Endothelin-1 Protects Human Melanocytes from the Photodamaging Effects of Ultraviolet Radiation by Activating the MAP Kinases JNK and P38

Endothelin-1 Protects Human Melanocytes from the Photodamaging Effects of Ultraviolet Radiation by Activating the MAP kinases JNK and p38

A dissertation submitted to the

Division of Research and Advanced Studies of the University of Cincinnati

in partial fulfillment of the requirements for the degree of

DOCTORATE OF PHILOSOPHY (Ph.D.)

in the Department of Cancer and Cell Biology of the College of Medicine

2014

Anne Marie von Koschembahr

A.B., Microbiology Miami University, Oxford, OH, 2007

Committee Chair: Zalfa Abdel-Malek, Ph.D.

ABSTRACT

Melanoma, the deadliest form of skin cancer, is derived from the malignant transformation of melanocytes, epidermal cells that produce the pigment melanin. The main etiological factor for all skin cancers, including melanoma, is exposure to solar ultraviolet (UV) radiation that results in DNA damage. If incorrectly repaired, DNA damage can cause mutations that malignantly transform melanocytes by deregulating critical signaling pathways that control proliferation and survival. Somatic mutations induced by UV cooperate with heritable mutations that increase an individual’s susceptibility to melanoma. With the steady rise in the incidence of melanoma over the past 30 years, there is a critical need to better understand the protective mechanisms that reduce the genotoxic effects of solar UV on melanocytes, thus inhibiting melanoma formation. A complex network of paracrine factors in the skin modulates the response of melanocytes to UV.

One important keratinocyte-derived paracrine factor is endothelin-1, which has mitogenic, melanogenic and survival effects in human melanocytes. Here, we investigate the role of endothelin-1 and its signaling pathways in reducing the genotoxic effects of UV. We report that endothelin-1 reduces the generation and enhances repair of UV-induced DNA photoproducts in cultured human melanocytes. Reduction in UV-induced DNA damage and apoptosis by endothelin-1 is not due to increased melanin content or proliferation. Treatment with endothelin-

1 activates the MAP kinases JNK and p38, and activating transcription factor-2 (ATF-2), and enhances their UV-induced phosphorylation. Inhibition of intracellular calcium mobilization, an important component of the endothelin-1 signaling pathway, markedly abrogates these effects.

Activation of both JNK and p38 are required for reducing DNA photoproducts, but only JNK phosphorylation is critical for activation of ATF-2 and inhibition of apoptosis. Activation of

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ATF-2 is not sufficient for reducing UV-induced DNA damage by endothelin-1. We propose that endothelin-1 restores genomic stability of human melanocytes by reducing the genotoxic effects of UV through its ability to activate the stress-activated MAP kinase signaling pathways

(specifically JNK and p38), thus preventing melanoma. Future studies will further delineate the activation of DNA repair pathways by endothelin-1. Understanding the mechanisms by which endothelin-1 counteracts the photodamaging effects of UV might identify new targets for melanoma prevention.

ACKNOWLEDGEMENTS

“Permanence, perseverance and persistence in spite of all obstacles, discouragements, and impossibilities: It is this that in all things distinguishes the strong soul from the weak.” - Thomas

Carlyle

First and foremost, I want to thank God for endowing me with the blessings of perseverance, knowledge and talents to successfully accomplish this experience.

I want to thank my Mom and Dad for their support, patience and always believing in me. Thank you for being an inspiration of dedication and hard work by installing me with a strong work ethic and a good heart. To my sister, Lauren, I cannot thank you enough for your love and continual support. I am so lucky to have you as my sister.

To my extended family, thank you for supporting me during this journey. I have always appreciated your interests in my educational experience. Thank you for your words of encouragement and rallying behind me.

I would like to thank Dr. Zalfa Abdel-Malek for serving as my advisor during this educational experience. I am very grateful for having the opportunity to learn and work in her laboratory, where I could develop as a student and scientist. To Dr. David Plas and Dr. William Miller, thank you for your support and always challenging me to think critically, which is a fundamental skill needed for success in this field. I have learned so much from both of you: from the classroom, to my qualifying exam and finally, serving on my thesis committee! To Dr. Chunying

Du and Dr. Sohaib Khan, thank you for your invaluable insight and feedback throughout this process. I want to give my utmost appreciation to all these members who served on my committee. Your commitment to my success has been much appreciated and I value each and

vi every one of you for your unique and respected contributions. Thank you for your guidance, patience and support.

I want to thank all of the staff involved in the Biomedical Sciences FLEX option program in the

College of Medicine, who accepted me as a student in 2008. To Dr. Robert Highsmith, thank you for believing in me and seeing my unique potential to succeed as a graduate student. To Mary Jo

Petersman and Laura Hildreth, thank you for answering all my questions and reassuring my concerns. I could not have made it past my first year without you!!

I want to acknowledge the funding sources that provided me with financial support to proceed as a graduate student. To the University of Cincinnati College of Medicine and the Biomedical Flex program, thank you for accepting me and supporting me throughout my search to find a laboratory where I could complete my research. I want to give a big thanks of gratitude to Dr.

Peter Stambrook for accepting me as a pre-doctoral training fellow on his training grant (NIEHS

T32-ES007250). This opportunity was invaluable to my success as a graduate student, as I would not have been able to continue without this support. Special thanks to the Department of

Dermatology who also contributed towards my educational stipend.

I wish to give a very big thank you to all the faculty, students and staff of the Cancer and Cell

Biology department. I have been very fortunate to work with many of you over the years. Thank you for allowing me to be a contributing member to your program. Special thanks to the program coordinators, past and present, for facilitating emails and information.

To the Department of Dermatology (past and current members), I have been very fortunate to work with all of you throughout the years. You have taught me many things and I have enjoyed sharing in your traditions and fellowship. Special thanks to Dr. Diya Mutasim, Dr. Brian Adams,

vii

Marcia Miladinov, Sara Deem and Christy Bailey for their unique contributions to my success. I am very humbled by your generosity and support.

To all members of the Abdel-Malek lab, both past and present: thank you for allowing me to train as a student in this environment. I have learned a lot from each of you over the past years.

