Copper Transporter 2 (CTR2) As a Regulator of Cisplatin Accumulation and Sensitivity

UNIVERSITY OF CALIFORNIA SAN DIEGO

Copper Transporter 2 (CTR2) as a Regulator of Cisplatin Accumulation and Sensitivity

A dissertation submitted in partial satisfaction of the requirements for the degree

Doctor of Philosophy

Biomedical Sciences

Brian G. Blair

Committee in charge:

Professor Stephen B. Howell, Chair Professor Daniel J. Donoghue Professor Paul A. Insel Professor Stanley J. Opella Professor Nicholas J. Webster

2009

The dissertation of Brian G. Blair is approved, and it is acceptable in quality and form for publication on microfilm and electronically:

Chair

University of California, San Diego

2009

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DEDICATION

To my parents and grandparents for their hard work and sacrifices that made my

education possible, and for teaching me to follow my passions.

To Erin, my loving and supportive wife, for encouraging me to be the best that I can

be in everything that I do.

TABLE OF CONTENTS

Signature Page ……………………………………………………………… iii

Dedication ……………………………..……………………………….…. iv

Table of Contents …………………………………………………………. v

List of Figures ……………………………………………………………... viii

List of Tables ………………………………………………………………. xi

Acknowledgements ………………………………………………………… xii

Curriculum Vitae …………………………………………………………… xx

Abstract of the Dissertation………………………………………………….. xxiii

Chapter 1: Introduction ……………………………………………………… 1

Platinum Based Chemotherapy ……………………………………… 1 Resistance to Platinum-Drugs ……………………………………….. 4 Platinum-Drug Transport ...………………………………………….. 5 Copper Homeostasis ...……………………………………………….. 7 Copper Transporter 2 ..…………….…………………………………. 16 Summary ...…………………………………………………………… 19 Hypothesis ...…………………………………………………………. 19 Acknowledgements ………………………………………………….. 20

Chapter 2: Effect of the Loss of CTR2 on Pt-drug Sensitivity and Accumulation in Vitro ……………………………………………. 21

Introduction …………………………………………………………... 21 Results ..…………….……………………………………………….... 21 Discussion ..……………….…………………………………………... 32 Materials and Methods ..………….…………………………………… 34 Acknowledgements ..…………………………………………………. 39

Chapter 3: Effect of the Loss of CTR2 on Pt-drug Sensitivity and Accumulation in Vivo ……………………………………………. 41

Introduction …………………………………………………………... 41

Results ..…………….……………………………………………….... 42 Discussion ..……………….…………………………………………... 50 Materials and Methods ..………….…………………………………… 53 Acknowledgements …………………………………………………… 56

Chapter 4: The Regulation of CTR2 Expression and Degradation By Copper and Platinum .....……………………………………… 57

Introduction …………………………………………………………... 57 Results ..…………….……………………………………………….... 58 Discussion ..……………….…………………………………………... 70 Materials and Methods ..………….…………………………………… 76 Acknowledgements …………………………………………………… 81

Chapter 5: CTR2 is Partially Localized in the Nucleus ………………………. 82

Introduction …………………………………………………………... 82 Results ..…………….……………………………………………….... 83 Discussion ..……………….…………………………………………... 88 Materials and Methods ..………….…………………………………… 90 Acknowledgements …………………………………………………… 92

Chapter 6: CTR2 Limits CTR2 Accumulation Through the Inhibition of Endocytosis ……………………………………………………. 93

Introduction …………………………………………………………... 93 Results ..…………….……………………………………………….... 95 Discussion ..……………….…………………………………………... 105 Materials and Methods ..………….…………………………………… 108 Acknowledgements …………………………………………………… 114

Chapter 7: Conclusions and Future Directions ..……………………………… 115

Summary ..………………………………………………………...... 115 Effect of decreased CTR2 levels on accumulation and sensitivity of DDP, CBDCA and Cu ..…………………………………………… 116 In Vivo Studies ..…………………………………………………...... 118 Regulation of CTR2 by Copper and Cisplatin ..……………………… 120 Regulation of CTR2 by Atox1 ..……………………………………… 121 Nuclear Localization of CTR2 ………………………..……………… 123

CTR2 as a Regulator of Endocytosis …..…………………………….. 125 Clinical Implications ...……………………………………………….. 127 Conclusions …..……………………………………………………..... 131 Acknowledgements …………………………………………………… 130

References ……………………………………………………………………. 132

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LIST OF FIGURES

Chapter 1

Figure 1-1. Chemical structures of cisplatin, carboplatin, and oxaliplatin …… 3

Figure 1-2. The Cu transport pathway ………………………………………. 10

Figure 1-3. Structure of hCTR1 protein ……………………………………... 12

Figure 1-4. Solution structure of Atox1 ……………………………………... 14

Figure 1-5. Structure of hCTR2 ……………………………………………… 18

Chapter 2

Figure 2-1. Relative CTR2 mRNA and protein levels in parental and knockdown cells measured by qRT-PCR and Western blot analysis ….…….. 23

Figure 2-2. Inhibition of growth as a function of concentration …………….. 26

Figure 2-3. Cell accumulation of DDP in CTR1 +/+ , CTR1 +/+ CTR1kd , CTR1 -/-, CTR1 -/- CTR1 kd cells ……………………………………………. 27

Figure 2-4. Cell accumulation of [ 14 CBDCA] and 64 Cu in CTR1 +/+ , CTR1 +/+ CTR1 kd , CTR1 -/-, CTR1 -/- CTR1 kd cells ………………. 28

Figure 2-5. Pt accumulation in DNA following 1 h exposure to 30 M DDP as measured by ICP-MS …………………………………………. 30

Chapter 3

Figure 3-1. Innoculation into mice …………………………………………… 43

Figure 3-2. Immunohistochemical characterization of proliferation and apoptosis in CTR1 -/- and CTR1 -/- CTR1 kd cells …………………………..... 44

Figure 3-3. Immunohistochemical characterization of angiogenesis in CTR1 -/- and CTR1 -/- CTR1 kd cells …………………………..... 45

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Figure 3-4. Steady state copper levels in CTR1 -/- and CTR1 -/- CTR1 kd grown in vitro or in vivo ………………………………………………… 46

Figure 3-5. Tumor accumulation of DDP in CTR1 -/- and CTR1 -/- CTR1 kd tumors 1 h following a single intraperitoneal injection of 10 mg/kg DDP …………………………………………………... 47

Figure 3-6. Effect of knocking down CTR2 on responsiveness to DDP in vivo ……………………………………………………… 48

Figure 3-7. Relationship between CTR2 expression and DDP sensitivity in ovarian carcinoma cell lines …………………………………….. 50

Chapter 4

Figure 4-1. Measurement of CTR2 in 2008 cells following 1h pre-treatment with either 200 M CuSO 4, 100 M BCS, 30 M DDP or drug-free media ………………………………………………….. 59

Figure 4-2. Representative deconvolution micrographs of 2008 cells stains for CTR2 following 1 h pre-treatment with either 200 M CuSO 4, 100 M BCS, 30 M DDP or drug-free media ………… 60

Figure 4-3. Measurement of CTR2 half-life by Western blot ……………….. 62

Figure 4-4. Deconvolution microscopy of 2008 cells following exposure to drug-free control media or 50 mM bortezomib in the presence or absence of 100 M BCS ………………………………………… 63

Figure 4-5. Effect of Cu and BCS on DDP uptake and sensitivity ………….. 65

Figure 4-6. Deconvolution microscopic images of Atox1 +/+ and Atox1 -/- cells following 1 h pre-treatment of either 200 M CuSO 4, 100 M BCS, 30 M DDP or drug-free media ………………………………… 68

Figure 4-7. CTR2 expression in Atox1 +/+ and Atox1 -/- cells ………………… 70

Figure 4-8. Model of CTR2 regulation by Atox1 …………………………… 75

Chapter 5

Figure 5-1. Deconvolution microscopic images of CTR2 in 2008 cells …….. 83

Figure 5-2. Subcutaneous mouse tumor section at 20x and 40x focal planes .. 84

Figure 5-3. Effect of Cu and BCS on nuclear CTR2 …………………………. 86

Figure 5-4. Western blot analysis of CTR2 in the cytosolic and nuclear fractions of 2008 cells following 1 h pre-treatment with either 200 M CuSO 4, 100 M BCS, 30 M DDP or drug-free media ………………….. 87

Chapter 6

Figure 6-1. Pt accumulation in DNA and vesicles following exposure to DDP. 96

Figure 6-2. Pt content as a function of efflux time ………………………….. 97

Figure 6-3. Whole cell accumulation of Texas red labeled dextran ………… 99

Figure 6-4. Dextran content as a function of efflux time ……………………. 100

Figure 6-5. Whole cell Pt accumulation in CTR1+/+, CTR1+/+ CTR2kd, CTR1-/- and CTR1-/- CTR2kd cells ……………………………. 102

Figure 6-6. 1 h accumulation of 30 µM DDP with and without PDGF pre-treatment …………………………………………….. 103

Figure 6-7. CTR2 activates the GTPases that control macropinocytosis ……. 104

Chapter 7

Figure 7-1. Stabilization of macropinocytosis by activated Rac1 and cdc42 .. 126

LIST OF TABLES

Chapter 2

Table 2-1. Accumulation of DDP, CBDCA, and Cu ……………………… 31

ACKNOWLEDGEMENTS

I would like to thank and acknowledge the following people:

Jesse Bertinato for providing personal knowledge on CTR2 as well as the

antibody used in these experiments;

Sandra Schmid for providing discourse and input with regards to the study of

endocytosis;

Martin Hetzer and Jessica Tsalmas for providing the NPC antibody;

Matt Niederst for providing Akt reagents and antibodies;

Minji Jo for input and resources for the study of GTPases;

Kersi Pestonjamasp for providing training and assistance in collecting

microscopy data;

Nissi Varki for providing assistance in this analysis of pathology samples;

Laarni Gapuz and the UCSD Cancer Center Histology and

Immunohistochemistry Shared Resource for their technical assistance;

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Gina Butcher and Leanne Nordeman of the Program in Biomedical Sciences for their assistance and support;

Ben Ho Park and Kurt Bachman for setting me off on this crazy career called science. Thank you for taking an inept college grad and turning him into a scientist. You truly are two kings;

Drs. Daniel J. Donoghue, Paul A. Insel, Stanley J Opella and Nicholas J.

Webster for taking the time to serve on my dissertation committee. Your valuable input, guidance and support is greatly appreciated and served to assist my growth as a scientist;

All the members of the Howell Laboratory, past and present, who have assisted in the composition of this dissertation;

Alison Holzer and Goli Samimi for laying the groundwork that made this work possible;

Christopher Larson for assistance and contributions made through the entirety of these studies;

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Preston Adams for assistance in the collection of uptake and in vivo data, and for providing valued discourse during the course of these studies;

Catherine Pesce for assistance in the harvesting of in vivo samples;

Paolo Abada for feedback and discourse regarding CTR2 in the nucleus and as a mediator of endocytosis;

Gerald “Jack” Manorek for valued discourse during the course of these studies;

Xinjin Lee for valued feedback;

Sakura Moua, Nichole Chung and Phillip Chang for providing valuable assistance;

Angela Robles for assistance, editing and support;

My mentor, Stephen B. Howell, for his guidance and advice. Thank you for your counseling, commitment, hard work and dedication through the years.

Thank you for trusting me to find my way through the roadblocks and being there to help navigate when things got too rough. Most importantly, thank you for teaching me how to think like a true scientist. I have enjoyed my time here

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and appreciate all the time and effort you have put into helping me succeed in

my graduate career.

Finally, I’d like to thank those who have helped contribute to my studies away from the lab:

I’d like to thank my parents, Barbara and G. Blair, for their constant love and

support. Thank you for allowing me to find my own way so many miles away.

Most importantly, thank you for teaching me to use my abilities to the fullest

and for instilling in me the values of hard work, patience and love.

To my brother, Christopher Blair for his friendship, love and support. Thanks

for being available to help distract me from the stress of lab – I can always

count on your being awake despite the time difference!

To the rest of my family, thank you for your patience, love and support

through these years away from home.

To my in-laws, Alan and Rose Marie Drenning, thank you for allowing me to

take your daughter across the country and for always praying for our success.

Thank you to all of my friends, old and new, who have helped this time fly by

and kept me grounded and smiling this whole time: Matt, Bill, Perry, Mike,

Dave, Rach, Eilis, the Phelpses, the “Lisas”, Ken and Dave, Alara, Kylie and

all the “law friends”, Julie and the choir, Chris, Alicia, Catherine, Angie,

Paolo, Jack, Preston and Shiobhan – (thanks for making sure we never went

thirsty).

Most importantly, I’d like to thank my wonderful wife, the piece that makes

me whole. Thank you for talking me into taking this opportunity, for always

being there no matter what, for helping me through my struggles, being my

favorite editor and for making me happy each and every day with your love.

You are all that I could ever need.

Acknowledgements for individual chapters are as follows:

Chapter 1: Brian G. Blair was the primary author of this chapter. Stephen B.

Howell supervised the writing of this chapter.

Chapter 2: A majority of the content of Chapter 2 has been published in

Clinical Cancer Research (Blair BG, Larson CA, Safaei R, Howell SB. Copper

Transpoter 2 Regulates the Cellular Accumulation and Cytotoxicity of Cisplatin and

Carboplatin. Clin Cancer Res. 2009 Jul 1;15(13):4312-21.) Brian G. Blair was the

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primary researcher and author for this chapter. Stephen B. Howell supervised and directed the research in this chapter. Christopher A. Larson assisted in the measurement of Pt and Cu accumulation and provided helpful feedback. Roohangiz

Safaei provided helpful discussion. The author would like to thank Dr. Dennis Thiele for generously providing the CTR1 +/+ and CTR1 -/- mouse embryo fibroblasts, Dr.

Jessie Bertinato for the anti-CTR2 antibody and Ms. Sakura Moua for technical assistance.

Chapter 3: A majority of the content of Chapter 3 has been submitted for publication. The remaining sections of this chapter have been adapted from published results in Clinical Cancer Research (Blair BG, Larson CA, Safaei R, Howell SB.

Copper Transpoter 2 Regulates the Cellular Accumulation and Cytotoxicity of

Cisplatin and Carboplatin. Clin Cancer Res. 2009 Jul 1;15(13):4312-21.) Brian G.

Blair was the primary researcher and author for this chapter. Stephen B. Howell supervised and directed the research in this chapter. Christopher A. Larson and

Preston L. Adams assisted in the in vivo studies as well as the measurement of Pt and

Cu accumulation and provided helpful feedback. Catherine Pesce and Gerald Manorek assisted in tumor and organ harvesting. Roohangiz Safaei provided helpful discussion.

Dr. Dennis Thiele for provided the CTR1 +/+ and CTR1 -/- mouse embryo fibroblasts, and Dr. Jessie Bertinato provided the anti-CTR2 antibody.

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Chapter 4: The contents of this chapter have been submitted for publication in full. Brian G. Blair was the primary researcher and author for this chapter. Stephen B.

Howell supervised and directed the research in this chapter. Christopher A. Larson and

Preston L. Adams assisted in the measurement of Pt accumulation and provided helpful feedback. Roohangiz Safaei provided helpful discussion. The author would like to thank Dr. J. D. Gitlin for generously providing the Atox1 +/+ and Atox1 -/- mouse embryo fibroblasts, Dr. Jessie Bertinato for the anti-CTR2 antibody.

Chapter 5: The majority of the contents of this chapter have been submitted for publication. Brian G. Blair was the primary researcher and author for this chapter.

Stephen B. Howell supervised and directed the research in this chapter. Paolo B.

Abada independently confirmed the Western blots contained in this chapter.

Christopher A. Larson and Preston L. Adams assisted in preparation of microscope slides, harvesting of mouse tumors and provided feedback. Roohangiz Safaei provided helpful discussion. Patholgy for this chapter was provided by Laarni Gapuz and Dr.

Nissi Varki of the UCSD pathology core. Kersi Pestonjamasp provided training and valuable input on microspopic technique. The author would like to thank Dr. Jessie

Bertinato for the providing the anti-CTR2 antibody and Dr. Martin Hetzer for the NPC antibody.

Chapter 6: A portion of the content of Chapter 6 has been published in

Clinical Cancer Research (Blair BG, Larson CA, Safaei R, Howell SB. Copper

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Transpoter 2 Regulates the Cellular Accumulation and Cytotoxicity of Cisplatin and

Carboplatin. Clin Cancer Res. 2009 Jul 1;15(13):4312-21.) Brian G. Blair was the primary researcher and author for this chapter. Stephen B. Howell supervised and directed the research in this chapter. Christopher A. Larson and Preston L. Adams provided feedback. Roohangiz Safaei provided helpful discussion. Dr. Sandra Schmid provided valuable insight and suggestions as to the understanding of endocytosis. The author would like to thank Dr. Jessie Bertinato for the providing the anti-CTR2 antibody, Dr. Alexandra Newton for the anti-Akt antibodies and Dr. Minji Jo for the anti-Rac1 and anti-cdc42 antibodies.

Chapter 7: Brian Blair was the primary author of this chapter. Stephen B.

Howell supervised the writing of this chapter.

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CURRICULUM VITAE

EDUCATION

1997-1998 Villanova University , Villanova, PA Honors Biology Major

1998-2001 Syracuse University , Syracuse, NY Bachelor of Science, Biology Magna Cum Laude

2003-2004 Johns Hopkins School of Medicine , Baltimore, MD Biotechnology Masters Program

2004-2009 University of California San Diego San Diego, CA Doctor of Philosophy in Biomedical Sciences December 2009

EXPERIENCE

2000-2001 Syracuse University , Syracuse, NY Undergraduate Research Assistant Scott A. Heckathorn, Ph.D. Laboratory

2002-2004 Johns Hopkins School of Medicine , Baltimore, MD Research Technician Ben Ho Park, M.D., Ph.D. Laboratory

2004-2006 Salk Institute , San Diego, CA Outreach Lead Instructor Mobile Science Laboratory

2004-2009 University of California San Diego , San Diego, CA Graduate Student Researcher Stephen B. Howell, M.D. Laboratory

HONORS AND AWARDS

1997-1998 Villanova University Honors Society Member

1998-2001 Syracuse University Honors Society Member

2001 Golden Key National Honor Society Inductee

2009 Scholar-in-Training Award Finalist – American Association for Cancer Research Annual Meeting

2009 Speaker at Novel Mechanisms of Drug Resistance Mini- symposium – American Association for Cancer Research Annual Meeting PUBLICATIONS

Bachman KE, Blair BG, Brenner K, Bardelli A, Arena S, Zhou S, Hicks J, De Marzo AM, Argani P, Park BH. p21(WAF1/CIP1) Mediates the Growth Response to TGF- beta in Human Epithelial Cells. Cancer Biol Ther. 2004 Feb;3(2):221-5.

Bachman KE, Argani P, Samuels Y, Silliman N, Ptak J, Szabo S, Konishi H, Karakas B, Blair BG, Lin C, Peters BA, Velculescu VE, Park BH. The PIK3CA Gene is Mutated with High Frequency in Human Breast Cancers. Cancer Biol Ther. 2004 Aug;3(8):772-5. Erratum in: Cancer Biol Ther. 2005 Feb;4(2):133.

Keen JC, Zhou Q, Park BH, Pettit C, Mack KM, Blair B, Brenner K, Davidson NE. Protein Phosphatase 2A Regulates Estrogen Receptor Alpha (ER) Expression Through Modulation of ER mRNA Stability. J Biol Chem. 2005 Aug 19;280(33):29519-24.

Huang Y, Keen JC, Pledgie A, Marton LJ, Zhu T, Sukumar S, Park BH, Blair B, Brenner K, Casero RA Jr, Davidson NE. Polyamine Analogues Down-Regulate Estrogen Receptor Alpha Expression in Human Breast Cancer Cells. J Biol Chem. 2006 Jul 14;281(28):19055-63.

Abukhdeir AM, Blair BG, Brenner K, Karakas B, Konishi H, Lim J, Sahasranaman V, Huang Y, Keen J, Davidson N, Vitolo MI, Bachman KE, Park BH. Physiologic Estrogen Receptor Alpha Signaling in Non-Tumorigenic Human Mammary Epithelial Cells. Breast Cancer Res Treat. 2006 Sep;99(1):23-33.

Karakas B, Weeraratna A, Abukhdeir A, Blair BG, Konishi H, Arena S, Becker K, Wood W 3rd, Argani P, De Marzo AM, Bachman KE, Park BH. Interleukin-1 Alpha Mediates the Growth Proliferative Effects of Transforming Growth Factor-Beta in p21 Null MCF-10A Human Mammary Epithelial Cells. Oncogene. 2006 Sep 7;25(40):5561-9.

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Abukhdeir AM, Vitolo MI, Argani P, De Marzo AM, Karakas B, Konishi H, Gustin JP, Lauring J, Garay JP, Pendleton C, Konishi Y, Blair BG, Brenner K, Garrett-Mayer E, Carraway H, Bachman KE, Park BH. Tamoxifen-Stimulated Growth of Breast Cancer Due to p21 Loss. Proc Natl Acad Sci U S A. 2008 Jan 8;105(1):288-93.

