroblasts demonstrate increased phosphorylation of p130cas, paxillin, and FAK [Zheng,

2013].

PTPN12 is important in the regulation of Rho GTPase, which is important in the coupling of membrane protusions and tail retraction during cell migration [Zheng, 2013].

PTPN12-deficient fibroblasts have enhanced membrane protusions at the leading edge and long, unretracted tails in the rear, and are unable to migrate [Zheng, 2013]. The exhaggerated membrane protusions at the leading edge are mediated by enhanced

Rac1 activity, while the long, unretracted tails in the rear are mediated by decreased

RhoA activity [Zheng, 2013].

PTPN12 is involved in the regulation of Src, Abl, Wiskott-Aldrich syndrome protein

(WASP), and filamin-A. PTPN12 also interacts with C-terminal Src kinases (Csk), which phosphorylates Src Y527, inhibiting Src activation [Zheng, 2013]. Overexpression of

PTPN12 results in the inactivation of c-Src via dephosphorylation of a positive regulatory tyrosine [Zheng, 2013]. C-Abl is important in membrane ruffling in response to growth factor stimulation [Zheng, 2013]. It also antagonizes migration [Zheng, 2013]. PSTPIP connects PTPN12 to c-Abl, resulting in the dephosphorylation of c-Abl by PTPN12

[Zheng, 2013]. C-abl phosphorylates PSTPIP, a substrate of PTPN12, and PTPN12

61 dephosphorylates PSTPIP [Zheng, 2013]. In addition, PSTPIP binds to WASP and the activation of WASP results in actin nucleation and the formation of filopodia and lamellipodai, both of which are important in cell migration [Zheng, 2013].

Shc is regulated by PTPN12. PTPN12 associates with Shc, which is important in growth factor signaling and motility [Zheng, 2013]. Insulin has been shown to induce the association between Shc and PTPN12 [Zheng, 2013].

As discussed earlier, FAK is implicated in the formation of focal adhesions, and the inhibition of FAK promotes cell migration, invasion, and metastasis [Angers-Loustau,

1999]. Overexpression of PTPN12 results in the dephosphorylation of FAK at Y397, promotion of disassembly and turnover of focal adhesions, and promotion of Ras- induced migration [Zheng, 2013], while overexpression of catalytically inactive mutants

PTPN12 C231A and PTPN12 D199A did not result in the dephosphorylation of FAK at

Y397 [Zheng, 2013]. p130Cas is a substrate for PTPN12 [Garton, 1994]. Inactivated p130Cas is located in the cytosol, while phosphorylated p130Cas is located in focal adhesions [Garton, 1994].

P130Cas has been shown to interact with FAK [Garton, 1994]. P130Cas is phosphorylated during integrin-mediated cell adhesion [Garton, 1994]. PTPN12 dephosphorylates p130Cas, which is important in inhibiting the transforming effects of oncogenes [Garton, 1994]. p50Csk, a 50 kDa cytoplasmic tyrosine protein kinase, is expressed ubiquitously, and contains an SH3 region, an SH2 domain, and a catalytic domain [Davidson, 2001]. It is

62 responsible for inhibiting several cellular responses [Davidson, 2001]. It phosphorylates the inhibitory carboxyl-terminal tyrosine of Src family tyrosine protein kinases, which promotes the association of the carboxyl terminus and the SH2 domain of Src-related enzymes to repress their enzymatic activity [Davidson, 2001]. Csk has been shown to inhibit G protein-mediated activation of MAPK, and it has been shown to bind tyrosine- phosphorylated focal adhesion proteins including paxillin and p125FAK [Davidson,

2001]. The SH3 domain of Csk binds a proline-rich sequence of PTPN12 [Davidson,

2001]. Together, PTPN12 and p50csk inhibit intracellular tyrosine protein phosphorylation [Davidson, 2001].

Regulation of PTPN12-

PTPN12 activity can be regulated at the transcriptional level by gene deletion or mutation, or via post-translational modifications such as phosphorylation, oxidation, and caspase-dependent cleavage or proteolysis [Zheng, 2013].

Phosphorylation of PTPN12 affects its catalytic activity and access to substrates [Zheng,

2013]. Activation of protein kinase C (PKC) and cAMP-dependent protein kinases results in the phosphorylation of PTPN12 at S435 in the non-catalytic domain and S39 in the catalytic domain [Zheng, 2013]. This phosphorylation of PTPN12 results in the downregulation of enzymatic activity [Zheng, 2013]. Garton et al. studied reversible phosphorylation of PTPN12 and identified Ser39 on PTPN12 as an inhibitory phosphorylation site [Garton, 1994]. Protein kinases PKA and PKC, which are known to have overlapping substrate specificities, phosphorylate Ser39 in vitro, but the sequence

63 surrounding Ser39 (RRLSTK) is not that of a typical PKA phosphorylation site [Garton,

1994].

PTPN12 can also be regulated by oxidation via dephosphorylation of a catalytically essential cystein residue that is highly sensitive to oxidation [Zheng, 2013]. Oxidation blocks nucleophilic activity of PTPN12 and inactivates it in a reversible manner [Zheng,

2013]. PTPN12 inhibition by oxidation relives the inhibitory effects on paxillin, which is one of the mechanisms by which reversible oxidation may promote tumor migration and invasion [Zheng, 2013].

Proteolysis is important in the regulation of PTPN12 activity [Zheng, 2013]. Apoptosis is important in cancer development and progression, and PTPN12 may be involved in apoptosis amplification [Zheng, 2013]. PTPN12 has been shown to be regulated by proteolysis during apoptosis [Zheng, 2013].

PTPN12 and Focal Adhesions-

While FAK and Src have been shown to promote focal adhesion formation, PTPN12 has been shown to be involved in the breakdown of focal adhesions and stress fibers

[Angers-Loustau, 1999]. Plating COS-1 cells transfected with PTPN12 on FN resulted in the movement of PTPN12 to the membrane periphery, and this is thought to occur after integrin activation [Angers-Loustau, 1999]. In addition, PTPN12 dephosphorylated several focal adhesion-associated proteins, including p130Cas [Angers-Loustau, 1999].

Furthermore, in PTPN12-/- cells, several focal adhesion proteins were hyperphosphorylated, including FAK, paxillin, and p130Cas, and there was an increase in

64 spreading rate on FN [Angers-Loustau, 1999]. In PTPN12 -/- fibroblasts, there was an increase in focal adhesions when compared to the parental fibroblasts [Angers-Loustau,

1999]. A study using phenylarsine oxide, a PTP inhibitor that reacts with two thiol groups of cystine residues in the active site of the phosphatase, showed that treatment of cells with this inhibitor resulted in increased stress fiber formation even after starvation [Angers-Loustau, 1999].

PTPN12 and Migration-

PTPN12 plays a role in all steps of migration including membrane protrusions, tail retraction, and regulation of focal adhesions [Zheng, 2013]. It binds to and regulates proteins in the migration complex [Zheng, 2013]. PTPN12 can have both positive and negative effects on migration [Zheng, 2013]. It regulates the actin cytoskeleton polymerization through its interaction with WASP via PSTPIP, and by the formation of lamelliposia and cell retraction through Rac1 and RhoA via the regulation of upstream effects such as RhoGAB [Ayoub, 2013]. Furthermore, PTPN12 -/- cells show an increase in focal adhesion formation, which might explain why migration of these cells was inhibited in a wound healing assay [Angers-Loustau, 1999]. Furthermore, PTPN12+/- cells are round and demonstrate punctuate focal adhesions at the tips of their membranes, which is suggestive of migration [Angers-Loustau, 1999]. Overexpression of

PTPN12 in rat fibroblasts results in a decrease in cell migration and an increase in the time of cellular spreading on FN [Angers-Loustau, 1999; Davidson, 2001].

PTPN12 and Cancer Overview-

PTPN12 is implicated in cancer. It has been associated with cancer growth, proliferation, and metastasis [Wu, 2013]. It has been investigated in colon cancer, esophageal ovarian cancer, squamous cell carcinoma, breast cancer [Wu, 2013], ovarian cancer, and prostate cancer [Xunyi, 2002].

PTPN12 and Breast Cancer-

Studies have shown that loss of PTPN12 can lead to malignant transformation of human mammary epithelial cells, and restoration of PTPN12 into these cells inhibits proliferation, tumorigenicity, and metastatic potential [Xunyi, 2002]. In addition,

PTPN12 is down-regulated in breast cancer cells when compared to normal breast tissue

[Sun, 2011; Wu, 2013; Xunyi, 2013]. In one study, PTPN12 expression was inhibited in

32% of breast cancer cases compared to 11.3% in paired adjacent non-tumor tissues

[Xunyi, 2002]. Another study showed that 22.6% of breast cancer tumors have a deletion in PTPN12 and 37% of invasive breast cancers have low PTPN12 expression

[Sun, 2011; Zheng, 2013]. PTPN12 was also undetectable in 9.1% of HER2 amplified tumors and 60.4% of TNBC tumors [Sun, 2011]. Amino acid mutations at positions I322,

A573, and K709 on PTPN12 were found in primary human breast tumors and in breast cancer cell lines [Zheng, 2013].

Low PTPN12 expression is positively associated with lymph node status, stage, and distant metastatic relapse in breast cancer [Wu, 2013]. In addition, Yuan et al. report that low PTPN12 expression is associated with increased tumor and lymph node size, lymph node metastasis, distant metastasis, and histological grade [Xunyi, 2013; Xunyi,

2002]. Patient outcomes after surgery are poorer in patients with low PTPN12 expressing tumors than patients with high PTPN12 expressing tumors [Xunyi, 2002].

Furthermore, loss of PTPN12 leads to acinar morphogenesis and cellular transformation in mammary epithelial cells [Sun, 2011]. Silencing of PTPN12 has been shown to result in the inhibition of the tumor suppressor phosphatase and tensin homolog (PTEN) [Villa-

Moruzzi, 2013]. In addition, the PI3K/AKT signaling pathway, which is linked to enhanced migration, is enhanced in PTPN12 silenced cells [Villa-Moruzzi, 2013].

Furthermore, PTPN12 has been linked to migration of breast cancer cells via FAK signaling [Villa-Moruzzi, 2013]. FAK is commonly upregulated in cancer cells, and can act as both a kinase and a platform to recruit signaling molecules to focal adhesions [Villa-

Moruzzi, 2013]. FAK undergoes cycles of activation and inactivation by PTPN12 during the migration of cancer cells [Villa-Moruzzi, 2013]. Results from experiments conducted by Sun et al. suggest that PTPN12 is a tumor suppressor [Sun, 2011]. The restoration of

PTPN12 in breast cancer cells inhibits proliferation, tumorigenicity, and metastatic potential [Sun, 2011]. In addition, overexpression of PTPN12 is associated with longer disease free survival [Wu, 2013].

PTPN12 inactivation may occur through two mechanisms- genetic change and epigenetic change [Xunyi, 2002]. Aberrant methylation of CpG islands is common in tumor suppressor gene silencing and leads to transcription repressor binding, compressed chromatin, and transcription silencing [Xunyi, 2002]. Promoter CpG island hypermethylation occurs more frequently in cell lines and tissue specimens with low

PTPN12 expression [Xunyi, 2002].

While many studies suggest that PTPN12 deletion results in increased cell migration, one study found that both overexpression and inhibition of PTPN12 results in decreased cell motility [Zheng, 2013]. This suggests that there is a balance of PTPN12 function that is required for the regulation of cell adhesion and focal adhesion turnover [Zheng,

2013].

Deletions in PTPN12, defective PTPN12 sequence variants, or loss of expression of

PTPN12 are frequent in TNBC [Sun, 2011]. Based on the results from a study by Albeck et al., 60% of TNBC do not express PTPN12 [Albeck, 2011]. Decreased PTPN12 expression is associated with lymph node status and distant metastatic relapse in TNBC

[Wu, 2013]. Furthermore, according to a study conducted by Wu et al., 60.2% of non-

TNBC tumors overexpress PTPN12, while only 39.3% of TNBC tumors overexpress

PTPN12 [Wu, 2013].

Studies show that PTPN12 suppresses TNBC cell transformation by interacting with EGFR and HER2 [Zheng, 2013]. PTPN12 silencing leads to EGFR hyperactivity in HER2-negative,

TNBC [Xunyi, 2002]. PTPN12 may oppose the activity of the EGFR via dephosphorylation

[Zheng, 2013].

PTPN12 has been linked to the EGFR through binding to the SH3 domain of Grb3 via its first proline rich domain and to Shc via its NPLH domain [Ayoub, 2013]. It is a suppressor of mammary epithelial cell transformation and of breast cancer cell proliferation and metastasis through its interaction with the EGFR and HER2 [Faraldo, 2001]. PTPN12 blocks EGFR function by dephosphorylation [Faraldo, 2001]. Inhibition of the EGFR is a

68 method of controlling cell proliferation, survival, and tumorigenesis [Sun, 2011].

Inactivation of PTPN12 in HER2-negative breast cancer is common- 96% of PTPN12 deficient breast cancers are HER2-negative [Sun, 2011]. Another study found that only

9% of HER2 positive cancers are PTPN12-negative [Albeck, 2011]. PTPN12 is involved in a network of serial negative regulation with alternating oncogenes and tumor suppressors consisting of b-TRCP, TEST, and miR-124, which inhibits receptor tyrosine kinases such as EGFR [Sun, 2011]. Sun et al. show that PTPN12 inactivation leads to

HER2/EGFR hyperactivity and cellular transformation in breast cancer [Sun, 2011].

Normally, PTPN12 interacts with HER2 and EGFR receptors, so loss of PTPN12 function leads to hyperactivation of the HER2 receptor signaling [Sun, 2011]. PTPN12 inactivation leads to mammary epithelial cell transformation that is partially dependent on HER2 and

EGFR [Sun, 2011], as HER2 activity is inhibited in PTPN12-deficient breast cancers when

PTPN12 function is restored [Sun, 2011]. PTPN12 may play an important role in the treatment of HER2-negative tumors [Sun, 2011].

A potential mechanism of EGFR activation-induced tumor cell migration involves

PTPN12 [Zheng, 2013]. Activated EGFR has reduced FAK activity mediated by PTPN12 dephosphorylation, decreasing the number of focal contacts, resulting in motile cells

[Zheng, 2013].

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Chapter 2

Aim 1

Abstract

Stimulation of estrogen receptor (ER)-negative human breast cancer cells with 17β- estradiol (E2β) results in fibronectin (FN) matrix assembly and release of heparan bound-epidermal growth factor (HB-EGF) via the G-protein-coupled estrogen receptor,

GPER. This mechanism of action results in the recruitment of FN-engaged integrin α5β1 to fibrillar adhesions and the formation of integrin α5β1-Shc adaptor protein complexes.

Here, we show that GPER stimulation of murine 4T1 or human SKBR3 breast cancer cells promotes the formation of focal adhesions, actin stress fibers, and results in increased cellular adhesion and haptotaxis on FN, but not collagen. These actions are also induced by the xenoestrogen, bisphenol A, and the ER antagonist, ICI 182, 780, but not the inactive stereoisomer, 17α-estradiol (E2α). In addition, we show that GPER stimulation of breast cancer cells allows for FN-dependent, anchorage-independent growth and FN fibril formation in hanging drop assays, indicating that these GPER-mediated actions occur independently of adhesion to solid substrata. Stable expression of Shc mutant

Y317F lacking its primary tyrosyl phosphorylation site disrupts E2β-induced focal adhesion and actin stress fiber formation, and abolishes E2β-enhanced haptotaxis on

FN, and anchorage-dependent growth. Collectively, these data demonstrate that E2 action via GPER enhances cellular adhesivity and FN matrix assembly and allows for anchorage-independent growth, cellular events that may allow for cellular survival and tumor progression.

Introduction

Fibronectin (FN) plays a major role in cellular adhesion, growth and survival, and it is important in processes such as wound healing [Stoffels, 2013], vascular growth [Astrof,

2009], and embryonic development [Francis, 2002]. FN is synthesized in a soluble form as a dimeric glycoprotein that is assembled into an insoluble fibrillar matrix in a complex, dynamic cell-mediated process that is initiated by its specific recognition via individual Arg-Gly-Asp (RGD) binding sites on each monomer by integrin α5β1 that facilitates integrin clustering. Upon FN engagement, integrin α5β1 undergoes conformational alterations associated with increased receptor affinity [Faull, 1993]. FN- occupied integrin α5β1 is then recruited to sites of close cell-matrix contact known as focal adhesions that are enriched in tyrosyl phosphorylated proteins and actin stress fibers where robust anchorage to FN occurs. The local concentration of integrin-bound

FN increases, allowing bound FN molecules to more readily interact with one another and form short FN fibrils between cells. Conversion of soluble FN to insoluble fibrils proceeds when cryptic FN-binding sites are exposed along the length of bound FN by contractile forces that stretch FN by pulling on their FN-bound integrin receptors

[Hocking, 2000] and partially unfolding FN, unmasking cryptic FN-binding sites [Ingham,

1997; Klotzsch, 2009], and allowing nearby FN molecules to associate. This FN-FN interaction enables the soluble, cell-associated fibrils to branch and stabilize into an insoluble FN matrix. Fragmentation of FN uncovers nonRGD binding sites leading to enhanced integrin α5β1 adhesion and FN matrix contractility [Valenick et al, 2005], illustrating the influence of matrix proteases on provisional FN matrix assembly.

Altered expression of FN or perturbations in the specific recognition of FN by integrin

α5β1 has been associated with the development of cancer and fibrosis. A number of studies have shown that FN is critical to normal homeostasis of the mammary gland and is associated with the development of breast cancer. The addition of exogenous FN negatively impacts acinar differentiation in the mammary gland and creates a microenvironment conducive to the growth of mammary epithelia [Williams, 2008].

Integrin α5β1 and FN are prominently expressed in the mammary gland and their basal expression is increased during active proliferation of mammary gland tissue in mice suggesting that this FN-integrin interaction may be required for hormone-dependent proliferation in the mammary gland [Woodward, 2001]. Transgenic mice expressing dominant-negative integrin β1 show disrupted mammary gland development that is associated a loss of AKT activation and Shc-dependent extracellular regulated kinase-1 and -2 (Erk-1/-2) activation [Faraldo, 2002]. Moreover, successful implantation of human mammary tumor xenografts in immunocompromised mice is facilitated by coadministration of exogenous FN, indicating a survival advantage for tumor cells that interact with FN [Price, 1996]. This observation is supported by studies that have shown that mammary adenocarcinoma cells are capable of converting soluble FN into fibrils

[Saulnier, 1996] resulting in increased responsiveness to growth factors and enhanced anchorage-independent growth [Qiao, 2000].