To Dr. Viki Swope, I wanted to say how much I enjoyed getting to learn from you and the time we connected at the IPCC in France. That experience will stay with me always. To Josh

Jameson, I enjoyed all the time we spend together. You will make an amazing doctor and I look forward to seeing you successful in your future role.

Special thanks to Dr. George Babcock, Dr. Phil Hexley and Karen Domenico for all your help with the flow cytometer. Thank you to Dr. Dorothy Supp and Kevin McFarland, for teaching me how to perform mRNA studies and allowing me to conduct a portion of my research in the Supp laboratory. I wish to send my appreciation to Dr. Steven Boyce, who permitted me to use his fluorescent microscope.

To Dr. Raymond Boissy, thank you for all your advice and mentoring. You always see the potential in me and your support has been much appreciated! To the past and present members of the Boissy lab, I have been fortunate to have worked with you! You have taught me critical skills and insight during this experience, all with patience and understanding. Dr. Jody Ebanks, you were such a blessing to help me navigate the department and were a great student role model. I was fortunate to have learned from you and I value our friendship very much. Thank you for never giving up on me! Let’s see what the future has in store for us. To Amy Koshoffer, I was so thankful to learn various scientific techniques from you. You were always patient and open- minded with me as a learning student. I wish you all the best in your new position. Finally, to

viii

JàNay Woolridge, your support these last few years have been invaluable to me. You will go far in this experience and I look forward to hearing about your successful defense and graduation!

To Dr. Ana Luisa Kadekaro, there is so much I want to say. God really blessed me when he put you in my life. You welcomed me into the department with open arms and made me feel valued from day one. Thank you so much for all of your valuable insight, wisdom, passion and critiques, all of which were shared in order to make me a better scientist. Thank you for serving as a strong mentor and seeing so many possibilities in me! Obrigada!

To all my friends in the College of Medicine: we have shared so many experiences and grown together through learning. I am grateful to have met this group of highly talented individuals. I look forward to learning about your successes.

To all of my former professors and educators: Your passion for education and learning is an inspiration to all students. Thank you for encouraging me to achieve anything I wanted to become. You have played a vital role in my educational success, which I truly appreciate.

To my closest friends: you have served as the rock on which I stand and your unwavering support makes me incredibly humble and thankful! Even though time passes between our visits over the years, your constant support has been steadfast and true. Each one of you has played a special role in my life, especially throughout this experience. I look forward to sharing this success with you. Amanda Hayden and Mickel Murillo Ràmon, you were the first to know and support me in my decision to attend graduate school. Lauren Fleming, my favorite Francophone and fellow Miamian, whose friendship and love has been so important to me. To my former

Northside crew (Daniela, Rob, Megan, Alex, Amy, Jestin, Mike): we have all transitioned in and out of Cincinnati, but the memories we shared together in our neighborhood will live on in my

ix heart. Let’s see what the future holds in store for this group! To John Stegall, my “older brother”, thank you for your consideration and support! Many thanks to David Abad, who pulled me up when I was down and helped me find my way around. Your technical experience has been vital to my graduate experience, more than I can put into words! To the recently new Dr. Fabiola

Bittencourt and Dr. Marthe-Sandrine Eiymo Mwa Mpollo, your friendship and encouragement have been much appreciated over the years. To Novelle and Max Kimmich, our friendship has grown strong over these past couple years. You have supported me unconditionally and made me feel like one of the family. To Annie and Didier Cazejust, thank you for welcoming me with open arms during my visits back to Paris. You are my French family and I am very lucky to know you.

Finally, to Bertin Ondja’a: I cannot put fully into words how thankful I am for you. You have taught me many things, and supported me in good and bad times. I have cherished all the memories we have shared together and I look forward to the future memories we will create together. I will be here to help support you through the remainder of your studies and I look forward to sharing in your success when you have successfully obtained your PhD. I care about you very much and I hold you close in my heart.

“Education is the most powerful weapon you can use to change the world.” – Nelson Mandela

DEDICATION

This dissertation is dedicated to my parents, Theresa M. and Alan D. Smith, for their never-

ending love and support. I love you both with all my heart.

TABLE OF CONTENTS

ABSTRACT ...... iii ACKNOWLEDGEMENTS ...... vi DEDICATION ...... xi TABLE OF CONTENTS ...... 1 LIST OF TABLES ...... 3 LIST OF FIGURES ...... 4 LIST OF ABBREVIATIONS ...... 6 CHAPTER 1: INTRODUCTION ...... 12 1. Introduction ...... 13 1.1 The Skin ...... 14 1.1.1. Human skin anatomy ...... 14 1.2. The melanocyte (MC) ...... 16 1.2.1. Melanin ...... 17 1.3. Skin pigmentation ...... 19 1.4. UV irradiation ...... 22 1.4.1. Biological effects of UV in the skin ...... 23 1.5. Paracrine responses following UV irradiation of the skin ...... 26 1.5.1. Endothelin-1 (ET-1) ...... 27 1.6. UV activation of the MAP kinases ...... 32 1.7. Significance of the project to melanoma prevention ...... 41 CHAPTER 2: ENDOTHELIN-1 PROTECTS HUMAN MELANOCYTES FROM UV- INDUCED DNA DAMAGE BY ACTIVATING P38 AND JNK SIGNALING PATHWAYS . 42 2. Endothelin-1 Protects Human Melanocytes from UV-Induced DNA Damage by Activating p38 and JNK Signaling Pathways ...... 43 2.1. Abstract ...... 44 2.1.1. Keywords ...... 44 2.2. Introduction ...... 45 2.3. Materials and Methods ...... 47

2.4. Results ...... 52 2.5. Discussion ...... 57 2.6. Acknowledgements ...... 63 2.7. Author Contributions ...... 64 2.8. References ...... 65 2.9. Figures ...... 72 2.10. Supplemental Figures ...... 78 CHAPTER 3: CONCLUSION ...... 94 3. Conclusion: Major Findings, Summary, Significance and Future Directions ...... 95 3.1. Major Findings ...... 95 3.2. Summary ...... 96 3.3. Significance of the study to MC biology and melanoma ...... 101 3.4. Future Directions ...... 102 REFERENCES ...... 110 APPENDIX ...... 150 AI: Additional Data for Chapter 2 ...... 151