Larson CA, Blair BG, Safaei R, Howell SB. The role of the Mammalian Copper Transporter 1 in the Cellular Accumulation of Platinum-Based Drugs. Mol Pharmacol. 2009 Feb;75(2):324-30.

Safaei R, Maktabi MH, Blair BG, Larson CA, Howell SB. Effects of the Loss of Atox1 on the Cellular Pharmacology of Cisplatin. J Inorg Biochem. 2009 Mar;103(3):333-41.

Blair BG, Larson CA, Safaei R, Howell SB. Copper transporter 2 Regulates the Cellular Accumulation and Cytotoxicity of Cisplatin and Carboplatin. Clin Cancer Res. 2009 Jul 1;15(13):4312-21.

Blair BG, Larson CA, Adams PL, Abada PB, Safaei R, Howell SB. Regulation of CTR2 Expression by Copper and Cisplatin in Human Ovarian Carcinoma Cells. Submitted in full. Oct. 2009.

Blair BG, Larson CA, Adams PL, Pesce C, Safaei R, Howell SB. Loss of Copper Transporter 2 (CTR2) confers DDP Sensitivity in Vivo . Submitted in full . Nov. 2009.

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ABSTRACT OF THE DISSERTATION

Copper Transporter 2 (CTR2) as a Regulator of Cisplatin Accumulation and Sensitivity

Brian G. Blair

Doctor of Philosophy in Biomedical Sciences

University of California, San Diego, 2009

Professor Stephen B. Howell, Chair

Platinum(Pt)-containing cancer drugs are highly polar molecules that do not diffuse across lipid membranes; thus, their uptake into tumor cells must involve a transport process. Cells selected for resistance to these drugs uniformly exhibit impaired drug accumulation. The copper (Cu) transport pathway has been demonstrated to be responsible for the majority of Pt-drug accumulation and cellular trafficking.

The overall goal of this dissertation was to determine whether Cu transporter 2

(CTR2) plays a role in the cellular accumulation of cisplatin (DDP), and if so, whether it influences the sensitivity of cells to DDP. This was accomplished through the study

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of a CTR2 knockdown model system. It was discovered that the loss of CTR2 protein expression leads to increased DDP accumulation and sensitivity both in vitro and in vivo. Additionally, it was determined that CTR2 is required for optimal tumor growth, as CTR2 kd tumors demonstrated increased the frequency of apoptotic cells and reduced vascular density. Once CTR2 was established as a regulator of DDP accumulation and sensitivity, the investigations went on to focus on how CTR2 expression and degradation is controlled by DDP, and Cu. Cu and DDP exposure were shown to increase CTR2 levels. This increase was associated with an increase in

CTR2 mRNA and prolongation of CTR2 half-life. Cu starvation triggered rapid degradation of CTR2, which was dependent on proteosomal activity and the status of the copper chaperone Atox1. Consistent with the observations previously made, reduction of CTR2 by Cu starvation also enhanced DDP uptake and cytotoxicity.

During the course of these studies, the unique observation was made that CTR2 is partially localized in the nucleus of cells. Finally, the mechanism by which decreased

CTR2 levels lead to increased accumulation of DDP was explored. CTR2 knockdown did not change the rate of efflux of or the amount of vesicular DDP. Decreased CTR2 levels, due to knockdown or degradation, triggered the up-regulation of cellular macropinocytosis and activation of the GTPases necessary for endocytosis. Inhibition of endocytosis blocked the increased accumulation of DDP in CTR2 kd cells, suggesting that CTR2 limits Pt-drug accumulation through the regulation of endocytosis.

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Chapter 1:

Introduction

Platinum-based chemotherapy

The chemotherapeutic benefit of Platinum (Pt) based constructs, originally demonstrated in 1965, derived from the demonstration that Pt was an inhibitor of bacterial cell division (Rosenberg, Vancamp et al. 1965). Following this discovery, the potential for Pt-derived compounds serve as anti-cancer agents has been extensively explored. The following sections provide background information on two of the Pt compounds currently used in the clinic.

Cisplatin

Cisplatin (DDP) is among the most widely and effective anti-cancer agents that have been used since the 1970s (Chu 1994; Jordan and Carmo-Fonseca 2000; Fuertes,

Alonso et al. 2003; Rabik and Dolan 2007). DDP is bound by plasma proteins upon entering the bloodstream and is relatively unreactive in this form. When taken up by cells, the chloride atoms are displaced from DDP (Figure 1-1). This is due to the low cytosolic chloride concentration (2-30 mM). Displacement of the chloride atoms renders the DDP molecule active with the potential to interact with nucleophilic sites on numerous intracellular molecules including: nuclear and mitochondrial DNA,

1 2

RNA, cysteine-, histidine- and methionine-containing proteins and phospholipids. DDP forms adducts on the purine bases of nuclear DNA (Johnson,

Laub et al. 1997; Sedletska, Giraud-Panis et al. 2005; Zorbas and Keppler 2005); however, it is not clear how DDP traffics through the cell to reach the nucleus. These adducts are thought to be the primary source of DDP-induced cytotoxicity through the inhibition of DNA synthesis and transcription. The adducts are typically 1,2d(GG) and

1,2-d(ApG) intrastrand DNA crosslinks; however, interstrand and 1,3-intrastrand crosslinks have also been observed (Johnson, Laub et al. 1997; Sedletska, Giraud-

Panis et al. 2005; Zorbas and Keppler 2005). The formation of these DNA adducts is thought to trigger a DNA damage response program leading to cell cycle arrest and/or apoptosis. It has been proposed that the cytotoxic response to DDP-induced DNA damage depend upon the mismatch repair system (Gong, Costanzo et al. 1999; Lin and

Howell 1999; Strathdee, Sansom et al. 2001).

Carboplatin

At clinically effective doses DDP exhibits a high level of toxicity to a large number of organ systems and thereby produces neuropathy, nephrotoxicity and myelosuppression (Rabik and Dolan 2007). As a result several other Pt-based drugs have been developed in an attempt to improve the therapeutic utility of DDP.

Carboplatin (CBDCA) is a DDP analog that produces similar DNA adducts in

DDP. However, CBDCA has a different pattern of toxicity than that of DDP. CBDCA

4 is more stable in aqueous environments due to a single 1,1 cyclobutanedicarboxylate leaving group instead of the two chlorides of DDP (Figure1-1). This change slows the aquation rate of CBDCA and reduces reactivity. However, due to its similarities to

DDP, CBDCA shares many of the same drawbacks. Cells resistant to DDP are cross- resistant to CBDCA and vice versa .

Resistance to Pt drugs

Pt-based chemotherapeutics are among the most widely used and effective anti-cancer agents. However, patients often develop resistance during the course of therapy (Schabel, Skipper et al. 1980; Andrews and Howell 1990; Gately and Howell

1993; Muggia and Los 1993; Walker and Walker 1999; Niedner, Christen et al. 2001).

This resistance is thought to result from the selective survival of tumor cells that have sustained mutations that reduce drug uptake or otherwise interfere with the cytotoxic mechanisms of the drug. DDP is a mutagen and even a single exposure to the drug can result in the generation of cells resistant to both DDP itself and other classes of chemotherapeutic agents (Lin, Kim et al. 1999). Once formed, the selective pressure created by additional DDP treatment results in the resistant cells becoming the dominant population in the tumor. Even a modest 1.5 to 3-fold drop in sensitivity to

DDP at the cellular level can lead to treatment failure in vivo (Inoue, Mukaiyama et al.

1985; Wilson, Ford et al. 1987; Muggia and Los 1993). Resistance can emerge very quickly and even low levels of resistance, as measured in tissue culture, are sufficient to produce clinical failure (Andrews and Howell 1990; Muggia and Los 1993).

Unfortunately, the mechanisms that account for Pt-drug resistance have not been fully identified. Changes in drug influx and efflux, deficiencies in the mismatch repair pathway and down-regulation of the apoptotic cascade are all among the possible mechanisms that have been proposed (Chu 1994; Crul, Schellens et al. 1997;

Manic, Gatti et al. 2003; Sedletska, Giraud-Panis et al. 2005; Kuo, Chen et al. 2007).

However, there is a strong correlation between drug sensitivity and drug accumulation, with DDP-resistant cells accumulating less drug than their sensitive counterparts

(Metcalfe, Cain et al. 1986; Waud 1987; Teicher, Holden et al. 1991; Kelland, Mistry et al. 1992; Twentyman, Wright et al. 1992; Gately and Howell 1993; Oldenburg,

Begg et al. 1994; Johnson, Shen et al. 1996; Song IS 2004).

Pt drug Transport

Due to their polar nature, Pt-agents cannot readily diffuse across the cell membrane and therefore must be taken up by transport mechanisms such as via a pump or channel (Andrews, Mann et al. 1988; Andrews and Albright 1991; Mann,

Andrews et al. 1991). DDP uptake is pH-sensitive as well as potassium (K +) ion- dependent (Atema, Buurman et al. 1993; Amtmann, Zoller et al. 2001; Marklund,

Andersson et al. 2004; Chen, Jiang et al. 2005). The presence of reducing agents such as ascorbate and dithioreitol has been shown to enhance Pt-drug uptake, suggesting that uptake is charge-dependent (Sarna and Bhola 1993; Chiang, Song et al. 1994;

Zhang, Zhong et al. 1994; Blasiak, Kadlubek et al. 2002).

It is important to note that while impaired drug accumulation has been observed in Pt-drug resistant cells and tumors, it is unknown whether this decrease in accumulation is due to a failure of drug uptake or enhanced efflux and the published studies are conflicting on this point. Waud et al . observed a decrease in the rate of

DDP uptake and adduct accumulation but no difference in efflux in DDP-resistant murine leukemia cells (Waud 1987). Teicher et al . also demonstrated a decrease in whole cell and nuclear Pt content in 5 different DDP-resistant cell lines (Teicher,

Holden et al. 1991). Several further studies suggest the importance of DDP influx in resistance (Troger, Fischel et al. 1992; Johnson, Shen et al. 1996; Helleman, Burger et al. 2006). However, several other studies indicate that some resistant lines show increase in DDP efflux (Parker, Eastman et al. 1991; Fujii, Mutoh et al. 1994; Chau and Stewart 1999). This disparity may be due to the inability of many early studies to detect a large portion of DDP efflux due to technical limitations. It was later discovered that much of DDP efflux occurs very rapidly (Mann, Andrews et al. 1990).

As discussed in detail in this chapter, the Cu transporter CTR1 has been linked to Pt-drug uptake and accumulation (Holzer, Samimi et al. 2004; Larson, Blair et al.

2008; Blair, Larson et al. 2009). Additionally, the organic cation transporter family

(OCT) has also been reported to mediate DDP uptake and accumulation (Briz, Serrano et al. 2002; Yonezawa, Masuda et al. 2006; Zhang, Lovejoy et al. 2006). However, details of the mechanism by which these transporters function remain unknown and despite intense study for over 20 years, little is still known about the intracellular trafficking of the Pt-based drugs.

Copper Homeostasis

Linkage between the cellular pharmacology of Cu and the Pt drugs was first suggested by the finding that cells selected for resistance to Cu were cross-resistant to the Pt drugs and vice versa (Naredi, Heath et al. 1994; Fukuda, Ohe et al. 1995; Rixe,

Ortuzar et al. 1996; Shen, Pastan et al. 1998). Furthermore, several studies have provided evidence that the Pt-containing drugs are taken up, shuttled within the cell and exported by the proteins involved in Cu homeostasis. The following sections review current knowledge regarding the role of Cu homeostasis proteins in the transport of Pt-drugs.

Background

Cu (Cu) is an essential trace metal necessary for the activity of several intracellular enzymes including superoxide dismutase, cytochrome-c oxidase, lysyl oxidase and dopamine β-hydrolase. Cu’s ability to undergo reversible oxidation from

Cu(I) to Cu(II) under physiologic conditions is essential for its role in cellular functions such as electron transport and the detoxification of reactive oxygen (Linder and Hazegh-Azam 1996). However, this oxidation process produces reactive oxygen species that can cause severe damage to DNA and other components of the cell including lipids and several proteins (Linder and Hazegh-Azam 1996). High levels of intracellular Cu can also have a toxic effect on cells. Cu can displace other integral

8 metal cofactors from their interacting proteins, thereby disrupting normal function

(Pena, Lee et al. 1999).

Copper Transport

Eukaryotic cells have developed a complex system of Cu transporters and chaperones that protect Cu(I) during its influx and distribute it throughout the intracellular space (Camakaris, Voskoboinik et al. 1999; O'Halloran and Culotta 2000;

Huffman and O'Halloran 2001). The proteins that participate in Cu homeostasis are characterized by unique domains, that are rich in cystine, methionine or histidine, and called metal binding sequences (MBS). As Cu enters and is trafficked around the cell, it is shuttled from one MBS-containing protein to the next so that virtually no Cu (less than 10 −18 M) remains free in the cell (Pufahl, Singer et al. 1997; Hamza, Schaefer et al. 1999; Lippard 1999; Rae, Schmidt et al. 1999).

Figure 1-2 is a schematic of the current level of understanding of Cu homeostasis and the system of transporters, chaperones and enzymes involved in cellular Cu metabolism. Cu(II) is delivered to the cell surface bound to ceruloplasmin and is reduced at the cell surface by the reductases FRE1 and FRE2 (Hassett and

Kosman 1995; Georgatsou and Alexandraki 1999). Upon reduction, Cu is passively transported across the plasma membrane by the trimeric channel-transporter CTR1 in an energy-independent manner (Pena, Lee et al. 1999). It is then transferred to the chaperones Atox1, Cox17 and Ccs. Atox delivers Cu to the Golgi apparatus proteins

ATP7A and ATP7B (Klomp, Lin et al. 1997; Huffman and O'Halloran 2000). Cox 17

9 shuttles Cu to the mitochondria and cytochrome c oxidase (Amaravadi, Glerum et al.

1997). Ccs transports Cu to superoxide dismutase (SOD1) (Culotta, Klomp et al.

1997). ATP7A and ATP7B sequester Cu into the trans -Golgi network where it is then loaded onto ceruloplasmin and other Cu-dependent enzymes (Dierick, Adam et al.

1997; Suzuki and Gitlin 1999). ATP7A and ATP7B are thought to function to limit the toxic effect of Cu. Excess Cu within the cell causes ATP7A and ATP7B to relocate away from the trans -Golgi to either the plasma membrane or vesicular compartments (Petris, Mercer et al. 1996; Petris and Mercer 1999). This translocation is thought to be necessary for the efflux of Cu either directly by ATP7A and ATP7B or by a process downstream of these two proteins (Camakaris, Petris et al. 1995;

Roelofsen, Wolters et al. 2000).

Copper Transporter 1 (CTR1)

Cu Transporter 1 (CTR1, SLC31A1) is the major plasma membrane Cu transporter, though Cu is believed to be taken up in a less efficient manner by other transporters such as Nramp1, DCT1 and DMT1 (Gunshin, Mackenzie et al. 1997;

Sharp 2003). The hCtr1 gene is located on chromosomal region 9q31–32 and generates a 28 kDa protein which is subsequently glycosolated to form the mature 35 kDa form (Petris 2004). Mammalian CTR1 is expressed in all tissues with highest levels in the liver, kidney and heart, followed by the intestine, and with the lowest

11 expression occurring in the brain and muscle (Zhou and Gitschier 1997). CTR1 was first identified as a Cu transporter on the basis of homology to the known yeast Cu transporters yCtr1 and yCtr3 t (Zhou and Gitschier 1997). Human CTR1 was shown to enhance the uptake of Cu when expressed in yeast (Moller, Petersen et al. 2000).

Figure 1-3 depicts the structure of CTR1. The CTR1 protein contains 3 transmembrane regions and exists in the membrane as a homotrimer where it forms a channel (Lee, Prohaska et al. 2000; Aller and Unger 2006; Nose, Rees et al. 2006; De

Feo, Aller et al. 2009). hCTR1 features a 69 amino acid extracellular N-terminal tail that contains three methionine-rich metal binding domains (MBD). hCTR1 also contains a 48 amino acid intracellular loop, situated between the first and second transmembrane domains, the function of which is not well defined. The second transmembrane region of hCTR1 also contains a fourth MBD. It is these MXXM and

MXXXM regions, and more specifically the third and fourth MBD, that are primarily responsible for the binding and transport of Cu (Dancis, Yuan et al. 1994; Eisses and

Kaplan 2005; De Feo, Aller et al. 2009). The C-terminal tail contains only 14 amino acids and C189 has been shown to be necessary for the multimerization of CTR1 (Lee,

Howell et al. 2007). Cu associates with the hCTR1 metal binding domains and is transported into the cytosol through the channel formed by the hCTR1 trimer. Cu is then handed off to the various chaperones to be trafficked throughout the cell.

Atox1

Atox1 (HAH1) was first characterized by the ability of its yeast homologue

Atx1 to rescue the SOD1 knockout phenotype. Mammalian Atox1 was found to serve as a Cu chaperone (Klomp, Lin et al. 1997). Human Atox1 is a 68 amino acid cytosolic protein containing a critical MXXM metal binding domain. The structure of

Atox1 is depicted in Figure 1-4. Atox1 is thought to accept Cu from CTR1 and shuttle it to intracellular targets (Xiao and Wedd 2002). However, the exact nature of this interaction between CTR1 and Atox1 is still unclear. Once in the Cu- bound state,

Atox1 serves as a chaperone to deliver Cu to its target proteins, including ATP7A and

ATP7B among others. Atox1 is necessary for the delivery of Cu to the trans-Golgi network, as well as the Cu-mediated relocation of ATP7A and ATP7B from the trans-

Golgi to more peripheral locations (Hamza, Prohaska et al. 2003; Lutsenko,

Tsivkovskii et al. 2003; Strausak, Howie et al. 2003; Miyayama, Suzuki et al. 2009).

Atox1 has also been linked to the regulation of several Cu proteins such as ATP7B,

SOD1 and CCS (Lutsenko, Tsivkovskii et al. 2003; Jeney, Itoh et al. 2005; Miyayama,

Suzuki et al. 2009).

Recently, Atox1 has been found to act as a nuclear transcription factor by Itoh et al. (Itoh, Kim et al. 2008). Atox1 -/- cells were unable to demonstrate Cu-dependent cyclin D1 and S phase activation. Atox1 was found to translocate to the nucleus and bind to and activate the cyclin D1 promoter when cells were stimulated with Cu (Itoh,

Kim et al. 2008; Muller and Klomp 2008). Atox1 was later found to also regulate transcription of the SOD complex (Itoh, Ozumi et al. 2009).

Copper Transporters and Pt drug Pharmacology

The Cu homeostasis proteins were first linked to Pt-drug transport and resistance when it was observed that DDP resistance could be caused by increased

ATP7B levels (Komatsu, Sumizawa et al. 2000). Later, cross-resistance was noted between Cu and DDP (Katano, Kondo et al. 2002), and DDP-resistant cell lines were shown to be deficient in both Cu and DDP accumulation (La Fontaine, Firth et al.

1998).

ATP7B and ATP7A play a role Pt-drug efflux. Over-expression of ATP7B is associated with a reduced capacity to accumulate DDP and CBDCA (Komatsu, Ikeda et al. 2001; Katano, Safaei et al. 2003). ATP7B over-expression also enhances the efflux of Pt-drugs, and DDP can interact and affect the relocation of ATP7B within the cell (Katano, Safaei et al. 2003; Katano, Safaei et al. 2004). ATP7A is up-regulated in some resistant tumors and the forced expression of ATP7A decreases DDP

15 accumulation (Samimi, Varki et al. 2003; Samimi, Safaei et al. 2004). However, the action of ATP7A and ATP7B alone has not been able to account for the observed changes in drug accumulation in DDP-resistant cells. It is very likely that the observed decrease of drug accumulation associated with resistance is primarily due to changes taking place in the drug influx mechanism.

hCTR1 plays an important role in the cellular pharmacology of DDP and probably that of carboplatin and oxaliplatin as well. Loss of CTR1 function rescues yeast and mouse cells from the toxic effects of DDP and this rescue was correlated with decreased uptake of drug (Ishida, Lee et al. 2002; Lin, Okuda et al. 2002).

Decreased hCTR1 mRNA expression correlates with increased resistance to DDP

(Katano, Kondo et al. 2002). However, in one study over-expression of CTR1 in human ovarian cancer cells did not increase sensitivity to the cytotoxic effects of

DDP, and the formation of DNA adducts remained equal to that of parental cells

(Holzer, Samimi et al. 2004). This suggests that while CTR1 levels play an important role in acquired resistance, they are not the only determinant of drug sensitivity.

However, cells containing homozygous deletions of murine CTR1 exhibit decreased

Pt drug accumulation although some drug still enters these cells at a lower rate

(Holzer, Samimi et al. 2004; Larson, Blair et al. 2008). This observation suggests that

DDP can enter the cell through transporters other than CTR1. The Pt-containing drugs are thought to interact with the metal binding domains of hCTR1 in a manner similar to that of Cu. Current evidence leaves open the question of whether the Pt-containing drugs pass through the pore formed by CTR1 or via macropinocytosis as hCTR1

16 disappears from the plasma membrane within the first few minutes of DDP exposure

(Holzer, Samimi et al. 2004). If the Pt-containing drugs do enter via an endocytotic process, how the DDP disassociates from hCTR1, escapes the endosome and is trafficked throughout the cell are as-yet undefined.