In general, parenchymal cells require integrin-dependent attachment to solid substrata for their survival as their detachment results in rapid apoptosis, a process referred to as anoikis [Frisch, 1996; Francis, 2002]. In contrast, tumor cells are often capable of growing in an anchorage-independent manner, an attribute that best predicts their metastatic potential upon transplantation in mice [Quinn, 2009]. The survival of tumor cells under these imposed in vitro growth conditions is partly reflective of their capacity to assemble a provisional extracellular matrix [Saulnier, 1996] as well as their capacity to circumvent death signals sensed by mechanosensors that report reduced tensile forces

[Discher, 2005]. As measured in a two-dimensional environment, ligation of integrin

α5β1 to FN-coated substrata is sufficient to promote intracellular signals associated with cellular growth and survival, including activation of Src, focal adhesion kinase (FAK),

B/AKT, and Erk-1/-2 [Francis, 1994; Frisch, 1996; Green, 2009]. Stimulation of adherent cells with serum-derived factors results in the recruitment of tyrosyl phosphorylated proteins (FAK, src, vinculin and paxillin) into focal adhesion plaques and the formation of

RhoA-dependent actin stress fibers [Ridley, 1992]. However, matrix engagement by integrin is not sufficient to promote subsequent cell growth responses and FN polymerization is a critical requirement for measurable adhesion-dependent growth on planar surfaces [Sottile, 1998]. Regulatory roles for phosphatidylinositol 3-OH kinase,

FAK, Src-like kinases and phospho-paxillin in FN matrix assembly have been suggested

[Wierzbicka-Patynowski, 2002; Wierzbicka-Patynowski, 2007]. Studies evaluating FN assembly in a three-dimensional environment have shown that exogenous FN can facilitate fibrillogenesis [Mao, 2005], but that this cellular activity is not sufficient for

84 anchorage-independent growth by mammary adenocarcinoma cells, as they also must become responsive to external growth factors [Saulnier, 1996; Qiao, 2000; Accornero,

2010].

The adaptor protein Shc is intrinsically involved in intracellular signaling events that determine growth factor responsiveness and bidirectional integrin signaling

[Ravichandran, 2001]. Shc is expressed as three isoforms that share a carboxyl terminal, are mapped to a single gene locus, and designated as p66Shc, p52Shc, and p46Shc that occur as a result of differential ribosomal initiation. Shc proteins are substrates for receptor and nonreceptor tyrosine kinases, and each possesses both PTB and SH2 phosphotyrosine binding domains that act to dictate its interaction with signaling effectors and form signaling complexes. Whereas p52Shc facilitates survival and proliferative signals, primarily through Ras recruitment and activation, p66Shc is best known as a proapoptotic protein. Both p52Shc and p66Shc traffic to focal adhesion plaques; however, p66Shc recruitment to focal adhesions is associated with RhoA- dependent anoikis [Ma, 2007]. This observation is consistent with the fact that p66Shc is poorly expressed in hematopoietic lineage cells that are considered to be anchorage- independent and insensitive to substrate stiffness [Migliaccio, 1997; Bonati, 2000;

Zhang, 2003; Giles, 2009]. Moreover, lung carcinoma cell lines that lack p66Shc display aggressive metastatic behavior and bypass anoikis [Ma, 2010]. Finally, patients whose breast or colon cancers express higher ratios of tyrosyl phosphorylated Shc to p66Shc

85 are linked to poor prognosis [Frackelton, 2006; Grossman, 2007], further suggesting that failure to regulate Shc is associated with more advanced cancer.

Tumors that arise from the mammary gland exhibit biological behaviors that are described dichotomously as either estrogen- or growth factor-dependent. This categorization is largely derived from analysis of known receptors for estrogen receptors

(ERs) and epidermal growth factor receptors (EGFR) in breast tumor biopsies, and the fact that there is a strong inverse relationship between expression of ER and EGFR

[Filardo, 2006]. G-protein-coupled estrogen receptor, GPER, represents a newly appreciated estrogen receptor whose expression in primary breast tumors is directly linked to tumor size and metastasis [Filardo, 2006; Arias-Pulido, 2010; Ignatov, 2011;

Steiman, 2013], a relationship diametrically opposed to the one shared between ER and these same tumor progression variables. This observation suggests that GPER plays a distinct role from ER in breast cancer biology and is consistent with the fact that GPER and ER are structurally distinct receptors that promote estrogen-mediated signals measured with different metrics and kinetics. While ER bears structural homology shared by the members of the nuclear steroid hormone receptor superfamily and functions as a hormone-inducible transcription factor, GPER belongs to the most broadly studied class of cell surface receptors, the G-protein-coupled receptor (GPCR) superfamily. GPER promotes rapid signals attributed to this receptor class, including stimulation of adenylyl cyclase [Filardo, 2002] and EGFR activation via the release of membrane-tethered HB-EGF [Filardo, 2000]. We have previously shown that EGFR

86 transactivation by GPER requires activation of the FN receptor, integrin α5β1, in breast cancer cells, as measured by its recruitment to fibrillar adhesions, the conversion of soluble FN to a detergent- insoluble form, and the association of integrin α5β1 with the signaling adaptor, Shc [Quinn, 2009].

Here, the influence of this GPER-integrin α5β1-Shc dependent signaling mechanism on breast cancer cell adhesion was further evaluated by measuring its influence on cancer cell cytoarchitecture, adhesion and haptotactic responses on immobilized FN, and to evaluate their role in fibrillogenesis and growth in anchorage-independent conditions.

Materials and Methods

Cell Culture- SKBR3 (ERα-, ERβ-, GPER+) breast cancer cells were obtained from the

American Type Culture collection (ATTC) (Manassas, VA). SKBR3 variants expressing dominant-negative Shc and dominant-negative GPER were generated as described previously [Quinn, 2009]. ER-negative murine 4T1 breast cancer cells were obtained from the ATCC. 4T1 cells expressing dominant-negative GPER-1 was generated as described previously [Quinn, 2009]. All cells were grown in phenol red-free (PRF)

DMEM/Ham’s F12 media (1:1) with 5% fetal bovine serum and 25 µg/ml gentamicin.

Growth factors, estrogens, antiestrogens, and matrix proteins- Water-soluble 17β- estradiol (E2β), 17α-estradiol (E2α), and angiotensin II (ATII) were purchased from Sigma

(St. Louis, MO). Bisphenol A (BPA) was a kind gift from the Hixon Lab at Brown

University (Providence, RI). ICI 182, 780 was purchased from Tocris Bioscience (Ellisville,

MO). Bovine, human, and rat FN were purchased from EMD Millipore (Milford, MA).

Antibodies- mAB IC3 specific for rat FN was a kind gift from the Schwarzbauer Lab at

Princeton University (NJ) and has been previously described [Sechler, 1996].

Phosphotyrosine-specific mAB, 4G10, was purchased from Upstate Biotechnology, Inc.

Rabbit polyclonal antibodies (AB1949) specific for the cytoplasmic tail of integrin α5 subunit protein was purchased from Chemicon. Inhibitory rat anti-mouse integrin α5β1 monoclonal antibody, clone BMB4 was purchased from Millipore. Alexa fluor dye-

88 conjugated secondary antibodies were purchased from Molecular Probes, Inc. (Eugene,

OR)/Invitrogen.

Cellular Stimulation and protein extraction- Conditions for quiescence, cell stimulation, and protein extraction were discussed previously [Filardo, 2000].

Immunofluorescence- Focal adhesions were visualized in 4T1 and SKBR3 cells that were seeded onto glass coverslips in PRF-DMEM/F12 medium containing FN-reduced serum and allowed to adhere overnight at 37C. The following day, serum was removed by washing in PRF-DMEM/F12. Cells were then cultured in the same media in the absence of serum for an additional 30hrs. Serum-starved cells were fed 2 µg/ml rat FN in the absence or presence of ligand (10 nM E2α, 10 nM E2β, 1 µM ICI 182, 780, 10 nM BPA) for 2 hr. Cells were then washed, fixed for 5 min in 4% paraformaldehyde, permeabilized in 0.05% Triton for 60 sec, and blocked in 5% BSA/PBS for 30min. Cells were incubated with phosphotyrosine specific 4G10 antibody diluted 1:500 in PRF-DMEM/F12 containing 5% BSA for 60 min. Coverslips were washed in PRF-DMEM/F12, and cell- associated antibodies were detected using Alexa 594-conjugated anti-mouse IgG diluted

1:1000 and delivered in PRF-DMEM/F12 containing 5% BSA for 30 min. After staining, coverslips were washed and mounted on glass slides in Vectashield/4’6-diamidino-2- phenylindole (vector laboratories, Inc., Burlingame, CA). FN fibril formation was examined in SKBR3, SKBR3Δ154, and SKBR3 Shc317Y/F, and 4T1 cells that were seeded onto glass coverslips in PRF-DMEM/F12 medium containing FN-reduced serum. Starved

89 cells were fed rat plasma FN (25 µg/ml) in PRF-DMEM/F12 medium in the presence of ligand (10 nM E2α, 10 nM E2β, 1 µM ICI 182, 780, 10 nM BPA) for 18 hr and then fixed and prepared for immunostaining as above. Fixed cells were stained with IC3 ascites diluted 1:1000 and delivered in PBS containing 1% BSA for 60 min. IC3 mAB was detected by staining with Alexa 594 conjugated anti-mouse IgG (1:1000) and processed for microscopy as above. Integrin α5β1 and stress fibers were visualized in SKBR3,

SKBR3Δ154, and SKBR3 Shc317Y/F cells that were seeded onto glass coverslips in PRF-

DMEM/F12 medium containing FN-reduced serum and allowed to adhere overnight at

37C. After adhesion, serum was removed by washing 3x with PRF-DMEM/F12, and the cells were then cultured in the same media in the absence of serum for an additional 30 hr. Serum-starved cells were fed 2 µg/ml rat FN in the absence or presence of E2β (10 nM) for 2 hr. Cells were then washed, fixed for 5 min in 4% paraformaldehyde, permeabilized in 0.05% Triton for 60 sec, and blocked in 5% BSA/PBS for 30 min. Cells were incubated with rabbit polyclonal antibodies (AB1949) specific for the cytoplasmic tail of integrin α5 subunit protein diluted 1:500 and TRITC-phalloidin diluted 1:500 in

PRF-DMEM/F12 containing 5% BSA for 60 min. Coverslips were washed in PRF-

DMEM/F12, and cell-associated antibodies were detected using Alexa 594-conjugated anti-rabbit IgG diluted 1:1000 and delivered in PRF-DMEM/F12 containing 5% BSA for 30 min. After staining, coverslips were washed and mounted on glass slides in

Vectashield/4’6-diamidino-2-phenylindole (vector laboratories, Inc., Burlingame, CA). All immunofluorescent images were visualized with a Nikon Eclipse 80i microscope (Nikon,

Inc., Melville, NY) equipped with a Nikon Plan Fluor 100x0.5-1.3 Oil Iris with differential

90 interference contrast and epifluorescent capabilities. Digital images were captured using a QImaging Retiga 2000R digital camera and Nikon imaging software (Elements Basic

Research 3.0).

Adhesion Assay- 48-well plates were coated with 200 µl of PRF-DMEM/F12 containing 2

µg/ml human FN or 10 µg/ml collagen overnight. Wells were blocked with 5% BSA PRF-

DMEM/F12 for 1 hr. SKBR3, SKBR3Δ154, and SKBR3 Shc317Y/F cells were seeded in triplicate, left untreated or treated with 10nM E2β, and allowed to adhere for 2 hr.

Nonadherent cells were gently washed away with PRF-DMEM/F12. Adherent cells were fixed and stained with 0.4% crystal violet and 4% ethanol in water for 5 min, and then washed 2x in large volumes of water. Crystal violet dye was extracted with 10% acetic acid and absorbance was measured spectrophotometrically at 550nm. All data points were determined from triplicate assays and expressed as the mean +/- the standard deviation. Nonspecific adhesion was subtracted as determined from cells that were seeded in BSA in the absence of substratum.

Boyden Chamber Migration Assay- Haptotaxis assays were conducted using modified

Boyden chambers consisting of a porous polycarbonate membrane (6.5 µm thickness, 8

µm pores; Transwells, CoStar corporation, Cambridge, MA) [Filardo, 1995]. The lower surfaces of the Transwell membrane were coated by adding 200 µl of serum free, PRF-

DMEM/F12 containing 2 µg/ml human FN, or 10 µg/ml collagen to the lower reservoir overnight. The underneath surface of the membrane was then blocked in 5% BSA in for

1 hr. SKBR3, SKBR3Δ154, and SKBR3 Shc317Y/F (105) cells were placed in the upper reservoirs of the Transwell in serum free, PRF-DMEM/F12 and left untreated or treated with 10 nM E2β and allowed to migrate overnight at 37C. Non-migrated cells were removed from the upper surface of the membrane using a Q-tip and cells remaining attached to the lower surface were fixed in ethanol and stained with 0.4% crystal violet in sodium borate buffer, pH 9.2, for 5 min, and then washed 2x in large volumes of water. Dye was eluted from the migrant cells using acetic acid and measured spectrophotometrically at 550 nM. Each data point was measured in triplicate and measured as the mean plus or minus the standard deviation. Nonspecific migration was subtracted as determined from cells that were seeded in BSA in the absence of substratum.

Anchorage-independent growth- SKBR3, SKBR3 GPERΔ154, SKBR3 Shc317Y/F, 4T1 vector and 4T1 GPERΔ154 cells (104) were seeded into PRF DMEM-F12 media in 0.35% agarose in the absence or presence of E2β (10 nM) and 10% FBS which was FN-depleted using gelatin-conjugated sepharose and supplemented with exogenous FN (2 µg/ml). Cells were grown for 10 days at 37C in a humidified chamber at 5% CO2. In some assays, inhibitory rat anti-integrin mouse α5β1 monoclonal antibody, clone BMB4 from

Millipore or control nonimmune rat antibodies were incorporated in the agar overlay.

Cultures were weighed every 2 days and evaporated water was replaced as needed.

Images of colonies were captured at 100x magnification (Brightfield). Colonies of greater than 20 cells were enumerated by direct counting.

Anchorage independent fibrillogenesis- Anchorage-independent fibrillogenesis was determined from hanging drop assays as previously described [Foty, 2011; Bartosh,

2014; Archacka, 2014]. Briefly, 4T1 vector or 4T1 GPERΔ154 cells were placed in suspension in serum-free media supplemented with or without agonist (10 nM E2α, 10 nM E2β, 10 nM ATII, 1 μM ICI 182, 780, RGD, RGE) and rhodamine-labeled bovine FN (30

μg/ml). For each treatment, 10 x 15 µl aliquots were distributed onto the underside of a

100 mm petri dish lid in a humidified chamber. Hanging drop cultures were incubated for 18 hr at 37C. Cells were fixed, stained with DAPI, and transferred to a glass slide.

Images were captured using a Nikon 80i inverted fluorescent microscope fitted with a

Retiga color camera at 100x magnification. Multiple Z-axis sections were reconstructed into 3D images using Nikon imaging software.

Results

E2β-induced mobilization of α5β1 into focal adhesions, the formation of actin stress fibers, and FN fibril formation require GPER and the Shc signaling adapter protein.

Src-like kinases, integrin α5β1, and Shc have been identified as integral components of a signaling pathway leading to E2β-mediated transactivation of the EGFR and FN matrix assembly [Filardo, 2000; Quinn, 2009]. In the latter study, we showed that stimulation of human SKBR3 cells with E2β results in the recruitment of FN-occupied integrin α5β1, into fibrillar adhesions at the cell periphery [Quinn, 2009], but did not address the role of GPER or Shc in the formation of focal adhesion plaques, presumed precursory adhesion structures which give rise to fibrillar adhesions. To gain knowledge as to whether GPER action was necessary for E2β-induced clustering of integrin α5β1 into focal adhesions and the formation of actin stress fibers, these cellular activities were compared in vector control SKBR3 cells and in a derivative line of SKBR3 cells expressing a dominant-negative form of GPER (GPERΔ154) (Figure 1). In these experiments, SKBR3 and SKBR3 GPERΔ154 cells were seeded onto coverslips, serum-starved, and stimulated with E2β in the presence of exogenous FN for 2 hr. Cells were fixed and stained with integrin α5β1 specific antibodies (green) and rhodamine-conjugated phalloidin to identify polymerized actin stress fibers (red). While parental SKBR3 cells demonstrated prominent actin stress fibers that co-localized with integrin α5β1 in focal adhesions, neither actin stress fibers nor integrin α5β1 enriched focal adhesions were observed in unstimulated cells (Figure 1A). Similarly, SKBR3 cells expressing a dominant-negative

94 form of GPER (GPERΔ154) that were stimulated with E2β were also unable to form actin stress fibers or concentrate integrin α5β1 into focal adhesions (Figure 1A), suggesting that GPER was required for the cellular activation events which recruit integrin α5β1 to focal adhesions and promote actin stress fiber formation. As previously demonstrated,

GPER stimulation with E2β resulted in the formation of FN fibrils (red), while dominant- negative GPER compromised FN fibril formation (Figure 1A).

To establish the requirement of Shc in E2β-induced integrin α5β1 recruitment to focal adhesions and the formation of actin stress fibers and FN fibril formation, these cellular events were evaluated in SKBR3 cells expressing control vector or a mutant Shc protein lacking its primary tyrosyl phosphorylation site, Shc317Y/F (Figure 1B). Focal adhesions and actin stress fibers were not measured in quiescent, unstimulated control or

Shc317Y/F cells. Following exposure to E2β, integrin α5β1 was recruited into prominent focal adhesions that co-aligned with the termini of actin stress fibers, while SKBR3

Shc317Y/F cells showed an impaired ability with regards to these integrin activation events. Expression of Shc317Y/F in SKBR3 cells negatively affected E2β induced clustering of integrin α5β1 into focal adhesions and showed less prominent focal adhesion plaques that appeared to be disordered with regards to their alignment with the termini of focal adhesions (Figure 1B). Likewise, E2β-induced FN fibril formation observed in control SKBR3 cells was prohibited in SKBR3 Shc317Y/F cells.

Collectively, these results indicate that Shc signaling following GPER activation is required to promote the recruitment of integrin α5β1 to focal adhesions and to induce actin stress fiber formation and consequent FN fibril formation. We have previously shown that Shc317Y/F accumulates relative to Shc wild-type protein on integrin α5β1 and cells expressing this mutant Shc protein fail to form fibrillar adhesions [Quinn,

2009]. Thus, our current observations may further suggest that the primary tyrosyl phosphorylation site on Shc is not required for entry of integrin α5β1 into focal adhesions, but that this site is required for its subsequent recruitment to fibrillar adhesions.

A SKBR3 SKBR3Δ154 B SKBR3 SKBR3 Shc317Y/F

α Stress Fibers Stress

Integrin 100 μm FN Fibrils FN

Figure 1- E2β-induced recruitment of integrin α5β1 to focal adhesions and the formation of actin stress fibers and FN fibrils are

GPER- and Shc- dependent. (A, top panel) SKBR3 cells expressing vector or GPERΔ154 and (B, top panel) SKBR3 cells expressing vector or Shc317Y/F were seeded onto coverslips, serum starved, and stimulated with E2β (10 nM) and exogenous rat FN (2 μg/ml).

Cells were incubated for 2 hr, fixed with 4% paraformaldehyde, permeabilized with detergent, and stained with integrin α5β1- specific antibodies (green) and Alexa-594-phalloidin (red). Nuclei were stained with DAPI (blue). (A, bottom panel) SKBR3 cells expressing vector or GPERΔ154 and (B, bottom panel) SKBR3 cells expressing vector or Shc317Y/F were seeded onto coverslips, serum starved, and stimulated with E2β (10 nM) and exogenous rat FN (25 μg/ml). Cells were incubated for 18 hr, fixed with 4% paraformaldehyde, and stained with rat FN specific mAB, IC3 (red). Nuclei were stained with DAPI (blue).

The influence of the xenoestrogen, bisphenol A, or the ER antagonist, ICI 182, 780, on the recruitment of tyrosyl phosphorylated proteins to focal adhesions and the production of

FN fibrils.