LIST OF TABLES Table Page

Table A1. Synergistic effect of ET-1 and α-MSH on melanocyte proliferation. 151

LIST OF FIGURES

Figure Page

1.1. Detailed cross-section of human skin. 15

1.2. Schematic representation of the epidermal melanin unit. 19 1.3. Melanoma risk is associated with constitutive skin pigmentation. 21 1.4. Radiation spectrum of UV and its biological effects in human skin. 22 1.5. Overview of ET-1 synthesis. 28 1.6. Activation of the MAP kinases JNK and p38 signaling pathways. 34 2.1. ET-1 protects HM from UV-induced DNA damage and apoptosis. 72 2.2. Role of intracellular Ca2+ and PKC signaling pathways in mediating the activation of JNK and p38 and the downstream target ATF-2 by UV and ET-1. 73 2.3. Significance of JNK and p38 for activation of ATF-2 in response to UV and ET-1. 75 2.4. Effect of JNK or p38 inhibition on UV-induced CPD and apoptosis. 76 2.S1. Effect of duration of ET-1 pre-treatment on CPD induction and repair, and the kinetics of CPD repair in ET-1 treated HM. 78 2.S2. Dose-dependent response of ET-1 on CPD induction and repair. 80 2.S3. Effect of 1 nM ET-1 on cAMP production. 81 2.S4. Effect of ET-1 on CPD induction and repair in HM expressing LOF MC1R. 82 2.S5. Effect of ET-1 on HM pigmentation. 83 2.S6. Effect of ET-1 on HM proliferation. 84 2.S7. Activation of the MAP kinases JNK and p38 and their downstream target ATF-2 by UV and ET-1. 86 2.S8. Effect of UV and ET-1 on ERK activation. 88 2.S9. Verification of efficacy of JNK, p38 and PKC pharmacological inhibitors. 89 2.S10. Reciprocal effects of MAP kinase inhibition on activation of JNK and p38. 91 2.S11. Schematic representation of the effects of ET-1 on the UV response of HM. 92

A2. Regulation of XPA, cyclin D1 and MC1R genes by ET-1 and UV. 152 A3. Revised summary of study findings. 153

LIST OF ABBREVIATIONS

2-APB 2-aminoethoxydiphenylborate

6,4-PP pyrimidine (6-4) pyrimidone photoproduct

8-OHdG/8-oxo-dG 8-hydroxydeoxyguanosine

α-MSH α-melanocyte stimulating hormone

ACTH adrenocorticotropic hormone

Akt v-akt murine thymoma viral oncogene homolog/protein kinase B

ALDH aldehyde dehydrogenase

ANOVA analysis of variance

AP-1 activator protein 1

Apaf-1 apoptotic peptidase activating factor 1

APC allophycocyanin

APE-1/Ref-1 apurinic/apyrimidinic endonuclease 1

AREG amphriregulin

ASK1 apoptosis signal-regulating kinase 1

ASP agouti signaling protein

ATF-2 activating transcription factor 2

ATP adenosine triphosphate

ATPase adenosinetriphosphatase

Bad Bcl2-associated agonist of cell death

BAPTA/AM 1,2-bis(o-Aminophenoxy)ethane-N,N,Nʹ,Nʹ-tetraacetic Acid Tetra(acetoxymethyl) Ester

Bax Bcl2-associated X protein

Bcl-2 B-cell CLL/lymphoma 2

Bcl-xL B-cell lymphoma-extra large

BER base excision repair bFGF basic fibroblast growth factor

Bid BH3 interacting domain death agonist

Bim Bcl2-like 11 (apoptosis facilitator)

BPE bovine pituitary extract

BrdU bromodeoxyuridine

Brg1 Brahma-related gene 1

Ca2+ calcium

CaM-K Ca2+/calmodulin-dependent protein kinase cAMP cyclic adenosine monophosphate

Cdc25b cell division cycle 25b

CDK cyclin-dependent kinase

CDKN1A cyclin-dependent kinase inhibitor 1A (p21) cDNA complementary DNA c-Fos FBJ murine osteosarcoma viral oncogene homolog c-Jun jun proto-oncogene

CPD cyclobutane pyrimidine dimer

CREB cAMP response element-binding protein

CYCS cytochrome c, somatic

DAG diacylglycerol

DDB2 DNA-damage binding protein 2

DEJ dermal-epidermal junction

DMBA 7,12-dimethylbenz(a)anthracene

DNA deoxyribonucleic acid

ECE endothelin converting enzymes

ECM extracellular matrix

ER endoplasmic reticulum

ERK extracellular signal-related kinase

ET-1/EDN1 endothelin-1

ET-2 endothelin-2

ET-3 endothelin -3

ETBR/EDNBR endothelin B receptor

FAK focal adhesion kinase

Fas Fas cell surface death receptor

FB fibroblast

FOXO forkhead box O

GADD45 growth arrest and DNA damage 45

GAPDH glyceraldehyde-3-phosphate dehydrogenase

GPX glutathione peroxidase

H2O2 hydrogen peroxide

HAT histone acetyltransferase

HCl hydrochloric acid

HBEGF heparin-binding EGF-like growth factor

HGF/SF hepatocyte growth factor/scatter factor

HM human melanocyte

HRP horseradish peroxidase

IκB nuclear factor of kappa light polypeptide enhancer in B-cells inhibitor

IP3 inositol-1,4,5-triphosphate

JNK c-Jun NH2-terminal protein kinase

JunD jun D proto-oncogene

KC keratinocyte

KILLER tumor necrosis factor receptor superfamily, member 10b

LOF loss of function

MAP/MAPK mitogen activated protein (kinase)

MAP2K mitogen activated protein kinase kinase

MAP3K mitogen activated protein kinase kinase kinase

MAPKAP2 mitogen-activated protein kinase-activated protein kinase 2

MARCKS myristoylated alanine-rich protein kinase C substrate

MC melanocyte

MC1R melanocortin 1 receptor

Mcl-1 myeloid cell leukemia 1

Mdm2 mouse double minute 2 p53 binding protein homolog

MEF murine embryonic fibroblast

MITF microphthalmia-associated transcription factor mRNA messenger RNA

NAC N-acetylcysteine

NER nucleotide excision repair

NF-κB nuclear factor kappa-light-chain-enhancer of activated B cells

NGF nerve growth factor

OGG1 8-oxoguanine DNA glycosylase p16 cyclin-dependent kinase inhibitor 2A p53 tumor protein p53 p73 tumor protein 73 p90rsk p90 ribosomal S6 kinase