Copper Transporter 2

Structure and Characterization

CTR2 is a Cu transport protein with a great deal of structural homology to

CTR1. While containing a truncated N-terminal domain, CTR2 has 3 transmembrane domains and a large cytosolic loop (Puig, Lee et al. 2002; Rees and Thiele 2007).

Though sharing only 41% amino acid homology, it is important to note that both hCTR1 and hCTR2 share the third N-terminal MXXXM metal binding motif, as well as the MBD located in the highly conserved second transmembrane domain (Petris

2004; Rees and Thiele 2007). Mutational analysis of CTR1 has shown that these are the only two MBD absolutely necessary to transport Cu, suggesting that CTR2 is fully functional as a Cu transporter (Puig, Lee et al. 2002; Nose, Rees et al. 2006).

CTR2 was identified as a Cu transporter on the basis of its homology to yCtr2, and its ability to rescue the Cu-deprived phenotype of ctr1 ∆ctr3 ∆ mutants (Rees and

Thiele 2007). In yeast, Ctr2 and its S. pombe orthologue Ctr6 are localized in vacuoles with the C-terminal tail oriented toward the cytosol. It has been shown that yCtr2

17 releases Cu from intercellular stores under conditions of Cu starvation and delivers Cu to various chaperones (Kampfenkel, Kushnir et al. 1995; Portnoy, Schmidt et al. 2001;

Rees and Thiele 2007). The ability of yCtr2 to transport Cu is dependent on the action of the iron reductase FRE6 (Rees and Thiele 2007). Furthermore, the Ctr2-1 mutant of yeast Ctr2, which partially mislocalizes to the plasma membrane, mediates Cu transport in a manner similar to that of Ctr1 {Rees, 2004 #7027).

Mammalian CTR2

Little is known about human CTR2, the structure of which is diagramed in

Figure 1-5; however, it is believed that hCTR2 plays a similar role to that of yCtr2 and is primarily responsible for release of Cu from intracellular vesicle stores. CTR2 is primarily localized to late endosomes and lysosomes, although it has been reported to localize on the plasma membrane in some cells {van den Berghe, 2007 #9392;

Bertinato, 2007 #9593}. Akin to CTR1, CTR2 forms multimers, some of which co- localize with CTR1 (van den Berghe, Folmer et al. 2007). Mammalian CTR2 increases Cu influx in cells in which it localizes to the plasma membrane (Bertinato,

Swist et al. 2007), although its affinity for Cu is less than that of CTR1 (Bertinato,

Swist et al. 2007; van den Berghe, Folmer et al. 2007). Like CTR1, CTR2 is able to transport silver, but not zinc, iron and manganese (van den Berghe, Folmer et al.

2007). Changes in CTR2 expression do not affect Cu efflux, suggesting that it functions primarily as an influx transporter or in the intracellular storage of Cu (van den Berghe, Folmer et al. 2007). It has also been shown that CTR2 can inhibit SOD

18 protein expression, suggesting that CTR2 may be integral to the regulation of other Cu metabolism proteins (van den Berghe, Folmer et al. 2007). Given that CTR1 transports

DDP and that CTR2 has substantial structural similarity to CTR1, it appears likely that

CTR2 also transports DDP and may mediate the residual DDP influx observed in

CTR1 knockout cells, as well as regulation of Pt drug release from intracellular compartments.

Summary

The chemotherapeutic Pt-containing drugs DDP and CBDCA are used to treat a wide variety of tumors. However, the rapid development of resistance to these drugs limits their clinical effectiveness. There is a strong correlation between development of Pt drug resistance and decreased drug uptake. It has been established that Pt- containing drugs are taken in, shuttled and exported by the transporters and chaperones that mediate Cu homeostasis. Specifically, the Cu transporter CTR1 has been identified as a major Pt drug influx transporter. However, CTR1 alone does not account for the total influx of drug, supporting the hypothesis that there must be other mechanisms of Pt drug uptake.

Hypothesis

CTR2 is similar in structure to CTR1, whose ability to regulate the accumulation of the Pt-containing drugs is now well established. The objective of this thesis research was to define the role of CTR2 as a regulator of Pt drug accumulation and as a determinant of sensitivity to the cytotoxic effect of DDP. The hypothesis tested was that CTR2 is a major regulator of the accumulation of DDP and its clinically important analogs, and that defects in the function of this transporter lead to clinically relevant changes in the sensitivity of tumors to these drugs.

Acknowledgments

Brian G. Blair was the primary author of this chapter. Stephen B. Howell directed and supervised the writing of this chapter.

Chapter 2:

The Effect of the Loss of CTR2 on Pt drug Sensitivity and Accumulation in Vitro

Introduction

The goal of the experiments described in this chapter was to determine whether

CTR2, like CTR1, functions as a transporter for the Pt-containing chemotherapeutic agents and whether it modulates sensitivity to the cytotoxic effects of these drugs. The role of CTR2 was examined by knocking down CTR2 expression in an isogenic pair of mouse embryo fibroblasts consisting of a CTR1 +/+ line and a CTR1 -/- knockout line.

CTR2 levels were determined by qRT-PCR and Western blot analysis. DDP was quantified by measuring cellular levels of Pt by ICP-MS and 64 Cu and 14 C-carboplatin

(CBDCA) accumulation by γ and scintillation counting, respectively. Drug sensitivity was determined by changes in growth rate as measured by staining with sulforhodamine B.

Results

Knockdown of mCTR2 in mouse embryo fibroblasts.

21 22

A wild-type (CTR1 +/+ ) mouse embryo fibroblast cell line and an isogenic line, in which both alleles of CTR1 had been knocked out (CTR1 -/-), were used to examine the effect of disabling the function of CTR2. This system allowed examination of disabling CTR2 in cells that were either proficient or deficient in CTR1 function. The

CTR1 +/+ and CTR1 -/- cells were infected with lentivirus expressing a shRNA targeted to mCTR2, and individual colonies were selected using 5 uM puromycin. Knockdown of mCTR2 expression was analyzed by quantitative reverse transcription-PCR (qRT-

PCR) and Western blot analysis, and a single clone was chosen for further study and expanded to form a subline. As shown in Figure 2-1, CTR2 mRNA and protein expression was reduced by 88.5 ± 3.8% (SEM) and 86 ± 2.2% (SEM), respectively, in the CTR1 +/+ /CTR2 kd subline. CTR2 knockdown did not affect CTR1 levels as measured by qRT-PCR (data not shown). Figure 2-1 also demonstrates that CTR2 mRNA and protein expression was reduced by 81.8 ± 6.4% and 87.1 ± 4.6%, respectively, in CTR1 -/-/CTR2 kd cells.

Reduction of CTR2 expression increases sensitivity to DDP and CBDCA.

The CTR1 +/+ /CTR2 +/+ , CTR1 +/+ /CTR2 kd , CTR1 -/-/CTR2 +/+ and CTR1 -/-/CTR2 kd cells were exposed to increasing concentrations of DDP for five days, and the change in growth rate was quantified by staining the remaining cells with sulforhodamine B.

Figure 2-2A shows the concentration-survival curves for each of the cell lines. Loss of

CTR1 function rendered the CTR1 -/-/CTR2 +/+ cells 2.6-fold more resistant to DDP

+/+ +/+ relative to the CTR1 /CTR2 cells. The mean (± SEM) IC 50 values were 2.1 ± 0.02

µM and 5.5 ± 0.2 µM for the two cell lines, respectively (p = 0.002). In contrast, the knockdown of CTR2 rendered cells hypersensitive to DDP irrespective of whether

CTR1 was expressed or not. Knockdown of CTR2 in the CTR1 +/+ /CTR2 +/+ cells reduced the IC 50 by 69% to 0.7 ± 0.01 µM (p = 0.0001). Likewise, knockdown of

-/- +/+ CTR2 in the CTR1 /CTR2 cells reduced the DDP IC 50 by 51% to 2.7 ± 0.2 µM (p

= 0.0002). Thus, loss of mCTR2 expression caused a 3.2-fold increase in DDP sensitivity in wild-type cells and a 2.0-fold increase in cells lacking mCTR1.

A similar effect on cell growth was observed when the knockdown cells were exposed to CBDCA (Figure 2-2B). The mean ± SEM IC 50 values for CBDCA were as follows: CTR1 +/+ /CTR2 +/+ cells, 79.6 ± 0.3 µM; CTR1 +/+ /CTR2 kd cells, 38.4 ± 1.1

µM; CTR1 -/-/CTR2 +/+ cells, 197.8 ± 7.1 µM; and, CTR1 -/-/CTR2 kd cells, 95.9 ± 2.4

µM. Thus, reduction of mCTR2 expression caused a 1.9-fold increase in CDCBA sensitivity in the parental wild-type cells (p = 0.0006) and a 2.1-fold increase in cells lacking mCTR1 (p = 0.002). In contrast, as shown in Figure 2-2C, while loss of CTR1 function rendered the cells 2.6-fold resistant to Cu (p = 0.009), reduction in the

25 expression of CTR2 had no discernible effect on the sensitivity to Cu in either the

+/+ -/- CTR1 or CTR1 background. The mean ± SEM IC 50 values for Cu were:

CTR1 +/+ /CTR2 +/+ cells, 243.9 ± 20.9 µM; CTR1 +/+ /CTR2 kd cells 309.6 ± 23.0 µM;

CTR1 -/-/CTR2 +/+ cells, 93.1 ± 9.2 µM; and, CTR1 -/- CTR2 kd cells, 95.8 ± 25.6 µM.

Thus, knockdown of CTR2 produced a similar effect on sensitivity to DDP and

CBDCA; however, there was a clear difference in the effect of knocking down CTR2 expression on sensitivity to these two Pt-containing drugs and the effect on sensitivity to Cu.

Reduction of CTR2 expression increases whole cell Pt drug accumulation.

To determine whether the change in sensitivity to DDP was linked to changes in drug accumulation, total (whole cell) Pt accumulation was measured following either a 5 min or 1 h exposure to 30 µM DDP in all four cell lines by ICP-MS. The data were normalized to the content of sulfur as measured by ICP-OES as a surrogate for total cellular protein. Figure 2-3 shows that, in both the CTR1 +/+ and CTR1 -/-

27 backgrounds, reduction in the expression of CTR2 increased the whole cell accumulation of DDP. Reduction of CTR2 expression in the CTR1 +/+ background increased initial accumulation, determined by 5 min exposure, by 2.2-fold (p = 0.004), whereas in the CTR1 -/- background the increase was 2.8-fold (p = 0.006). After 1 h of

DDP exposure, the accumulation was 2.1-fold higher in CTR1 +/+ /CTR2 kd cells than in

CTR1 +/+ /CTR2 +/+ cells (p = 0.003.); likewise the uptake was 3.5-fold higher in CTR1 -/-

/CTR2 kd cells than in CTR1 -/-/CTR2 +/+ cells (p = 0.03) (Figure 2-3). As shown in

Figure 2-4, a similar, although more muted, change in accumulation was observed for

CBDCA after a 1 h period of drug exposure.

As expected, deletion of CTR1 reduced the Cu accumulation at 1 h to 70% of control. Knockdown of CTR2 expression in the CTR1 +/+ background caused 1.4-fold increase in Cu uptake (p = 0.01) (Figure 2-4). Knockdown of CTR2 in the CTR1 -/-

28 background had little effect. These results indicate that CTR2 has greater effects on the cellular pharmacology of the Pt-containing drugs than did Cu. In wild-type cells, knockdown of CTR2 increased Cu uptake, suggesting that CTR2 functions to efflux

Cu. Under circumstances where Cu uptake was severely impaired due to loss of CTR1 function, knockdown of CTR2 had little further effect. In contrast, knockdown of

CTR2 substantially increased DDP uptake irrespective of the status of CTR1. This implies that the interaction of DDP with CTR2 is independent of the function of

CTR1. The fact that similar effects were observed on both the initial and subsequent phases of uptake indicates that either CTR2 functions to suppress influx, or it affects an efflux system that operates much more rapidly than previously appreciated.

Loss of mCTR2 expression increases DNA adduct formation.

In a prior study conducted in human ovarian cancer cells, forced over- expression of CTR1 increased DDP accumulation but failed to increase cytotoxicity or

DNA adduct formation (Holzer, Samimi et al. 2003). To determine whether the increased influx of DDP that accompanies the knockdown of CTR2 led to more drug reaching the nucleus and critical targets that mediate cytotoxicity, the extent of DNA adduct formation was measured in each of the four cell lines after a 1 h exposure to 30

µM DDP. Figure 2-5 shows that knockdown of CTR2 in both the CTR1 +/+ and CTR1-/- background increased DNA adduct formation. Knockdown of CTR2 in the CTR1 +/+ cells increased DNA adduct formation by 2.1-fold (p = 0.0002), whereas in the CTR1 -

/- cells, it increased adduct formation by 3.2-fold (p = 0.001). The close parallel between the increase in DNA adduct formation closely and the increase in whole cell accumulation indicates that the enhancement of drug accumulation was not simply due sequestration of drug in intracellular vesicles. Instead, the increased Pt represented a pool of drug available for trafficking to the nucleus and reacting with DNA.

Table 2-1. Accumulation of DDP, CBDCA, and Cu.

CTR1 +/+ /CTR2 +/+ CTR1 +/+ /CTR2 kd CTR1 -/- CTR1 -/- /CTR2 +/+ /CTR2 kd DDP uptake at 0.63 ± 0.10 1.31 ± 0.12 0.31 ± 0.07 0.86 ± 0.02 5 min* DDP uptake at 2.53 ± 0.70 5.33 ± 0.12 1.22 ± 0.05 4.46 ± 0.31 1 h* CBDCA uptake at 1 99.5 ± 17.7 128.7 ± 27.8 55.5 ± 5.6 63.9 ± 11.8 h** Cu uptake at 1 4.40 ± 0.18 5.76 ± 0.17 3.21 ± 0.26 3.78 ± 0.03 h* DNA adduct formation, pM 0.14 ± 0.02 0.30 ± 0.01 0.10 ± 0.01 0.32 ± 0.03 Pt/ug DNA Vesicle accumulation, 0.50 ± 0.01 0.51 ± 0.02 0.49 ± 0.01 0.48 ± 0.04 ng Pt/ µg sulfur *ng/ µg sulfur **cpm/ug protein

Discussion

The results of this chapter indicate that CTR2 is an important determinant of both sensitivity to the cytotoxic effect of DDP and its intracellular pharmacology. To study the effect of CTR2 on the cellular pharmacology of the Pt drugs, we took advantage of a very powerful model and knocked down CTR2 expression in both

CTR1 +/+ and CTR1 -/- mouse embryo fibroblasts. Elimination of CTR1 substantially increased resistance to DDP and CBDCA, as we have previously reported (Holzer,

Manorek et al. 2006; Larson, Blair et al. 2008). A decrease in CTR2 by ~85% was achieved in each cell line by shRNA lentiviral infection. These reductions in CTR2 protein level led to a 2.0 to 3.2-fold increase in sensitivity to DDP irrespective of the presence or absence of CTR1. A similar result was observed for CBDCA. Thus, the effect of reducing CTR2 expression was not dependent on the CTR1 status of the cells. This is in contrast to its effect on sensitivity to the cytotoxic effect of Cu.

Elimination of the expression of CTR1 produced the anticipated response in sensitivity to Cu; however, knockdown of CTR2 in the CTR1 -/- cells had no further effect on sensitivity. These results support two conclusions. First, in the case of the Pt drugs, the effect of knocking down CTR2 appears to be independent of the status of CTR1.

Second, CTR2 functions differently than CTR1 with respect to Cu and the Pt drugs.

To explore the mechanism by which loss of CTR2 increased cell sensitivity to the Pt drugs, we measured whole cell drug accumulation at 5 min and 1 h and the extent of DNA adduct formation by ICP-MS. Consistent with our prior studies

(Larson, Blair et al. 2008), deletion of both alleles of CTR1 reduced the influx of DDP

33 when measured at both 5 min and 1 h, and this was accompanied by a proportional decrease in DNA adduct formation. Reduction of CTR2 expression had the opposite effect. Knockdown of CTR2 led a ~2.1 to 3.5-fold increase in whole cell Pt accumulation and DNA adduct formation, and it did so irrespective of whether CTR1 was expressed or not. The increase in whole cell Pt accumulation and DNA adduct formation was similar to the magnitude of the change in cytotoxicity, suggesting that the hypersensitivity caused by loss of CTR2 was directly linked to increased accumulation. Knockdown of CTR2 produced a very similar change in cytotoxicity and drug accumulation for CBDCA, indicating that, despite the differences in the structure of DDP and CBDCA and their rates of aquation and reaction with nucleophilic targets, these drugs are affected similarly by CTR2.

As noted with respect to cytotoxicity, the knockdown of CTR2 had somewhat different effects on the cellular accumulation of DDP and CBDCA versus Cu. Several points are noteworthy. First, complete loss of CTR1 expression only reduced whole cell Cu accumulation at 1 h by 31%, implicating existence of a mechanism for Cu accumulation in addition to CTR1. Second, unlike what occurs for DDP, the effect of knocking down CTR2 on Cu accumulation was dependent on CTR1: Knockdown of

CTR2 increased Cu accumulation only when CTR1 was expressed. In the absence of

CTR1 expression, the reduction in CTR2 produced only a small additional increase in uptake. Interestingly, the increase in DDP accumulation that accompanied knockdown of CTR2 was associated with an increase in cytotoxicity, whereas this was not true for

Cu. These observations imply that CTR2 interacts differently with DDP than with Cu.

The results of this chapter indicate that CTR2 is a potent regulator of DDP and

CBDCA initial uptake and accumulation. Loss of CTR2 will lead to increased accumulation of these Pt drugs and thereby enhance their cytotoxic effect.

Materials and Methods

Drugs and reagents.

Platinol AQ was a gift from Bristol-Myers Squibb (Princeton, NJ); it contains

3.33 mM DDP in 0.9% NaCl. [ 14 C]-CBDCA was purchased from Amersham

Biosciences (Pittsburgh, PA). The drugs were diluted into OptiMEM Reduced Serum

Media (Gibco, 31985-070) to produce final concentrations of 10, 30 and 100 M.

Bradford reagent was purchased from BioRad Laboratories, Inc. (Hercules, CA), sulforhodamine B was obtained from Sigma-Aldrich (St. Louis, MO) and 0.4% sulforhodamine B (w/v) was solubilized in 1% (v/v) acetic acid solution.

Cell types, culture and engineering .

Parental mouse embryonic fibroblasts containing wild-type alleles of CTR1

(CTR1 +/+ ) and an isogenic line in which both copies of CTR1 had been somatically knocked out (CTR1 -/-) were a gift from Dr. Dennis Thiele (Lee, Petris et al. 2002). The

CTR2 kd sublines were constructed by infecting the CTR1 +/+ and CTR1 -/- cells with lentivirus expressing a shRNA targeting mouse CTR2 mRNA purchased from Sigma-

Aldrich (St. Louis, MO). The shRNA sequences used were:

CCGGGCCTTGGAACACATGAGGATTCTCGAGAATCCTCATGTGTTCCAAGG

CTTTTTG and CCGGCCCACTTCTCAACATGACTTACTCGAGTA

AGTCATGTTGAG AAGTGGGTTTTTG. Knockdowns were selected in media containing 5 µM puromycin. Cell survival following exposure to increasing concentrations of drugs was assayed using the sulforhodamine B assay system

(Monks, Scudiero et al. 1991). Five thousand cells were seeded into each well of a 96- well tissue culture plate. Cells were incubated overnight at 37°C, 5% CO 2 and then exposed to varying drug concentrations in 200 l complete medium. Cells were allowed to grow for 5 days, after which the media was removed, the protein precipitated with 50% trichloroacetic acid and stained using 100 l of 0.4% sulforhodamine B in 1% acetic acid at room temperature for 15 minutes. Following washing, the absorbance of each well at 515 nm was recorded using a Versamax

Tunable Microplate Reader (Molecular Devices, Sunnyvale, CA). All experiments were repeated at least three times using three cultures for each drug concentration.

Western blotting .

Whole-cell lysates were dissolved in lysis buffer (150 mM NaCl, 5 mM EDTA,

1% Triton X-100, and 10 mM Tris, pH 7.4) and were subjected to electrophoresis on 4 to 15% gels using ~30 µg protein per lane. Protein levels were first determined by

Bradford assay (Bio-Rad, Richmond, CA). A Bio-Rad trans-blot system was used to transfer the proteins to Immobilin-P membranes (Millipore, Billerica, MA). Blots were

36 incubated overnight at 4°C in 4% dry nonfat milk in Tris-buffered saline (150 mM

NaCl, 300 mM KCl, 10 mM Tris, pH 7.4, 0.0% Tween 20). Blots were incubated for 1 h at room temperature in CTR2 antibody at 1:400 dilution (generous gift from Dr.

Bertinato). A horseradish peroxidase-conjugated secondary antibody (GE Healthcare,

Chalfont St. Giles, Buckinghamshire, UK) was dissolved in 4% milk in the Tris- buffered saline buffer and incubated with the blot for 1 h at room temperature. After four 5-min washes, blots were exposed to the PIERCE ECL reagent (Thermo

Scientific, Wilmington, DE) and detected on X-ray films (HyBlot CL; Denville

Scientific, Inc. Metuchen, NJ).

qRT-PCR .