Xenoestrogens and ER antagonists act as GPER agonists [Filardo, 2000]. To determine whether these estrogenic steroids also influence GPER dependent formation of focal adhesions and FN fibrils, these integrin activation events were assessed in SKBR3 cells expressing vector or GPERΔ154 protein (Figure 2A). For these experiments, cells were made quiescent by seeding them onto glass coverslips in FN-reduced serum followed by serum-deprivation. Quiescent cells were left untreated or exposed to E2β, its inactive stereoisomer (E2α), the xenoestrogen, bisphenol A (BPA), or the ER antagonist, ICI 182,

780, and focal adhesions were assessed by measuring the clustering of tyrosyl phosphorylated proteins by immunostaining with the phosphotyrosine-specific monoclonal antibody, 4G10. Stimulation with E2β, BPA, or ICI 182, 780 resulted in the recruitment of homogenously distributed tyrosyl phosphorylated proteins into focal adhesion plaques (Figure 2A). Focal adhesions were similarly measured in murine 4T1 breast cancer cells that were stimulated with the ER antagonist, ICI 182, 780, or the xenoestrogen, BPA (data not shown). Focal adhesions were neither measured in quiescent cells nor in cells that were stimulated with the inactive stereoisomer E2α

(Figure 2A) nor EGF (data not shown). These results suggest that E2β-stimulated enrichment of tyrosyl phosphorylated proteins to focal adhesion plaques occurs independently of the ER and does not require EGF stimulation.

Previously, we have shown that stimulation of human SKBR3 cells with E2β results in the recruitment of FN-occupied integrin α5β1, to the cell periphery into fibrillar adhesions, specialized adhesion structures at which FN fibrils form [Quinn, 2009]. Here, we addressed the capacity of SKBR3 cells to form FN fibrils in response to stimulation with

ICI 182,780 or BPA. As shown in Figure 2B, either ICI 182, 780 or BPA, as well as E2β resulted in the formation of FN fibrils that were detected at the periphery of SKBR3 cells. FN fibril formation was not observed in SKBR3 cells that were left untreated or stimulated with E2α (Figure 2B) or EGF (data not shown). Similar observations were measured in murine 4T1 breast cancer cells. In both cell types, expression of GPERΔ154 protein prohibited FN fibril formation (Figure 2B and data not shown), demonstrating that these estrogenic hormones are capable of promoting FN fibril formation

(fibrillogenesis) in ER-negative breast cancer cells that are attached to planar surfaces coated with this extracellular matrix protein.

A UT E2α E2β

100 μm

BPA ICI Focal Adhesions Focal

B UT E2α E2β

BPA ICI FN Fibrils FN

Figure 2- E2β stimulation of human ER-negative breast cancer cells induces the formation of focal adhesions and FN fibrils. (A)

Human SKBR3 cells grown on glass coverslips were serum-starved and stimulated with either E2β (10 nM), its inactive stereoisomer,

E2α (10 nM), the xenoestrogen, BPA (10 nM), or the ER antagonist, ICI 182, 780 (1 µM), and incubated for 2 hr in the presence of exogenous rat FN (2 μg/ml). Following incubation, cells were fixed in 4% paraformaldehyde, permeabilized with detergent, and focal adhesions were detected using phosphotyrosine-specific antibodies (red). Nuclei were stained with DAPI (blue). (B) SKBR3 cells grown on glass coverslips were serum-starved and stimulated as described above. Cells were incubated for 18 hr in the presence of exogenous rat FN (25 μg/ml), fixed in 4% paraformaldehyde, and FN fibrils were detected using rat FN-specific mAB, IC3 (red). Nuclei were stained with DAPI (blue).

Stimulation of GPER selectively enhances the adhesivity of human breast cancer cells for

FN-coated substrata by a mechanism that requires the primary tyrosyl phosphorylation site on Shc.

In many instances, agonists that employ GPCRs promote enhanced cellular adhesive interactions by modulating the affinity of integrins for their cognate extracellular matrix

99 proteins and also by inducing the recruitment of integrins to focal adhesion plaques, a process referred to as inside-out integrin signaling [reviewed in Ginsberg, 2005; Luo,

2007]. To examine the influence of GPER stimulation by E2β on the adhesion of SKBR3 breast cancer cells for immobilized adhesive ligand, SKBR3 or SKBR3 GPERΔ154 cells were detached and exposed to E2β or left untreated and seeded into polystyrene wells coated with various concentrations of FN or collagen I (COLL) (Figure 3). Following a 2 hour incubation time at 37C, cells that were not firmly attached were gently washed away, adherent cells were fixed, cellular attachment was assessed by staining the remaining adherent cells with crystal violet, and attachment was measured as a function of eluted dye recovered from the resulting adherent cells. Unstimulated SKBR3 cells adhered to both FN- and COLL-coated substrata in a dose-dependent fashion with maximum cell adhesion measured at coating concentrations of 20 and 10 µg/mL of FN and COLL, respectively (data not shown). A two-fold increase (p=0.034) in the capacity of E2β-stimulated versus unstimulated SKBR3 cells to adhere to wells coated with suboptimal concentrations of FN (2 µg/mL) was observed (Figure 3A), which was associated with increased cellular spreading. In contrast, more modest differences in enhanced E2β-mediated adhesion to COLL- coated substrata were measured with no discernible difference in cellular spreading (data not shown). E2β increased adhesivity was eliminated in SKBR3 cells expressing GPERΔ154 suggesting that E2β promotes adhesion of SKBR3 breast cancer cells in a GPER-dependent manner. Similarly, Shc was tested for its involvement in GPER-enhanced adhesivity by comparing the relative capacity of SKBR3 and SKBR3 Shc317Y/F cells to adhere to immobilized FN or COLL

100

(Figure 3B). No significant increase (p=0.46) in adhesion between the E2β-stimulated and untreated SKBR3 Shc317Y/F cells was observed, suggesting that Shc is also involved in the signaling events that lead to increased adhesion to FN.

Cellular adhesion to planar surfaces is greatly strengthened by cell spreading [Lotus,

1989] and is often associated with increased cellular motility as measured in haptotactic responses on immobilized adhesive ligands. To examine whether GPER and Shc promote increased migration on FN-coated substrata, SKBR3 vector control, SKBR3 GPERΔ154, or

SKBR3 Shc317Y/F cells were seeded in the presence of 10 nM E2β (or left untreated) into the upper reservoirs of modified Boyden chambers containing a porous polycarbonate membrane (10 µm thickness, 8 µm pore) whose undersurface was coated with adhesive ligand (2 µg/mL FN or 10 µg/mL COLL) (Figure 4). Cell migration was measured by determining the number of cells that were capable of migrating from the upper reservoir across the membrane to its undersurface. On FN coated membranes,

SKBR3 vector control cells stimulated with E2β showed a 6.5-fold increase in their capacity to migrate compared with unstimulated cells in this assay (Figure 4A). SKBR3 cells that were plated onto membranes that were coated with higher concentrations of

FN (5-20 mg/mL) did not show increased haptotaxis when stimulated with E2β (data not shown), suggesting that GPER enhanced migration was the product of increased recruitment of FN receptors to cellular adhesion sites (data not shown). E2β enhanced haptotaxis on FN was abrogated in SKBR3 GPERΔ154 cells demonstrating that this migratory response was dependent upon GPER action (Figure 4A). Likewise, expression

101 of Shc317Y/F also impeded E2β enhanced haptotaxis on FN (Figure 4B). SKBR3 cells migrated equally well on COLL coated substrata independent of E2β stimulation and

Shc317Y/F had no impact on COLL migration suggesting that GPER signaling did not enhance migration on this extracellular matrix protein (data not shown).

A 0.18 UT 0.16 0.14 E2β 0.12 0.10 0.08 0.06

Absorbance 0.04 0.02 0.00 SKBR3 SKBR3Δ154 B 0.18 0.16 0.14 0.12 0.10 0.08 0.06

Absorbance 0.04 0.02 0.00 SKBR3 SKBR3 Shc317Y/F

Figure 3- GPER stimulation promotes SKBR3 cell adhesion onto FN-coated, but not collagen-coated, substrata in a Shc-dependent manner. (A) SKBR3 cells expressing vector or GPERΔ154 and (B) SKBR3 cells expressing vector or Shc317Y/F were seeded onto 48- well plates coated with FN (2 μg/ml) or collagen (10 μg/ml) and allowed to attach for 2 hr at 37C in the absence or presence of E2β

(10 nM). After adhesion, unattached cells were removed by gentle washing, and the remaining adherent cells were fixed and stained with crystal violet. Excess crystal violet was washed away and cell-associated crystal violet was extracted with 10% acetic acid.

Absorbance was measured at 550 nm. Each data point represents the mean +/- the standard deviation of triplicate samples.

Nonspecific adhesion as measured on BSA-coated wells was subtracted.

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A SKBR3 SKBR3Δ154 400 UT 350

UT 300 E2β

250

500 μm 200

150

100 E2β 50

0 Number of Cells Migrated Cells of Number SKBR3 SKBR3Δ154

SKBR3 B SKBR3 Shc317Y/F 400

350 UT 300 250

200

150

100

E2β 0 Number of Cells Migrated Cells of Number SKBR3 SKBR3 Shc317Y/F

Figure 4- GPER stimulation enhances haptotaxis of human SKBR3 breast cancer cells on FN-coated, but not collagen-coated, substrata in a Shc-dependent manner. (A) SKBR3 vector or GPERΔ154 cells and (B) SKBR3 vector or Shc317Y/F cells were left untreated or treated with E2β (10 nM) and seeded into the transwells of modified Boyden chambers containing a porous polycarbonate membrane (10 µm thickness, 8 µm pore) that were left untreated or coated with either FN (2 μg/ml) or collagen (10

μg/ml) and incubated overnight at 37C. Non-migrant cells were removed from the top of each chamber and migrant cells on the lower surface of the membranes were fixed and stained with crystal violet. The number of migrated cells were counted and images were taken at 40x magnification.

GPER promotes anchorage-independent growth by promoting fibrillogenesis in a three- dimensional environment.

Since the conversion of soluble FN into fibrils by mammary adenocarcinoma cells has been linked to enhanced anchorage-independent growth [Saulnier, 1997] and GPER action results in fibrillogenesis on planar surfaces (Figures 1 and 2) [Quinn, 2009], the possibility that GPER-mediated fibrillogenesis is required for anchorage-independent

103 growth was evaluated (Figures 5 and 6). To address this hypothesis, human SKBR3 or murine 4T1 breast cancer cells were seeded into semi-solid media supplemented with

FN-depleted fetal bovine serum in the presence of increasing amounts of exogenous FN.

Under conditions of FN depletion, neither SKBR3 nor 4T1 cells were able to form colonies whereas both cell types readily formed colonies in the presence of 10% fetal bovine serum, which had not been FN depleted (data not shown and Figure 6A).

However, either cell line readily formed colonies in FN reduced conditions with 2 µg/mL exogenous FN (Figure 5 and 6), provided that E2β was also present. Neither cell line grew in the presence of exogenous E2α and FN. Expression of GPERΔ154 in either cell background effectively prohibited E2β-dependent, anchorage-independent growth

(Figures 5A and 6B), suggesting that these growth properties are GPER-dependent. The specificity of GPERΔ154 for inhibiting GPER action in this assay was demonstrated by the observation that the substitution of E2β for exogenous angiotensin II (ATII), which acts through its dedicated cognate GPCR, could restore FN-dependent, anchorage- independent growth (data not shown and Figure 6B). Expression of Shc317Y/F in the

SKBR3 cell background had an inhibitory effect on colony formation suggesting that Shc is also required for E2β-dependent growth in semi-solid media (Figure 5B). Inclusion of inhibitory anti-integrin α5β1 antibodies in semi solid media containing murine 4T1 cells showed that integrin α5β1 engagement is necessary for anchorage-independent growth under these conditions of limiting FN and exogenous E2β (Figure 6C). Collectively, these data indicate that GPER stimulation and exogenous FN is required for E2β-dependent, anchorage-independent growth.

104

To directly address the capacity of GPER to promote fibrillogenesis in a three- dimensional environment, murine 4T1 or 4T1 GPERΔ154 breast cancer cells were cultured in hanging drops assays in which rhodamine-labelled FN was incorporated. As demonstrated in Figure 7, 4T1 cells that were cultured in suspension (in the absence of a substratum) overnight in the presence of E2β were capable of forming FN fibrils in this anchorage-independent assay (Figure 7A). Similarly FN fibrils were also measured by 4T1 cells that were stimulated with ATII. FN fibril formation was not measured in untreated or E2α treated cells. GPERΔ154 specifically inhibited fibrillogenesis in hanging drop cultures of 4T1 cells that were stimulated by E2β, but had no effect on ATII-induced fibrils (Figure 7A). The ER antagonist, ICI 182,780 also induced the formation of FN fibrils in the hanging drop assay (Figure 7B) and this cellular activity was inhibited by

GPERΔ154 (data not shown). Moreover, hanging drop cultures of E2β-stimulated 4T1 breast cancer cells that were supplemented with soluble RGD peptide were unable to form FN fibrils; however, control RGE peptide did not have a negative effect on anchorage-independent fibrillogenesis (Figure 7B). Taken together the data in Figures 5-

7, suggests that GPER promotes anchorage-independent growth through its ability to synthesize FN fibrils.

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A SKBR3 SKBR3Δ154 100 90 UT 80 E2β UT 70 60 200 μm 50 40 30 20

10 Number of Colonies of Number E2β 0 SKBR3 SKBR3Δ154

SKBR3 B SKBR3 Shc317Y/F 100 90 UT 80 70 60 50 40 30 20

10 Number of Colonies of Number E2β 0 SKBR3 SKBR3 Shc317Y/F

Figure 5- GPER enhances FN-dependent, anchorage-independent growth of SKBR3 cells in a Shc-dependent manner. (A) SKBR3 vector or GPERΔ154 and (B) SKBR3 vector or Shc317Y/F cells were seeded into phenol red-free DMEM-F12 media containing 2% FN reduced serum in 0.35% agarose in the absence or presence of E2β (10 nM) and supplemented with exogenous FN (2 µg/ml). Cells were grown for 10 days at 37C in a humidified chamber. Cultures were weighed every 2 days and water was replaced as needed.

Images of colonies were captured at 10x magnification (brightfield). Examples shown above are representative views of multiple experiments.

106

A UT E2 C

200 μm

B UT E2 ATII

4T1 No No FN 200 μm

4T1 Δ154

4T1 FN

4T1 Δ154

Figure 6- E2β stimulation alters colony morphology of ER-negative mouse breast cancer cells grown in soft agar. (A) 4T1 cells were seeded into phenol red-free DMEM-F12 media containing 2% FN reduced serum in 0.35% agarose in the absence or presence of E2β

(10 nM) and supplemented with exogenous FN (2 µg/ml). Cells were grown for 10 days at 37C in a humidified chamber. Cultures were weighed every 2 days and water was replaced as needed. Images of colonies were captured at 10x magnification (brightfield).

Examples shown above are representative views of multiple experiments. (B) 4T1 vector or 4T1 GPERΔ154 cells were seeded into phenol red-free DMEM-F12 media containing 2% serum in 0.35% agarose and left untreated or stimulated with E2β (10 nM) or ATII

(10 nM) in the presence or absence of exogenous FN (2 μg/ml). Growth conditions were the same as above. (C) 4T1 cells were seeded into phenol red-free DMEM-F12 media containing 2% serum in 0.35% agarose and left untreated or stimulated with E2β (10 nM), ICI 182, 780 (1 μM), IgG and E2β (10 nM), or anti-integrin α5bβ and E2β (10 nM) in the presence or absence of exogenous FN

(2μg/ml). Growth conditions are the same as above.

107

A UT E2α E2β ATII

4T1

100 μm

4T1Δ154

B ICI E2β + RGD E2β + RGE

4T1

Figure 7- GPER promotes FN fibril formation of mouse 4T1 breast cancer cells cultured in hanging drops . (A) 4T1 vector or

GPERΔ154 cells were placed in suspension in serum-free media supplemented with or without E2α (10 nM), E2β (10 nM), or angiotensin II (ATII) (10 nM), and rhodamine-labeled bovine FN (30 μg/ml). For each treatment, 10 x 15 µl aliquots were distributed onto the underside of a 100 mm petri dish lid in a humidified chamber. Hanging drop cultures were incubated for 18 hr at 37C. Cells were fixed, stained with DAPI and transferred to a glass slide. Images were captured using a Nikon 80i inverted fluorescent microscope fitted with a Retiga color camera at 100x magnification. Multiple Z-axis sections were reconstructed into 3D images using Nikon imaging software. Examples shown above are representative views of multiple experiments. (B) 4T1 vector cells were placed in serum free media supplemented with or without ICI 182, 780 (1 μM), or E2β (10 nM) with RGD or RGE, and rhodamine- labeled bovine FN (30 μg/ml). Hanging drops were incubated and analyzed as described in (A).

108

Discussion

Evidence is provided here that estrogenic hormones act via GPER to enhance integrin

α5β1-dependent adhesion of breast cancer cells to FN via a signaling mechanism that requires tyrosyl phosphorylation of the Shc adaptor protein. Specifically, we show that

GPER stimulation promotes: i) the formation of focal adhesions leading to the reorganization of actin stress fibers; ii) enhanced cellular adhesivity and haptotaxis on immobilized FN; iii) anchorage-independent FN fibril formation; and iv) FN-dependent, anchorage-independent growth. We have previously reported that integrin α5β1 and

Shc are integral components of a signaling pathway leading to E2-mediated EGFR transactivation and FN matrix assembly on planar surfaces [Quinn, 2009]. Collectively, these data support the idea that GPER-1 coordinates two key cellular events required for the survival of breast cancer cells that escape the confines of glandular epithelia and invade the surrounding tissue parenchyma, namely, responsiveness to soluble growth factors and the capacity to form a provisional extracellular matrix. Our findings are consistent with studies in mice that have shown a requirement for exogenous FN for efficient tumor cell implantation [Price, 1975] and integrin β1 and Shc for homeostasis of the mammary gland [Faraldo, 2003].

Upon engagement of their adhesive ligands, integrins promote intracellular signals and are recruited to focal adhesion plaques, events which result in increased integrin affinity and avidity, and are necessary for cell survival and tissue homeostasis. External cues from soluble mediators also may modulate integrin affinity by a process that is referred

109 to as inside-out integrin signaling [Ginsburg]. In many instances, these extrinsic factors modulate integrin affinity signal through GPCRs. For example, affinity upregulation of integrin αllbβ3 for its ligand fibrinogen, a key event in thrombus formation, occurs in response to stimulation of platelets with ADP, epinephrine, or thrombin, whose receptors are GPCRs [Cosemans, 2008]. Likewise, β1 and β2 integrins on leukocytes exhibit increased affinity for their adhesive proteins in response to a broad array of immunomodulatory substances that act through GPCRs, including, but not limited to, f-

Leu-Met-Phe, chemokine, and complement cascade products [Oberyszyn, 1998].

Similarly, our findings here, indicate that E2 action via GPER, activates integrin α5β1 resulting in its recruitment to focal adhesions (Figures 1, 2), increased adhesion and haptotaxis on planar surfaces coated with FN (Figures 3, 4), and FN matrix assembly in two- and three-dimensional environments (Figures 1, 2, 5-7) by breast cancer cells.

Since the experiments presented here do not directly address conformational alterations in the external domains of integrin α5β1 associated with FN affinity, it is not possible to formally conclude whether GPER enhanced cellular adhesivity occurs via inside-out signaling. However, our findings are consistent with prior observations that have shown that GPCR activation promotes allosteric changes within the ligand binding domains of β1, β2 and β3 integrins resulting in enhanced adhesive function [Oberyszyn,

1998; Phillips, 2001; Cowan, 2000]. Enhanced integrin α5β1 adhesive action of breast cancer cells for FN does not appear to be solely relegated to signaling by GPER, as angiotensin II stimulation of breast cancer cells also enhanced fibrillogenesis (Figure 6).