PAR2 protease-activated receptor 2

PBS phosphate buffer saline

PCR polymerase chain reaction

PERP PERP, TP53 apoptosis effector

PI propidium iodide

PI3K phosphatidylinositol-4,5-bisphosphate 3-kinase

PIP2 phosphatidylinositol-4,5-bisphosphate

PKA protein kinase A

PKC protein kinase C

PLC-β phospholipase C-β

PMAIP1 phorbol-12-myristate-13-acetate-induced protein 1 (aka Noxa)

POMC pro-opiomelanocortin

PUMA p53 upregulated modulator of apoptosis qRT-PCR quantitative real-time reverse transcription PCR

RIPA radioimmunoprecipitation assay

RNA ribonucleic acid

RNAi RNA interference

RNA-seq RNA sequencing

RPA replication protein A

ROS reactive oxygen species

SB 202190 4-[4-(4-fluorophenyl)-5-(4-pyridinyl)-1H-imidazol-2-yl]-phenol

SB 203580 4-(4-fluorophenyl)-2-(4-methylsulfinylphenyl)-5-(4-pyridyl)-imidazole

SB 242235 1-(4-piperidinyl)-4-(4-fluorophenyl)-5-(2-methoxy-4-pyrimidinyl)- imidazole

SEER Surveillance Epidemiology and End Results

SEM standard error of the mean

SFN stratifin (aka 14-3-3 σ) shRNA short hairpin RNA

SNK Student-Newman-Keuls

SP 600125 anthra[1,9-cd]pyrazol-6(2H)-one, 1,9-pyrazoloanthrone

SWI/SNF switching/sucrose non-fermenting

TP63 tumor protein p63

TIGAR TP53-inducible glycolysis and apoptosis regulator

TPA 12-O-tetradecanoylphorbol-13-acetate

TYRP1 tyrosinase related protein-1

TYRP2 tyrosinase related protein-2

USF-1 upstream signaling transcription factor 1

UV ultraviolet (radiation)

UVA ultraviolet A

UVB ultraviolet B

UVC ultraviolet C

XP Xeroderma pigmentosum

XPA Xeroderma pigmentosum, complementation group A

XPB Xeroderma pigmentosum, complementation group B

XPC Xeroderma pigmentosum, complementation group C

XPD Xeroderma pigmentosum, complementation group D

XPE Xeroderma pigmentosum, complementation group E

XPF Xeroderma pigmentosum, complementation group F

XPG Xeroderma pigmentosum, complementation group G

CHAPTER 1

INTRODUCTION

1. Introduction

Malignant melanoma, a neoplasm caused by the malignant transformation of melanocytes

(MCs), is the most fatal form of skin cancer. Solar ultraviolet (UV) radiation is the main etiological factor for skin cancers, including melanoma (Elwood and Jopson, 1997; Gilchrest et al., 1999; Miller and Mihm, 2006). The strongest evidence for the carcinogenic effects of UV is provided by the disease Xeroderma Pigmentosum (XP), characterized by extreme sensitivity to

UV-induced DNA damage and increased risk for melanoma and non-melanoma skin cancers

(Daya-Grosjean, 2008). Clinical data suggest that having a few severe sunburns during the childhood years increases the risk for development of melanoma (Lew et al., 1983), and that this risk doubles after having five or more sunburns during an individual’s lifetime (Pfahlberg et al.,

2001). Experimental evidence that UV exposure is causal for melanomagenesis was demonstrated in a transgenic hepatocyte growth factor/scatter factor (HGF/SF) murine model, where exposure to a single severe dose of UV during neonatal life resulted in melanoma development (Noonan et al., 2001).

The National Cancer Institute’s Surveillance Epidemiology and End Results (SEER) Cancer

Statistics Review estimates that over 76,000 new cases of melanoma will be diagnosed and almost 10,000 melanoma-related deaths will occur in the US during the year 2014. While melanomas consist of less than 10% of all skin cancer incidences, they account for almost 80% of all skin cancer-related fatalities (Gilchrest et al., 1999; Abdulla et al., 2005; Pfeifer and

Besaratinia, 2012; Howlander et al., 2013). Melanoma incidence has been increasing across age groups over the past several decades in the United States, especially in the younger population

(D’Orazio et al., 2013b; Howlander et al., 2013). According to the SEER, the mortality rate for malignant melanoma has increased by about 2% annually since the 1960s. With increasing

13 incidence and mortality, malignant melanoma is truly an emerging public health concern.

Therefore, it is critical to understand the molecular mechanisms of UV exposure in MCs in order to develop strategies for effective photoprotection and prevention of MC transformation.

1.1 The Skin

The skin is the largest organ in the human body and provides the main barrier between the internal organs and the external environment (Haake et al., 2001; Tobin, 2006). The main function of the skin is to serve as a protective barrier against harmful environmental insults, including exposure to solar UV radiation (Tobin, 2006; Proksch et al., 2008). The skin also functions to regulate internal body temperature and protect from excessive water loss (Lin and

Fisher, 2007). The diverse properties of human skin make it a complex and highly developed sensory system that responds to various environmental stimuli and provides a mechanism to maintain the body’s homeostasis, thus promoting and organism’s survival (Slominski and

Wortsman, 2000; Chilcott, 2008; Slominski et al., 2008, 2012 ).

1.1.1. Human skin anatomy

Human skin is a complex multilayered organ that is comprised of three structural layers: the hypodermis, dermis and epidermis (Figure 1.1) (Haake et al., 2001). Interaction between these different layers is important in human development and maintenance for overall homeostasis

(Haake et al., 2001).

Figure 1.1. Detailed cross-section of human skin. Modified from http://cosmetique.ch/skin-care/skin- cross-section.jpg

The innermost layer is the hypodermis (or subcutis), which is primarily composed of fat cells known as adipocytes. This layer functions to control thermoregulation, provide energy and cushions the skin against bone and other internal organs (Katinakis, 2000; Haake et al., 2001).