CTR2 mRNA levels were measured using a qPCR method of detection of relative amounts of first-strand cDNA. cDNA was generated from mRNA isolated using Trizol (Invitrogen, Carlsbad, CA). Purified mRNA was converted to cDNA using Oligo(dT) 20 priming and the SuperScript III First-Strand Kit (Invitrogen). qPCR was performed on a Bio-Rad MyIQ qPCR machine (Hercules, CA). The forward and reverse primers for hCTR2, mCTR2, and mouse ß-actin were, respectively: mCTR2 forward – tccaggtagtcatcagct; mCTR2 reverse – tggcagtgctctgtgatgtc; ß-actin forward

– aggtgacagattgcttctg; ß-actin reverse – gctgcctcaacacctcaac. Reactions were prepared using iQ SYBR Green Supermix (Bio-Rad, Hercules, CA), according to manufacturer’s recommendations. Samples were prepared in quadruplicate with three

37 independent sample sets being analyzed. Analysis was done using the Bio-Rad iQ5 system software (Hercules, CA).

Measurement of drug accumulation into whole cells and DNA .

CTR1 +/+ , CTR1 +/+ CTR2 kd , CTR1 -/- and CTR1 -/- CTR2 kd cells were grown to

90% confluence in T-150 tissue culture flasks. Cells were then harvested using trypsin, and 7.5 x 10 5 cells were placed into each well of 6-well tissue culture plates and allowed to grow overnight in 2.5 ml of media at 37°C in 5% CO 2. The next day, medium was removed by aspiration and the cells were exposed to 500 l of drug- containing OptiMEM medium (Invitrogen, Carlsbad, CA) at 37°C for either 0, 5 or 60 min, after which the drug-containing medium was removed, the plates were washed the times with ice-cold PBS and were then placed on ice. In the case of the time zero samples, the drug-containing medium was aspirated within 15 sec of the start of drug exposure. Two hundred and fifteen l of concentrated (50-70%) nitric acid was added to each well and the plate was incubated overnight at room temperature. The following day the acid was moved into Omni-vials (Wheaton, Millville, NJ) and incubated at room temperature overnight to dissolve all cellular debris. The following day, the nitric acid was diluted with 3 ml of buffer (0.1% Triton X-100, 1.4% nitric acid, 1 ppb

In in ddH 2O). Pt concentration was measured using a Perkin-Elmer Element 2 ICP-MS located at the Analytical Facility at Scripps Institute of Oceanography at UCSD. As a method of normalization, total sulfur was measured using a Perkin-Elmer ICP-OES also located at SIO at UCSD. Samples that were previously prepared for the ICP-MS

38 were then introduced into the ICP-OES where total µg of sulfur was measured. All data presented are the means of at least three independent experiments each performed with six wells per concentration tested.

For measurement of Pt in DNA, cells were lysed and DNA harvested using

DNAzol (Invitrogen) according to the manufacturer’s protocol. As a method of normalization, DNA was measured prior to addition of nitric acid using a Nanodrop

1000 spectrophotometer (Thermo Scientific, Wilmington, DE). The microsome and

DNA samples were digested in nitric acid prior to measurement of Pt by ICP-MS as described above.

Measurement of [ 14 C]-CBDCA and 64 Cu accumulation.

Cells were seeded at 7.5 x 10 5 per well in 6-well tissue culture plates and allowed to grow overnight in 2.5 ml of media at 37°C in 5% CO 2. For measurement of

[14 C]-CBDCA accumulation, 500 µl of 50 µM [ 14 C]-CBDCA was added to the cells

and incubated at 37°C in 5% CO 2 for 60 min. At the end of the incubation period, the plates were placed on ice, and the wells were rinsed three times with 3 ml of ice-cold

PBS. Cells were harvested in 200 µl lysis buffer (150 mM NaCl, 5 mM EDTA, 1%

Triton X-100, and 10 mM Tris, pH 7.4) and transferred to tubes containing 3 ml of scintillation buffer (National Diagnostics, Atlanta, GA). [ 14 C]-CBDCA was quantified by scintillation counting. Total protein as measured by Bradford assay was used for

64 64 normalization of values. For measurement of Cu accumulation 2 µM CuSO 4 was

added to the plates and incubated at 37°C in 5% CO 2 for 60 min. At the end of the

39 incubation period, the plates were placed on ice and the wells were rinsed three times with 3 ml of ice-cold PBS. Cells were harvested in 215 µl concentrated nitric acid and transferred to tubes containing 3 ml of buffer as described above for counting on a

Beckman Gamma 5500B (Beckman Coulter, Fullerton, CA). Total sulfur was used for normalization as described earlier. All data presented are the means of at least three independent experiments each performed with six wells per concentration tested.

Statistical Analysis.

All data represents at least three independent experiments, presented with the standard error from the mean (SEM). Statistical comparisons were done using a two- tailed t-test.

Acknowledgements

A majority of the content of Chapter 2 has been published in Clinical Cancer

Research (Blair BG, Larson CA, Safaei R, Howell SB. Copper Transpoter 2 Regulates the Cellular Accumulation and Cytotoxicity of Cisplatin and Carboplatin. Clin Cancer

Res. 2009 Jul 1;15(13):4312-21.) Brian G. Blair was the primary researcher and author for this chapter. Stephen B. Howell supervised and directed the research in this chapter. Christopher A. Larson assisted in the measurement of Pt and Cu accumulation and provided helpful feedback. Roohangiz Safaei provided helpful discussion. The author would like to thank Dr. Dennis Thiele for generously providing the CTR1 +/+

40 and CTR1 -/- mouse embryo fibroblasts, Dr. Jessie Bertinato for the anti-CTR2 antibody and Ms. Sakura Moua for technical assistance.

Chapter 3

The Effect of the Loss of CTR2 on Cisplatin Sensitivity and

Accumulation in Vivo

Introduction

The previous chapter demonstrated that CTR2 limits the accumulation of DDP in cell line models. This chapter reports on studies directed at examining whether

CTR2 is an important determinant of the responsiveness to DDP in vivo . Specifically, the effect of knocking down the expression of CTR2 in malignant mouse embryo fibroblasts on the accumulation of DDP and the ability of DDP to slow tumor growth was examined. Tumors derived from cells in which CTR2 had been knocked down grew more slowly than those derived from wild-type cells, and this was associated with an increased frequency of apoptotic cells and decreased vascular density.

However, the tumors in which CTR2 was knocked down accumulated more Pt following injection of DDP and exhibited a much greater response to treatment. These observations suggest that selective inhibition of CTR2 expression or function may be a useful strategy for enhancing the effectiveness of DDP chemotherapy.

41 42

Results

Effect of CTR2 Knockdown on Tumor Growth Rate.

To determine the dependence of tumor growth on CTR2 in vivo we utilized a mouse embryo fibroblast cell line in which both alleles of CTR1 had been deleted in order to remove any confounding effects of CTR1. The expression of CTR2 in these

CTR1 -/- cells was knocked down using a lentiviral vector that expressed a shRNAi directed to the CTR2 mRNA. As noted in the previous chapter, the level of expression of CTR2 protein in the cell line before tumor inoculation was reduced to 87.1± 4.6 % of that in the parental CTR1 -/- cells. The CTR1 -/- and CTR1 -/- CTR2 kd cells were inoculated subcutaneously into nu/nu mice; both types of cells formed tumors with equal frequency. Immunohistochemical analysis of sections from these tumors demonstrated robust expression of CTR2 in the CTR1 -/- tumors, but no detectable

CTR2 expression in the CTR1 -/- CTR2 kd tumors (Figure 3-1A). As shown in Figure 3-

1B, CTR1 -/- tumors grew 5.8-fold more rapidly than CTR1 -/- CTR2 kd tumors.

Effect of CTR2 on Proliferation and Apoptosis in Vivo .

To examine the basis for the difference in growth rate, CTR1 -/- and CTR1 -/-

CTR2 kd tumors were harvested and preserved in paraffin blocks. The tumors were accessed for Ki67 staining to determine the effect of knocking down CTR2 on proliferation rates. CTR2 kd cells demonstrate 24.3 ± 10.3% (p < 0.02) fewer Ki67 stained cells than CTR1 -/- tumors (Figure 3-2A). The tumors were sectioned and the frequency of apoptotic cells measured by TUNEL assay (Figure 3-2B). The average number of TUNEL positive nuclei per high power field was determined for each tumor type. CTR1 -/- tumors had an average 42.8 ± 6.2 TUNEL positive nuclei per high power field (Figure 3-2B). In contrast CTR1 -/- CTR2 kd tumors had an average of 81.8

± 10.8 TUNEL positive nuclei per high power field. Thus, the frequency of apoptotic cells in the CTR1 -/- CTR2 kd tumors was 1.9-fold higher than in the CTR1 -/- tumors suggesting that the death rate of tumor cells was increased when CTR2 was knocked down.

Effect of CTR2 on Vessel Density in Vivo .

Cu is essential for angiogenesis, and adequate vascularization is required for tumor growth. To determine whether knockdown of CTR2 altered the extent of angiogenesis in tumors, subcutaneously implanted CTR1 -/- and CTR1 -/- CTR2 kd

45 tumors were harvested and frozen in O.C.T. compound. Tumors were sectioned and stained with an antibody to the endothelial cell marker CD31. Figure 3-3 shows a reduced density of CD31-expressing cells in the CTR1 -/- CTR2 kd tumors. The number of vessels per square mm was 83.7 ± 7.0 in the CTR1 -/- tumors but was reduced to

57.3 ± 3.5 in the CTR1 -/- CTR2 kd tumors (Figure 3-3). Thus, the vessel density was

1.5-fold higher in the CTR1 -/- tumors (p < 0.001) indicating that CTR2 has a substantial effect on tumor vessel formation.

Effect of CTR2 on Cu Content In Vitro and In Vivo .

The exact role of CTR2 in Cu homeostasis is not defined. Knockdown of

CTR2 increased the steady-state level of Cu in the CTR1 -/- CTR2 kd cells when grown in vitro . The level in CTR1 +/+ cells was 1.10 ± 0.02 ng Cu/ug sulfur when grown in standard tissue culture medium (Figure 3-4A). The level in the CTR1 -/- cells did not significantly differ being 0.90 ± 0.10 ng Cu/ug sulfur (Figure 3-4A). Knockdown of

CTR2 in the CTR1 -/- cells increased the steady-state Cu level by 2.1-fold to 1.89 ±

0.01 ng Cu/ug sulfur (p < 0.01) (Figure 3-4A). To determine whether similar differences were observed when the CTR1 -/- and CTR1 -/- CTR2 kd cells were grown in vivo , untreated CTR1 -/- and CTR1 -/- CTR2 kd subcutaneous tumors were harvested and dissolved in nitric acid and the Cu levels were assayed by ICP-MS. There was no significant difference in steady state Cu levels which were 552.2 ± 18.3 ng Cu/mg sulfur in the CTR1 -/- tumors and 535.9 ± 36.0 ng Cu/mg sulfur in the CTR1 -/- CTR2 kd tumors (p = 0.7) (Figure 3-4B). Thus, despite the clear effect of knocking down CTR2 on cellular Cu levels when grown in vitro , when grown in vivo the knockdown of

CTR2 did not alter Cu levels.

Effect of CTR2 knockdown on DDP Accumulation In Vivo.

Nu/nu mice with subcutaneous CTR1 -/- and CTR1 -/- CTR2 kd tumors were injected intraperitoneally with 10 mg/kg DDP and 1 h later the mice were sacrificed

47 and tumors harvested. The Pt level in each tumor was determined by ICP-OES. As shown in Figure 3-5, the average Pt level in the CTR1 -/- tumors was 2.26 ± 0.36 ng

Pt/mg sulfur. The average Pt level in the CTR1 -/- CTR2 kd tumors was 20.62 ± 3.53 ng

Pt/mg sulfur. Thus, the CTR1 -/- CTR2 kd tumors accumulated 9.1-fold more Pt at 1 h after injection of DDP than the CTR1 -/- tumor (p = 0.006). This is a very large difference in Pt accumulation compared to the 3.5-fold difference in uptake observed for these cells when grown in vitro and what is generally observed in DDP-sensitive and DDP-resistant cell lines.

Effect of CTR2 Knockdown on Responsiveness to DDP In Vivo.

As shown in Figure 3-6A, a single intraperitoneal injection of the maximum tolerated dose of DDP (10 mg/kg) produced little slowing of the growth of CTR1 -/- tumors relative to the growth rate of the untreated control tumors (p = 0.75).

However, the same dose of DDP clearly slowed the growth of CTR1 -/- CTR2 kd tumors

(p <0.0009) (Figure 3-6B). The average volume of the DDP-treated CTR1 -/- CTR2 kd tumors was 74% smaller than the untreated CTR1 -/- CTR2 kd tumors by week 7. Many of these tumors shrank in size and remained smaller than they were before treatment.

Six of the 16 tumors became undetectable and did not regrow during the period of observation. The remaining tumors either stopped growing or grew at a much slower rate than the CTR1 -/- tumors. Thus, consistent with the effect of CTR2 on DDP accumulation, CTR2 is a major determinant of therapeutic efficacy of DDP in vivo .

Relationship between hCTR2 expression and DDP sensitivity in ovarian cancer.

Given that a reduction in CTR2 was associated with a 2.0- to 3.2-fold increase in drug sensitivity in the mouse embryo CTR1 +/+ and CTR1 -/- fibroblasts, it was of interest to determine whether human ovarian cell lines that vary in sensitivity to DDP differ in their expression of CTR2. CTR2 protein expression levels were analyzed in six established human ovarian carcinoma cell lines of varying sensitivity to DDP. The

IC 50 for DDP was determined for each cell line by quantifying inhibition of growth by staining with sulforhodamine B. CTR2 mRNA expression was quantified using qRT-

PCR with β-actin as a loading control. CTR2 expression was quantified by densitometry on triplicate Western blots on which α-tubulin expression was used as a loading control (Figure 3-7A). Figures 3-7B and 3-7C show that there was a significant correlation between CTR2 expression at both the mRNA (r 2 = 0.97, p =

0.0003) and protein levels (r 2 = 0.71, p < .04) and resistance to the cytotoxic effect of

DDP. The higher the expression of hCTR2, the greater was the observed DDP IC 50 .

This result suggests that CTR2 expression may be one of the parameters that determine differences in DDP sensitivity in human ovarian carcinomas.

Discussion

The experiments reported in this chapter sought to investigate the effect of

CTR2 on tumor growth rate, DDP accumulation and responsiveness in vivo . Since the results from the previous chapter indicated that the effect of CTR2 on DDP uptake and sensitivity are independent of CTR1 status when the cells are grow n in vitro , in the current experiments I opted to use CTR1 -/- cells as the parental cells for knockdown of

CTR2 in part because this removed any possibility of compensating effects on Cu homeostasis that might be mediated by CTR1 when CTR2 was knocked down.

The first finding of importance was that the growth rate of the CTR1 -/- CTR2 kd tumors was substantially slower than that of the CTR1 -/- tumors. The finding of a higher frequency of apoptotic cells, slower proliferation and a lower density of CD31- positive capillaries in the CTR1 -/- CTR2 kd tumors provides a reasonable explanation for why the growth rate was impaired, but how the loss of CTR2 produces these effects remains unknown. There was a 2.1-fold higher level of Cu in the CTR1 -/-

CTR2 kd cells when grown in vitro that was not observed when they were grown in vivo . Since the whole tumor was harvested, which contains mouse derived endothelial cells and mesenchymal cells in addition to tumor cells, it is conceivable that the difference in the Cu content of the actual tumor cells was missed. However, the fact that there was such a large difference in Pt levels suggests that the extent of contamination of the tumor cells by normal mouse cells was not large. There is a substantial body of data indicating that Cu is required for angiogenesis (reviewed in

(Finney, Vogt et al. 2009)) but the fact that no differences were detected in steady- state Cu levels between the two types of tumors leaves open the possibility that, despite its important role in Cu homeostasis, CTR2 modulates the angiogenic response via effects on other pathways. It is of interest that CTR1 has been shown to regulate signaling via the FGF receptor pathway during embryonic development of neuroectoderm in Xenopus (Haremaki, Fraser et al. 2007). Given the structural

52 similarity of CTR1 and CTR2, it is conceivable that CTR2 regulates the synthesis or release of angiogenic factors from tumor cells.

CTR2 is an important determinant of the sensitivity of DDP when cells are grown in tissue culture; however, results obtained using cultured cells do not always extrapolate to the in vivo setting. Gratifyingly, the results of the current chapter show that CTR2 is a major determinant of both DDP accumulation and its therapeutic effectiveness in vivo as well as in vitro. Given the fact that CTR2 also regulates sensitivity to carboplatin in vitro , it is likely that CTR2 expression is also important to the tumor pharmacology and responsiveness of this drug in vivo as well.

The magnitude of the effect of knocking down CTR2 on DDP accumulation in vivo (9.1-fold) was larger than the difference in uptake observed when the cells were grown in vitro (3.5-fold), and is very large relative to the ~50% difference in DDP uptake typically observed in isogenic pairs of DDP-sensitive and resistant cell lines

(Andrews and Howell 1990; Muggia and Los 1993). This suggests that CTR2 is particularly important to DDP accumulation in the complex in vivo environment in which rates of drug delivery, protein binding and aquation are different from those obtained when cells are exposed to DDP in vitro . Although the cellular level of Cu modulates the uptake of DDP, the similarity of the Cu levels in the CTR1 -/- and

CTR1 -/- CTR2kd tumors makes it unlikely that Cu availability accounts for the difference in DDP accumulation.

Given that small changes in CTR2 expression produced relatively large changes in sensitivity to the cytotoxic effect of DDP, we were interested in whether

53 differences in CTR2 expression in human tumor cell lines were linked to differences in intrinsic sensitivity to DDP. In a panel of six human ovarian carcinoma cell lines, we found a significant correlation between DDP sensitivity and both CTR2 mRNA level (r 2 = .97) and CTR2 protein level (r 2 = 0.71). This provides the impetus for further studies focused on how CTR2 levels change during the acquisition of resistance that accompanies repeated exposure to DDP in vitro and in vivo .

Materials and Methods

Drugs and Reagents .

Platinol AQ was a gift from Bristol-Myers Squibb (Princeton, NJ); it contains

DDP at a concentration of 3.33 mM in 0.9% NaCl. The anti-CTR2 antibody was a gift from Dr. Jessie Bertinato (Health Canada, Ottawa, ON).

Cell Types.

Parental mouse embryonic fibroblasts in which both copies of CTR1 had been somatically knocked out (CTR1 -/-) were a gift from Dr. Dennis Thiele (Lee, Petris et al. 2002). The CTR2 kd sublines were constructed by infecting the CTR1 -/- cells with lentivirus expressing a shRNAi targeting mouse CTR2 mRNA purchased from Sigma-

Aldrich (St. Louis, MO) as previously described (Blair, Larson et al. 2009).

Western Blotting .

Cell lysates were prepared and subsequent Western blots were conducted as previously described.

Immunohistochemistry and Chemiluminescent Immunoblotting.

Upon harvesting, tumors were either imbedded in paraffin or frozen in O.C.T. compound as previously described (Holzer, Manorek et al. 2006). Sections were stained according to the protocol outlined in the Catalyzed Signal Amplification

System (cat. #K1500; DAKO, Carpinteria, CA). Endogenous biotin was blocked by first overlaying the slides with 0.1% avidin for 15 min and washing 3 times with 0.1%

Triton-X in PBS. Slides were then overlaid with 0.01% biotin for 15 min followed by another 3 washes with 0.1% Triton-X in PBS. Nonspecific protein binding was blocked by immersion of the slides in 1% BSA in PBS for 20 min. Slides were incubated with anti-hCTR2 antibody at a dilution of 1:200 in 1% BSA in PBS overnight at 4º C. As a negative control, parallel sections were incubated with non- immune rabbit IgG1 sera (prediluted; DAKO). Slides were also stained with anti-Ki67 to determine the fraction of S phase nuclei and anti-CD31 antibody (BD Pharmingen,

San Jose, CA) to quantify vascular density and apoptotic nuclei were detected by

TUNEL assay (Latimer, Menchaca et al. 2009).

Quantification of vascular density.

The mean vascular density (vessels per square millimeter) for each tumor was calculated as previously described (Lucidarme, Kono et al. 2006). Five light

55 microscope pictures at 40X were taken at different locations in each tumor sample.

The total count of CD31-stained vessels was divided by the area of the five fields to obtain the mean vascular density. Eight to 10 tumors of each cell type were scored by two blinded observers; the values reported are the mean of the ratio for the two types of tumors.

Measurement of Drug Accumulation into Tumors.

One hour following intraperitoneal injection of 10 mg/kg DDP tumors were harvested and digested in 70% nitric acid overnight, diluted to a final 5% nitric acid

(0.1% Triton X-100, 1.4% nitric acid, 1 ppb In in ddH2O). Pt concentration was measured using a Perkin-Elmer Element 2 ICP-OES located at the Analytical Facility at Scripps Institute of Oceanography at UCSD. As a method of normalization, total sulfur was also measured by ICP-OES in each sample. Cu levels were measured using the same instrument. All data presented are the means of at least 3 independent experiments each performed with 6 cultures per concentration tested.