110

Shc is a signaling adaptor that couples integrins, receptor tyrosine kinases (RTKs),

GPCRs, and a variety of other receptors to the ras-mek-erk signaling cascade via its interaction with Grb2/Sos [Ravichandran, 2001]. Activation of Shc occurs as a result of its phosphorylation at a conserved tyrosine which, in turn, produces an SH2 binding site for Grb-2 [Ravichandran, 2001]. Upon outside-in integrin activation by ligation to their respective ECM ligands, Shc becomes phosphorylated and is selectively recruited to integrins α5β1 (FN) and αvβ3 (vitronectin), but not to integrins α2β1 or α6β1 which bind collagen or laminin, respectively [Wary, 1996]. Similar to our results presented here, others have also shown that Shc is required for integrin-mediated spreading and haptotaxis on FN, but not collagen [Sweet, 2012]. Moreover, these authors found that

Shc may represent an integration point for vEGF receptor and ECM signaling. Consistent with this idea, we found that expression of a Shc mutant lacking its primary tyrosyl phosphorylation site, Shc317Y/F, or blockade of integrin α5β1 ligation to FN with soluble

RGD peptides inhibited GPER-mediated tyrosyl phosphorylation of the EGFR by E2β, but not by exogenous EGF-related ligands [Quinn, 2009], suggesting that Shc and integrin

α5β1 are signaling intermediaries necessary for activation of the EGFR-to-erk signaling axis by GPER. While each of the three Shc isoforms, p66Shc, p52Shc and p46Shc have been shown to localize to focal adhesions, it is important to note that the p66Shc isoform appears to play a unique role in sensing cell adhesion as p66Shc promotes anoikis via RhoA activation in detached cells [Ma, 2007]. Likewise, under conditions of oxidative stress, p66Shc uncouples the ras-mek-erk signaling cascade [Aray, 2008].

These findings are consistent with studies which have shown that lung cancer cells

111 lacking p66Shc display traits associated with advanced cancer, including anchorage- independent growth [Ma, 2010]. However, GPER enhanced adhesivity to FN does not simply appear to be a result of preferential recruitment of the p52Shc or p46Shc isoforms to integrin α5β1 as human breast cancer cell lines that express p66Shc remain competent with regards to their capacity to promote integrin α5β1-dependent EGFR transactivation and enhanced FN adhesivity and growth in soft agar [Quinn, Magruder and Filardo, unpublished results].

The primary tyrosyl phosphorylation site on Shc (Tyr317 on p52Shc) serves as an SH2 binding site that not only facilitates the activation of ras-mek-erk, but also promotes the recruitment of proteins that concentrate in focal adhesions, including tyrosine kinases such as FAK and Src-like kinases [Charest, 1996]; tyrosine phosphatases, PTPN12, SHP-2

[Habib, 1994]; and tyrosine phosphorylated proteins such as paxillin, talin, and vinculin

[Petita]. Prior studies have shown that Shc couples with integrin during matrix engagement [Habib, 1994; Wary, 1996; Deshmukh, 2010] and that Shc localizes to focal adhesions in attached cells [Angers, 1999], events associated with increased contractility tension at integrin anchorage points [Ma, 2007; Debnath, 2010]. In response to shear stress, αvβ3 and β1 integrins associate with Shc, which results in cell attachment to substratum [Chen, 1999]. Our prior results indicate that tyrosyl phosphorylation of Shc is not a prerequisite for its physical association with integrin α5β1 following GPER stimulation [Quinn, 2009], which is consistent with results reported by others examining the requirements for Shc activation and complex formation with other integrins

112

[Phillips, 2001; Cowan, 2000]. In fact, expression of Shc317Y/F did not block GPER- enhanced association of endogenous Shc isoforms with immunopurified integrin α5β1, but rather resulted in its accumulation relative to these endogenous isoforms, suggesting a possible role for protein tyrosine phosphatases (PTPs) for the release of Shc from integrin α5β1. PTPs regulate tyrosine phosphorylation as antagonists to tyrosine kinases [Sun, 2011]. Many PTPs have been shown to play a role in the regulation of focal adhesion disassembly, cell spreading, and migration [Zheng, 2013; Yu, 1998].

Specifically, dominant-negative SHP2 mutants altered integrin-dependent activation of the Erk pathway, focal adhesion turnover, cell spreading, and migration [Zheng, 2013].

Furthermore, overexpression of a dominant-negative mutant of PTP1B resulted in a defect in integrin-mediated adhesion and signaling in fibroblasts [Arregui, 1998]. In addition, PTP-α was shown to regulate the activity of Src and this regulated cell-stratum adhesion [Harder, 1998]. The role of PTPs in GPER signaling is an area for further examination. However, this idea is consistent with the concept that focal adhesion formation is a dynamic process and may suggest that disassembly and re-assembly of

Shc-integrin complexes is necessary for the formation of more ordered focal adhesions and actin stress cables that may be aligned through SH2 binding proteins [Barberis,

2007]. Inhibition of the β1-integrin function in the mammary gland of mice results in a decrease in Shc phosphorylation, failure to recruit Grb2, and failure to activate the Ras pathway [Faraldo, 2003]. Further evidence that the association of Shc with integrin is that ECM engagement by integrin β1 is required for muscarinic acid receptor-mediated activation of FAK and paxillin [Fetuccia, 2002], two proteins that are recruited to focal

113 adhesions. Tyrosyl phosphorylation of FAK and paxillin are associated with integrin signaling [Katz, 2003; Ren, 2000]. Shc-/- fibroblasts show a decrease in activation of Erk in response to ECM ligands [Faraldo, 2001]. However, we have shown that tyrosine phosphorylation of FAK at its autophosphorylation site is not increased following E2 stimulation [Filardo and Quinn, unpublished results]. One possible explanation for this is that GPER stimulation in human breast cancer cells does not generate a sufficient number of integrin α5β1-FN bonds to exceed a signaling threshold that permits FAK activation [Quinn, 2009]. Here, we show that Shc317Y/F also prevents GPER enhanced adhesion as measured in cellular adhesion or haptotaxis on FN-coated planar surfaces

(Figures 3 and 4), suggesting that tyrosyl phosphorylation of Shc317 is required for activation of integrin α5β1-dependent adhesive responses.

FN matrix assembly is a cell-mediated process that depends on the binding of FN to integrins [Wu, 1993]. In addition to promoting FN-FN interactions outside the cell, integrins link to the actin cytoskeleton through their cytoplasmic domains and coordinate FN fibril formation with intracellular signaling [Wierzbicka, 2003]. Integrins affect intracellular processes through signaling molecules such as paxillin, vinculin, talin,

FAK, and Src [Miranti, 2002]. These signaling molecules are then incorporated into focal and fibrillar adhesions [Geiger, 2001], which are associated with cellular survival.

Integrin binding to FN induces the activation of FAK [Friedland, 2009]. Cell adhesion and spreading, migration, cell survival, and proliferation depend on integrin signaling through FAK, which then interacts with downstream signaling molecules such as Src,

114

PI3-K, and Grb7 [Schlaepher, 1999]. FN matrix assembly occurs in a three-dimensional environment, which is required for anchorage independent growth. Anchorage- independent growth is a predictor of metastasis and is linked to increased responsiveness to growth factors. Saulnier et al. show that treatment of SP01 breast cancer cells with TGF-β and HGF results in significant colony growth, suggesting these growth factors may be affecting the expression or secretion of FN, supporting colony growth [Saulnier, 1996]. Furthermore, the ability of mammary adenocarcinoma cells to convert soluble FN into fibrils results in increased responsiveness to growth factors and anchorage-independent growth [Saulnier, 1996; Qiao, 2000]. In another study, implantation of mammary tumor xenografts in immunocompromised mice is enhanced with the introduction FN [Price, 1996]. We show that stimulation of GPER enhanced FN- dependent, anchorage-independent growth in a Shc-dependent manner (Figures 5 and

6). As discussed above, p66SShc promotes anoikis in unattached cells [Ma, 2007]. SKBR3 cells lack the p66Shc isoform, but we have previously shown that MDA-MB-231 cells, which express p66Shc, undergo GPER mediated adhesion and anchorage-independent growth [Quinn, Magruder, Filardo, unpublished results], suggesting that absence of p66Shc is not a requirement for growth in soft agar. We further show that integrin α5β1 is required for this effect (Figure 6C), specifically the ability of integrin to signal in an inside-out manner (Figure 7B).

Estrogens regulate homeostasis of the mammary gland, and anti-estrogens and environmental estrogens are known to directly influence the behavior of mammary

115 adenocarcinoma cell lines [Filardo, 2006]. Here, we show that bisphenol A, or the ER antagonist, ICI 182,780, also promote GPER-mediated recruitment and clustering of integrin α5β1 into focal adhesions (Figures 1 and 2). These findings support prior studies, which have shown that tamoxifen induces changes in cytoarchitecture [Ehlers,

1999; Sapino, 1986]. Interestingly, although expression of Shc317Y/F did block GPER mediated FN fibril formation [Quinn, 2009], it did not completely impede GPER enhanced recruitment of integrin α5β1 into focal adhesions as integrin α5β1 recruitment occurred in the context of mutant Shc, but resulted in the formation of poorly organized focal adhesions which were not properly aligned with actin stress fibers (Figure 1). Taken together, these data support the hypothesis that GPER signals via integrin α5β1 and Shc to coordinate growth factor responsiveness and anchorage- independent growth, events critical for breast cancer progression. GPER expression in primary breast tumors directly varies with tumor size and metastasis, markers of advanced disease, which is a relationship that is diametrically opposed to that shared by

ER and these same prognostic variables [Filardo, 2006]. The more recent finding that

GPER expression in triple negative breast cancer directly varies with disease progression underscores a potential role for this newly appreciated estrogen receptor in advanced disease [Steiman, 2013]. Our work here defining the cell biological influence of GPER stimulation on FN matrix assembly and adhesion to this extracellular matrix protein provides insight into the cellular mechanisms by which estrogens may promote the survival of metastatic breast cancer cells.

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Chapter 3

Aim 2

128

Abstract

GPER. This mechanism of action results in the recruitment of FN-engaged integrin α5β1 to fibrillar adhesions and the formation of integrin α5β1-Shc adaptor protein complexes.

It also promotes the formation of focal adhesions and actin stress fibers resulting in increased cellular adhesion and haptotaxis on FN in a Shc-dependent manner.

Furthermore, GPER stimulation of breast cancer cells allows for FN-dependent anchorage independent growth. These are all events that may allow for cellular survival and tumor progression. PTPN12, a phosphatase known to interact with Shc, plays a role in focal adhesion formation and fibrillogenesis. Here, we show that overexpression of

PTPN12 inhibits GPER-mediated focal adhesion and FN fibril formation, adhesion and haptotaxis on FN, and anchorage independent growth. Moreover, PTPN12 knockdown positively influences these events associated with cellular survival.

129

Introduction

Estrogen (E2) is a steroid hormone that promotes rapid or pregenomic events, which occur within minutes of E2 exposure, and genomic transcriptional responses, which occur over several hours after exposure [Filardo, 2006]. E2 is important in the development and homeostasis of the mammary gland, and the growth of mammary tumors [Filardo, 2006]. It signals via the commonly known nuclear estrogen receptor

(ER) [Thomas, 2005], but studies in animal and cell models demonstrating that E2 promotes rapid biochemical effects that are not related to its known actions via the ER have led to the recognition of G protein-estrogen receptor (GPER) as a novel estrogen receptor [Filardo, 2006]. GPER belongs to the G protein-coupled receptor (GPCR) superfamily. GPER promotes the rapid stimulation of adenylyl cyclase [Filardo, 2000] and EGFR transactivation via the release of membrane-bound HB-EGF [Filardo, 2000].

GPER has significance in breast cancer based on studies conducted in human breast tumors and breast cancer cell lines. GPER expression in primary tumors is positively associated with tumor size, HER2 expression, and the presence of extramammary metastases [Filardo, 2006; Arias-Pulido, 2010; Ignatov, 2011; Steinman, 2013].

Furthermore, in ER-negative, GPER-positive SKBR3 human breast cancer cells, E2 promotes GPER-dependent EGFR transactivation and stimulation of adenylyl cyclase

[Albanito, 2007]. This was inhibited when GPER antisense oligonucleotides were used

[Albanito, 2007]. In another cell model, GPER-null MDA-MB-231 cells did not stimulate adenylyl cyclase. However, overexpression of GPER in MDA-MB-231 cells restores the activation of adenylyl cyclase by E2 [Filardo, 2005]. In addition, GPER-dependent E2

130 action results in the activation of integrin α5β1 and increased FN matrix assembly, which are significant in advanced disease because these activities allow for cell survival of metastatic cancer cells [Quinn, 2009].

FN plays a major role in cell adhesion, growth, and survival. It is synthesized in a soluble form and then assembled into an insoluble fibrillar matrix in a cell-mediated process that is initiated by its recognition via Arg-Gly-Aso (RGD) binding sites on each monomer by integrin α5β1. Integrin α5β1 is the major receptor in FN matrix assembly [Pankov,

2000]. It is present in both focal and fibrillar adhesions, and FN matrix assembly is dependent on the translocation of activated integrin α5β1 to these adhesions [Pankov,

2000]. Focal adhesions are where anchorage to FN occurs, and they are enriched in tyrosyl phosphorylated proteins. Altered expression of FN or alterations in the specific recognition of FN by integrin α5β1 is associated with the development of cancer. Several studies have shown that FN is associated with the development of breast cancer.

Exogenous FN negatively impacts acinar differentiation in the mammary gland and creates an environment conducive to the growth of mammary epithelia [Williams,

2008]. Studies have shown that mammary adenocarcinoma cells are capable of converting soluble FN into fibrils [Saulnier, 1996], resulting in increased responsiveness to growth factors and enhanced anchorage-independent growth [Qiao, 2000].

Anchorage-independent growth is an attribute that best predicts metastatic potential

[Quinn, 2009]. Successful implantation of human mammary tumor xenografts in immunocompromised mice is facilitated by the addition of exogenous FN, suggesting a survival advantage for tumors that interact with FN [Price, 1996]. The expression of

131 integrin α5β1 and FN is increased during proliferation of mammary gland tissue in mice suggesting that a FN-integrin interaction may be required for hormone-dependent proliferation in the mammary gland [Woodward, 2001]. We have previously shown that

EGFR transactivation by GPER requires activation of the integrin α5β1 in breast cancer cells as measured by its recruitment to fibrillar adhesions, the conversion of soluble FN to a detergent- insoluble form, and the association of integrin α5β1 with the signaling adaptor, Shc [Quinn et al, 2009]. Shc is involved in intracellular signaling events that determine growth factor responsiveness and bidirectional integrin signaling

[Ravichandran, 2001]. Shc is expressed as three isoforms (p66, p52, p46) that share a carboxyl terminal. Shc proteins are substrates for receptor and nonreceptor tyrosine kinases, and each possesses both PTB and SH2 phosphotyrosine binding domains that act to dictate its interaction with signaling effectors and form signaling complexes.

PTPN12 is a phosphatase that interacts with the p52 and p66 Shc proteins, but not the p46 isoform [Habib, 1994]. Shc binds to the NPLH sequence of PTPN12 [Faisal, 2002]. It appears that PTPN12 is a negative regulator of Shc because overexpression and antisense experiments show that PTPN12 down-regulates lymphocyte stimulation by dephosphorylation of Shc and downregulating the Ras pathway [Faisal, 2002].

Furthermore, PTPN12 antagonizes pro-mitogenic EGFR signaling by displacing Shc from the EGFR [Zheng, 2013]. PTPN12 blocks EGFR function by dephosphorylation [Faraldo,

2001], and inhibition of the EGFR is a method of controlling cell proliferation, survival, and tumorigenesis [Sun, 2011]. PTPN12 is also important in the regulation of proteins involved in focal adhesion regulation such as FAK, p130Cas, and paxillin [Zheng, 2013].

132

These proteins become tyrosine-phosphorylated in response to the activation of the integrin signaling pathways and dephosphorylated in response to the activation of

PTPN12 [Zheng, 2013]. Thus, PTPN12 controls the rate of focal adhesion turnover and cell migration [Zheng, 2013]. PTPN12 deficient fibroblasts demonstrate increased phosphorylation of p130Cas, paxillin, and FAK [Zheng, 2013]. FAK is implicated in the formation of focal adhesions, and the inhibition of FAK promotes cell migration, invasion, and metastasis [Angers-Loustau, 1999]. Overexpression of PTPN12 results in the dephosphorylation of FAK at Y397, promotion of disassembly and turnover of focal adhesions, and promotion of Ras-induced migration [Zheng, 2013].

PTPN12 is down-regulated in breast cancer cells when compared to normal breast tissue

[Sun, 2011; Wu, 2013; Xunyi, 2002]. Low PTPN12 expression is positively associated with lymph node status, stage, and distant metastatic relapse in breast cancer [Wu, 2013]. In addition, Xunyi et al. report that low PTPN12 expression is associated with increased tumor and lymph node size, lymph node metastasis, distant metastasis, and histological grade [Xunyi, 2002]. Here, we show that PTPN12 plays a role in events associated with cellular survival. Overexpression of PTPN12 in human SKBR3 breast cancer cells inhibits

GPER-mediated focal adhesion and FN fibril formation, adhesion and haptotaxis on FN, and anchorage independent growth. Moreover, PTPN12 knockdown positively

133 influences the ability of GPER to promote the formation of focal adhesions and FN fibrils, adhesion and haptotaxis on FN, and anchorage independent growth.

134

Materials and Methods

Cell Culture- SKBR3 (ERα-, ERβ-, GPER+) breast cancer cells were obtained from the

American Type Culture collection (Manassas, VA). SKBR3 cells expressing wild-type (WT)

PTPN12, phosphatase-dead (PD) PTPN12, mutant NPLH (Shc binding site) PTPN12, and

PTPN12 shRNA were generated as described below. GE11β1 mouse epithelial cells were generated as described by Quinn [Quinn, 2009]. B11WT PTPN12+/+and B14EV PTPN12-/- primary embryonic fibroblasts were a kind gift from the Tremblay Lab at McGill

University [Cheng, 2001]. shRNAs and cDNAs- A GIPZ lentiviral PTPN12 shRNA (V2LHS-170948) was obtained from the Hannon-Elledge shRNA collection (Open Biosystems). WT PTPN12 and PD PTPN12 cDNAs were a kind gift from the Westbrook lab at Baylor College of Medicine (Houston,

TX) [Sun, 2011]. PD PTPN12 was generated by the Westbrook lab by mutating amino acid C231 to S [Sun, 2011]. cDNAs were cloned into pDEST26 vector (Life Technologies) using Gateway Technology (Life Technologies). PTPN12 cDNA with a mutation in its Shc binding site was generated by altering the cDNA encoding full-length human PTPN12 protein from the Westbrook lab. To accomplish this, a PCR product encoding three point mutations in the NPLH region of PTPN12 (N599I, P600A, H602L), which are known to inhibit Shc binding to PTPN12 [Sun, 2011], was synthesized using forward 5’-

CTGATGACTCAGACTCAGATGAAAG-3’ and reverse 5’-

AGAGAAGTGCAATAGTAAAACTGAGG-3’ oligonucleotide primers. The PCR-amplified fragment was subcloned into pDEST26.