The middle layer of the skin is the dermis, comprised mainly of fibroblasts (FBs) that secrete a collagenous extracellular matrix (ECM). The dermis functions in regulating internal temperature, providing tensile strength, receiving and processing external stimuli, preventing water loss

(Haake et al., 2001), as well as providing nutritional and physical support to the epidermis

(McLafferty et al., 2012). The dermis is separated from the epidermis by the dermal-epidermal junction (DEJ), a complex, multilayered membrane that is synthesized by keratinocytes (KCs) and FBs. This dynamic interface allows for the exchange of metabolic products while giving mechanical support to the epidermis (Burgeson and Christiano, 1997; Kanitakis, 2002). It is at this basement membrane (basal layer) where epidermal MCs are located and reside permanently

(Nordlund and Boissy, 2001). The epidermis is stratified squamous epithelium that makes up the external portion of the skin. Ninety percent of the epidermis is comprised of KCs, the structural cells which undergo differentiation (keratinization or cornification) as they migrate in the direction from the basal layer to the external surface (Haake et al., 2001; McLafferty et al.,

2012). KCs produce keratins, structural proteins that give the skin its strength and create a waterproof barrier. Maintenance of cells in the epidermis depends on a balance of cell proliferation and cell death (differentiation and apoptosis) of KCs (Eckert et al., 1997). In order to maintain epidermal homeostasis, KCs interact with other neighboring cells, mainly

Langerhans cells, the epidermal antigen presenting cells (macrophages); Merkel cells, the sensory receptor cells; and MCs, the pigment producing cells (Haake et al., 2001; McLafferty et al, 2012).

1.2. The melanocyte (MC)

MCs make up about 8% of all epidermal cells and are responsible for the producing the pigment melanin, which contributes to skin pigmentation and protects against exposure to UV. They are derived from the neural crest during embryogenesis, transitioning from neural stem cells to melanoblasts, the precursors of MCs (Buac and Pavan, 2007; White and Zon, 2008; Kawakami and Fisher, 2011). Melanoblasts undergo extensive migration to reach their destination, which is

16 predominately in the hair follicles and basal layer of the skin (Goding, 2007). Once established within the basal layer, the vast majority of MCs are fully differentiated and rarely divide, just like other cells derived from neural crest origin (Haake et al., 2001; Cichorek et al., 2013).

Unlike KCs that have a short life span, which are regenerated by the highly proliferative basal

KCs and undergo a well-defined differentiation program, MCs survive for decades in the human epidermis, where they are resistant to apoptosis and have poor proliferation capacity (Quevedo et al., 1969; Plettenberg et al., 1995).

1.2.1. Melanin

As previously mentioned, one of the main functions of the skin is to protect the body from harmful UV exposure. UV-induced damage is reduced by the presence of chromophores in the skin, such as DNA and proteins, which absorb the radiation (Young, 1997). The MCs in the skin produce the pigment melanin, the main photoprotective mechanism, which shields skin cells from UV rays. Human MCs synthesize two forms of melanin: eumelanin and pheomelanin.

Eumelanin is the black-brown pigment, which is found in abundance in dark-skinned individuals.

Pheomelanin is the red-yellow pigment, which is best evident in red or blonde hair (Prota, 1992;

Vincensi et al., 1998; Ito et al., 2000; Brenner and Hearing, 2008). Production of melanin, particularly eumelanin, is limited by the activity of tyrosinase, the rate limiting enzyme, which catalyzes 3 steps in the melanin synthetic pathway that converts tyrosine to melanin, as well as the amount of tyrosinase and the tyrosinase related proteins TYRP-1 and TYRP-2 (Halaban et al., 1983).

Melanin is made in specialized organelles called melanosomes (Marks and Seabra, 2001;

Dell’Angelica, 2003). Melanosome biogenesis starts in the MC cytoplasm, with the maturation process occurring within the dendrites. This maturation can be broken down into four stages.

Stage I and II are referred to as the “pre-melanosome”, as there is no actual pigment within the organelle. Stage III contains a low level of pigment, with Stage IV containing the most melanin

(Marks and Seabra, 2001; Dell’Angelica, 2003; Costin and Hearing, 2007). Melanosomes that contain eumelanin are large, elliptical structures, while pheomelanin-containing melanosomes are small and round (Brenner et al., 2008). MCs are dendritic cells that use their dendrites to transfer melanosomes to the surrounding KCs. Dendrite formation is a dynamic process that is stimulated by UV exposure and biochemical melanogenic factors, and is usually indicative of increased melanin synthesis, as dendrites serve as conduits of melanosome transfer to KCs

(Gibbs et al., 2000; Haake et al., 2001; Nordlund and Boissy, 2001; Katinakis, 2002; Scott, 2002;

McLafferty et al., 2012; Cichorek et al., 2013). UV exposure increases the levels of protease- activated receptor 2 (PAR2) on KCs, which facilitates uptake of melanosomes (Scott et al.,

2001). A single MC interacts with 32-36 KCs, forming what is the “epidermal melanin unit”, which insures the even pigmentation of human skin (Figure 1.2) (Haake et al., 2001; Nordlund and Boissy, 2001; Kanitakis, 2002).

Figure 1.2. Schematic representation of the epidermal melanin unit. MCs reside at the basal layer and interact with 30-40 KCs through their formation of dendrites, thus forming the epidermal melanin unit. These dendrites promote the transfer of melanosomes to surrounding KCs, conferring protection from UV exposure. Modified from Cichorek et al., 2013

1.3. Skin pigmentation

Constitutive pigmentation is defined as the amount of total melanin (eumelanin and pheomelanin) in an individual’s epidermis in the absence of the effects of external stimuli, like

UV. Constitutive pigmentation of human skin is genetically determined by: (a) the rate of synthesis of melanin, (b) the relative ratio of eumelanin and pheomelanin synthesized, which is mainly regulated by the melanocortins, the melanocortin 1 receptor (MC1R), and agouti signaling protein (ASP), (c) the rate of transfer of mature melanosomes to KCs, and (d) the distribution of melanin in suprabasal layers of the skin (Scott et al., 2001; Costin and Hearing,

2007; Yamaguchi et al., 2007). Defects in any of these steps cause aberrant pigmentation, resulting in a variety of different skin pigmentary disorders (e.g. albinism, Griscelli’s syndrome)

(Costin and Hearing, 2007; Yamaguchi et al., 2007; Yamaguchi and Hearing, 2009).