Determination of Drug Sensitivity in Vivo.

To grow the various types of cells as xenografts, 3 x 10 6 cells in 100 µl were inoculated at 4 subcutaneous sites into 20 g female nu/nu mice. Cell types were randomized between shoulder and hip, left and right, ensuring that there were always tumors of the same type on left and right. Tumors were allowed to grow until they were 2 mm in diameter, at which point each mouse received a single dose of DDP 10

56 mg/kg by intraperitoneal injection. Tumor size was monitored 3 times per week for 7 weeks. Tumor volume was estimated using the equation (length x width2)/2.

Statistical Analysis.

All data were derived from at least 3 independent experiments, and are presented as mean values with standard errors (SEM). A total of 23 mice were used for tumor experiments (12 DDP-treated, 11 untreated) each with 4 tumors distributed as described above. Statistical comparisons were performed using a two-tailed t-test with the assumption of unequal variance.

Acknowledgements

A majority of the content of Chapter 3 has been submitted for publication. The remaining sections of this chapter have been adapted from published results in Clinical

Cancer Research (Blair BG, Larson CA, Safaei R, Howell SB. Copper Transpoter 2

Regulates the Cellular Accumulation and Cytotoxicity of Cisplatin and Carboplatin.

Clin Cancer Res. 2009 Jul 1;15(13):4312-21.) Brian G. Blair was the primary researcher and author for this chapter. Stephen B. Howell supervised and directed the research in this chapter. Christopher A. Larson and Preston L. Adams assisted in the in vivo studies as well as the measurement of Pt and Cu accumulation and provided helpful feedback. Catherine Pesce and Gerald Manorek assisted in tumor and organ harvesting. Roohangiz Safaei provided helpful discussion.

Chapter 4

The Regulation of CTR2 Expression and Degradation by

Copper and Pt

Introduction

The results of the studies described in the previous chapters established that

CTR2 can regulate Pt drug accumulation and sensitivity both in vitro and in vivo. How

CTR2 expression and localization is regulated has not yet been explored. Cu and DDP can quickly down-regulate the expression of CTR1 (Holzer and Howell 2006). Within

15 minutes of DDP exposure, nearly all CTR1 is removed from the plasma membrane in many types of cells in a process that involves macropinocytosis, ubiquitination and subsequent degradation of CTR1 by the proteosome (Holzer and Howell 2006; Jandial and Howell 2008). However, within ~30 min following the removal of DDP, the amount of plasma membrane bound CTR1 returns to normal (Holzer and Howell

2006). No information is available regarding the regulation of CTR2. Given its similarity to CTR1 with respect to structure and Cu transport, it is likely that CTR2 is also regulated by DDP and Cu availability.

57 58

The goal of the experiments described in this chapter was to determine how the expression of CTR2 is affected by Cu and DDP. The experiments revealed that CTR2 mRNA and protein levels increase when mammalian cells are exposed to either Cu or

DDP. Additionally, they demonstrated that CTR2 protein is rapidly degraded when cells are starved for Cu. Similar to the effect of knocking down CTR2 expression using shRNAi, the loss of CTR2 caused by depletion of cellular Cu leads to increased

DDP accumulation and cytotoxicity. Finally, we present evidence that the regulation of CTR2 due to the availability of Cu is mediated by Atox1.

Results

Regulation of CTR2 Expression by Cu and DDP .

Human ovarian carcinoma 2008 cells were exposed to 200 µM CuSO 4 for 1 h followed by the isolation of mRNA and cellular proteins to assess the effect of Cu on

CTR2 expression. At 1 h, the level of CTR2 mRNA increased by 2.4-fold (p = 0.01) as measured by qRT-PCR (Figure 4-1A), and this was accompanied by a 2.4-fold (p =

0.02) increase in CTR2 protein, as quantified by Western blot analysis (Figure 4-1B).

To confirm that the availability of Cu affects CTR2 levels, the 2008 cells were treated with 100 µM BCS for 1 h to deplete intracellular Cu. As shown in Figure 4-1, this brief period of Cu starvation produced a 29% decrease in CTR2 mRNA (p = 0.052) but a more marked 90% decrease in CTR2 protein level (p = 0.0006).

To determine whether DDP was able to regulate CTR2 expression in a manner similar to that of Cu, cellular CTR2 mRNA and protein levels were measured

59 following exposure of the 2008 cells to 30 µM DDP for 1 h. DDP treatment increased the CTR2 mRNA level by 1.4-fold (p = 0.057) (Figure 4-1A) and produced a similar

1.4-fold increase in protein level (p = 0.063) (Figure 4-1B). Thus, at the concentration tested, DDP caused only modest changes in CTR2 mRNA and protein expression compared to those produced by Cu alone.

To further document the ability of Cu and DDP to regulate protein expression of CTR2, 2008 ovarian carcinoma cells were treated for 1 h with either 200 µM

CuSO 4, 100 µM BCS, 30 µM DDP or drug-free control media and were then stained with an anti-CTR2 antibody and a fluorescently tagged secondary antibody and visualized by deconvolution microscopy. As shown in Figure 4-2, exposure to 200 µM

CuSO 4 greatly increased the cellular level of CTR2 protein, and Cu starvation produced by a 1 h treatment with 100 µM BCS resulted in near total loss CTR2 protein. In contrast, exposure to 30 µM DDP for 1 h produced only a modest increase in CTR2 staining.

Effect of Cu and DDP on CTR2 Half-life.

To determine whether the increase in CTR2 level following DDP and Cu treatment was due to increased protein stability, 2008 cells were treated with cycloheximide to block new CTR2 synthesis and the CTR2 level was determined as a function of time in the presence or absence of either DDP or Cu. The data presented in

Figure 4-3 demonstrate that both Cu and DDP increased CTR2 stability. The half-life of CTR2 was 14.5 ± 2.2 min in the absence of either Cu or DDP. The addition of 30

µM DDP for 1 h increased CTR2 half-life to 22.7 ± 0.7 min, or by 1.6-fold (p =

0.016). Exposure to Cu increased the half-life to 42.6 ± 4.0 min, or by 3.1-fold (p =

0.001). Thus, Cu and DDP can regulate the expression of CTR2 at the post- transcriptional level.

CTR2 is degraded by the proteosome.

Copper starvation quickly leads to the down-regulation of CTR2 protein. To determine if this decrease in CTR2 occurred via proteosomal degradation, cells were pretreated with bortezomib (BTZ), which blocks proteosomal degradation. Cells treated with orwithout BTZ were subjected to copper starvation. As indicated in

Figure 4-4, treatment with BTZ alone does not affect CTR2 protein concentration or localization. In the absence of BTZ treatment, copper-starved cells quickly lose CTR2 protein as previously observed. However, in the presence of BTZ, copper-starved cells

63 no longer degrade CTR2. This result implies that the degradation of CTR2 caused by copper starvation is dependent on proteosomal activity.

Cu Starvation Enhances DDP Uptake and Sensitivity.

Our previous studies in mouse embryo fibroblasts indicated that knockdown of mCTR2 enhanced the cellular accumulation and cytotoxicity of DDP (Blair, Larson et al. 2009). To determine whether reduction of CTR2 expression mediated by Cu depletion produced the same effect, and whether this occurred in human tumor cells,

2008 ovarian carcinoma cells were treated with 200 µM CuSO 4, 100 µM BCS or drug- free control media for 1 h and then exposed to increasing concentrations of DDP for

15 min. The Pt content of the cells was then measured by ICP-MS. The data presented in Figure 4-5A indicate that pre-treatment with Cu did not significantly change the

64 whole cell accumulation of DDP at 15 min. However, pre-treatment with BCS increased whole cell uptake at 15 min by 2.2-fold (p<0.0003).

To assess the effect of Cu exposure or depletion on the cytotoxicity of DDP, the cells were treated with either 200 µM CuSO 4, 100 µM BCS or drug-free control media for 1 h and the effect on growth rate over the ensuing 5 days was assessed using a sulforhodamine B assay. Figure 4-5B demonstrates that pre-treatment with Cu did not significantly change the sensitivity of the 2008 cells to DDP. The mean (± SEM)

IC 50 was 100.4 ± 4.5 µM in the absence of Cu pre-treatment and 101.8 ± 0.9 µM when the cells were pre-treated with Cu. However, as shown in Figure 4-5C, BCS-mediated

Cu starvation significantly increased sensitivity to DDP (2.0-fold). The mean (±

SEM) IC 50 value was 100.4 ± 4.5 µM in the absence of BCS pre-treatment and 49.1 ±

14.2 µM with BCS pre-treatment (p = 0.03). Thus, consistent with the effect of knocking down mCTR2 in mouse embryo fibroblasts, reduction in CTR2 expression by depletion of intracellular Cu enhanced both the cellular accumulation and cytotoxicity of DDP in 2008 human ovarian carcinoma cells.

Atox1 Influences the Expression of CTR2.

If CTR2 levels are regulated by the availability of intracellular Cu, then defects in the network of Cu chaperones that control the distribution of Cu might modulate

CTR2 levels as well. This possibility was addressed using a mouse embryo fibroblast sub-line in which both alleles of Atox1 had been knocked out (Atox1 -/-). In these cells there is a failure to deliver Cu to the secretory pathway for export from the cells and their steady-state Cu level is 3.2-fold higher than that of parental cells (Hamza,

Prohaska et al. 2003). The parental Atox1 +/+ and Atox1 -/- cells were treated with 200

µM CuSO 4, 30 µM DDP, 100 µM BCS or drug-free control media for 1 h and were then fixed and stained with antibody against CTR2. As shown in Figure 4-6, loss of

Atox1 greatly increased the steady state level of CTR2. However, this change in steady state level did not appear to alter the subcellular localization of CTR2 protein

(red). Cu and DDP mediated up-regulation of CTR2 protein was similar in the

Atox1 +/+ cells to that observed in 2008 cells. Interestingly, the level of CTR2 protein in Atox1 -/- cells did not significantly change when cells were treated with DDP and

Cu. Furthermore, whereas exposure to 100 µM BCS markedly reduced CTR2 expression in Atox1 +/+ cells, BCS failed to down-regulate CTR2 expression in the

Atox1 -/- cells. The requirement for Atox1 was further examined by Western blot analysis. The data presented in Figure 4-7 confirms that the CTR2 protein level in untreated Atox1 -/- cells was 3.1-fold higher than that in untreated Atox1 +/+ cells (p =

0.003). Again, while DDP or Cu exposure affected CTR2 levels in Atox1 +/+ cells in a similar fashion to that in 2008 cells, DDP and Cu did not significantly alter CTR2

67 protein expression in Atox1 -/- cells. Exposure to BCS down-regulated the expression of CTR2 in the Atox1 +/+ cells but failed to do so in the Atox1 -/- cells. BCS pre- treatment of the Atox1 +/+ cells caused a near total loss of CTR2 protein (12.5%, p

<0.0001) whereas in the Atox1 -/- cells there was no discernible difference in expression.

To determine how Atox1 influences CTR2 mRNA levels, CTR2 expression was quantified by qRT-PCR in the Atox1 +/+ and Atox1 -/- cells with and without drug

69 pre-treatment. Consistent with the protein expression data, the steady-state level of

CTR2 mRNA in Atox1 -/- cells was 3.5-fold higher than that in the Atox1 +/+ cells

(Figure 4-7). The magnitude of the change in CTR2 mRNA produced by exposure to

200 µM CuSO 4 (1.8-fold increase, p = 0.03), 30 µM DDP (1.6-fold increase, p<0.04) or 100 µM BCS (32% decrease, p<.07) in Atox1 +/+ cells was similar to that observed in 2008 cells. Interestingly, while the CTR2 protein level did not significantly change in the Atox1 -/- cells in response to DDP or Cu exposure, CTR2 mRNA expression in

Atox1 -/- cells changed in a manner similar to that observed in Atox1 +/+ and 2008 cells.

Exposure to 200 µM CuSO 4 led to a 1.9-fold increase in CTR2 mRNA (p<0.01) and cells exposed to 30 µM DDP had a 1.4-fold increase in CTR2 mRNA (p = 0.02). Cu starvation due to 100 µM BCS treatment resulted in a 43% decrease in CTR2 mRNA expression.

Discussion

The level of expression of CTR2 is a major determinant of sensitivity to the cytotoxic effect of DDP (Blair, Larson et al. 2009). To study the regulation of CTR2,

I measured the effect of Cu, DDP and Cu starvation on CTR2 mRNA and protein expression. Treatment of 2008 human ovarian carcinoma cells with 200 µM CuSO 4 for 1 h led to a 2.4-fold increase in both CTR2 mRNA expression and protein level.

On the other hand, starving the 2008 cells of Cu by treating with 100 µM BCS for 1 h

71 depleted CTR2 mRNA and reduced the protein to nearly undetectable levels. Similar results were obtained in mouse embryo fibroblast lines. Thus, Cu regulates CTR2 expression and the close relationship between the changes in mRNA and protein levels support the conclusion that this regulation occurs, at least partially, at the transcriptional level. Having previously shown that the expression of CTR2 has a large effect on both the accumulation and cytotoxicity of DDP (Blair, Larson et al. 2009), it was of interest to determine whether DDP also modulates the expression of this protein. Exposure to 30 µM DDP for 1 h increased CTR2 mRNA expression and protein levels by 1.4-fold. This indicates that, at a concentration known to be cytotoxic, DDP up-regulates CTR2, and thus that DDP induces the expression of a protein that limits its own uptake into tumor cells. It is noteworthy that the two Cu transporters, CTR1 and CTR2, have opposite effects on the cellular accumulation of

DDP, with CTR1 enhancing and CTR2 limiting uptake (Blair, Larson et al. 2009;

Larson, Blair et al. 2009), and that DDP and Cu have opposite effects on the level of expression of these two transporters. DDP causes rapid degradation of CTR1 (Holzer and Howell 2006; Jandial and Howell 2008) whereas the same concentration produces a modest increase in the expression of CTR2. Immunocytochemical and deconvolution microscopic examination confirmed the results of the qRT-PCR and

Western blot analyses with respect to the ability Cu and DDP to increase, and BCS to deplete, CTR2 in the 2008 ovarian cancer cells. Similar effects were also observed in mouse embryo fibroblasts. The immunocytochemical analysis disclosed that neither

Cu nor DDP produced a major change in the subcellular localization of CTR2.

To determine whether Cu and DDP regulate CTR2 by altering its degradation,

CTR2 half-life and proteosomal degradation were measured following Cu exposure or starvation. Cycloheximide inhibits protein synthesis, and the disappearance of a protein during cycloheximide exposure is commonly used to estimate its half-life.

Using this approach, the half-life of CTR2 under basal conditions was ~14.5 min but following 1 hr exposure to 200 µM CuSO 4 or 30 µM DDP , the half-life increased to

~42.5 min or ~22.7 min, respectively. Thus, the increase in the level of CTR2 protein following exposure to Cu or DDP likely results from regulation at both the transcriptional and post-transcriptional level.

CTR1 down-regulation by DDP and Cu is mediated by proteosomal degradation (Holzer and Howell 2006 ; Jandial and Howell 2008) and the current results indicate that Cu starvation triggers down-regulation of CTR2 in a similar manner, since the degradation of CTR2 was blocked by the proteosome inhibitor bortezomib. Thus, both CTR1 and CTR2 depend on the proteosome for degradation despite that fact that different signals trigger down-regulation. CTR1 is known to become polyubiquitinated in response to signals that trigger its degradation (e.g.,

DDP exposure) (Safaei, Maktabi et al. 2009); whether CTR2 becomes polyubiquinated in response to a degradation signal (e.g., byCu starvation) remains to be determined.

Knockdown of CTR2 expression using RNA inference results in a large increase in the cellular accumulation and cytotoxicity of DDP (Blair, Larson et al.

2009). In addition, increased CTR2 levels have been identified as a potential marker of

DDP resistance (Blair, Larson et al. 2009). Since Cu starvation leads to a rapid and near total loss of CTR2, and Cu exposure quickly up-regulates CTR2 levels, it was of interest to determine whether these Cu-induced changes in CTR2 also affect DDP accumulation and cytotoxicity. Whole cell Pt levels were measured by ICP-MS, and the extent of cell killing by SRB assay, after a 15 min exposure to DDP that was preceded by treatment with either Cu or BCS. Cu treatment did not significantly alter

Pt uptake or enhance cytotoxicity; however, treatment with BCS increased whole cell

Pt accumulation 2.2-foldand increased cytotoxicity 2-fold. These results indicate that

DDP accumulation is increased by depletion of CTR2 irrespective of how this decrease is attained. These data offer additional evidence that reduction of CTR2 hypersensitizes cells to DDP, as was previously observed in cells in which CTR2 was constitutively knocked down with a shRNAi vector. Furthermore, they establish that

BCS can be utilized as a tool to reduce CTR2 levels in future studies and raise the possibility that Cu chelators that can be given to patients might be used to enhance tumor sensitivity to the Pt-containing drugs.

Atox1 is the Cu chaperone that transfers Cu to the efflux transporters ATP7A and ATP7B. Atox1 is also a Cu-dependent transcription factor capable of activating the expression of cyclin D1 and SOD1 (Itoh, Kim et al. 2008; Muller and Klomp

2008). Analysis of the CTR2 promoter region with the Genomatix software suggested that it contains several potential Atox1 binding motifs and thus raised the possibility that the regulation of CTR2 mRNA expression by Cu, DDP and Cu starvation could be due to different transcriptional activation by Atox1. Under steady-state conditions

74 the Atox1 -/- cells were found to express 3.5-fold more CTR2 mRNA and 3.1-fold more protein than Atox1 +/+ cells. This result suggests that Atox1 may play a role in inhibiting CTR2 expression under basal conditions at either the transcriptional or post- translational level. It is possible that the observed increase in CTR2 in Atox -/- cells is due to changes in the copper levels rather than direct Atox1 transcriptional regulation.

Consistent with this concept, Atox1 -/- cells have been reported to have higher endogenous Cu levels than Atox +/+ cells. Furthermore, the magnitude of the changes in

CTR2 mRNA in response to excess Cu, DDP or starvation were similar in the

Atox1 +/+ and Atox1 -/- cells, indicating that Atox1 was not required for these perturbations and therefore is unlikely to directly regulate CTR2 transcription. Atox1 has been reported to directly regulate CTR1 proteosomal degradation stimulated by

DDP and Cu (Safaei, Maktabi et al. 2009).While not essential for the changes in mRNA level, Atox1 was essential to the ability of Cu starvation to down-regulate

CTR2 protein. When cells are starved for Cu, CTR2 is quickly degraded by the proteosome. It is likely therefore that the interaction of Atox1 with Cu is necessary for the stabilization of CTR2, and the apo-form of Atox1 is necessary for CTR2 degradation. Figure 4-8 presents a model of how CTR2 might be regulated by Atox1.

Atox 1 appears to be necessary for the Cu-dependent regulation of CTR2 at the post- transcriptional level either through direct interaction with CTR2 or a downstream effect on a pathway that controls CTR2 stability. We suggest that Atox1 in the apo- conformation, is necessary for the degradation of CTR2, as evidenced by the fact that both Atox1 and copper starvation are necessary for post-transcriptional regulation of

CTR2. Presence of the unbound form of Atox1 is needed for CTR2 degradation, since both Cu and loss of Atox1 stabilize CTR2 protein.

In summary, CTR2 has a large effect on DDP accumulation and sensitivity in human ovarian carcinoma cells. As such, study of its protein regulation remains tremendously important. CTR2 levels are regulated by the availability of Cu as well as by DDP at the transcriptional and post-transcriptional level. The question of how

Atox1 controls CTR2 expression merits further investigation in an effort to identify strategies by which the effectiveness of DDP can be enhanced.

Materials and Methods

Drugs and reagents .

Platinol AQ was a gift from Bristol-Myers Squibb (Princeton, NJ); it contains

DDP at a concentration of 3.33mM in 0.9% NaCl. CuSO 4 and BCS were purchased from Sigma-Aldrich (St. Louis, MO). The drugs were diluted into OptiMEM Reduced

Serum Media (Gibco, 31985-070) to produce final concentrations. Bradford reagent was purchased from BioRad Laboratories, Inc. (Hercules, CA), sulforhodamine B was obtained from Sigma-Aldrich (St. Louis, MO) and 0.4% sulforhodamine B (w/v) was solubilized in 1% (v/v) acetic acid solution.

Cell types, culture and engineering .

Parental mouse embryonic fibroblasts containing wild-type alleles of Atox1

(Atox1 +/+ ) and an isogenic line in which both copies of Atox1 had been somatically knocked out (Atox1 -/-) were a generous gift from Dr. J.D. Gitlin (Washington

University, St. Louis, MO) (Hamza, Prohaska et al. 2003). Ovarian carcinoma 2008 cells were obtained from Dr. Phillip Disaia (Disaia, Sinkovics et al. 1972).

Cell Survival Assay.