135

Transfections and Selection of Stable Cell Lines- SKBR3 or GE11β1 cells were mock transfected or transfected with PTPN12 shRNA using lipofectamine (GIBCO-BRL). Three days after transfection, 3 µg/ml Puromycin was added to the growth medium. The resulting population of Pyromycin-resistant cells was used for further study. SKBR3 or

GE11β1 cells were transfected with pDEST26 vector control, WT PTPN12, PD PTPN12, or mutant NPLH PTPN12 using lipofectamine (GIBCO-BRL). Three days after transfection, 1

µg/ml Neomycin was added to the growth medium. The resulting population of

Neomycin-resistant cells was used for further study.

Growth Factors, Estrogens, and Matrix Proteins- Water-soluble E2β and 17α-estradiol

(E2α) were purchased from Sigma (St. Louis, MO). Bovine, human, and rat FN were purchased from EMD Millipore (Milford, MA).

Antibodies- mAB IC3 specific for rat FN was a kind gift from the Schwartzbauer Lab at

Princeton University (NJ) and has been previously described [Sechler, 2001].

Phosphotyrosine-specific mAB, 4G10, was purchased from Upstate Biotechnology, Inc.

Alexa fluor dye-conjugated secondary antibodies were purchased from Molecular

Probes, Inc. (Eugene, OR)/Invitrogen.

Immunoblotting- SKBR3 PTPN12 variant cells, B11WT, or B14EV cells were seeded onto petri dishes and allowed to adhere overnight. Cells were lysed and equivalent amounts of total protein were electrophoresed through 8% SDS-polyacrylamide gels and transferred to PVDF. Membranes were blocked in phosphate-buffered saline (PBS) plus

3% BSA. PTPN12 was detected by probing the membrane with PTPN12 antibodies

136 delivered in 3% BSA in PBS. Antibody-antigen complexes were detected using HRP- conjugated antirabbit secondary antibodies diluted 1:5000 and enhanced chemiluminescence.

Immunofluorescence- PTPN12 expression was visualized in SKBR3 and GE11β1 PTPN12 variant cells that were seeded onto glass coverslips in PRF-DMEM/F12 medium containing 5% FBS and allowed to adhere overnight at 37C. After adhesion, cells were washed, fixed for 5 min in 4% paraformaldehyde, permeabilized in 0.05% Triton for 60 sec, and blocked in 5% BAS/PBS for 30 min. Cells were incubated with PTPN12 specific antibody diluted 1:50 in PRF-DMEM/F12 containing 5% BAS for 60 min. Coverslips were washed in PRF-DMEM/F12, and cell-associated antibodies were detected using Alexa

594-conjugated anti-rabbit IgG diluted 1:1000 and delivered in PRF-DMEM/F12 containing 5% BSA for 30 min. After staining, coverslips were washed and mounted on glass slides in Vectashield/4’6-diamidino-2-phenylindole (Vector Laboratories, Inc.,

Burlingame, CA).

Focal adhesions were visualized in SKBR3 PTPN12 variant cells that were seeded onto glass coverslips in PRF-DMEM/F12 medium containing FN-reduced serum and allowed to adhere overnight at 37C. After adhesion, serum was removed by washing 3x with

PRF-DMEM/F12, and the cells were then cultured in the same media in the absence of serum for an additional 30 hr. Serum-starved cells were fed 2 µg/ml rat FN in the absence or presence of 10 nM E2β for 2 hr. Cells were then washed, fixed for 5 min in

4% paraformaldehyde, permeabilized in 0.05% Triton for 60 sec, and blocked in 5%

137

BSA/PBS for 30min. Cells were incubated with phosphotyrosine specific 4G10 antibody diluted 1:500 in PRF-DMEM/F12 containing 5% BSA for 60 min. Coverslips were washed in PRF-DMEM/F12, and cell-associated antibodies were detected using Alexa 594- conjugated anti-mouse IgG diluted 1:1000 and delivered in PRF-DMEM/F12 containing

5% BSA for 30 min. After staining, coverslips were washed and mounted on glass slides in Vectashield/4’6-diamidino-2-phenylindole (vector laboratories, Inc., Burlingame, CA).

FN fibril formation was examined in SKBR3 or GE11β1 PTPN12 variant cells that were seeded onto glass coverslips in PRF-DMEM/F12 medium containing FN-reduced serum.

Starved cells were fed rat plasma FN (25 µg/ml) in PRF-DMEM/F12 medium in the presence of 10 nM E2β for 18 hr and then fixed and prepared for immunostaining as described above. Fixed cells were stained with IC3 ascites diluted 1:1000 and delivered in PBS containing 1% BSA for 60 min. IC3 mAB was detected by staining with Alexa 594 conjugated anti-mouse IgG (1:1000) and processed for microscopy as described above.

All immunofluorescent images were visualized with a Nikon Eclipse 80i microscope

(Nikon, Inc., Melville, NY) equipped with a Nikon Plan Fluor 100x0.5-1.3 Oil Iris with differential interference contrast and epifluorescent capabilities. Digital images were captured using a QImaging Retiga 2000R digital camera and Nikon imaging software

(Elements Basic Research 3.0).

Transactivation Assay- B11WT and B14EV cells were seeded onto petri dishes and allowed to adhere overnight at 37C. After adhesion, serum was removed by washing 3x with PRF-DMEM/F12, and the cells were then cultured in the same media in the absence

138 of serum for an additional 3 days. Serum-starved cells were left unstimulated, stimulated with E2β for 5, 15, or 30 min, or stimulated with EGF for 15 min. Cells were lysed and equivalent amounts of total protein were electrophoresed through 8% SDS- polyacrylamide gels and transferred to PVDF. Membranes were blocked in PBS plus 3%

BSA. Phosphorylation of the EGFR was detected by probing the membrane with 4G10 antibodies delivered in 3% BSA in PBS. Antibody-antigen complexes were detected using

HRP-conjugated anti-mouse secondary antibodies diluted 1:5000 and enhanced chemiluminescence.

Adhesion Assay- 48-well plates were coated with 200 µl of serum free PRF-DMEM/F12, 2

µg/ml human FN, or 10 µg/ml collagen overnight. Wells were blocked with 5% BSA in serum free PRF-DMEM/F12 for 1 hr. SKBR3 PTPN12 variant cells were seeded in triplicate, left untreated or treated with 10nM E2β, and allowed to adhere for 2 hr. Cells that did not adhere were gently washed away with water. Adherent cells were fixed and stained with crystal violet (20ml water, 0.8ml ethanol, 0.04g crystal violet) for 5 min, and then washed 2x in large volumes of water. Crystal violet was extracted with 200 µl

10% acetic acid and absorbance was measured at 550nm.

Boyden Chamber Migration Assay- The bottom portion of transwell plates were coated with serum free PRF-DMEM/F12, 2 µg/ml human FN, or 10 µg/ml collagen overnight.

Transwells were blocked in 5% BSA in serum free PRF-DMEM/F12 for 1 hr. SKBR3

PTPN12 variant cells were placed in suspension in serum free PRF-DMEM/F12 and left untreated or treated with 10nM E2β. Cells were then plated onto transwells and

139 allowed to migrate overnight at 37C. Cells were fixed and stained with crystal violet (20 ml borate buffer and 0.04g crystal violet) for 5 min, and then washed 2x in large volumes of water. Cells that did not migrate were removed from the top of each chamber and images of membranes were captured. Each data point was measured in triplicate and measured as the mean plus/minus standard deviation.

Soft Agar- SKBR3 PTPN12 variant cells (104) were seeded into phenol red-free DMEM-

F12 media containing 2% FN reduced serum in 0.35% agarose in the absence or presence of E2β (10 nM) and supplemented with exogenous FN (2 µg/ml). Cells were grown for 10 days at 37C in a humidified chamber. Cultures were weighed every 2 days and water was replaced as needed. Images of colonies were captured at 10x magnification (brightfield).

140

Results

PTPN12 expression in human SKBR3 breast cancer or GE11β1 cells transfected with

PTPN12 shRNA, WTPTPN12, PDPTPN12, or mutant NPLH PTPN12

PTPN12 shRNA, WT PTPN12 cDNA, and PD PTPN12 cDNA were obtained from outside sources. PD PTPN12 was generated by mutating amino acid C231 to S [Sun, 2011].

PTPN12 cDNA with a mutation in the Shc binding site was generated by altering the cDNA encoding full-length human PTPN12 protein. Three point mutations were made in the NPLH region of PTPN12. PCR amplified cDNAs were subcloned into a pDEST26 vector and transfected into SKBR3 or GE11β1 cells using lipofectamine. Stable transfection of

SKBR3 and GE11β1 cells with PTPN12 shRNA (Figure 1A), and WT PTPN12, PD PTPN12, and mutant NPLH PTPN12 (Figure 1B) was confirmed by western blot analysis and immunofluorescence (Figure 2 and Figure 3). A decrease in PTPN12 expression was observed in cells with PTPN12 shRNA, and an increase in PTPN12 expression was observed in cells with WT PTPN12, PD PTPN12, or mutant NPLH PTPN12 when compared to vector control SKBR3 cells.

141

A SKBR3 B SKBR3 PTPN12 SKBR3 SKBR3 mut NPLH SKBR3 shRNA SKBR3 WTPTPN12 PDPTPN12 PTPN12 PTPN12 PTPN12

Actin Actin

Figure 1- PTPN12 expression in human SKBR3 breast cancer cells stably transfected with PTPN12 shRNA, WT PTPN12, PD PTPN12, or mutant NPLH PTPN12. Equivalent amounts of cellular protein prepared from SKBR3 breast cancer cells transfected with (A)

PTPN12 shRNA and (B) WT PTPN12, PD PTPN12, or mutant NPLH PTPN12 were immunoblotted with PTPN12 antibodies and actin

(control).

A SKBR3 SKBR3 PTPN12 shRNA PTPN12

B SKBR3 SKBR3 WTPTPN12 SKBR3 PDPTPN12 PTPN12

Figure 2- PTPN12 expression in human SKBR3 breast cancer cells stably transfected with PTPN12 shRNA, WT PTPN12, or PD

PTPN12. PTPN12 expression was measured in SKBR3 cells that were stably transfected with (A) PTPN12 shRNA and (B) WT PTPN12 or PD PTPN12 by immunofluorescence using PTPN12-specific antibodies.

142

GE11β1 GE11β1 PTPN12 shRNA

Figure 3- PTPN12 expression in GE11β1 cells transfected with PTPN12 shRNA. PTPN12 expression was measured in GE11β1 cells that were stably transfected with PTPN12 shRNA by immunofluorescence using PTPN12-specific antibodies.

Overexpression of PTPN12 inhibits E2-induced, GPER-mediated focal adhesion formation

Integrin α5β1 and Shc are integral components of E2-induced, GPER-mediated EGFR transactivation and FN fibrillogenesis [Filardo, 2000; Quinn, 2009; Magruder, in preparation]. Stimulation of human SKBR3 cells with E2 results in the recruitment of integrin α5β1 and Shc into focal adhesions [Magruder, in preparation] and fibrillar adhesions at the cell periphery [Quinn, 2009]. PTPN12 interacts with Shc [Habib, 1994] and is important in the regulation of proteins involved in focal adhesions [Zheng, 2013].

These proteins become phosphorylated in response to activation of integrin signaling and dephosphorylated in response to activation of PTPN12 [Zheng, 2013]. To determine whether PTPN12 is involved in E2β-induced, GPER-mediated focal adhesion formation,

SKBR3 PTPN12 variants were seeded onto coverslips, starved, and left untreated or stimulated with E2β for 2 hr in the presence of exogenous FN. Cells were fixed and stained with anti-phosphotyrosine antibodies (Figure 4A and Figure 4C) and anti-FAK antibodies (Figure 4B and Figure 4D) to identify focal adhesion formation. While E2β-

143 stimulated vector control SKBR3 cells demonstrated prominent focal adhesion formation, focal adhesions were not observed in unstimulated cells. This E2β-induced effect was enhanced in SKBR3 PTPN12 shRNA, SKBR3 PD PTPN12, and SKBR3 mutant

NPLH cells, and interestingly there was an increase in focal adhesion formation in these cells that were left unstimulated when compared to unstimulated vector control SKBR3 cells. SKBR3 cells overexpressing WT PTPN12 that were stimulated with E2β were unable to form focal adhesions suggesting that PTPN12 is a negative regulator of E2β- induced, GPER-mediated focal adhesion formation. Overexpression of PTPN12 may result in the inability of PTPN12 to offload from Shc, inhibiting activation of integrin

α5β1 and focal adhesion formation. Inhibition of PTPN12 activity or inhibition of

PTPN12-Shc binding results in aberrant activation of integrin α5β1 and focal adhesion formation.

144

A SKBR3 SKBR3 PTPN12 shRNA B SKBR3 SKBR3 PTPN12 shRNA

UT UT

E2 E2

C SKBR3 SKBR3 WTPTPN12 SKBR3 PDPTPN12 SKBR3 mut NPLH PTPN12

D SKBR3 SKBR3 WTPTPN12 SKBR3 PDPTPN12 SKBR3 mut NPLH PTPN12

Figure 4- Overexpression of PTPN12 inhibits the formation of focal adhesions in a GPER-dependent manner. (A, B) SKBR3 vector control or SKBR3 PTPN12 shRNA cells, and (C, D) SKBR3 vector control, SKBR3 WT PTPN12, SKBR3 PD PTPN12, or SKBR3 mutant

NPLH PTPN12 breast cancer cells grown on glass coverslips were serum-starved for 30 h and left untreated or stimulated for 2 h with

E2β (10 nM) in the presence of exogenous rat FN (2 μg/ml). Cells were fixed in 4% paraformaldehyde, permeabilized with detergent, and focal adhesions were detected using (A, C) phosphotyrosine-specific antibodies or (B, D) FAK-specific antibodies (red). Nuclei were stained with DAPI (blue).

Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated formation of FN fibrils

145

Previously, we have shown that Shc is a requirement for E2β-induced, GPER-mediated

FN fibril formation [Magruder, in preparation]. PTPN12 has been shown to be a negative regulator of Shc signaling, and has been shown to be important in the regulation of proteins involved in focal adhesion formation and migration [Zheng, 2013], but its role in GPER-mediated FN fibril formation has not been elucidated. Here, we addressed the capacity of SKBR3 or GE11β1 PTPN12 variants to form FN fibrils in response to stimulation with E2β. SKBR3 or GE11β1 PTPN12 variants were seeded onto coverslips, starved, and left untreated or stimulated with E2β in the presence of exogenous rat FN.

Cells were fixed and FN fibrils were then detected using anti-rat FN antibodies (Figure 5 and Figure 6). As shown in Figures 5 and 6, an increase in FN fibril formation was observed in E2β-treated vector control SKBR3 or GE11β1 cells when compared to untreated cells. This E2β-induced effect was enhanced in SKBR3 or GE11β1 PTPN12 shRNA, PD PTPN12, and mutant NPLH cells. Interestingly, there was an increase in FN fibril formation in untreated SKBR3 or GE11β1 PTPN12 shRNA, PD PTPN12, and mutant

NPLH PTPN12 cells when compared to vector transfected cells. There was an inhibition of FN fibril formation in E2β-treated SKBR3 or GE11β1 WT PTPN12 cells when compared to E2β-treated vector control cells. These results suggest that overexpression of PTPN12 results in the inability of PTPN12 to offload from Shc, inhibiting activation of integrin

α5β1 and FN fibril formation. Inhibition of PTPN12 activity or inhibition of PTPN12-Shc binding results in aberrant activation of integrin α5β1 and FN fibril formation.

146

Furthermore, we show the requirement of GPER in E2β-induced FN fibril formation.

GE11β1 vector control and GE11β1Δ154 cells were stimulated with E2 and exogenous rat FN as described above (Figure 6A). GE11β1 cells were able to form FN fibrils, while

GE11β1Δ154 cells were not.

SKBR3 SKBR3 WTPTPN12 SKBR3 PDPTPN12 SKBR3 mut NPLH PTPN12

Figure 5- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated formation of FN fibrils. SKBR3 vector control, SKBR3 WT

PTPN12, SKBR3 PD PTPN12, or SKBR3 mutant NPLH PTPN12 were seeded onto coverslips, serum starved, and left untreated or stimulated with E2β (10 nM) in the presence of exogenous rat FN (25 μg/ml) for 18 h. Cells were fixed with 4% paraformaldehyde and stained with rat FN specific mAB, IC3 (red). Nuclei were stained with DAPI (blue).

147

A GE11β1 GE11β1Δ154

GE11β1 GE11β1 GE11β1 mut B GE11β1 WTPTPN12 PDPTPN12 NPLH PTPN12

Figure 6- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated formation of FN fibrils in GE11β1 cells. (A) GE11β1 cells transfected with vector control or GPERΔ154, and (B) GE11β1 cells transfected with vector control, WT PTPN12, PD PTPN12, or mutant NPLH PTPN12 were seeded onto coverslips, serum starved, and left untreated or stimulated with E2β (10 nM) in the presence of exogenous rat FN (25 μg/ml) for 18 h. Cells were fixed with 4% paraformaldehyde and stained with rat FN specific mAB,

IC3 (red). Nuclei were stained with DAPI (blue).

Inhibition of PTPN12 function results in constitutive activation of EGFR

PTPN12 is a phosphatase that interacts with Shc proteins [Habib, 1994]. Shc and PTPN12 are both important in EGFR signaling and have been shown to interact with the EGFR

[Zheng, 2013]. PTPN12 antagonizes pro-mitogenic EGFR signaling by displacing Shc from the EGFR [Zheng, 2013]. A novel position for Shc, upstream of the EGFR, in GPER- mediated signaling has recently been described [Quinn, 2009], suggesting that PTPN12 may also play a role in GPER-mediated signaling upstream of the EGFR. To determine the role of PTPN12 in GPER-mediated EGFR transactivation, B11WT PTPN12+/+ and

B14EV PTPN12-/- (figure 7A) primary embryonic fibroblasts were starved and left

148 untreated or stimulated with E2β for 5, 15, or 30 min, or EGF for 30 min. Cells were lysed and equivalent amounts of total protein were electrophoresed. Phosphorylation of the EGFR was detected by probing the membrane with phosphotyrosine-specific antibodies (Figure 7B). Increased tyrosyl phosphorylation of the EGFR was observed in

B11WT cells stimulated with E2β when compared to unstimulated B11WT cells.

Equivalent levels of tyrosyl phosphorylation of the EGFR was observed in unstimulated and E2β stimulated cells, suggesting that in cells lacking PTPN12, the EGFR is constitutively activated.

A B11 WT B14 EV

PTPN12

B B11WT B14 EV

p-EGFR

UT E2 E2 E2 EGF UT E2 E2 E2 EGF (5) (15) (30) (15) (5) (15) (30) (15)

Figure 7- Inhibition of PTPN12 results in constitutive activation of the EGFR. (A) Equivalent amounts of total cellular protein prepared from B11WT and B14EV cells were immunoblotted with PTPN12 specific antibodies. (B) EGFR tyrosyl phosphorylation was measured in B11WT and B14EV cells left untreated, stimulated with E2β for 5, 15, or 30 min, or stimulated with EGF for 15 min.

Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated cell adhesion onto FN- coated substrata

149

Adhesion is important in migration. In order for a cell to migrate, adhesion between cells must loosen [Ruoslahti, 1999], and upon contact with a distant site in the body, the cell must have the ability to adhere to this new site. FN is a major cell adhesion protein that mediates anchorage through integrin α5β1 [Ruoslahti, 1999]. GPER has been shown to promote enhanced cellular adhesion interactions by modulating the affinity of integrin α5β1 for FN and also by inducing the recruitment of integrins to focal adhesion plaques [Quinn, 2009; Magruder, in preparation; Ginsberg, 2005; Luo, 2007]. PTPN12 has been shown to be important in the regulation of proteins involved in adhesion. To examine the influence of PTPN12 in E2β-induced, GPER-mediated adhesion of SKBR3 breast cancer cells for immobilized adhesive ligand, SKBR3 PTPN12 variants were detached and exposed to E2β or left untreated and seeded into polystyrene wells coated with various concentrations of FN or collagen. Following a 2 hour incubation time, cells that were not firmly attached were gently washed away and adherent cells were fixed, and cellular attachment was assessed by staining the remaining adherent cells with crystal violet. Attachment was measured as a function of eluted dyed recovered from the resulting adherent cells (Figure 8). Unstimulated SKBR3 cells adhered to both FN- and collagen- coated substrata in a dose-dependent fashion with maximum cell adhesion measured at coating concentrations of 20 µg/ml and 10 µg/ml of FN and collagen, respectively. There was an increase in the capacity of E2β-stimulated versus unstimulated SKBR3 cells to adhere to wells coated with suboptimal concentrations of FN (2 µg/ml), which is associated with increased cellular spreading. In

150 contrast, more modest differences in enhanced E2β-mediated adhesion to collagen- coated substrata were measured with no discernible difference in cellular spreading.

There was enhanced adhesivity in E2β-stimulated PTPN12 shRNA, PD PTPN12, and mutant NPLH PTPN12 SKBR3 cells when compared to untreated cells. E2β-induced increased adhesivity was eliminated in SKBR3 cells overexpressing WT PTPN12 suggesting that PTPN12 is important in GPER-mediated adhesion of SKBR3 breast cancer cells.

A 1.60 1.40 UT E2 1.20 1.00 0.80 0.60

0.40 Absorbance 0.20 0.00 SKBR3 SKBR3 PTPN12 shRNA

B 0.8 0.7 UT E2 0.6 0.5 0.4 0.3 0.2 Absorbance 0.1 0 SKBR3 SKBR3 SKBR3 SKBR3 mut WTPTPN12 PDPTPN12 NPLH PTPN12

Figure 8- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated cell adhesion onto FN-, but not collagen-, coated substrata. (A) SKBR3 vector control or SKBR3 PTPN12 shRNA, and (B) SKBR3 vector control, SKBR3 WT PTPN12, PD PTPN12, or mutant NPLH PTPN12 cells were seeded onto 48-well plates coated with FN (2 μg/ml) or collagen (10 μg/ml) and allowed to attach for 2 hr at 37C in the absence or presence of E2 (10 nM). After adhesion, unattached cells were removed by gentle washing, and the remaining adherent cells were fixed and stained with crystal violet. Excess crystal violet was washed away and cell-associated crystal violet was extracted with 10% acetic acid. Absorbance was measured at 550 nm. Each data point represents the mean +/- the standard deviation of triplicate samples. Nonspecific adhesion as measured on BSA-coated wells was subtracted.

151

Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated haptotaxis on FN- coated, but not collagen-coated substrata

Cellular adhesion is often associated with increased cellular motility as measured in haptotatic responses on immobilized adhesive ligands. Shc has been shown to be an important mediator in the promotion of GPER-dependent, E2β-induced haptotaxis of

SKBR3 breast cancer cells [Magruder, in preparation]. Furthermore, PTPN12 has been shown to be involved in the breakdown of focal adhesions and stress fibers [Angers,

1999], and is a regulator of motility. PTPN12 silencing enhances migration [Villa-

Moruzzi, 2013], and overexpression of PTPN12 inhibits motility [Zheng, 2013]. To examine whether PTPN12 inhibits GPER-mediated migration on FN-coated substrata,

SKBR3 PTPN12 variants were seeded in the presence or absence of E2β into the upper reservoirs of modified Boyden chambers containing porous polycarbonate membrane

(6.5 µm thickness, 8 µm pore) whose undersurface was coated with adhesive ligand (2

µg/ml FN or 10 µg/ml collagen). Cell migration was measured by determining the number of cells that were capable of migrating from the upper reservoir across the membrane to its undersurface (Figure 9). On FN-coated membranes, SKBR3 vector control cells stimulated with E2β showed an increase in their capacity to migrate compared with the unstimulated cells in this assay. SKBR3 vector control cells that were plated onto membranes that were coated with higher concentrations of FN (5-10 µg/ml) did not show increased haptotaxis when stimulated with E2β (data not shown),

152 suggesting that GPER enhanced migration was the product of increased recruitment of

FN receptors to cellular adhesion sites (data not shown). This result was observed in cells expressing PD PTPN12, or PTPN12 with a mutant Shc binding site. However, E2β- enhanced haptotaxis on FN was abrogated in SKBR3 cells overexpressing WT PTPN12 demonstrating that PTPN12 plays a negative regulatory role in GPER-dependent migration. All cells migrated equally well on collagen coated substrata independent of

E2β stimulation, and PTPN12 has no impact on collagen migration, suggesting that GPER signaling did not enhance migration on this extracellular matrix protein (data not shown).

153

SKBR3 SKBR3 SKBR3 mut SKBR3 WTPTPN12 PDPTPN12 NPLH PTPN12

200 180 UT E2 160 140 120 100 80 60 40

Number of Cells of Number 20 0 SKBR3 SKBR3 SKBR3 SKBR3 mut WTPTPN12 PDPTPN12 NPLH PTPN12

Figure 9- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated haptotaxis on FN-coated, but not collagen-coated, substrata. SKBR3 vector control, SKBR3 WT PTPN12, SKBR3 PD PTPN12, or SKBR3 mutant NPLH PTPN12 were left untreated or treated with E2β (10 nM) and seeded into Transwells that were left untreated or coated with either FN (10 μg/ml) or collagen (10

Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated anchorage-independent growth

The conversion of soluble FN into fibrils by mammary adenocarcinoma cells results in enhanced anchorage-indepdendent growth [Saulnier, 1997]. We have previously shown that GPER action promotes FN matrix assembly [Quinn, 2009], and that this GPER- mediated fibrillogenesis is required for anchorage-indepdendent growth in a Shc-

154 dependent manner [Magruder, in preparation]. Because PTPN12 binds to Shc and inhibits cellular events associated with the phosphorylation of Shc, we hypothesized that overexpression of PTPN12 inhibits GPER-mediated, anchorage-independent growth. To address this hypothesis, human SKBR3 cells were seeded into semi-solid media supplemented with FN-depleted fetal bovine serum in the presence of increasing amounts of exogenous FN (Figure 10). Under conditions of FN depletion, SKBR3 vector control cells were unable to form colonies, but these cells were able to form colonies in the presence of 10% fetal bovine serum which had not been depleted (data not shown).

However, SKBR3 cells formed colonies in FN reduced conditions in the presence of 2

µg/ml of exogenous FN and E2β. Overexpression of WTPTPN12 in these cells prohibited

E2β-dependent colony formation, and expression of PTPN12 shRNA, PD PTPN12, or

PTPN12 with a mutant Shc binding site resulted in enhanced E2β-dependent anchorage- independent growth. These results suggest that PTPN12 is an important mediator of

E2β-depedent growth in soft agar.

155

SKBR3 A SKBR3 PTPN12 shRNA 60 UT E2

50 UT 40

E2 Colonies of Number 10

0 SKBR3 SKBR3 PTPN12 shRNA

SKBR3 SKBR3 SKBR3 mut B SKBR3 WT PTPN12 PD PTPN12 NPLH PTPN12

100 UT E2 UT 80

20 E2 Colonies of Number 0 SKBR3 SKBR3 SKBR3 SKBR3 mut WTPTPN12 PDPTPN12 NPLH PTPN12

Figure 10- Overexpression of PTPN12 inhibits E2β-induced, GPER-mediated anchorage-independent growth. (A) SKBR3 or SKBR3

PTPN12 shRNA cells, and (B) SKBR3, SKBR3 WT PTPN12, SKBR3 PD PTPN12, or SKBR3 mutant NPLH PTPN12 cells were seeded into phenol red-free DMEM-F12 media containing 2% FN-reduced serum in 0.35% agarose in the absence or presence of E2(10 nM) and supplemented with exogenous FN (2 g/ml). Cells were grown for 10 days at 37C in a humidified chamber. Cultures were weighed every 2 days and water was replaced as needed. Images of colonies were captured at 10x magnification (brightfield).

Examples shown above are representative views of multiple experiments.

156

Discussion

In this study, we show that overexpression of PTPN12, which is known to interact with

Shc, inhibits focal adhesion and FN fibril formation, adhesion and haptotaxis on FN, and anchorage-independent growth. Moreover, we show that ablation of PTPN12 expression or function positively influences the ability of GPER to promote the formation of focal adhesions and FN fibrils, adhere and haptotax on FN, and grow in an anchorage-independent manner. We have previously shown that stimulation of ER- negative human breast cancer cells with E2β results in FN matrix assembly and transactivation of the EGFR via GPER. This mechanism of action results in the recruitment of FN-engaged ingegrin α5β1 to fibrillar adhesions and the formation of integrin α5β1-Shc adaptor protein complexes. It also promotes the formation of focal adhesions and actin stress fibers resulting in increased cellular adhesion and haptotaxis on FN in a Shc-dependent manner. Furthermore, GPER stimulation of breast cancer cells allows for FN-dependent, anchorage-independent growth. These are all events that may allow for cellular survival and tumor progression.

The phosphorylation and dephosphorylation of proteins on tyrosine residues are important in normal and malignant cell regulation [Habib, 1994]. Shc is a substrate for tyrosine kinases following treatment of cells with growth factors and hormones [Habib,

1994]. It is responsible for transmitting activation signals from the receptor or tyrosine kinases to downstream signaling components [Faisal, 2002]. We have shown that Shc is an important mediator in events associated with cellular survival [Magruder, in

157 preparation], and because PTPN12 is a phosphatase that interacts with the p52 and p66

Shc proteins, but not the p46 isoform [Habib, 1994], we hypothesized that PTPN12 is a negative regulator of Shc- and GPER-mediated events associated with cellular survival.

Others have demonstrated that PTPN12 is a negative regulator of Shc. Overexpression and antisense experiments show that PTPN12 down-regulates lymphocyte stimulation by dephosphorylation of Shc and downregulating the Ras pathway [Faisal, 2002].

FN matrix assembly is important in anchorage-independent growth, which is a predictor of metastasis [Filardo, 2006]. Integrin α5β1 is the major receptor in FN matrix assembly that is able to convert soluble FN to its insoluble form [Pankov, 2000]. Integrin α5β1 is present in both focal and fibrillar adhesions, and FN matrix assembly is dependent on the translocation of activated integrin α5β1 to these adhesions [Pankov, 2000]. FN matrix assembly occurs when integrin α5β1 binds soluble FN and converts it into fibrils that make up the matrix [Ruoslahti, 1999]. FN and integrins are expressed in the mammary gland and are affected by E2 [Quinn, 2009]. As sited by Quinn, the ability of mammary adenocarcinoma cells to convert soluble FN into fibrils results in increased responsiveness to growth factors and anchorage-independent growth [Quinn, 2009].

Integrins bind to the RGD cell attachment sequence on FN, but treatment of breast cancer cells with alternative peptides containing the RGD sequence results in antimetastatic effects [Ruoslahti, 1999]. Many PTPs, including SHP1, SHP2 (PTPN11),

PTPα, and PTPN12, are involved in focal adhesion disassembly and cell migration [Zhang,

2009]. Specifically, PTPN11 regulates focal adhesions, cell spreading, and migration

[Zheng, 2013]. Dominant-negative SHP2 mutants have altered receptor tyrosine kinase

158 signaling and integrin-dependent activation of the Erk pathway, gene activation, focal adhesion and stress fiber turnover, cell spreading and migration, and cell proliferation

[Zheng, 2013]. Furthermore, EGF-induced Ras activation is inhibited in SHP2-null cells

[Zheng, 2013]. The increased stress fiber and focal adhesion formation in SHP2-null cells may be due to an increase in Rho activity [Zheng, 2013]. A transmembrane PTP, LAR, has been shown to translocate to focal adhesions and promote their disassembly [Angers-

Loustau, 1999]. Yersinia YopH, a bacterial PTP, has been shown to dephosphorylate p130Cas and FAK, resulting in disruption of focal adhesion formation [Angers-Loustau,

1999]. Most interestingly, PTPN12 is important in the regulation of proteins involved in focal adhesion formation. These proteins become tyrosine-phosphorylated in response to the activation of the integrin signaling pathways and dephosphorylated in response to the activation of PTPN12 [Zheng, 2013]. Thus, PTPN12 controls the rate of focal adhesion turnover and cell migration [Zheng, 2013]. Plating COS-1 cells transfected with

PTPN12 on FN resulted in the movement of PTPN12 to the membrane periphery

[Angers, 1999]. Furthermore, in PTPN12-/- cells, several focal adhesion proteins were hyperphosphorylated, including FAK, paxillin, and p130CAS, and there was an increase in spreading rate on FN [Angers, 1999]. In PTPN12-/- fibroblasts, there was an increase in focal adhesions when compared to the parental fibroblasts [Angers, 1999]. These observations support our results that PTPN12 overexpression results in the inhibition of

GPER-mediated focal adhesion and FN fibril formation, while inhibition of PTPN12 function results in an increase in GPER-mediated focal adhesion and FN fibril formation

(Figures 4 and 5).

159

Integrins and growth factor receptors are both linked to cancer independently, are present in cell adhesions, and have been shown to interact with eachother [Geiger,

2011]. Integrins are required for growth factor receptor signaling and growth factor receptors are important in the regulation of integrin activation [Brizzi, 2012].

EGFR transactivation occurs following the release of HB-EGF from the cell surface via a

Gβγ-Src-Shc signaling pathway [Thomas, 2004]. Details of this Gβγ signaling have not been sorted out yet, but it has been linked to the activation of matrix metalloproteinases (MMPs), which may be responsible for the cleavage and release of

HB-EGF from the cell surface [Filardo, 2005]. In addition, a PTP may be involved in signaling events associated with growth factor release. Shc and PTPN12 have been shown to play a role in EGFR signaling. Feedback phosphorylation of Shc requires

160

PTPN12, which occupies the Shc PTB domain through the NPLH motif [Zheng, 2013].

PTPN12 antagonizes pro-mitogenic EGFR signaling by displacing Shc from the EGFR and by dephosphorylating its Grb2 binding XYN motifs [Zheng, 2013]. We have provided evidence that Shc may also promote EGFR transactivation from a novel location, upstream of the EGFR [Quinn, 2009]. Here, we further show that inhibition of PTPN12 results in the constitutive activation of the EGFR (Figure 7), but the sequence of events of this GPER-mediated EGFR activation still need to be elucidated.

Many PTPs have been shown to play a role in the regulation of cell adhesion and migration [Zheng, 2013; Yu, 1998]. Overexpression of a dominant-negative mutant of

PTP1B resulted in a defect in integrin-mediated adhesion and signaling in fibroblasts

[Arregui, 1998]. In addition, PTP-α was shown to regulate the activity of Src and this regulated cell-stratum adhesion [Harder, 1998]. The role of PTPs in events associated with cellular survival is an area for further examination; however, our results here support the observation that overexpression of PTPN12 inhibits motility [Zheng, 2013].

We show that overexpression of PTPN12 inhibits GPER-mediated adhesion and haptotaxis on FN (Figures 8 and 9). We further show that inhibition of PTPN12 function result in an increase in GPER-mediated adhesion and haptotaxis on FN.

FN matrix assembly is important in anchorage-independent growth, which is a predictor of metastasis [Filardo, 2006]. Implantation of mammary tumor xenografts in immunocompromised mice is enhanced with the introduction FN [Price]. We show that stimulation of GPER-1 enhanced FN-dependent, anchorage-independent growth in a

161

Shc-dependent manner [Magruder, in preparation]. Because we have demonstrated that Shc plays a role in anchorage-independent growth, we investigated whether

PTPN12 also plays a role in a cell’s ability to grow in an anchorage-independent manner.

We found that overexpression of PTPN12 results in the inhibition of human breast cancer cells to grow in a GPER-dependent, anchorage-independent manner (Figure 10).

We further found that inhibition of PTPN12 function results in enhanced GPER- mediated, anchorage-independent growth.

Our results support the findings that PTPN12 is a negative regulator of events associated with cellular survival in cancer. Studies have shown that loss of PTPN12 can lead to malignant transformation of human mammary epithelial cells, and restoration of

PTPN12 into these cells inhibits proliferation, tumorigenicity, and metastatic potential

[Xunyi, 2002]. In addition, PTPN12 is down-regulated in breast cancer cells when compared to normal breast tissue [Sun, 2011; Wu, 2013; Xunyi, 2013]. Low PTPN12 expression is positively associated with lymph node status, stage, and distant metastatic relapse in breast cancer [Wu, 2013]. In addition, Yuan et al. report that low PTPN12 expression is associated with increased tumor and lymph node size, lymph node metastasis, distant metastasis, and histological grade [Xunyi, 2013; Xunyi, 2002].

Furthermore, loss of PTPN12 leads to acinar morphogenesis and cellular transformation in mammary epithelial cells [Sun, 2011]. The restoration of PTPN12 in breast cancer cells inhibits proliferation, tumorigenicity, and metastatic potential [Sun, 2011]. In addition, overexpression of PTPN12 is associated with longer disease free survival [Wu, 2013].

Taken together, our results support the hypothesis that PTPN12 may have importance in

162

GPER-mediated progression of breast cancer cells. Breast cancer cells use GPER to regulate focal adhesion and FN fibril formation, adhesivity and haptotaxis on FN, and anchorage independent growth.

163

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169

Chapter 4

Overview, Future Directions, Clinical Implications

170

Overview

There is overwhelming evidence that GPER, a novel estrogen receptor, acts independently of the ER and via an alternative mechanism. Expression of the ER and

GPER is observed independently in both breast tumors and in breast cancer cells lines

[Filardo, 2012]; ER and GPER have different binding affinities for their ligands [Filardo,

205]; ER antagonists act as GPER agonists [Filardo, 2005]; the activities associated with

E2 signaling via ER and GPER are different; ER- and GPER-null nice each exhibit different phenotypes [Filardo, 2005]; and ER and GPER are associated with different clinicopathological markers of human breast cancer [Filardo, 2006].

In elucidating the signaling mechanism of GPER, several groups have shown that stimulation of GPER with E2 is associated with activated G proteins and the upregulation of adenylyl cyclase activity [Thomas, 2005; Filardo, 2005]. GPER also promotes proHB-

EGF release, EGFR transactivation, and Erk-1/-2 activation when stimulated with E2

[Filardo, 2005; Thomas, 2005]. Furthermore, it has recently been shown that GPER- dependent E2 action results in the activation of integrin α5β1, the primary FN receptor, and increased FN matrix assembly, which is significant in advanced disease because these activities allow for survival of metastatic cancer cells [Quinn, 2009]. The adaptor protein Shc is intrinsically involved in intracellular signaling events that determine growth factor responsiveness and bidirectional integrin signaling [Ravichandran, 2001]; therefore, the role of Shc was investigated in GPER-mediated signaling. Previously

[Quinn, 2009] and here we show that Shc is required for GPER-mediated events

171 associated with cellular survival such as focal adhesion and FN fibril formation, adhesion and haptotaxis on FN, and anchorage independent growth. Furthermore, because the phosphatase PTPN12 interacts with Shc and has been shown to regulate the disassembly of focal adhesions [Zheng, 2013], we investigated the role of this phosphatase in GPER-mediated signaling. Here, we show that PTPN12 overexpression inhibits GPER-mediated events associated with cellular survival, suggesting that PTPN12 is acting as an inhibitor of Shc and integrin α5β1 in GPER-mediated signaling.