Differences in constitutive pigmentation are not due to differences in the number of MCs in the skin of individuals with different melanin contents (Yamaguchi and Hearing, 2009). The differences between light and dark-skin individuals are due to number, size and degradation of melanosomes, as well as the relative amounts of eumelanin and pheomelanin synthesized by

MCs (Costin and Hearing, 2007; Kadekaro et al., 2003; Ebanks et al., 2011). Difference in skin pigmentation is mainly due to eumelanin content, since pheomelanin content can be equivalent in individuals with different pigmentary phenotypes (Hennessy et al., 2005; Wakamatsu et al.,

2006). Eumelanin blocks the penetration of UV radiation through the skin layers (Kaidbey et al.,

1979). Eumelanin has also been shown to neutralize UV-induced reactive oxygen species (ROS), hence reducing oxidative DNA damage (Bustamente et al., 1993; Picardo et al., 1999; Maresca et al., 2008).

While eumelanin has better photoprotective properties, pheomelanin is easily photodegraded

(Prota, 1992; Vincensi et al., 1998; Ito et al., 2000; Brenner and Hearing, 2008). Data suggest that pheomelanin is capable of generating oxidative stress and promoting melanomagenesis by generating free radicals, both in the presence and absence of UV, thus rendering MCs susceptible to transformation (Prota, 2000; Mitra et al., 2012; Denat et al., 2014; Meierjohann, 2014;

Panzella et al., 2014).

Constitutive skin pigmentation is the major defense mechanism against the damaging effects of

UV, and is a determinant of the tanning response to UV. The Fitzpatrick scale (ranging from I to

VI) classifies skin phototypes based on an individual’s constitutive pigmentation and ability to tan following UV exposure (Figure 1.3) (Walker et al., 2003; Brenner and Hearing, 2008). Skin phototypes I-II are those that burn rather than tan when exposed to UV, while skin phototypes

III-VI tan readily and rarely burn.

Figure 1.3. Melanoma risk is associated with constitutive skin pigmentation. Skin phototype is estimated by the Fitzpatrick scale, where low number individuals are lightly pigmented and higher number individuals have darker pigmented skin. Fair-skinned individuals, as depicted by phototype I and II, tend to burn rather than tan following UV exposure and therefore have increased risk for developing melanoma, compared to phototypes III-VI. Modified from D’Orazio et al., 2013b

The extent of UV-induced DNA damage correlates inversely with skin pigmentation, based on the Fitzpatrick scale (Kobayashi et al., 1993, 2001). There is an overwhelming amount of epidemiological data indicating that incidence of skin cancers is highest in individuals with fair skin and poor tanning ability (Gilchrest et al., 1999; Wagner et al., 2002; Miller and Mihm,

2006; Christian et al., 2011; D’Orazio et al., 2013b). In vivo studies have shown that individuals with fair skin have more UV-induced DNA photoproducts compared to dark skin individuals

(Tadokoro et al., 2003; Yamaguchi et al., 2006). Similarly, in vitro, MCs with a low melanin content encounter more DNA photoproducts than their counterparts with a high melanin content

21 in response to the same dose of UV (Hauser et al., 2006) and are more susceptible to oxidative

DNA damage (Maresca et al., 2008).

1.4. UV irradiation

Solar UV radiation is made of electromagnetic rays spanning the wavelengths 100-400nm and can be divided into UVA (320 – 400 nm), UVB (280 – 320 nm) and UVC (100 – 280 nm)

(Figure 1.4) (Diffey, 2002).

Figure 1.4. Radiation spectrum of UV and its biological effects in human skin. Solar UV radiation can be broken down into UVA, UVB and UVC. The ozone layer absorbs UVC and ~90% of UVB, thus

22 allowing UVA and ~10% UVB to reach the Earth’s atmosphere. UV penetrates the skin in a wave-length dependent manner: longer wavelength UVA penetrates into the dermis, while UVB is mainly absorbed by the epidermis. UVA is known for generating oxidative stress response that indirectly damages DNA, while UVB is more associated with direct DNA damage formation. Modified from D’Orazio et al., 2013

The atmospheric ozone layer absorbs UVC rays and a large percentage (~90%) of UVB rays

(Roy et al., 1998). UVA and the remaining UVB rays not absorbed by the ozone layer are able to reach the Earth’s surface. Exposure to this environmental factor can have deleterious biological effects in the skin, such as inflammation, degenerative aging and cancer (Setlow, 1974; Elwood and Jopson, 1997; D’Orazio et al., 2013). The depth of penetration through the skin and biological effects differ between UVB and UVA, with UVB being more energetic and mutagenic than UVA, and UVA having the capacity to penetrate deeper in the skin, reaching the dermal layer (Pfeifer and Besaratinia, 2012). UVB is more potent than UVA in inducing inflammation, the underlying cause for erythema (redness) and sunburn, DNA damage and tanning (Soehnge et al., 1997; Kadekaro et al., 2003; Abdulla et al., 2005; Besaratinia et al., 2008). Exposure to

UVA, such as commercial tanning beds, is highly associated with advanced photoaging (due to its ability to reach FBs in the dermis), immunosuppression and wrinkles (Krutmann, 2001;

D’Orazio et al., 2013b).

1.4.1. Biological effects of UV in the skin

As previously mentioned, UVB is more energetic and mutagenic than UVA. Acute exposure to

UVB induces ertythema (redness), due to inflammation. Exposure to UVB stimulates melanin synthesis and distribution, resulting in skin darkening (ie., tanning response) (Soehnge et al.,

1997; Kadekaro et al., 2003; Abdulla et al., 2005; Besaratinia et al., 2008; Pfeifer and

Besaratinia, 2012). UVB radiation penetrates the epidermis and is directly absorbed by the DNA, which can result in the formation of UV photolesions in the form of cyclobutane pyrimidine dimers (CPD) and pyrimidine (6-4) pyrimidone photoproducts (6,4-PP) (Freeman et al., 1989;

Mitchell et al., 1991; Sarasin, 1999; Ichihashi et al., 2003). If not repaired, these lesions will result in mutations due to C to T or CC to TT transitions, which are commonly referred to as

“UV signature” mutations (Sarasin, 1999; Sinha and Häder, 2002; Ichihasi et al., 2003; Lo et al.,

2005). CPDs are the major form of UV-induced DNA damage and are considered to be the major contributor to mutations. 6,4-PPs tend to be induced 5-10 fold less than CPD and are repaired at a faster rate (Mitchell, 1988; Eveno et al., 1995; Soehnge et al., 1997; Nakagawa et al., 1998; de Lima-Bessa et al., 2008).