Cell survival following exposure to increasing concentrations of drugs was assayed using the sulforhodamine B assay system (Monks, Scudiero et al. 1991). Five thousand cells were seeded into each well of a 96-well tissue culture plate. Cells were incubated overnight at 37°C, 5% CO 2 and then exposed to varying drug concentrations in 200 l complete medium. Cells were allowed to grow for 5 days, after which the media was removed, the protein precipitated with 50% trichloroacetic acid and stained

77 using 100 l of 0.4% sulforhodamine B in 1% acetic acid at room temperature for 15 min. Following washing, the absorbance of each well at 515 nm was recorded using a

Versamax Tunable Microplate Reader (Molecular Devices, Sunnyvale, CA). All experiments were repeated at least three times using three cultures for each drug concentration.

Western blotting .

Whole-cell lysates were prepared, Western blots were run and analyzed as previously described. The primary antibodies used for Western Blot analysis were: anti-CTR2 (provided by Dr. Jesse Bertinato), anti-CTR2 (Novus Biologicals,

Littleton, CO) and anti-tubulin (Santa Cruz Biotechnology, Inc. Santa Cruz, CA).

qRT-PCR .

CTR2 mRNA levels were measured using a qRT-PCR method of detection of relative amounts of first-strand cDNA. cDNA was generated from mRNA isolated using Trizol (Invitrogen, Carlsbad, CA). Purified mRNA was converted to cDNA using Oligo(dT) 20 priming and the SuperScript III First-Strand Kit (Invitrogen). qRT-

PCR was performed on a Bio-Rad MyIQ qPCR machine (Hercules, CA). The forward and reverse primers used were: mCTR2 forward – tccaggtagtcatcagct; mCTR2 reverse

– tggcagtgctctgtgatgtc; ß-actin forward – aggtgacagattgcttctg; ß-actin reverse – gctgcctcaacacctcaac; Gapdh forward – tcaccaccatggagaaggc; Gapdh reverse – gctaagcagttggtggtgca; mAtox1 forward - ctgggaggagtggagttcaa ; mAtox1 -

78 gccaaggtaggaaacagcct. Reactions were prepared using iQ SYBR Green Supermix

(Bio-Rad), according to the manufacturer’s recommendations. Samples were prepared in quadruplicate with three independent sample sets being analyzed. Analysis was done using the Bio-Rad iQ5 system software .

Measurement of drug accumulation into whole cells .

Cells were grown to 90 percent confluence in T-150 tissue culture flasks. Cells were then harvested using trypsin; 7.5 x 105 cells were placed into each well of 6-well tissue culture plates and allowed to grow overnight in 2.5 ml of media at 37°C in 5 percent CO 2. The next day, medium was removed by aspiration and the cells either pre-treated with CuSO 4, BCS or untreated control media for 1h. The media was then removed and the cells were exposed to 500 l of Cisplatin-containing OptiMEM medium (Invitrogen) at 37°C for either 0 or 60 min, after which the drug-containing medium was removed, the plates were washed three times with ice-cold PBS and were then placed on ice. In the case of the time zero samples, the drug-containing medium was aspirated within 15 sec of the start of drug exposure. 214 l of concentrated (50-

70%) nitric acid was added to each well and the plate was incubated overnight at room temperature. The following day the acid was moved into Omni-vials (Wheaton,

Millville, NJ) and incubated at room temperature overnight to thoroughly dissolve all cellular debris. The following day, the nitric acid was diluted with 3 ml of buffer

(0.1% Triton X-100, 1.4% nitric acid, 1 ppb In in ddH 2O). Pt concentration was measured using a Perkin-Elmer Element 2 ICP-MS located at the Analytical Facility at

Scripps Institute of Oceanography at UCSD. As a method of normalization, total sulfur was measured using a Perkin-Elmer ICP-OES, also located at SIO at UCSD.

Samples that were previously prepared for the ICP-MS were then introduced into the

ICP-OES where total µg of sulfur was measured. All data presented are the means of at least three independent experiments each performed with six wells per concentration tested.

Measurement of CTR2 Half-life.

2008 cells were pre-incubated with CuSO 4 and BCS for 1 h as previously described and were then exposed to 100 µg/mL cycloheximide for 0, 5, 10, 20, 30 or

45 min. Cells were then washed 3 times with PBS and lysates were harvested for

Western blotting as previously described in Chapter 2 .

Deconvolution Microscopy.

Cells were grown on 8-well microscope chamber slides (Waltham, MA). Upon reaching ~60% confluence, the media was removed from each chamber. The chambers were then treated with either 300 µl DMEM RS media alone (Invitrogen) or DMEM

RS containing either 200µM CuSO 4, 100 µM BCS or 30 µM DDP for 1 h. Following

1h drug exposure, the media was removed and the slide was treated and then washed 3 times with PBS. Cells were fixed with 3.7% formalin in PBS for 30 min followed by three 10 min PBS washes. Cells were then permeabilized with 0.3% Triton X in PBS followed by three 1 min PBS washes. The slides were blocked for 1 h with 5% BSA in

PBS and then treated with 20 µM anti-CTR2 antibody (provided by Dr. Jesse

Bertinato) overnight at 4ºC followed by three 10 min PBS washes. The slides were then exposed to 1:1000 anti-Rabbit Texas Red for 1 h, washed 3 times in PBS and viewed using a Deltavision deconvolution microscope (Applied Percision, Inc.

Issaquah, Washington). Other primary/secondary antibodies used include: anti-

Nuclear Pore Complex (NPC) proteins 414 (Abcam, Cambridge, MA); anti-mouse

FITC secondary antibody (Invitrogen).

Proteosomal Degradation Assay.

2008 cells were plated on chamber slides and treated with 50 nM bortezomib

(BTZ) for 4 h or left as untreated controls. During the last hour of BTZ exposure, selected wells were treated with 100 µM BCS. The cells were then fixed and prepared for microscopy as previously described.

Statistical Analysis.

All data were derived from at least 3 independent experiments, and presented with the standard error from the mean (SEM). Statistical comparisons were performed using a two-tailed t-test.

Acknowledgements

The contents of this chapter have been submitted for publication in full. Brian

G. Blair was the primary researcher and author for this chapter. Stephen B. Howell supervised and directed the research in this chapter. Christopher A. Larson and

Chapter 5

CTR2 is Partially Localized in the Nucleus

Introduction

Previous studies of yeast cells have shown that yCtr2 is associated with the vacuole and that the C-terminal tail of yCtr2 is oriented towards the cytosol (Rees,

Lee et al. 2004). Tagged mammalian CTR2 associates with vesicular compartments as well as the plasma membrane when expressed in mammalian cells (Bertinato, Swist et al. 2007; van den Berghe, Folmer et al. 2007). The goal of the studies described in this chapter was to determine if CTR2 is found elsewhere in the cell in its native state.

The results of the studies conducted in the previous chapters revealed that CTR2 is found not only on intracellular membranes but also in the nucleus and that exposure to either Cu or DDP increases the level of CTR2 in the nucleus as well as the cytoplasm.

82 83

Results

CTR2 is Associated with the Nucleus

Initial deconvolution microscopy images of human ovarian carcinoma 2008 cells stained for CTR2 suggested that it was found throughout the cytoplasm of cells but also in the DAPI-stained areas of cells, suggesting that CTR2 may also be located in the nucleus. To test directly if CTR2 is found in the nucleus, 2008 cells were co- stained with antibodies to CTR2 and the nuclear pore complex (NPC). As shown in

Figure 5-1, staining for NPC alone (green) produced a distinct outline of the nuclear membrane surrounding the DAPI stained nucleus (blue) consistent with the known location of the NPC. When cells are additionally stained with antibody to CTR2 (red) and examined using a focal plane that cut through the middle of the nucleus, large amounts of CTR2 were found in a punctuate pattern within the perimeter defined by the NPC and thus in the interior of the nucleus.

CTR2 is Partially Localized in the Nuclei of Tumor Cells in vivo .

To determine whether CTR2 is found with the nuclei of cells grown in vivo , subcutaneously inoculated 2008 tumor xenografts were harvested from mice and embedded in O.C.T. compound. Sections cut from these tumor samples were stained with an anti-CTR2 antibody. As shown in Figure 5-2, the majority of CTR2 (brown) was found within the nuclei of the tumor cells.

Cu Starvation Reduces Nuclear CTR2 Levels.

As shown in Figure 5-1, CTR2 was found abundantly in the nucleus as well as the cytoplasm of 2008 cells. To determine whether nuclear as well as cytoplasmic levels of CTR2 were modulated by Cu and DDP, 2008 cells were pre-treated with 200

µM CuSO 4, 30 µM DDP, 100 µM BCS or drug-free control media for 1 h and stained with antibodies against CTR2 and the nuclear pore complex (NPC) proteins. Figure 5-

3A shows that CTR2 (red) was partially localized within the nucleus, appearing as discrete foci randomly distributed throughout the nucleus and not clearly associated with nucleoli. There appeared to be no co-localization of CTR2 with the nuclear envelope (green). Three dimensional images compiled from z-stack sections confirmed that CTR2 is present within the nucleus of these cells (Figure 5-3B).

Exposure of 2008 cells to 30 µM DDP for 1 h produced a small increase in nuclear CTR2 level (Figure 5-3). Exposure to Cu produced a more pronounced increase without changing the distribution of CTR2 within the nucleus itself, and causing only a small change in the ratio of cytoplasmic to nuclear staining. Depletion

87 of Cu with BCS caused almost complete disappearance of CTR2 from the nucleus and other cellular compartments. To further assess the effect of Cu and DDP, nuclei were isolated and subjected to Western blot analysis. As shown in Figure 5-4, CTR2 was readily detected in the nuclear fraction of 2008 cells. Incubation of the cells with 30

µM DDP or 200 µM CuSO 4 for 1 h increased the amount of nuclear CTR2 by 25%

(p<0.05) and 40% (p< 0.03), respectively. Consistent with its effect on the level of

CTR2 in the whole cell, incubation with 100 µM BCS completely eliminated CTR2 from the nuclear fraction. Cytosolic CTR2 mimicked the whole cell response: DDP and Cu increased cytosolic CTR2 by 1.3-fold and 2.4-fold, respectively (Figure 5-4).

Interestingly, as assessed by both immunofluorescent staining and Western blot analysis, the effect of DDP and Cu on nuclear CTR2 was muted in comparison to their effect on cytosolic CTR2.

Discussion

When co-stained with an antibody to a nuclear envelope protein, and particularly when the images were reconstructed in three dimensions, it was clear that a fraction of CTR2 resides inside the nucleus. This finding was verified by Western blot analysis of isolated nuclei. The microscopic observations were confirmed by use of two different anti-CTR2 antibodies. Additionally, tumor sections stained with anti-

CTR2 antibody also confirmed the presence of CTR2 in the nucleus in vivo . These observations provide strong evidence that a fraction of CTR2 is present in the interior of the nucleus.

Two additional points can be made regarding the nature of nuclear CTR2.

First, CTR2 did not co-localize with the NPC, as evidenced by the lack of yellow staining in cells by antibodies directed at both CTR2 and NPC, indicating that CTR2 does not associate with the nuclear envelope. Thus, CTR2 must reside inside the nucleus. Although this is an unexpected finding, since CTR2 is a transmembrane protein, there are other examples of transmembrane proteins that function at the plasma membrane, including EGFR (Liao and Carpenter 2007) and CD44 (Lee, Wang et al. 2009), that are also trafficked to the nucleus where they participate in transcriptional regulation. CTR2 may traffic to the nucleus in a similar manner and participate in transcriptional activation of Cu-responsive genes either directly or by regulating nuclear Cu. The second major observation that can be made from these observations is that the nuclear CTR2 was found in discrete but randomly distributed sites that yield a punctuate pattern similar to that of p-bodies. The function of CTR2 in

89 the nucleus, the mechanism by which it enters the nucleus and the cause of the formation of CTR2 aggregates is the subject of continuing research.

Immunocytochemical and deconvolution microscopic examination confirmed the results of the qRT-PCR and western blot analyses with respect to the ability Cu and DDP to increase, and BCS to deplete, CTR2 in the nucleus of 2008 cells. Similar effects were observed in mouse embryo fibroblasts. Neither Cu nor DDP produced a major change in the subcellular localization of CTR2. Akin to its effect on the level of

CTR2 in the cytosolic and membrane-bound fractions, DDP and Cu treatment increased the amount of CTR2 within the nucleus; however, this increase appeared to be less dramatic than in the rest of the cell.

The results of the experiments described in this chapter further demonstrate that CTR2 is associated with the nuclei of cells. All previous work on this protein has missed this crucial observation, only observing that CTR2 in non-nuclear cellular compartments . However, these previous studies were conducted in either yeast or in cells with over-expressed protein. Additionally, the only mammalian localization studies used a CTR2 that was tagged with GFP or vsvG on the C-terminus (Bertinato,

Swist et al. 2007; van den Berghe, Folmer et al. 2007). Mutational studies of CTR1 have shown that the C-terminal tail is essential for the correct multimerization and localization of CTR1 (Lee, Howell et al. 2007; De Feo, Aller et al. 2009). Due to their structural similarity, it is likely that the addition of C-terminal tags to CTR2 may inhibit or mask nuclear localization of CTR2.

Materials and Methods

Deconvolution Microscopy .

Cells were grown on 8-well microscope chamber slides (Waltham, MA). Upon reaching ~60-70% confluence, the media was removed from each chamber. The chambers were then treated with either 300 µl DMEM RS media alone (Invitrogen,

Carlsbad, CA) or DMEM RS containing either 200µM CuSO 4, 100 µM BCS or 30

µM DDP for 1 h. Following 1h drug exposure, the media was removed and the slide was treated then washed with PBS in triplicate. Cells were fixed with 3.7% formalin in

PBS for 30 min followed by three 10 min PBS washes. Cells were then permeabilized with 0.3% Triton X in PBS followed by three 1 min PBS washes. The slides were blocked for 1 h with 5% BSA in PBS and then treated with 20 µM anti-CTR2 antibody (provided by Dr. Jesse Bertinato) overnight at 4ºC followed by three 10 min

PBS washes. The slides were exposed to 1:1000 anti-Rabbit Texas Red for 1 h, washed 3 times in PBS and viewed using a Deltavision deconvolution microscope

(Applied Percision, Inc. Issaquah, Washington). Other primary/secondary antibodies used include: anti-Nuclear Pore Complex (NPC) proteins 414 (Abcam, Cambridge,

MA); anti-mouse FITC secondary antibody (Invitrogen).

Nuclear Fractionation .

Cells were grown to ~90% confluence, harvested and nuclei were extracted using NE-PER Nuclear and Cytoplasmic Extraction Reagents kit (Pierce) following the manufacturer’s protocols.

Western Blotting .

Whole-cell lysates were dissolved in lysis buffer (150 mM NaCl, 5 mM EDTA,

1% Triton X-100, and 10 mM Tris, pH 7.4) and were subjected to electrophoresis on 4 to 15% gels using ~30 µg of protein per lane. Protein levels were first determined by

Bradford assay (Bio-Rad, Richmond, CA). A Bio-Rad trans-blot system was used to transfer the proteins to Immobilin-P membranes (Millipore, Billerica, MA). Blots were incubated overnight at 4°C in 4% nonfat milk in Tris-buffered saline (150 mM NaCl,

300 mM KCl, 10 mM Tris, pH 7.4, 0.01% Tween 20). Blots were incubated with primary antibody for 1 h at room temperature. A horseradish peroxidase-conjugated secondary antibody (GE Healthcare, Chalfont St. Giles, Buckinghamshire, UK) was dissolved in 4% milk in the Tris-buffered saline buffer and incubated with the blot for

1 h at room temperature. After four 5-min washes, blots were exposed to the PIERCE

ECL reagent (Thermo Scientific, Wilmington, DE) and detected on X-ray films

(HyBlot CL; Denville Scientific, Inc. Metuchen, NJ). The primary antibodies used for

Western Blot analysis were: anti-CTR2 (provided by Dr. Jesse Bertinato), anti-CTR2

(Novus Biologicals, Littleton, CO) anti-tubulin (Santa Cruz Biotechnology, Inc. Santa

Cruz, CA) and anti-laminin B1 (Santa Cruz Biotechnology).

Acknowledgements

The majority of the contents of this chapter have been submitted for publication. Brian G. Blair was the primary researcher and author for this chapter.

Stephen B. Howell supervised and directed the research in this chapter. Paolo B.

Abada independently confirmed the result of the Western blot analysis presented in this chapter. Christopher A. Larson and Preston L. Adams assisted in preparation of microscope slides, harvesting of mouse tumors and provided feedback. Roohangiz

Safaei provided helpful discussion. Pathologic assessment was provided by Laarni

Gapuz and Dr. Nissi Varki of the UCSD Pathology Core. Kersi Pestonjamasp provided training and valuable input on microspopic technique. The author would like to thank Dr. Jesse Bertinato for the providing the anti-CTR2 antibody and Dr. Martin

Hetzer for the NPC antibody.

Chapter 6

CTR2 Limits DDP Accumulation Through the

Inhibition of Endocytosis

Introduction

The previous chapters presented data that establish CTR2 as a regulator of

DDP accumulation and sensitivity both in vitro and in vivo . Loss of CTR2 protein, either through shRNA knockdown or through post transcriptional degradation caused by Cu starvation, increases DDP sensitivity and uptake. This chapter explores several hypotheses as to the mechanism by which CTR2 regulates DDP uptake.

One mechanism by which CTR2 may enhance DDP uptake and cytotoxicity is by intracellular sequestration. In yeast, CTR2 is expressed in the vacuolar membrane

(Bellemare, Shaner et al. 2002; Rees, Lee et al. 2004), and in mammalian cells it is found in the mammalian equivalent of the yeast vacuole, which consists of the late endosomal and lysosomal compartments (Rees, Lee et al. 2004; Bertinato, Swist et al.

2007; van den Berghe, Folmer et al. 2007). Available evidence suggests that CTR2 functions primarily to efflux Cu from these structures under conditions of low environmental Cu (van den Berghe, Folmer et al. 2007). If CTR2 mobilizes DDP in a

94 similar manner, then the loss of CTR2 would lead to an increased amount of Pt trapped in the vesicles and an increase in total cellular Pt. A second mechanism by which CTR2 may regulate DDP uptake is through an effect on efflux. In addition to changes in initial uptake, increased drug accumulation may be caused by a loss of export. ATP7A and ATP7B are known to regulate Pt drug accumulation through effects on export (Samimi, Katano et al. 2003). If CTR2 also functions to export DDP then its loss would be expected to enhance total cellular DDP uptake.

A third mechanism may involve regulation of endocytosis by CTR2. Previous studies have reported a link between endocytosis and DDP sensitivity (Safaei, Katano et al.

2004; Shen, Liang et al. 2004; Shen, Su et al. 2004).

The experiments described in this chapter were designed explore the role of

CTR2 in endocytotic uptake of DDP and to provide new information on the mechanism by which CTR2 regulates DDP uptake and cytotoxicity. The findings lead to the conclusion that CTR2 does not regulate vesicular storage of DDP or the export of drug from cells. Instead, CTR2 regulates the rate of endocytosis. Preliminary data suggest that CTR2 up-regulates macropinocytosis through the activation of the endocytotic factors Rac1 and cdc42.

Results

Increased Pt accumulation in CTR2 knockdown cells is not due to enhanced vesicular accumulation.

In order to determine whether the changes in whole cell accumulation of DDP accompanying knockdown of CTR2 could be accounted for by enhanced accumulation in intracellular vesicles, all four cell lines (CTR1 +/+ , CTR1 +/+ CTR2 kd ,

CTR1 -/- and CTR1 -/- CTR2 kd ) were exposed to 30 µM DDP for 1 h. Intracellular vesicles were then isolated by sucrose gradient enrichment and their Pt content measured by ICP-MS. There was no difference in the concentration of Pt in the vesicles of any of the four cell types; the mean concentration ranged from 0.48 to 0.51 ng Pt/µg sulfur. However, as shown in Figure 6-1, loss of CTR1 and CTR2 produced quite large changes in the fraction of total intracellular Pt associated with the vesicles.

Loss of CTR1 decreased whole cell DDP uptake but nearly doubled the fraction of

DDP in the vesicular fraction (16.9 ± 0.9 to 31.7 ± 1.9% (SEM)) . In contrast, knockdown of CTR2 decreased the fraction in the vesicles in both the CTR1 +/+ and

CTR1 -/- cells. Knockdown of CTR2 in the CTR1 +/+ cells reduced the percentage by half to 8.3 ± 0.6%; in the CTR1 -/- cells it reduced the fraction by two thirds to 9.8 ±

1.1%. These results indicate that, while both CTR1 and CTR2 modulated whole cell

DDP uptake, neither had a significant effect on the absolute amount of DDP resident in the vesicular fraction.

Increased Pt accumulation in CTR2 knockdown cells is not due to a change of drug export.

To determine whether loss of drug export caused the increased DDP accumulation observed in CTR2 knockdown cells, both the initial and subsequent phases of drug export were examined. Cells were pretreated with 30 µM DDP for 1 h,

97 followed by replacement of the drug-containing media with media not containing drug. Total cellular Pt levels were measured by ICP-MS following 0, 0.5, 1, 2, 8, 30 and 240 min in drug-free medium. Figure 6-2 shows the percent of Pt remaining in each type of cell as a function of efflux time. While loss of CTR1 enhanced efflux, no change was detectable in either the early or late phases of DDP efflux when CTR2 was knocked down in either the CTR1 +/+ or CTR1 -/- cells.