Taken together, findings from other groups along with our results here suggest that upon GPER stimulation with E2, associated G proteins become activated, promoting the activation of integrin α5β1 following the phosphorylation of Shc upon the dissociation of

PTPN12 from Shc (Figure 1). The activation of integrin α5β1 then promotes the formation of focal adhesions and FN fibrils, cellular adhesion and haptotaxis, and anchorage independent growth. Events leading to the release of proHB-EGF and EGFR transactivation must still be elucidated, but it is hypothesized that an MMP may play a role in cleaving and releasing proHB-EGF from the cell surface.

172

FN Fibril Formation

E2 α5 β1 HB-EGF EGFR plasma GPER Timp-3 membrane MMP-3

b γ proHB-EGF b γ Shc Ca++ Src PTPN12 Kinase Cascades (minutes) Gene Transcription

Figure 1- Proposed mechanism of GPER action.

173

Future Directions

Aim 1- To evaluate the role of Shc in GPER-mediated events associated with tumor cell survival.

There are many experiments that can be done to further elucidate the role of Shc in

GPER-mediated events associated with tumor cell survival. Shc proteins are known to associate with integrins and localize to focal adhesions, but the role of Shc in integrin structures is not well understood [Ma, 2007]. To determine whether phosphorylated

Shc localizes to focal adhesions in response to GPER stimulation by E2β, immunofluorescence experiments using a phospho-Shc antibodies in addition to either integrin α5β1-specific or phosphotyrosine-specific antibodies should be used. Vector control SKBR3 cells and mutant Shc 317Y/F cells should be seeded onto coverslips in FN- reduced serum, starved following adhesion, and treated with exogenous FN and E2β for

2 hours. Cells should then be fixed, permeabilized, blocked, and incubated with phospho-Shc antibodies in combination with integrin α5β1 antibodies or phosphotyrosine antibodies to visualize focal adhesions. This experiment was already attempted, but there was a lack of phospho-Shc staining in all groups. There are several possible explanations for the inability to see phospho-Shc in focal adhesions. First, experimental error cannot be counted out. Based on a literature search, the specific phospho-Shc antibody used has not been used in immunofluorescne experiments before and we did not have a positive control to suggest that our conditions were optimized. Second, a more likely explanation is that phosphorylated Shc may be masked

174 by interacting proteins in immunofluorescence experiments. While using detergents used to permeabilize the cells, the phosphorylation site on Shc may be revealed and result in the dissociation of interacting proteins. To circumvent this problem, an immunoblot with cell lysates using a Triton-buffered detergent that does not interfere with Shc-integrin complexes as described by Quinn may be used [Quinn, 2009]. Another experimental alternative to visualize phospho-Shc in focal adhesions is to use total internal reflection fluorescence (TIRF) microscopy as described by Ma et al [Ma, 2007].

Ma et al used DsRed-zyxin to visualize focal adhesions and added a GFP tag to Shc [Ma,

2007]. Zyxin is an adaptor protein that is abundant protein in focal adhesions.

Shc’s role in anoikis is another area for further investigation. Anoikis is rapid apoptotic death of cells that are unable to grow in an anchorage-independent manner following detachment from a matrix. While each of the three Shc isoforms (p66Shc, p52Shc and p46Shc) have been shown to localize to focal adhesions, it is important to note that the p66Shc isoform appears to play a unique role in sensing cell adhesion, as p66Shc promotes anoikis via RhoA activation in detached cells [Ma, 2007]. Likewise, under conditions of oxidative stress, p66Shc uncouples the ras-mek-erk signaling cascade

[Aray, 2008]. These findings are consistent with studies which have shown that lung cancer cells lacking p66Shc display traits associated with advanced cancer, including anchorage independent growth [Ma, 2010]. However, GPER enhanced adhesivity to FN does not simply appear to be a result of preferential recruitment of the p52Shc or p46Shc isoforms to integrin α5β1 as human breast cancer cell lines that express p66Shc remain competent with regards to their capacity to promote integrin α5β1-dependent

175

EGFR transactivation and enhanced FN adhesivity and growth in soft agar [Quinn,

Magruder and Filardo, unpublished results]. SKBR3 cells lack the p66Shc isoform, but we have previously shown that MDA-MB-231 cells, which express p66Shc, undergo GPER- mediated adhesion and anchorage-independent growth [Quinn, Magruder, Filardo, unpublished results], suggesting that absence of p66Shc may not be a requirement for growth in soft agar. On the other hand, Ma et al have shown that vector control 293 cells form colonies in soft agar; however, when transfected with p66Shc, colony formation is inhibited [Ma, 2007]. It would be worthwhile to transfect SKBR3 cells with p66Shc to investigate whether the p66Shc isoform plays a role in GPER-mediated events associated with cellular survival including FN matrix assembly, adhesion onto FN, and anchorage-independent growth. In addition, cell death assays may provide additional information about the role of p66Shc in GPER-mediated adhesion onto FN. To carry out cell death assays, cells should be plated onto uncoated or FN coated low attachment 24- well plates. DNA fragmentation can be assessed using ELISA. Mitochondrial release of cytochrome c can be assessed by digitonin permeabilization [Ma, 2007]. Mitochondria can be harvested by centrifugation, and fractions can be immunoblotted for cytochrom c [Ma, 2007].

Another area for further investigation is Shc’s role in fibrillar adhesion formation.

Fibrillar adhesions are elongated matrix contacts that form when integrin α5β1 and tensin translocate away from focal adhesions [Geiger, 2011]. They bind FN parallel to actin bundles and tensin, and are lacking in focal adhesion proteins such as paxillin and vinculin [Pankov, 2000]. To our knowledge, Shc’s role in fibrillar adhesion formation has

176 not been investigated. To determine whether Shc interacts with integrin α5β1 during fibrillar adhesion formation, immunofluorescence costaining for phospho-Shc and

SNAKA-51 (an antibody that detects FN-occupied integrin α5β1 conformers [Quinn,

2009]) should be conducted in untreated and E2β-treated vector control and mutant

Shc 317Y/F SKBR3 cells.

Aim 2- To evaluate the role of PTPN12 in GPER-mediated events associated with tumor cell survival.

There are many experiments that can be done to further elucidate the role of PTPN12 in

GPER-mediated signaling. As a phosphatase, PTPN12 dephosphorylates its substrates.

One of the most important experiments to be done is an immunoprecipitation of

PTPN12 from lysates of SKBR3 cells transfected with PTPN12 shRNA, WT PTPN12, PD

PTPN12, and mutant NPLH PTPN12 and blot for phosphotyrosine. This would verify the loss of PTPN12 function in SKBR3 cells expressing PTPN12 shRNA, PD PTPN12, and mutant NPLH PTPN12. To carry out this experiment, SKBR3 PTPN12 variants should be seeded onto petri dishes and allowed to adhere overnight. Cells should be serum starved for 30 hours, and following starvation cells should be left untreated or treated with E2β for 15 min. Following stimulation, cellular proteins should be extracted in

Triton-buffered detergent as described by Quinn et al [Quinn, 2009]. Protein concentrations should be determined and 500 µg of protein should be incubated with protein G-agrose beads for 1 hour at 4C. Beads should be collected by centrifugation,

177 and lysate incubated with 2 µg/sample of PTPN12 antibody for 3 hours at 4C. Lysate and antibody should be incubated with fresh G-agarose beads overnight at 4C. Beads should then be washed with Triton lysis buffer, and incubated at 95C for 5 min with 2x Laemmli sample buffer containing 700mM β-mercaptoethanol. Samples should be centrifuged, and supernantant collected by pin-hole elution. Samples should be electrophoresed through 8% SDS-polyacrylamide gels and transferred to PVDF. Membranes should be blocked in PBS plus 3% BSA, and phosphorylated proteins can be detected by probing the membrane with phosphotyrosine 4G10 antibody delivered in 3% BSA in PBS.

Antibody-antigen complexes can be detected using HRP-conjugated secondary antibodies diluted 1:5000 and enhanced chemiluminescence. It is expected that there will be an increase in phosphorylated proteins in vector control SKBR3 cells treated with

E2β when compared to untreated cells. This effect is expected to be inhibited in SKBR3 cells overexpressing WT PTPN12, and enhanced in SKBR3 cells expressing PTPN12 shRNA, PD PTPN12, or PTPN12 with a mutant Shc binding site. Overexpression of

PTPN12 will result in an increased ability of cells to dephosphorylate proteins, while a decrease in PTPN12 expression or function will result in a decreased ability of cells to phosphorylate proteins. Cells expressing a form of PTPN12 with a mutation in its Shc binding site will lack to ability to bind to Shc, which will result in the inability to dephosphorylate proteins associated with focal adhesions.

After verifying the gain or loss of function of the PTPN12 variants, the ability of PTPN12 variants to dephosphorylate specific substrates should be tested. Because Shc is a necessary intermediate of E2β-induced GPER signaling, the ability of PTPN12 variants to

178 dephosphorylate Shc should be tested first. This can be accomplished by immunoprecipitating PTPN12 as described above and then probing for phosphorylated

Shc with a phospho-Shc antibody. It is expected that there will be an increase in phospho-Shc in vector control SKBR3 cells stimulated with E2β when compared to unstimulated cells. This effect is expected to be inhibited in SKBR3 cells overexpressing

WT PTPN12, and enhanced in SKBR3 cells expressing PTPN12 shRNA, PD PTPN12, or

PTPN12 with a mutant Shc binding site. Other PTPN12 substrates that should be tested include P130cas, Hefl, Sin, Csk, Grb2, Paxillin, and Hic5.

We have shown that stimulation of PTPN12-expressing B11WT mouse fibroblasts with

E2β results in EGFR transactivation, while phosphorylation of the EGFR was not observed in unstimulated cells. On the other hand, the EGFR of untreated PTPN12-null

B14EV mouse fibroblasts is constitutively activated in unstimulated cells. We would like to further examine EGFR transactivation of SKBR3 expressing PTPN12 shRNA, WT

PTPN12, PD PTPN12, or PTPN12 with mutation in the Shc binding site. Transactivation experiments should be conducted by seeding SKBR3 PTPN12 variants onto petri dishes and allowing them to adhere overnight. Cell should be serum-starved for 3 days, and following starvation, stimulated with E2β, EGF, or left untreated for 15 min. Cells should be lysed and equivalent amounts of total protein should be electrophoresed through 8%

SDS-polyacrylamide gels and transferred to PVDF. Membranes should be blocked in PBS plus 3% BSA. Phospho-EGFR can be detected by probing the membrane with phosphotyrosine antibodies delivered in 3% BSA in PBS. Antibody-antigen complexes can be detected using HRP-conjugated antimouse secondary antibodies diluted 1:5000

179 and enhanced chemiluminescence. It is expected that there will be an increase in phosphorylation of the EGFR in E2β-treated vector control SKBR3 cells when compared to untreated cells. This effect is expected to be inhibited in cells overexpressing WT

PTPN12, and enhanced in cells expressing PTPN12 shRNA, PD PTPN12, and PTPN12 with a mutant Shc binding site. EGFR phosphorylation is expected to be observed in all EGF- treated cells.

Affinity purification coupled to mass spectrometry has been used to study protein- protein interactions of kinases and phosphatases [Kean, 2012]. It can be performed on proteins following the use of immunprecipitating antibodies [Kean, 2012]. It identifies entire protein complexes and mixtures of multiple protein complexes [Kean, 2012]. It can also monitor dynamic changes in protein-protein interactions, which is important when studying the interactions of kinases and phosphatases [Kean, 2012]. It can also identify phosphorylation, providing information about signaling pathways. Affinity purification and mass spectrometry should be used to identify proteins in PTPN12 complexes and the phosphorylation of PTPN12 variants. Affinity purification coupled to mass spectrometry can be carried out using FLAG-tagged PTPN12 or by immunoprecipitating PTPN12. Cells should be lysed, and the immunoprecipitated

PTPN12 bait can be affinity purified. The protein mixture should be digested with trypsin and the peptides can be separated and fragmented. The mass to charge ratio can be measured by mass spectrometer. The identity of peptides and corresponding proteins can be determined by comparing acquired data with a database of theoretical spectra using a software program [Kean, 2012].

180

Aim 3- To determine the spatiotemporal relationships of MMP-3 and integrin α5β1 in

GPER-mediated fibrillogenesis, HB-EGF release, and EGFR transactivation.

Summary

Expression of the transmembrane estrogen receptor, GPER, in primary breast tumor biopsy specimens is associated with tumor progression variables that define advanced disease, including increased tumor size and the development of extra mammary metastases [Thomas, 2005]. Studies in vitro have shown that GPER action coordinates

FN matrix assembly [Filardo, 2005] and EGFR transactivation [Charest, 1996], two cellular activities associated with tumor cell survival. These activities require activation of integrin α5β1 from its default low affinity state and its subsequent translocation to fibrillar adhesions. They also require the adaptor protein, Shc [Quinn, 2009; Magruder, in preparation]. Other signaling effectors in GPER-mediated signaling are still being elucidated.

Matrix metalloproteinases (MMPs) play an important role during development and tumor progression by modifying the cellular microenvironment [Lochter, 1998]. Altering

MMP activity can enhance or inhibit signaling through ECM receptors with effects on migration and invasion [Lochter, 1998]. MMP-3 has been shown to be used by mammary tumor cells for invasion and has been cloned as a metastasis specific gene

[Lochter, 1998]. Furthermore, increased expression of MMP-3 in the tumor

181 microenvironment is associated with a poor prognosis in breast cancer [Sympson, 1994;

Duffy, 2000], and transgenic mice expressing autoactivated MMP-3 develop breast cancer [Sympson, 1995]. Given these results, MMP-3 may function as part of a transmembrane signaling complex with integrin α5β1 to promote GPER-mediated events associated with cellular survival.

While it is clearly understood that MMP-3 is involved in tumor cell invasion, the concept that it is involved in FN matrix assembly and remodeling is not appreciated. The experiments outlined here will provide a better mechanistic understanding of the post-

FN binding exoplasmic events that regulate integrin α5β1-mediated events associated with cell survival. To investigate whether MMP-3 functions as part of this transmembrane signaling complex with integrin α5β1 to promote GPER-mediated events, the spatiotemporal interaction between integrin α5β1, MMP-3, and tissue inhibitors of metalloproteinases (TIMPs) will be studied. Immunocytofluorescent and biochemical approaches [Quinn, 2009] will be used to measure codistribution and/or association of MMP-3 and integrin α5β1. Indices of cell survival will be measured in the presence or absence of E2β, and cancer cell invasion will be assessed. Furthermore, the effect on E2β-induced fibrillogenesis and EGFR transactivation will be determined.

TIMP-3 will be coexpressed with MMP-3 to address its influence on α5β1 activation, FN matrix assembly, release of growth factor, and activation of EGFR.

182

Proposed Methodology

Human SKBR3 breast cancer cells and mouse 4T1 breast cancer cells will be used to carry out the experiments in the aim. SKBR3 cells express low levels of MMP-3, while murine 4T1 cells express high levels of MMP-3 complexed with TIMP. All cell cultures will be grown in phenol red-free DMEM/Ham’s F12 media supplemented with 5% FBS and gentamicin.

To generate molecular clones of MMP-3, intact human MMP-3 will be expressed from pcDNA3.1Zeo(+). An autoactivated version of MMP-3 lacking the amino terminal pro- inhibitory domain will be generated by PCR. The amplified product will be digested with

EcoRI and NotI, and then subcloned into the same vector. Intact or autoactivated MMP-

3 will then be transfected into SKBR3 or 4T1 cells. Transfected cells will be selected by antibiotic resistance and expanded. Following expansion in culture, MMP-3 expression will be confirmed by immunofluorescent analysis and western blotting. MMP-2 will serve as a control. To generate molecular clones of TIMP-3, flag-tagged TIMP-3 will be transfected into SKBR3 or 4T1 cell sublines expressing MMP-3 [Quinn, 2009].

MMP-3 shRNAs will be used to measure the effect of inhibition of MMP-3 on measures of cellular survival. SKBR3 or 4T1 cells will be co-transfected with mRFP and commercially available shRNAs encoding GPER, ATIIR (control), MMP-3, or MMP-2

(control) (Santa Cruz Biotechnologies). Subpopulations of mRFP positive cells with reduced target protein expression will be selected by immunofluorescence using Alexa

183

488 secondary antibodies and a combination of flow cytometry and/or clonal selection using an inverted microscope. In the case that clonal populations are isolated, individual low expressing clones will be pooled. Following expansion in culture, knockdown will be confirmed by immunofluorescent analysis and western blotting.

MMP-3 activity will be measured by zymography and by the use of fluorigenic substrates.

The sub-cellular distribution of MMP-3 and TIMP-3 in focal or fibrillar adhesions and their association with integrin α5β1 will be visualized using dual and triple immunofluorescent analysis. Cells will be seeded onto glass coverslips in PRF-

DMEM/F12 medium containing FN-reduced serum and allowed to adhere. After adhesion, serum-starved cells will be fed rat plasma FN in the presence or absence of

E2β (10 nM) for 2 hours and then fixed. Cells will be incubated with specific antibodies that recognize integrin α5β1, phosphotyrosine, ligand-occupied α5β1 (SNAKA-51), FN, vinculin, or MMP-3, and detected using appropriate secondary antibodies derivatized with Alexa 350-, 488- or 594-fluorescent dyes. All immunofluorescent images will be visualized with a Nikon Eclipse 80i microscope equipped with a Nikon Plan Fluor

100x0.5-1.3 Oil Iris with differential interference contrast and epifluorescent capabilities. Digital images will be captured using a QImaging Retiga 200R digital camera and Nikon imaging software [Filardo, 2002].

184

The involvement of MMP-3 in fibrillogenesis will be visualized using immunofluorescent analysis. Cells will be seeded onto glass coverslips in PRF-DMEM/F12 medium containing

FN-reduced serum and allowed to adhere. After adhesion, serum-starved cells will be fed rat plasma FN in the presence or absence of E2β (10 nM) for 18 hours and then fixed. Cells will be incubated with specific antibodies that recognize rat FN and detected using appropriate secondary antibodies derivatized with Alexa 350-, 488- or 594- fluorescent dyes.

The involvement of MMP-3 in EGFR transactivation will be determined in serum starved cells that will be left untreated or treated with E2β (10nM) or EGF for 15 min. Cells will be lysed and equivalent amounts of total protein will be electrophoresed through 8%

SDS-polyacrylamide gels and transferred to PVDF. Membranes will be blocked in PBS plus 3% BSA, and phospho-EGFR will be detected by probing the membrane with phosphotyrosine antibodies delivered in 3% BSA in PBS. Antibody-antigen complexes will be detected using HRP-conjugated antimouse secondary antibodies diluted 1:5000 and enhanced chemiluminescence.

Cell invasion will be measured using BIOCOAT invasion chambers. Invasion into Collagen

I or Matrigel of cells stimulated with E2β, EGF, or vehicle will be determined. Cells seeded onto the upper surface of the gel will be removed and invading cells on the lower surface of the membrane will be viewed by epifluorescence.