Longer UVA wavelengths allow for deep penetration of UVA rays through the skin, reaching the dermis. UVA rays are the main culprit in photoaging through their effects on dermal FBs

(Krutmann, 2001; D’Orazio et al., 2013b; Pfeifer and Besaratinia, 2012). While UVA might not cause melanoma in humans, they interact synergistically with UVB to cause photocarcinogenesis, including melanoma (Jhappan et al., 2003; DeFabo et al., 2004; Noonan et al., 2012). Unlike UVB, UVA is not absorbed by the DNA, but rather by endogenous photosensitizers (i.e., porphyrins, bilirubin, pheomelanin, riboflavin) resulting in the generation of ROS, like hydrogen peroxide, superoxide and singlet oxygen (de Gruijl et al., 1999, 2001;

Wondrak et al., 2005; Song et al., 2009). Production of ROS induces cellular oxidative stress, which causes damage to DNA (single and double strand breaks, and DNA adducts) and proteins, as well as lipid peroxidation (Morliére et al., 1991; Nishigori et al., 2004; Cadet and Douki,

2011). The signature lesion for ROS-induced DNA damage is 8-hydroyxdeoxyguanosine (8-

OHdG or 8-oxo-dG), which is mutagenic (Kvam and Tyrell, 1997; Ichihashi et al., 2003). There

24 is also evidence that UVA is capable of inducing photolesions (CPD and 6,4-PP), however the mechanism of formation is still poorly understood (Mouret et al., 2006, 2010; Girard et al.,

2011).

CPDs and 6,4-PPs are known to be primarily repaired using the nucleotide excision repair (NER) pathway (Sinha and Häder, 2002; D’Orazio et al., 2013). The significance of this pathway in preventing cancer, mainly skin cancer, is best illustrated in patients with XP, a rare syndrome characterized by UV hypersensitivity due to mutations in the NER genes (XPA, XPB, XPC, XPD,

XPE, XPF and XPG), which encode proteins and enzymes that contribute to removal of DNA photoproducts (Daya-Grosjean, 2008). XP patients have a 1000 fold higher risk for all forms of skin cancer, including melanoma, than the remaining population due to inefficient NER capacity

(Kraemer et al., 1994; Daya-Grosjean, 2008). CPD, which are UV signature DNA lesions, are the main causes of non-melanoma skin cancers (basal cell carcinoma and squamous cell carcinoma) that originate from KCs (D’Orazio et al., 2013). Mutations in p53, which are very common in non-melanoma skin cancers, are typical of UV-signature mutations (Kanjilal et al.,

1993; Sato et al., 1993; Daya-Grosjean et al., 1995). Unrepaired DNA photoproducts result in mutations that are also found in melanoma (Sinha and Häder, 2002; D’Orazio et al., 2013,

2013b). Oxidative DNA damage is primarily removed by the DNA repair pathway known as base excision repair (BER) (Barzilai and Yamamoto, 2004). If not repaired, this form of damage can also be mutagenic, causing chromosomal instability and promoting tumor formation, leading to non-melanoma and melanoma skin cancers (Sander et al., 2003; Bickers and Athar, 2006;

Narendhirakannan and Hannah, 2013). From the above, it can be concluded that melanoma arises due to mutations caused by unrepaired UV-induced DNA photoproducts and oxidative DNA damage that is exacerbated by the oxidation of pheomelanin (Prota, 2000; Sinha and Häder,

2002; Mitra et al., 2012; D’Orazio et al., 2013, 2013b; Machado de Melo et al., 2013; Denat et al., 2014; Meierjohann, 2014; Panzella et al., 2014).

1.5. Paracrine responses following UV irradiation of the skin

The survival, function and homeostasis of MCs are maintained by biochemical mediators in their microenvironment (Swope et al., 1995; Halaban, 2000; Imokawa, 2004). It is established that a complex paracrine network exists in the epidermis, and that this network is activated following exposure to UV (Chakraborty et al., 1996; Gilchrest et al., 1996; Yamaguchi et al., 2007;

Berridge, 2012). KCs synthesize and secrete paracrine factors that bind to specific membrane surface receptors or nuclear receptors expressed by MCs, and production of these factors is increased upon UV exposure. Some of the KC-derived paracrine factors regulate the synthesis of melanin, promote survival and have been shown to activate DNA repair pathways of MCs

(Archambault et al., 1995; Bender et al., 1997; Halaban, 2000; D’Orazio et al., 2013). For example, the KC-derived nerve growth factor (NGF) reduces apoptosis and increases expression of the anti-apoptotic protein Bcl-2 in UV-irradiated MCs (Zhai et al., 1996). 1,25 dihydroxyvitamin D3 (1,25(OH)2 vitamin D3) reduces CPDs and increases p53 expression in

UV-irradiated mice in vivo (Dixon et al., 2005, 2011). A hallmark of UV exposure is tanning, resulting from increased melanin synthesis and its distribution in the epidermis. The melanocortins, α-melanocyte stimulating hormone (α-MSH) and adrenocorticotropic hormone

(ACTH) increase pigmentation, reduce apoptosis and enhance CPD repair (Böhm et al., 2005;

Kadekaro et al., 2005, 2010; Swope et al., 2014). α-MSH reduces ROS levels (Kadekaro et al.,

2005) and increases expression of key enzymes involved in BER, which removes oxidative DNA damage, specifically through α-MSH mediated accumulation of p53 (Kadekaro et al., 2012).

Endothelin-1 (ET-1), another KC-derived factor, promotes survival and reduces oxidative stress in MCs irradiated with UV (Tada et al., 1998; Kadekaro et al., 2005). These findings provide unequivocal evidence for the participation of epidermal-derived paracrine factors in the DNA damage response of MCs, and prevention of UV-induced genotoxicity and cytotoxicity. The expected outcome is maintenance of genomic stability and inhibition of melanoma.