Loss of CTR2 increases endocytosis.

To determine whether CTR1 or CTR2 effects endocytotic rate, the accumulation of Texas red-labeled dextran was measured in CTR1 +/+ , CTR1 +/+

CTR2 kd , CTR1 -/- and CTR1 -/- CTR2 kd cells (Figure 6-3A). The accumulation of

98 dextran is a well-validated measure of the rate of macropinocytosis (Chauhan, Liang et al. 2003; Holzer and Howell 2006). Loss of CTR1 did not significantly change the uptake of dextran: Following 30 min exposure to Texas red dextran, there was no difference in its accumulation in CTR1 -/- and CTR1 +/+ cells (p=0.5). However, loss of

CTR2 significantly increased the endocytosis of dextran. CTR1 +/+ CTR2 kd cells took up 2.0-fold more dextran over a 30 min period than CTR1 +/+ cells (p=0.006). This increase in endocytosis in CTR2 kd cells occurred independently of CTR1 status.

CTR1 -/- CTR2 kd cells took up 1.9-fold more dextran over a 30 min period than CTR1 -/- cells (p=0.04) and 1.6-fold more dextran than CTR1 +/+ cells (p=0.09).

CTR2 degradation due to copper starvation induces endocytosis.

Cu starvation by BCS results in a rapid and near total degradation of CTR2. To strengthen the link between CTR2 and endocytosis, cells were exposed to BCS for 1 h to reduce CTR2 levels by ~90-95%. As shown in Figure 6-3, BCS induced loss of

CTR2 increased dextran uptake in both CTR1 +/+ and CTR1 -/- cells: 1.9-fold increase in CTR1 +/+ cells (p = 0.001) and a 2.0-fold increase in the CTR1 -/- cells (p = 0.0006).

100

Differential dextran uptake is not due to changes in exocytotic rate .

Exocytosis of Texas-red dextran was measured in CTR1 +/+ , CTR1 +/+ CTR2 kd ,

CTR1 -/- and CTR1 -/- CTR2 kd cells. As shown in Figure 6-4, the majority (~55%) of

Texas-red dextran exits the cell in the first 2 minutes and there is no significant difference in the rate of Texas-red dextran exocytosis in CTR1 +/+ , CTR1 +/+ CTR2 kd ,

CTR1 -/- and CTR1 -/- CTR2 kd cells either during the initial or late phase of efflux.

Inhibition of Macropinocytosis blocks CTR2 dependent DDP accumulation.

As described in previous chapters, knockdown of CTR2 mediated by either shRNAi or Cu starvation substantially increases DDP uptake in vitro as well as in vivo . To determine whether this increase is due to enhanced endocytosis, DDP accumulation was measured in CTR1 +/+ , CTR1 +/+ CTR2 kd , CTR1 -/- and CTR1 -/-

CTR2 kd cells with and without inhibition of macropinocytosis by either amiloride or

101 wortmannin. As expected, knockdown of CTR2 in untreated cells caused a ~ 2.5-3.5- fold increase in DDP accumulation at 15 min and 1 h of DDP exposure (Figure 6-5).

Treatment with amiloride did not significantly change DDP uptake in either the

CTR1 +/+ or CTR1 -/- cells but completely blocked the increased DDP uptake found in the CTR2 kd cells (Figure 6-5A). When compared to untreated cells, amiloride pre- treatment produced a 71.0% (p = 0.001) and 65.3% (p = 0.002) decrease in DDP accumulation in CTR1 +/+ CTR2 kd and CTR1 -/- CTR2 kd cells, respectively.

Wortmannin has been used as a chemical inhibitor of macropinocytosis (Li,

D'Souza-Schorey et al. 1995; Dharmawardhane, Schurmann et al. 2000). CTR1 +/+ cells pretreated with wortmannin accumulated 1.8-fold (p = 0.02) more Pt over the course of an hour than untreated CTR1 +/+ cells (Figure 6-5B). Wortmannin had no effect on the accumulation of DDP in CTR1 -/- cells (Figure 6-5B). As observed with amiloride, wortmannin completely blocked the increased accumulation of DDP seen in untreated CTR1 +/+ CTR2 kd and CTR1 -/- CTR2 kd cells, producing 65.7% (p = 0.001) and 79.4% (p = 0.0001) decreases in DDP accumulation in CTR1 +/+ CTR2 kd and

CTR1 -/- CTR2 kd cells respectively (Figure 6-5B).

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Induction of macropinocytosis by PDGF increases DDP accumulation.

Inhibition of macropinocytosis blocked the increased DDP accumulation produced by knocking down CTR1. To further discern if the enhanced DDP uptake in

CTR2 kd cells was due to enhanced macropinocytosis, PDGF was used to up-regulate macropinocytosis in wild type cells. As observed previously, PDGF treatment leads to increased dextran uptake (Dharmawardhane, Schurmann et al. 2000). PDGF pre- treatment leads to a 1.8 ± 0.2 (p < 0.02) increase in Texas-red labeled dextran uptake

103 over 30 min. As demonstrated in Figure 6-6B, CTR1 +/+ cells pretreated with PDGF accumulated 2.1-fold (p < 0.001) more Pt following 1 h exposure to 30 µM DDP, consistent with the conclusion that DDP can enter cells through the endocytotic pathway.

Knockdown of CTR2 activates Rac1 and cdc42.

The active form of the Rho GTPases Rac1 and cdc42 are necessary for macropinocytosis. To determine whether CTR2 can regulate the activation of these proteins, the levels of active Rac1 and cdc42 were quantified by Western blot analysis using antibodies to the phosphorylated forms of these proteins in the CTR1 +/+ ,

CTR1 +/+ CTR2 kd , CTR1 -/- and CTR1 -/- CTR2 kd cells. Under basal conditions, neither the CTR1 +/+ nor CTR1 -/- cells contained detectable active Rac1 or cdc42. However,

104 knockdown of CTR2 resulted in a very strong activation of both Rac1 and cdc42 regardless of CTR1 status (Figure 6-7A). The total amount of Rac1 and cdc42 was similar in all four cell lines, suggesting that loss of CTR2 constitutively activates these

GTPases without changing their level.

Loss of CTR2 does not affect Akt activation.

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PI3K activation has been shown to activate Rac1 and cdc42. To determine whether the loss of CTR2 activates Rac1 and cdc42 via the PI3K pathway, the extent of Akt phosphorylation was measured in the CTR1 +/+ , CTR1 +/+ CTR2 kd , CTR1 -/- and

CTR1 -/- CTR2 kd cells (Figure 6-7B) since Akt becomes phosphorylated upon PI3K activation. The western blot analysis using an antibody specific for phosphorylated

S473 Akt demonstrated that the loss of either CTR1 or CTR2 did not change Akt phosphorylation (Figure 6-7B), thus suggesting that loss of CTR2 activates Rac1 and cdc42 downstream of PI3K.

Discussion

The results reported here provide only an outline of how CTR1 and CTR2 modulate the accumulation of DDP and CBDCA. Since DDP appears to trigger rapid macropinocytosis of CTR1 (Holzer and Howell 2006), one hypothesis is that DDP and

CBDCA can bind to the extracellular domain of CTR1 and enter via an endocytototic process that delivers them to intracellular vesicles. It is possible that CTR2 functions sequentially with CTR1 in the drug influx process by transporting DDP out of vesicles and into the cytoplasm. However, there are several problems with this model. First, one would expect knockdown of CTR2 to have little effect when CTR1 was not expressed and a limited amount of DDP was entering the cell by endocytosis; however, what was observed was that knockdown of CTR2 increased DDP uptake in

106 both CTR1 +/+ and CTR1 -/- cells. Second, knockdown of CTR2 had no effect on the absolute amount of DDP that accumulated in intracellular vesicles; changes in the fraction of DDP in the vesicular compartment were due to changes in uptake into other parts of the cell. Finally, how CTR2 might export DDP from intracellular vesicles is uncertain. The finding that knockdown of CTR2 enhanced the accumulation of DDP within 5 min suggests that CTR2 restrains initial influx, perhaps through regulation of the uptake at the plasma membrane or of trafficking to intracellular sites. The fact that CTR2 had no effect on DDP efflux implies that the major role of CTR2 is to mediate DDP influx rather than to restrain efflux.

DDP accumulation is partially controlled by membrane transporters; however, a portion of DDP uptake is not accounted for by these transporters (Troger, Fischel et al. 1992; Holzer, Manorek et al. 2006). The latter studies showed that DDP uptake was not saturable, suggesting the importance of a non-specific uptake mechanism such as endocytosis. There is strong indirect evidence that endocytotic processes account for some portion of DDP accumulation. The GTPases Rac1, RhoA and cdc42, which control clathrin- independent endocytosis, are down-regulated in several DDP- resistant cell lines (Shen, Su et al. 2004) as are cytoskeletal proteins necessary for endocytosis (Shen, Liang et al. 2004). Furthermore, CTR1 has been closely linked to macropinocytosis. As stated above, CTR1 degradation due to Cu and DDP exposure requires an endocytotic process (Holzer and Howell 2006).

The data presented in this chapter strengthen the link between endocytotic processes and DDP accumulation. The loss of CTR2 up-regulates the rate of DDP

107 accumulation and, in parallel, increases the rate of endocytosis. Uptake of dextran, a marker of endocytotic rate (Monti, Mercalli et al. 2003), was more extensive by cells in which expression of CTR2 was reduced by shRNAi-mediated knockdown or degradation by BCS. The observation that inhibition of endocytosis effectively blocked the increased DDP accumulation seen in cells deficient in CTR2 expression further strengthens the linkage between the rate of endocytosis and DDP uptake. This suggests that CTR2 acts to limit DDP accumulation by limiting endocytosis. The ability of wortmannin, and to a lesser extent, amiloride to increase DDP uptake in

CTR1 +/+ cells is most likely due to the inhibition of CTR1 down-regulation off the cell surface.

The mechanism by which loss of CTR2 up-regulates endocytosis remains unknown but may involve the activation of the Rho GTPases Rac1 and cdc42.

Clathrin-independent endocytosis, and specifically macropinocytosis, is a highly controlled and regulated process (reviewed in (Conner and Schmid 2003)).

Macropinocytosis is accompanied by membrane ruffling and the formation of membrane protrusions. This process is induced by PDGF signaling and is mediated by

Rac1 and cdc42. Activation of these proteins allows formation of the membrane protrusions necessary for macropinocytosis. The observations reported in this chapter demonstrate that PDGF-induced macropinocytosis significantly increases DDP uptake in manner similar to the knockdown of CTR2. This suggests that CTR2 protein may be downstream of the PDGF signaling cascade and may play an important role in the induction of macropinocytosis. Additionally, loss of CTR2 constitutively activates

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Rac1 and cdc42. This suggests that CTR2 limits the rate of endocytosis through the regulation of these GTPases. The lack of Akt activation in CTR2 kd cells suggests that

CTR2 controls the activation of the Rho family of GTPases without activating the entire PI3K cascade.

This chapter explored several hypotheses as to why cells with low CTR2 protein levels more readily accumulate DDP. The data support the conclusion that the increase in DDP accumulation is not due to a change in DDP export or accumulation of DDP in vesicular stores. Induction of endocytosis can clearly enhance DDP accumulation and CTR2 kd cells have an increased endocytotic rate whose inhibition blocks CTR2 mediated DDP uptake. Therefore it is likely that CTR2 can limit DDP accumulation through regulation of endocytotic signaling proteins.

Materials and Methods

Drugs and reagents.

Platinol AQ was a gift from Bristol-Myers Squibb (Princeton, NJ); it contains

DDP at a concentration of 3.33mM in 0.9% NaCl. CuSO 4 and BCS were purchased from Sigma-Aldrich (St. Louis, MO). The drugs were diluted into OptiMEM Reduced

Serum Media (Gibco, 31985-070) to produce final concentrations. Transferrin-Alexa

Fluor 546 and 70-kDa dextran-Texas red were purchased from Invitrogen (Carlsbad,

CA). Amiloride and wortmannin were purchased from Tocris Bilogicals (Ellisville,

Mo).Bradford reagent was purchased from BioRad Laboratories, Inc. (Hercules, CA).

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Sulforhodamine B was obtained from Sigma-Aldrich and 0.4% sulforhodamine B

(w/v) was solubilized in 1% (v/v) acetic acid solution.

Cell types, culture and engineering .

Parental mouse embryonic fibroblasts containing wild-type alleles of CTR1

(CTR1 +/+ ) and an isogenic line in which both copies of CTR1 had been somatically knocked out (CTR1 -/-) were a generous gift from Dr. D. Theile. CTR2 knockdown cells derived from these CTR1 +/+ and CTR1 -/- cells were generated as previously described (Blair, Larson et al. 2009). Ovarian carcinoma 2008 cells were obtained from Dr. Phillip Disaia (Disaia, Sinkovics et al. 1972).

Western blotting .

Whole-cell lysates and Western blots were prepared as previously described.

The primary antibodies used for Western Blot analysis were: anti-CTR2 (provided by

Dr Jesse Bertinato), anti-CTR2 (Novus Biologicals, Littleton, CO) anti-tubulin (Santa

Cruz Biotechnology, Inc. Santa Cruz, CA), anti-Rac1, anti-cdc42, anti-Akt, and anti pAkt S473 (Cell Signaling Technology, Inc. Boston, MA).

Measurement of DDP accumulation in vesicles.

For measurement of accumulation into vesicles, drug-exposed cells were harvested using trypsin and centrifuged at 2000 rpm for 10 min. Media was removed and cell pellets were combined. Vesicles isolated from lysed cells were separated by

110 sucrose gradient subcellular fractionation as described by Tjelle et al. (Tjelle, Brech et al. 1996). For measurement of Pt in DNA, cells were lysed and DNA harvested using

DNAzol (Invitrogen) according to the manufacturer’s protocol. As a method of normalization, DNA was measured prior to addition of nitric acid using a Nanodrop

1000 spectrophotometer (Thermo Scientific, Wilmington, DE). The microsome and

DNA samples were digested in nitric acid prior to measurement of Pt by ICP-MS as described above.

Measurement of drug export.

Cells were grown in 6-well plates as described above. One day after seeding, the medium was removed by aspiration and the cells were exposed to 500 l of 30 µM

DDP-containing OptiMEM medium (Invitrogen) at 37°C for 60 min. Cells were immediately rinsed with room temperature PBS, after which the drug-containing medium was replaced with drug free media for 0, 0.5, 1, 2, 8, 30 or 240 min. The plates were washed three times with ice-cold PBS and then placed on ice. Whole cell drug accumulation was determined as described above.

Measurement of Dextran Endocytosis.

CTR1 +/+ , CTR1 +/+ CTR2 kd , CTR1 -/- and CTR1 -/- CTR2 kd cells were plated in

96 well tissue culture dishes (~20-50,000 cells per well, 15 wells per cell type). 24 h later cells were treated with transferrin-Alexa Fluor 546 and 70-kDa dextran-Texas red for 30 min. The wells were immediately washed with ice cold PBS three times and

111 fixed using fixed with 3.7% formalin in PBS for 30 min followed by three 10 min PBS washes. Fluorescence was measured using an Infinite M-200 micro-plate reader

(Tecan US, Inc. San Jose, CA). The cells were then stained using 100 l of 0.4% sulforhodamine B in 1% acetic acid at room temperature for 15 min. Following washing, the absorbance of each well at 515 nm was recorded using a Versamax

Tunable Microplate Reader (Molecular Devices, Sunnyvale, CA) as a control for cell density. All experiments were repeated at least three times using three cultures for each drug concentration.

To access the effect of copper starvation on endocytosis, CTR1 +/+ and CTR1 -/- cells were pretreated with 100µM BCS for 1h or left untreated. Cells were then washed with 37°C PBS three times and Texas-red dextran endocytosis was measured as previously described.

Measurement of Dextran Exocytosis.

CTR1 +/+ , CTR1 +/+ CTR2 kd , CTR1 -/- and CTR1 -/- CTR2 kd cells were plated in

96 well tissue culture dishes (~20-50,000 cells per well, 15 wells per cell type). 24 h later cells were treated with transferrin-Alexa Fluor 546 and 70-kDa dextran-Texas red for 30 min. Cells were immediately washed with 37°C PBS three times and placed in normal media. 0,1,2,4,and 10 min following removal of dextran plates were fixed with

3.7% formalin in PBS for 30 min followed by three 10 min PBS washes. Fluorescence and absorbance were then measured. All conditions were repeated at least three times

112 using three cultures for each drug concentration. Retention of Texas-red dextran was assayed as a marker for rate of dextran exocytosis.

Measurement of drug accumulation into whole cells following PDGF stimulation.

Cells were grown to 90% confluence in T-150 tissue culture flasks. Cells were then harvested using trypsin, and 7.5 x 10 5 cells were placed into each well of 6-well tissue culture plates and allowed to grow overnight in 2.5 ml of reduced serum media at 37°C in 5% CO 2. The next day, medium was removed by aspiration and the cells either pre-treated with PDGF or untreated control media for 4h. The media was then removed and the cells were exposed to 500 l of Cisplatin-containing OptiMEM medium (Invitrogen) at 37°C for either 0 or 60 min. The drug-containing medium was removed, and the plates were washed 3 times with ice-cold PBS and then placed on ice. In the case of the time zero samples, the drug-containing medium was aspirated within 15 sec of the start of drug exposure. 215 l of concentrated (50-70%) nitric acid was added to each well and the plate was incubated overnight at room temperature.

The following day the acid was moved into Omni-vials (Wheaton, Millville, NJ) and incubated at room temperature overnight to thoroughly dissolve all cellular debris. The samples were then diluted with 3 ml of buffer (0.1% Triton X-100, 1.4% nitric acid, 1 ppb In in ddH 2O). Pt concentration was measured using a Perkin-Elmer Element 2

ICP-MS located at the Analytical Facility at Scripps Institute of Oceanography at

UCSD. As a method of normalization, total sulfur was measured using a Perkin-Elmer

ICP-OES also located at SIO at UCSD. Samples that were previously prepared for the

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ICP-MS were then introduced into the ICP-OES where total µg of sulfur was measured. All data presented are the means of at least three independent experiments each performed with six wells per concentration tested.

Inhibition of Endocytosis.

Cells were either pretreated with amiloride for 30 min or left untreated. Cells were then washed with 37°C PBS three times. The media was then removed and the cells were exposed to 500 l of Cisplatin-containing OptiMEM medium (Invitrogen,

Carlsbad, CA) at 37°C for either 0 or 15 minutes, after which the drug-containing medium was removed, the plates were washed the times with ice-cold PBS and were then placed on ice. Accumulation of Pt was measured as described previously.

Similar conditions were used to measure Pt accumulation following wortmannin pretreatment. In this case cells were pretreated with wortmannin for 1 h followed by addition of DDP for an additional hour. Cells were prepared for ICP-MS and ICP-OES as previously described.

Isolation of activated Rac1 and cdc42.

Cells were grown in serum free media for 24 h to ~80% confluence. Cells were washed 3x with PBS and immediately lysed in 1 ml of lysis buffer. Lysate was then incubated with 10 µg of PAK-1 PBD agarose (Millipore, Lake Placid, NY) for 1 h at

4º C. Beads were isolated and washed with lysis buffer 3x. Beads were resuspended in

Laemmli buffer and analyzed by Western blot as described previously.

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Statistical Analysis.

All data shown were derived from at least 3 independent experiments, and presented with the standard error from the mean (SEM). Statistical comparisons were performed using a two-tailed t-test.

Acknowledgements

A portion of the content of Chapter 6 has been published in Clinical Cancer

Research (Blair BG, Larson CA, Safaei R, Howell SB. Copper Transpoter 2 Regulates the Cellular Accumulation and Cytotoxicity of Cisplatin and Carboplatin. Clin Cancer

Safaei provided helpful discussion. Dr. Sandra Schmid provided valuable insight and suggestions as to the understanding of endocytosis. The author would like to thank Dr.

Jesse Bertinato for the providing the anti-CTR2 antibody, Dr. Alexandra Newton for the anti-Akt antibodies and Dr. Minji Jo for the anti-Rac1 and anti-cdc42 antibodies.

Chapter 7

Conclusions and Future Directions

Summary

The overall goal of the studies presented in this dissertation was to determine if the Cu transporter CTR2 plays a role in the cellular pharmacology of DDP, and if so, whether CTR2 influences the sensitivity of cells to this chemotherapeutic agent. The results of the experiments presented here establish that loss of CTR2 protein expression leads to increased DDP accumulation and cytotoxicity both in vitro and in vivo. Once CTR2 was established as a regulator of DDP accumulation and cytotoxicity, the investigations went on to focus on how CTR2 expression and degradation is controlled by DDP and Cu. Cu and DDP exposure were shown to increase CTR2 levels, whereas Cu starvation was found to trigger rapid degradation of this protein. Additionally, CTR2 was found to partially localize in the nucleus. Finally, the mechanism by which decreased CTR2 levels leads to increased accumulation of

DDP was explored. This increase in DDP accumulation is not due to a change in rate of efflux or amount of vesicular storage of DDP. The data presented suggest that loss of CTR2 triggers uptake of DDP through the up-

115 116 regulation of cellular macropinocytosis. This series of investigations has lead to a better understanding of the cellular pharmacology of DDP and the role CTR2 plays in regulation of Pt-based chemotherapeutics.