185

Cellular survival will be measured by clogenicity, mammospheroid formation, and anchorage-independent growth in soft agar assays. Clonogenicity will be examined by measuring the efficiency by which cells seeded at a single cell per well by limiting dilution grow to confluence in a 96-well cup. The relative efficiency of the clonogenicity of the manipulated cells will be compared to the appropriate controls as well as vector transfected controls. Clonogenicity will be measured as the percentage of wells that form colonies relative to the number of cells plated. Each experiment will be conducted in triplicate [Smalley, 2005].

Mammospheroid formation will be measured to determine the quantity of cells capable of in vitro cell renewal [Habib, 1994]. Cells will be cultured in suspension in serum-free

DMEM-F12. After two weeks, the number and size of mammospheres will be measured using Zeiss Axiovision software [Davidson, 2001]. Spheres will be collected by gentle centrifugation and dissociated enzymatically and mechanically to single cells. The dissociated cells will be sieved, analyzed microscopically for single-cellularity, and cell dispersion will be repeated as necessary. Dissociated cells will be used to determine the tumorigenicity of these cultures [Davidson 2001].

Anchorage-independent growth will be measured for cells suspended in defined media with or without E2β to simulate three different 3-D environments. The relative efficiency to form colonies will be compared in cells plated in 0.37% agarose in FN- depleted serum versus the same media reconstituted with low, medium, or high

186 concentrations of FN (2, 5, or 20 µg/ml). Where significant differences in colony formation are measured, mitotic indices and apoptotic rates will be determined from cells that are recovered from their 3-D environment as detailed below.

Mitotic index will be determined by embedded cells (5x103/mL) in 3-D gels (100 µl final volume) in 96-well plates and incubated in growth media. Cells will be pulsed for 48 hours with [3H]-thymidine (1 miCi/well). Gels containing cells will be removed from the plate and dialyzed against PBS for 24 hours to remove free [3H]-thymidine. Gels will then be lysed in SDS and radioactivity will be measured.

Apoptotic index will be measured by TUNEL. Apoptosis will be measured by in situ labeling. Gels will be fixed in 4% paraformaldehyde and then DMSO-methanol.

Apoptosis will be detected by Apoptag using Alexa 488-coupled digoxigen antibodies.

Apoptotic cells will be counted in 5 different high power (40x) microscopic fields and the apoptotic index will be expressed as follows: (number of positive apoptotic cells

(green)/number of total cells (red)/microscopic field) x 100%. After counting and image acquisition, the apoptotic index will also be quantified measuring the ratio of green:red fluorescence.

Expected Results and Discussion

187

Because endogenous MMP-3 protein is difficult to detect in SKBR3 breast cancer cells, intact MMP-3 containing its pro-inhibitory domain or amino terminally truncated MMP-

3 lacking this domain (autoactivated) will be overexpressed directly into SKBR3 cells. An expression cassette capable of expressing intact human MMP-3 has already been created and stably transfected into HEK-293 cells (Figure 2). HEK-293 cells stably transfected with MMP-3 express greater levels of MMP-3 protein when compared to

HEK-293 cells that were vector transfected or transfected with GPER (Figure 2). While transfection of MMP-3 into HEK-293 cells demonstrates that the method of transfection is effective, for consistency it is important that the same cell line used in the Shc and

PTPN12 experiments of Aims 1 and 2 is used in the MMP-3 experiments.

A HEK-293 HEK-293 GPER HEK-293 MMP-3

MMP-3

B HEK-293 GPER MMP-3 HEK-293 GPER MMP-3

MMP-3 MMP-2

Figure 2- Stably transfected HEK-293 cells expressing human MMP-3. HEK-293 cells stably transfected with MMP-3 express greater levels of MMP-3 protein when compared to HEK-293 cells that were untransfected or transfected with GPER as shown by (A) immunofluoresence and (B) western blot analysis.

188

To evaluate the effects of MMP-3 knockdown on measures of tumor cell survival MMP-3 shRNAs will be used. It is expected that MMP-3 shRNA will reduce MMP-3; however, there is a possibility that commercially available MMP-3 shRNAs may fail to significantly reduce MMP-3 target protein expression. If this is the case, siRNAs will be evaluated for knockdown activity and shRNAs will be modeled.

In determining the subcellular distribution of MMP-3 in focal and fibrillar adhesions, it is expected that MMP-3 will associate with active integrin α5β1-Shc complexes and be recruited to focal and fibrillar adhesions upon cellular stimulation with E2β. In the case that overexpression of intact or autoactivated MMP-3 results in the loss of E2β- dependent focal and fibrillar adhesion formation due to the expression of excess protease, MMP-3 from the conditioned media of HEK-293 cells engineered to express recombinant MMP-3 will be titrated on SKBR3 cells. Furthermore, MMP-3 association with low affinity integrin α5β1 either prior to or following E2β stimulation may require a stochiometric complex with TIMPs; thus, relevant TIMPs may need to be overexpressed and/or evaluated in SKBR3 cells and in HEK-293 supernatants. In the event that MMP-3 does not concentrate at focal or fibrillar adhesions or that an association between

MMP-3 and integrin α5β1 is not detected, MMP-2 will be investigated.

The role of MMP-3 in E2β-dependent fibril formation and EGFR transactivation has begun to be investigated. Human SKBR3 cells were transiently transfected with sense and antisense oligonucleotides derived from MMP-2 or MMP-3 or treated with an MMP

189 inhibitor, ilomastat, prior to stimulation with E2β or EGF. De novo synthesis of insoluble

FN or EGFR tyrosyl phosphorylation were then measured. It was determined that transient transfection of phosphothiorate antisense oligo-nucleotides specific for MMP-

3, but not MMP-2, abrogated GPER-mediated FN fibril formation and EGFR transactivation (Figure 3). In addition, SKBR3 breast cancer cells were left unstimulated or stimulated with E2β in the presence or absence of illomastat, an MMP-3 inhibitor.

E2β stimulated cells formed FN fibrils when compared to unstimulated cells, but this effect was inhibited in illomastat treated cells (Figure 4). To confirm these findings, FN fibril formation and EGFR transactivation will be investigated in cells transfected with intact or truncated MMP-3 and in cells transfected with MMP-3 shRNAs. It is expected that E2β stimulation will promote FN fibril formation and EGFR transactivation in cells with intact or truncated MMP-3, but not in cells with MMP-3 shRNAs.

A B E2 EGF

2AS 2S 3AS

- - -

2AS

3S 2S

3AS

- -

MMP

MMP MMP MMP

E2 : - + + + + +

MMP

None

P4C10

MMP MMP

P4C10

Unstimulated

MMP

GM6001 GM6001

Insoluble erbB1

IP:erbB1 blot:pptyr Soluble

Figure 3- Requirement of MMP-3 for GPER-mediated fibrillogenesis and EGFR transactivation. Human SKBR3 breast cancer cells were transiently transfected with sense and antisense oligonucleotides derived from MMP-2 or -3 or treated with an MMP inhibitor, illomastat (GM6001) or integrin β1 subunit specific mAB, P4C10 prior to stimulation with E2β or EGF. (A) De novo synthesis of insoluble FN or (B) EGFR tyrosyl phosphorylation were measured.

190

UT E2 FN E2 + FN

UT Ilomastat

Figure 4- Illomastat inhibits GPER-mediated FN fibril formation. Quiescent SKBR3 breast cancer cells were fed exogenous rat FN and treated with E2β for 18 hours. Cells were fixed and stained with FN-sepecific mAB, IC3. Nuclei were stained with DAPI.

The role of MMP-3 in GPER-mediated invasion and cell survival will be determined. It is expected that expression of MMP-3 will promote E2β-dependent invasion and cell survival as measured in matrix invasion assay, clogenicity, mammospheriod formation, and anchorage-independent growth assays. Cell invasion and cell survival will be reduced in MMP-3 shRNA expressing cells. Furthermore, mitotic index will be increased in E2β-stimulated cells expressing intact or autoactivated MMP-3, but reduced in cells expressing MMP-3 shRNA. On the other hand, apoptotic index will be reduced in cells expressing intact or autoactivated MMP-3, but increased in cells expressing MMP-3 shRNAs.

191

Aim 4- Mouse Model of E2β-induced metastatic growth.

Summary

It is widely appreciated that GPER promotes local release of EGF, FN matrix assembly, and anchorage-independent growth [Filardo, 2002]. These in vitro correlates suggest that GPER facilitates the survival of metastatic tumor cells after they seed a foreign tissue and these results are consistent with data that links GPER to the development of extra-mammary metastasis in human disease [Filardo, 2000]. To test this idea in a preclinical model, ER-negative murine 4T1 breast cancer cells will be employed.

Transplanted 4T1 cells spontaneously metastasize to lung, liver, lymph nodes, and brain in BALB/c mice and this animal model closely mimics stage IV human breast cancer

[Filardo 2002; Wang, 2008]. They are ideal for the studies outlined here because their capacity to infiltrate and form secondary tumor outgrowths in experimental metastasis models is dependent upon E2β [Filardo, 2006; Filardo, 2005]. Their capacity to colonize the lung is impaired in an ovariectomized (OVX) host and is restored upon supplementation with E2β, suggesting that their metastatic behavior is E2β-dependent.

It is likely that 4T1 cells metastasize because they express alternate estrogen receptors.

Recent evidence suggests that GPER is an alternate estrogen receptor that functions autonomously from the ER [Rios, 2001; Mitra, 2006; Kanda, 2004; Vivacqua, 2006].

Expression of carboxyl truncation mutant, GPERΔ154, selectively inhibits E2β-dependent

EGFR transactivation, FN matrix assembly, and anchorage-independent growth by breast cancer cells [Filardo, 2002]. In addition, preliminary data shows that this

192 interfering mutant also attenuates 4T1 metastasis (Figure 5). Here, the influence of exogenous E2β to promote GPER-dependent metastasis of 4T1 vector and 4T1

GPERΔ154 cells will be examined. To test the hypothesis that Shc and PTPN12 signaling effectors in GPER-mediated signaling play important roles in GPER-dependent breast tumor metastasis, 4T1 cells will be employed. The efficiency by which 4T1 vector versus

4T1 GPERΔ154 both transfected with (i) WT Shc, (ii) Shc Y317F, (iii) PTPN12 shRNA, (iv)

WT PTPN12, (v) PD PTPN12, or (vi) PTPN12 mutant NPLH cells metastasize in WT and

OVX BALB/c mice, and the influence of supplemental E2β on their metastatic behavior will be examined.

Proposed Methodology

Murine 4T1 breast cancer cells will be used in this aim because they are ER-negative, produce detectable levels of GPER, form E2β-dependent colonies in soft agar, and serve as a syngeneic model for E2β-induced stage IV human disease [Filardo, 2002; Wang,

2008]. All cell cultures will be grown as described above.

Shc (WT Shc, ShcY317F) and PTPN12 (WT PTPN12, PD PTPN12, PTPN12 mutant NPLH) molecular clones generated in aims 1 and 2 will be used. All forms of Shc and PTPN12 clones will be transfected into parental 4T1 cells or 4T1 HA-GPERΔ154 cells [Filardo,

2002] as described in aims 1 and 2. shRNAs for PTPN12 will be generated as described in aim 2. Shc and PTPN12 expression will be confirmed as described in aims 1 and 2.

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Eight week-old, female BALB/c mice and OVX BALB/c mice (Charles River Labs) will be maintained in accordance with Institutional Animal Care and Use Committee guidelines.

Mice will receive phytoestrogen-reduced chow and water ad libitum. Two weeks prior to tumor cell inoculation, mice will receive 30d E2β release pellets [Filardo, 2006;

Filardo, 2005]. Control mice will receive E2α pellets. 4T1 vector or 4T1 HA-GPERΔ154 derivative cells (105 in 0.1 ml of HBSS) will be injected intravenously into the tail vein or subcutaneously. Changes in body weight and the health of the mice will be monitored daily. Mice showing signs of distress or at intervals between 2-6 weeks following tumor implantation will be sacrificed and evaluated for tumor metastasis.

Mice will be evaluated for the presence of 4T1 tumor nodules. Small tumor nodules may be identified by fluorescent microscopy. Occult tumor cells will be measured as an index of drug resistance or fluorescence. Single cell suspensions will be made from select tissues. To detect drug resistant 4T1 tumor cells, portions of the tissue-derived cell suspension will be placed into culture vessels and treated with 6-thioguanine or hygromycin to allow for the selection of tumor cells. eGFP-tagged 4T1 tumor cells in cell suspensions will also be detected by flow cytometry, or cell suspensions containing suspect tumor cells will be treated with DAPI and then lysed. The metastatic index will be quantified by measuring the ratio of green fluorescence to blue fluorescence in lysates using the CytoFluor II fluorescent plate reader.

194

To determine mitotic index, two hours prior to sacrifice, mice will receive BUdR (1g/ kg) in saline. Secondary tumor nodules will be excised, cooled to –20C, and embedded in

OCT. Tumor tissue will be cryosectioned, fixed in acetonemethanol, and stained with

BUdR antibodies. Apoptotic index will be determined by TUNEL as described by Wang

[Wang, 2008].

To conduct Immunohistochemistry, Paraffin-embedded sections (4 µM) will be stained with antibodies as described previously [Filardo, 2002]. Sections will be incubated with antibodies specific for GPER, HA epitope, proHB-EGF, Shc, GST, PTPN12, EGFR/erbB1, integrin α5β1, and FN. Immunocomplexes will be visualized by the ABC peroxidase method (Vector Labs), and sections will be counterstained with hematoxylin.

The association between treatment groups, the occurrence, and location of a metastasis will be tested using a chi square test or a Fisher’s exact test as needed. Continuous variables will be tested for normality using the Kolmogorov-Smirnov test. Quantitative data resulting from measurements of the metastatic tumors will be compared between multiple treatment groups using the parametric one-way ANOVA test as described by

Wang [Wang, 2008].

Expected Results and Discussion

195

To determine whether the E2β-mediated effect is tumoral or extratumoral, endogenous

GPER signaling was inhibited by expressing an HA-tagged GPER interfering mutant (HA-

GPERΔ154) in preliminary studies. Expression of HA-GPERΔ154 selectively interferes with E2β-dependent fibrillogenesis and anchorage-independent growth, but has no influence on angiotensin II-mediated fibrillogenesis or EGFR transactivation (data not shown). Introduction of 4T1 breast cancer cells (105) via tail vein injection results in metastasis to the lung, liver, lymph nodes, and brain [Quinn, 2009; Wang, 2008]. Mice

(6/6) receiving vector transfected cells died of disease and developed pulmonary metastases with a mean overall survival time of 18 days (Figure 5). In contrast, mice receiving 4T1 HA-GPERΔ154 cells showed significantly longer survival times with half

(3/6) of the mice surviving 70 days after inoculation (mean overall survival time of 45.2 days). None of the surviving HA-GPERΔ154 mice showed signs of disease, while mice that died of delayed disease showed lung metastases. These data indicate that expression of functional GPER facilitates the outgrowth of pulmonary metastases and inactivation of GPER reduces the capacity of ER-negative tumor cells to assemble FN matrices, undergo anchorage-independent growth, and form lung metastases.

196

A B

Figure 5- Experimental pulmonary metastasis of 4T1 breast cancer cells. Vector- or HAGPERΔ154- transfected 4T1 cells (105) were injected into the tail vein of intact, 10 week-old female mice (6 mice per group). (A) Example of pulmonary lesions from a mouse receiving vector transfected cells. (B) Kaplan Meier curve plotting cumulative survival as a function of time. Mean overall survival times of 18 and 43 days were measured for mice receiving 4T1 cells with intact or inactivated GPER, respectively

Initial experiments will be required to determine the optimal dose of 4T1 cells necessary to measure E2β-induced metastasis. In terms of detecting secondary tumors, these studies will focus on pulmonary metastases; however, axillary and inguinal lymph nodes, bone, brain, and other visceral organs will be examined for the presence of 4T1 cells.

While 4T1 cells form grossly detectable secondary nodules (Figure 5), they are resistant to 6-thiogiuanine and this property has been previously exploited to measure micro- metastases [Filardo, 2002; Wang, 2008]. To further facilitate detection of metastatic 4T1 cells in situ and quantification of micro-metastases by flow cytometry, cells will be re- transfected with enhanced green fluorescent protein (eGFP) and then selected by forward sorting. In some experiments, the presence of occult tumor cells will be quantified by measuring GFP in cellular preparations by flow cytometry or in lysates by fluorimitry. Select secondary tumor masses will be evaluated by immunohistochemistry.

197

The work outlined here tests the hypothesis that GPER facilitates E2β-dependent metastasis of 4T1 cells and that Shc and PTPN12 play an important role in this signaling.

Based on prior work by others, it is expected that E2β will promote metastasis of 4T1 vector cells to a greater extent than HA-GPERΔ154 cells (Figure 5). In addition, metastasis will be reduced in 4T1 vector expressing Shc Y317F and WT PTPN12. It should be noted that because integrin α5β1 activation and FN matrix assembly occur upstream of proHB-EGF release and that the EGFR is not required for FN matrix assembly [Filardo,

2002], the effects of extratumoral GPER are likely to be partial on breast tumors cells expressing inactivated GPER. The experiments outlined here will provide a better understanding of the role that PTPN12 and Shc play in the development of E2β- mediated metastases through GPER signaling.

198

Clinical Implications

Breast cancer is the most common malignancy among women [Sun, 2011], with approximately one million new cases diagnosed each year [Xunyi, 2002]. Furthermore, it is the most common cause of cancer-related death in women [Bozza, 2013]. TNBC is an aggressive and deadly type of breast cancer [den Hollander, 2013]. Patients with TNBC generally present at more advanced stages and have a high rate of recurrence

[Chiorean, 2013]. The death rate at 5 years post-diagnosis in patients with TNBC is twice that of patients with non-TNBC [Steinman, 2013]. These tumors demonstrate molecular heterogeneity, which attributes to the lack of an effective target for treatment [den

Hollander, 2013]. In order to create an effective pharmaceutical agent for TNBC, a target that is present in the majority of TNBC tumors must be discovered [Steinman, 2013].

GPER expression in primary tumors is positively associated with tumor size, HER2 expression, and the presence of extramammary metastases [Filardo, 2006]. GPER may have significance in the treatment of TNBC and in the treatment of non-TNBC that fail to respond to ER antagonists. As discussed previously, patients with ER-positive breast tumors are often given tamoxifen or other ER antagonists to prevent reoccurrence of the tumor, but it has been shown that ER antagonists are GPER agonists in cultured breast cancer cell lines [Thomas, 2005]. One study showed that tamoxifen reduces recurrence in women with a history of lobular carcinoma by only 56% [den Hollander,

2013]. In addition, approximately 25% of patients with ER-positive tumors do not respond to tamoxifen based on data collected at 5-year follow-up [Filardo, 2006]. This

199 lack of response to ER antagonists could be due to E2β exerting its effects through

GPER. Furthermore, tamoxifen used to treat breast cancer has been shown to stimulate cell growth and proliferation in endometrial tissue [Ignatov, 2012]. Tamoxifen may be acting as a GPER agonist in endometrial tissue as well.

Some studies have demonstrated that aromatase inhibitors are more effective in the treatment of breast cancer, with an increased time to recurrence following treatment with aromatase inhibitors when compared to tamoxifen [den Hollander, 2013]. This supports the hypothesis that E2β-action through GPER may be responsible for lack of response to tamoxifen and suggest that aromatase inhibitors or combination therapy of

ER-antagonists with GPER-antagonists may be a more effective means of treatment.

Efforts to understand the specific biological mechanism of GPER action will serve as a foundation for the development of future therapeutic strategies that target this receptor and perhaps prevent metastatic disease.

200

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