Effect of decreased CTR2 levels on accumulation and sensitivity of DDP, CBDCA and Cu

The results reported here established that the level of expression of the Cu transporter CTR2 influences the accumulation and cytotoxicity of DDP, CBDCA and

Cu. In these studies an shRNAi was used to knockdown CTR2 expression in cells that had or lacked expression of CTR1. This system was chosen so that the effects of the loss of CTR2 could be studied relative to expression of CTR1. The findings revealed that regulation of DDP uptake by CTR2 appears to be independent of CTR1 expression, as knockdown of CTR2 produced a similar phenotype in both the

CTR1 +/+ and CTR1 -/- cells. The results of these studies offer strong evidence that decreased CTR2 protein levels sensitize cells to the effects of DDP and CBDCA. This sensitization was unexpected as previous studies of the CTR2 protein suggested that over-expression of this protein enhanced Cu uptake (Bertinato, Swist et al. 2008). The previous study found a portion of the over expressed CTR2 to be localized to the plasma membrane, a phenomena not seen in wild-type cells expressing just

117 endogenous CTR1. The re-localization of CTR2 to the plasma membrane may allow it to mimic the function of CTR1 and thereby enhance Cu uptake.

The results reported in this dissertation demonstrate that the cellular accumulation of DDP, CBDCA, and to a lesser extent Cu, is greatly enhanced when the level of CTR2 is decreased and this effect is irrespective of CTR1 status.

Conversely, decreased CTR1 levels diminished uptake of these drugs. Therefore, it appears that CTR2 and CTR1 regulate uptake in opposite fashions. Likewise, the affect of CTR2 and CTR1 expression on Pt drug sensitivity are the inverse of one another. While it is possible that these two proteins interact with one another, the current results show that CTR2 can limit the uptake and delivery of Cu and DDP independent of CTR1.

The Pt-containing drugs are thought to cause cell death by triggering apoptotic signals after DDP cross-links DNA. Decreased CTR2 protein levels enhance the amount of Pt associated with the DNA, thus suggesting that the cause of DDP hypersensitivity in these cells is due to increased DNA damage. This observation is important as increased whole-cell DDP accumulation does not always lead to enhanced killing. In fact, in one of the models that were studied the forced over- expression of CTR1 enhanced DDP uptake but had little effect on DDP cytotoxicity

(Holzer, Samimi et al. 2004). These CTR1 over-expressing cells accumulated 81% more DDP than wild type cells; however, no change was observed in the amount of

Pt-DNA adducts. The results of the studies reported in this dissertation demonstrate

118 that CTR2 knockdown not only enhances DDP accumulation but also increases delivery of DDP to its crucial targets.

In Vivo studies

Decreased CTR2 levels have a very significant effect on tumor response in mouse models. A single dose of DDP halted tumor growth in several of the test animals, and other tumors decreased in size often to the point of no longer being detectable. Responses of this degree suggest high clinical potential. The results discussed here confirm the potential for CTR2 as a target to enhance DDP sensitivity.

It was observed that CTR2 kd tumors accumulate ~9-fold more DDP than do tumors that express CTR2. As stated previously, as little as a 50% change in accumulation can be of clinical significance, suggesting that decreased CTR2 might greatly enhance DDP sensitivity in patients (Andrews and Howell 1990; Muggia and

Los 1993). The change in uptake observed in CTR2 kd tumors is much greater than the change seen in vitro. Furthermore, CTR2 kd tumors grow at a significantly slower rate than do those with CTR2 despite the fact that both cells types grew at similar rates in vitro . This suggests that CTR2 may be important to the tumor micro-environment and has effects on cells in vivo that are not fully reproduced when the cells are grown in vitro. This difference may be the focus of future research.

Several growth factor pathways, such as those triggered by FGF and EGF, are determinants of tumor growth. It is of interest that FGF signaling can be effected by

Cu (Di Serio, Doria et al. 2008). Also, angiogenesis is down-regulated in CTR2 kd

119 cells. Research can be conducted on these CTR2 kd cells to determine its effect on these pathways. For example, activation of ERK (MAPK), activation of Ras, regulation of cell division factors, rate of protein production and release of VEGF can all be measured in these cells in order to obtain a clearer picture of the effect of loss of

CTR2 on tumor growth and the microenvironment.

The role of CTR2 in Cu homeostasis is still unclear. While CTR2 kd tumors do not show a differential amount of steady state Cu, a difference may have been observed by the presence of Cu-rich inflammatory and endothelial cells in the tumor samples. There are several ways in which the question of whether the CTR2 kd tumor cells are physiologically deficient in intracellular Cu. Apo-ceruloplasmin is converted to holo-ceruloplasmin when Cu is pumped into the trans-Golgi by ATP7B (Hellman,

Kono et al. 2002). The two forms of ceruloplasmin migrate at separate rates on SDS-

PAGE gels. The relative levels of each form of ceruloplasmin can be determined, thereby allowing for analysis of the effect of CTR2 kd on ATP7B functionality. ATP7A is necessary for Cu incorporation into tyrosinase (Petris, Strausak et al. 2000).

Activation of tyrosinase can be measured by the colorimetric change in cells caused by the formation of DOPA when L-DOPA is oxidized. L-DOPA can be added to cultured cells as well as tumor slices to access the effect of CTR2 kd on ATP7A activity. In plants and yeast CTR2 silencing mutations impede SOD1 activity (Rees, Lee et al.

2004; Barhoom, Kupiec et al. 2008). The effect of decreased CTR2 on the ability of

CCS to deliver Cu to superoxide dismutase can be assayed in cell lysates using commercially available assays. Likewise the ability of COX17 to deliver Cu to the

120 mitochondria in CTR2 kd cells can be assayed by monitoring the oxidation of ferrocytochrome-c at 550 nm in mitochondria isolated from cultured cells (Johnson and Anderson 2008).

Regulation of CTR2 by Copper and Cisplatin

CTR1 expression and degradation is regulated by Cu and DDP availability

(Labbe, Zhu et al. 1997; Moller, Petersen et al. 2000; Holzer and Howell 2006). Due to their similar structure and ability to interact with similar molecules, CTR2 expression and degradation might also be regulated in a similar manner. The observations made in this dissertation suggest that CTR2 is very strongly regulated by the availability of Cu and DDP but in a manner opposite to that of CTR1. When cells were exposed to these molecules, CTR1 was quickly internalized and degraded by the proteosome. CTR2 reacts in an opposite fashion; its expression and half-life increase following Cu and DDP exposure. This observation is consistent with the finding that

CTR1 and CTR2 have opposite effects on the uptake of Cu and DDP and suggests that there are mechanisms that coordinate the expression of these two proteins to maintain Cu homeostasis.

Cu starvation was found to result in the rapid degradation of CTR2. This sharp decrease greatly enhanced the ability of cells to accumulate Cu and DDP. The degradation of CTR2 under conditions of Cu starvation appears to serve as a mechanism to enhance Cu availability under these harsh conditions. Since BTZ is able to block this Cu-dependent degradation of CTR2, one can infer that CTR2 degradation

121 is controlled by the proteasome. The proteasome is a large protein complex responsible for the breakdown of misfolded and unneeded proteins into reusable amino acids. The degradation process is initiated by ubiquitin ligases that add ubiquitin chains to lysines on the targeted protein (Banerjee and Wade 2002). If CTR2 is degraded by the proteasome under Cu starved conditions, it is likely that CTR2 becomes polyubiquitinated during Cu-induced depletion. CTR2 contains a lysine-rich cluster located in the intracellular loop of the protein (see Figure 1-5). This sequence is likely the site of ubiquitination; mutational analysis can be used to further elucidate the importance of this region for CTR2 degradation and Cu and DDP metabolism. To prove CTR2 is degraded in this fashion, CTR2 can be isolated from cells during the early stages of Cu starvation and subjected to western blot analysis with an anti- ubiquitin antibody (Rotin, Staub et al. 2000).

Regulation of CTR2 by Atox1

Experiments conducted as part of this dissertation demonstrated that Atox1 expression is necessary for the degradation of CTR2 in response to Cu starvation.

How exactly Atox1 regulates this process is still unknown. CTR1 is thought to interact with Atox1 directly and CTR2 may as well. Co-immunoprecipitation of the two proteins would provide evidence of such an interaction; however, the putative transchelation reaction between these proteins might be too transient to permit co-

122 immunoprecipitation. If it is found that they do interact, x-ray crystallography and/or

NMR using the C-terminal domain of CTR2 could be used to further explore the nature of this interaction. It is assumed that it is the holo-form of Atox1 that is necessary for the degradation of CTR2 as increased Cu exposure extends CTR2 half- life. To further verify this assumption, mutational analysis of CTR2 might be conducted. Currently, the Howell laboratory is pursuing the construction of Atox1 mutants lacking a proper CXXC metal binding domain. These mutants cannot bind Cu or DDP. Expression of this mutant form of Atox1 at normal levels would result in a situation where all the Atox1 present would be in the apo form. If CTR2 degradation is dependent on unbound Atox1, these mutants would be expected to have very low levels of CTR2.

Atox1 has also been shown to act as a transcription factor (Itoh, Kim et al.

2008). While the observations made in this dissertation suggest that Atox1 does not directly regulate CTR2 transcription, it is possible that Atox1 affects CTR2 transcription indirectly. Which promoters Atox1 interacts with and controls is the subject of current studies in the field. ChIP analysis of the CTR2 promoter region may elucidate the transcriptional regulators of CTR2, and the possible role of Atox1. Also,

Atox1 has been shown to regulate SOD1 both transcriptionally and post- transcriptionally (Itoh, Ozumi et al. 2009). As mentioned previously, CTR2 mutations can affect SOD1 activity. This suggests that there may be a link between Atox1 regulation of CTR2 and SOD1, and this possibility warrants further exploration.

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Nuclear Localization of CTR2

During the examination of cells stained with an anti-CTR2 antibody by deconvolution microscopy, the novel observation was made that CTR2 partially resides in the nucleus. Since CTR2 is a transmembrane protein this was unexpected and the goal of subsequent experiments was to prove the validity of this observation.

While there remains little doubt that CTR2 is partially localized in the nucleus, this conclusion leads to several more questions. Staining for CTR2 disclosed a punctuate distribution within the nucleus, suggesting that CTR2 is associated with sub-nuclear structures. What is the nature of these structures and what purpose do they serve?

Proteins that show a similar punctuate distribution have been found to function in

DNA repair and transcription. If CTR2 acts as a transcriptional regulator, several techniques may be employed to determine its target genes. CTR2 kd cells can be analyzed by mRNA microarray to elucidate possible targets based on changes in their expression when CTR2 is not present. If CTR2 binds DNA, ChIP-on-chip analysis can determine the CTR2 binding sites (Aparicio, Geisberg et al. 2004). Briefly, cells are fixed with formaldehyde to crosslink DNA-associated proteins to the DNA strands.

The DNA is then sheared and the protein of interest is immunoprecipitated. Finally the associated DNA fragments are analyzed by microarray to determine their identity and sequence.

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Methods are available to explore the nature of the structures that contain

CTR2. One approach would be to immunoprecipitate CTR2 from isolated nuclei following protein-protein crosslinking and then use proteomic analysis of tryptic digests to identify proteins that bind CTR2. However, this method can usually only verify interactions between suspected interaction partners. A yeast two-hybrid screen would be another way of identify interacting proteins (outlined in (Gietz, Triggs-Raine et al. 1997)).

The observation that CTR2 is in the nucleus raises the question of how it gets there. There are no obvious nuclear localization signals in the CTR2 protein. However,

CTR2 mutational analysis might lend new insight as to what controls the localization of CTR2. The C-terminal tail is required for CTR1 to properly multimerize and localize to the plasma membrane (Eisses and Kaplan 2002; Lee, Howell et al. 2007).

Also, C-terminally tagged CTR2 does not localize to the nucleus (Bertinato, Swist et al. 2007; van den Berghe, Folmer et al. 2007). Therefore, it is reasonable to suspect that the C-terminus of CTR2 is necessary for nuclear localization and that deletion of this domain prevents its trafficking to this site. CTR2 has been reported to sometimes appear on plasma membrane (Bertinato, Swist et al. 2007). To explore whether CTR2 is trafficked from the plasma membrane to the nucleus, biotin technology can be used.

Cell surface proteins can be tagged with biotin, washed then incubated for 24-48 h.

Nuclei from these cells can be extracted and assessed for biotinylated CTR2 that has translocated from the cell surface.

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CTR2 as a regulator of Endocytosis

CTR1 is most likely a direct transporter of DDP across lipid membranes either at the cell surface or in intracellular compartments. The mechanism by which CTR2 limits DDP accumulation is still unclear. The observations made in this dissertation demonstrate that the increased DDP uptake in CTR2 kd cells is not due to a change in drug efflux or microsomal storage of DDP. The experiments shown here provide evidence that loss of CTR2 up-regulates cellular endocytosis and that inhibition of endocytosis blocks CTR2- mediated enhancement in DDP accumulation. Clathrin- independent endocytosis is initiated by PDGF stimulation, and activation of this pathway was found to increase whole cell DDP accumulation. Taken together, these data suggest that the increased uptake of DDP in CTR2 kd cells is due to the observed up-regulation of endocytosis in these cells.

Endocytosis, and specifically macropinocytosis, is a tightly controlled process requiring activation of the GTPases Rac1 and cdc42 (Conner and Schmid 2003). Once activated, these GTPases stabilize the formation of actin and filamin projections into the extracellular space allowing the formation of endocytotic vesicles (see Figure 7-1).

Previous studies have shown that Rac1, cdc42, actin and filamin are all down- regulated in many DDP resistant cell lines (Shen, Liang et al. 2004; Shen, Su et al.

2004). Endocytosis has also been reported to be reduced in many DDP-resistant cells

126

(Chauhan, Liang et al. 2003; Liang, Mukherjee et al. 2006). CTR2 kd cells were found to have constitutively active Rac1 and cdc42, inferring that this mechanism may underlie up-regulated marcropinocytosis in these cells.

How does CTR2 limit the activation of Rac1 and cdc42? These GTPases are phosphorylated as a result of PI3K activation. CTR2 does not appear to regulate the

PI3K pathway as there is no change in Akt phosphrylation in CTR2 kd cells when compared to wild-type cells. This suggests that CTR2 might directly and specifically inhibit activation of the Rho-family GTPases. Co-immunoprecipitation or co- localization microscopy experiments may determine if CTR2 directly interacts with these proteins. There are several inhibitors and activators of Rac1 and cdc42 that may also prove to be the targets through which CTR2 modulates the activity of Rac1 and

127 cdc42 (reviewed in (Heasman and Ridley 2008)). For example, p27 kip1 inhibits the interaction of RhoA and Rac1 with its GEF (Besson, Gurian-West et al. 2004). As with CTR2, this protein is found in the cytosol and nucleus in a punctuate pattern.

Additionally, p27 kip1 is controlled by the availability of metals (Hoshino, Tomari et al.

2009). Relocalization of p27 kip1 out of the nucleus to the cytosol is necessary for inhibition of RhoA (Besson, Gurian-West et al. 2004). It is possible that CTR2 inhibits

Rac1 and cdc42 in a similar manner, or it might interact with p27 kip1 directly. There are several ways to access these possibilities, such as by determining whether CTR2 kd cells have lower levels of cytosolic p27 kip1 , determining whether Rac1/cdc42 inhibition is dependent on cytosolic levels of CTR2, and co-immunoprecipitation/co- localization of CTR2 with p27 kip1 .

Clinical Implications

The goal of the work presented in this dissertation was to evaluate the role

CTR2 as a determinant of the accumulation and cytotoxicity of the Pt-containing drugs. It was found that decreasing the level of CTR2 sensitizes cells to DDP and

CBDCA. How can these observations be taken further and lead to more effective chemotherapeutic treatments for patients?

The in vitro and in vivo observations indicate that CTR2 is a valid target through which to enhance Pt drug sensitivity. One strategy for disabling CTR2 is to use a gene silencing strategy similar to several that are currently the focus of clinical

128 research (Soutschek, Akinc et al. 2004; Castanotto and Rossi 2009; Siomi 2009).

However, there are several obstacles for the clinical use of siRNA technology. Poor cellular uptake and inadequate targeting of delivery of siRNA are the primary obstacles. Additionally, siRNA suffers from low stability, is oftentimes quickly degraded and demonstrates low permeability across the extra cellular matrix. If the siRNA is able to enter cells, other challenges include ineffective release from endosomes and separation from the carrier utilized for targeted delivery.

In addition to the difficulties facing effective delivery of siRNA therapies, there are side effects to siRNA gene therapy. Due to the ineffective delivery of these drugs, the dosage needed oftentimes requires the rapid injection of 2.5x the blood volume (Kim, Garg et al. 2009). Side effects include off target gene silencing, inhibition of native miRNA production and immunostimulation (Kim, Garg et al.

2009). To overcome these side effects and the other obstacles facing clinical development of siRNA therapies, much effort is being dedicated to the development of strategies to assist in targeted delivery and increased efficiency of siRNA. These strategies are outlined in Kim et al. (Kim, Garg et al. 2009) and include liposome, polymer and peptide-based delivery mechanisms As these strategies move targeted gene silencing closer to a reality, the studies outlined in this dissertation validate

CTR2 as a possible target for siRNA therapy. To further this finding, future studies can focus on developing carrier mechanisms for silencing CTR2 mRNA in vivo, as well as testing the effectiveness of these strategies on tumors in mouse models .

129

The experiments outlined in this dissertation also demonstrate that CTR2 can be down-regulated by Cu starvation, and that this also leads to increased Pt drug sensitivity. This observation also provides clinical potential. Several anti-cancer Cu chelating are undergoing clinical trials. These drugs include: d-penicillamine, tetrathiomolybdate, clioquinol, and trientine (Yoshii, Yoshiji et al. 2001; Pan, Kleer et al. 2002; Yu, Wong et al. 2006; Gupte and Mumper 2008). The data presented in this dissertation suggests that one effect of these drugs might be to decrease the level of cellular CTR2. Future studies may elaborate on this possibility in mouse models.

Likewise, the ability of these Cu chelators to sensitize tumors to DDP and CBDCA should be explored.

The ability of CTR2 to regulate the rate of endocytosis also has clinical promise. Knockdown of CTR2 enhances the rate of endocytosis and increases accumulation of DDP and CBDCA. However, endocytotic uptake is not a specific process and would not be limited to the Pt drugs. One general challenge for drug delivery is the inability of several drugs to efficiently enter cells. A decrease in CTR2 levels caused by either siRNA-mediated gene silencing or Cu starvation up-regulates endocytosis and this might enhance the accumulation of many classes of drug. Thus, modulating the expression of function of CTR2 has the potential of altering the clinical efficacy, and potentially the toxicity, of many types of chemotherapeutic agents.

The discoveries made during the course of this dissertation show promise for improving chemotherapeutic efficacy. However, as with every clinical strategy, there

130 are potential side-effects. Loss of CTR2 enhances endocytosis through the activation of the GTPases Rac1 and cdc42 and activation of these proteins controls several processes in addition to endocytosis. The same actin and filamin projections necessary for endocytosis and stabilized by Rac1 and cdc42 are also critical for cell migration. It is possible that decreasing CTR2 levels might not only activate endocytosis but also increase metastatic potential. The in vivo studies conducted as part of this thesis examined the effect on metastasis. However, this potential hazard must be examined in appropriate model systems before CTR2 siRNA gene therapy can be initiated.

Additionally, CTR2 is likely to play a key role in Cu homeostasis and this perturbation may have potential negative side effects. Disruption of Cu homeostasis might prove to be beneficial in certain cancer cells; however, no targeted therapy is perfect. Normal cells may be negatively affected by the loss of CTR2 and this potential side effect must be considered in the development of any clinical strategy.

Even if modulation of the expression or function of CTR2 proves impossible to use in patients, the discoveries made during the execution of this thesis have potential for benefiting patient care. The discovery that CTR2 expression regulates sensitivity to the Pt-containing drug suggests that measurement of the expression of CTR2 in pathological specimens may allow prediction of response to these drugs. Likewise, reassessment of how CTR2 levels change during therapy may allow modulation of dose intensity and quantification of the emergence of drug resistance.

131

Conclusions

I have established that CTR2 plays an important role in the transport of DDP and CBDCA into cells. CTR2 limits the uptake of Pt-containing drugs and decreasing the level of CTR2 increases drug accumulation and cytotoxicity. Upon Cu or DDP exposure, CTR2 levels quickly increase, and in Cu-starved conditions CTR2 is quickly degraded by the proteasome. CTR2 is found in the cytosol and nuclei of cells and

CTR2 can regulate cellular endocytosis. These findings provide insight into how the

CTR2 modulates the cellular pharmacology of the Pt drugs, and identify potentially novel strategies for enhancing their therapeutic effectiveness.

Acknowledgements

I wish to thank all the members of the Howell lab for insightful discussions.

Brian Blair was the primary author of this chapter. Stephen B. Howell supervised the writing of this chapter.

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