1.5.1. Endothelin-1 (ET-1)

Endothelins are signaling molecules made of 21 amino acid residues. There are three types of endothelin peptides, ET-1, ET-2 and ET-3, which are the product of three distinct genes that code for preproendothelin 1, 2 and 3. These proteins are targeted by furin-like proteases to make the big endothelins (big ET-1, big ET-2 and big ET-3), and finally by endothelin-converting enzymes (ECE) to yield the final active peptides (Figure 1.5) (Lüscher and Barton, 2000;

Saldana-Caboverde and Kos, 2010; Stow et al., 2011; Rosanò et al., 2013).

Figure 1.5. Overview of ET-1 synthesis. Structure of edn1 gene and mRNA, which translates into preproendothelin-1. PreproET-1 is processed in a proteolytic sequence to yield active ET-1. The fully mature structure of ET-1 contains 2 disulfide bridges and was rendered by Stow et al. (2011) from the RCSB Protein Data Bank (1T7H). Modified from Stow et al., 2011

ET-1 and ET-3 both play critical roles in MC development, survival and homeostasis. Signaling of the endothelin B receptor (EDNBR aka ETBR), particularly through FB derived ET-3, plays a critical role during embryonic development, as it is required for the migration and survival of neural crest derivatives after their commitment to the melanocytic lineage, and for the final differentiation stages of melanoblasts (Baynash et al., 1994). Mutations in either ET-3 or ETBR

28 result in spotted coat color and agangliolic megacolin in mice, causing a condition similar to

Hirchsprung’s disease (Hosoda et al., 1994). Mice deficient in ET-1 have craniofacial abnormalities and die due to respiratory complications (Kurihara et al., 1994).

Although ET-1 was initially described as a potent vasoconstrictive peptide for endothelial cells, it also is produced by KCs and functions as a mitogen and melanogenic factor for human MCs

(Yada et al., 1991; Imokawa et al., 1992, 1995; Tada et al., 1998). Additionally, KC secretion of

ET-1 is significantly increased in the skin following exposure to UV (Imokawa et al., 1992; Hara et al., 1995). ET-1 promotes MC dendrite formation, which enhances the transfer of melanosomes from MCs to KCs (Hara et al., 1995). In vivo studies on mouse skin revealed that increased expression of ET-1 in KCs is regulated transcriptionally by p53 (Hyter et al., 2013), and targeted deletion of the ET-1 gene in KCs results in MC apoptosis following UV exposure.

Cui et al. (2007) have reported that p53 upregulates expression of pre-opiomelanocortin

(POMC), the precursor to α-MSH and ACTH, in murine epidermis. Since p53 is activated by

DNA damage, we propose that increased ET-1 synthesis and POMC-derived peptides are part of the response to DNA damage. These findings underscore the significance of ET-1 in maintenance of MC survival.

The effects of ET-1 on MCs are mediated predominantly by binding to the ETBR, a membrane bound Gq protein-coupled receptor. The phosphoinositide cascade is mediated by the activation of phospholipase C-β (PLC-β), which cleaves the membrane bound phosphatidylinositol-4,5- bisphosphate (PIP2) into diacylglycerol (DAG) and inositol-1,4,5-triphosphate (IP3). DAG remains membrane-bound, while the generation of IP3 in the cytosol allows it to diffuse and bind to IP3 receptors, especially calcium channels in the smooth endoplasmic reticulum (ER). This results in the release of endogenous calcium (Ca2+) stores from the ER to mobilize in the cytosol.

Together, DAG and Ca2+ activate protein kinase C (PKC), leading to activation of the MAP kinase signaling pathway cascade, specifically the Raf/MEK/ERK pathway, promoting cell proliferation (Imokawa et al., 1996; Yada et al., 2001; Tada et al., 2002; Yogi et al., 2007; Chen et al., 2009). Downstream targets of this pathway include p90 ribosomal s6 kinase (p90rsk), cAMP response element binding protein (CREB) and cAMP-protein kinase A-CREB and the non-receptor tyrosine kinase, focal adhesion kinase (FAK) (Böhm et al., 1995; Imokawa et al.,

1996, 1997; Scott et al., 1997; Tada et al., 2002; Sato-Jin et al., 2008). Besides activating ERK, signaling of ET-1 through its receptor has been shown to activate other pathways in human MCs

(Rubanyi and Polokoff, 1994; Imokawa et al., 1996; Bouallegue et al., 2007; Saldana-Caboverde and Kos, 2010; Rosanò et al., 2013). Kadekaro et al. (2005) demonstrated that ET-1 participates in the activation of the PI3K/Akt signaling pathway in UV-irradiated human MCs, enhancing cell survival. Imokawa et al. (1996, 1997) reported that treatment of human MCs with ET-1 increased cAMP production, linking ET-1 to the cAMP/PKA signaling pathway, promoting melanogenesis and MC proliferation.

In human MCs in vitro, ET-1 interacts synergistically with α-MSH and basic fibroblast growth factor (bFGF) to promote proliferation, melanogenesis, and survival (Swope et al., 1995; Tada et al., 1998; Kadekaro et al., 2005). All of these effects require activation of multiple signaling pathways, namely increased intracellular Ca2+ concentration and activation of PKC (by ET-1), receptor tyrosinse kinases (by bFGF), and increased cAMP production (by α-MSH) (Abdel-

Malek et al., 1995; Halaban, 2000). Accordingly, these mitogens are used for maintenance and expansion of cultured MCs in vitro.

ET-1 participates in the DNA damage response of human MCs to UV, as evidenced by reduction of UV-induced apoptosis and decreased ROS generation in UV-irradiated MCs (Kadekaro et al.,

2005). In UV-irradiated MCs, ET-1 and α-MSH interact synergistically to promote survival by reducing apoptosis, particularly via activation of Akt and increasing the levels of the anti- apoptotic protein Bcl-2, as well as by enhancing repair of DNA photoproducts and reducing ROS levels (Kadekaro et al., 2005). The significance of ET-1 in maintaining MC homeostasis, particularly after UV exposure, was demonstrated in mice with targeted deletion of ET-1 in epidermal KCs, which exhibited reduced number of MCs and deficient repair of UV-induced

DNA damage (Hyter et al., 2013). This provides strong evidence for the protective role of ET-1 in promoting genomic stability following UV exposure. The mechanism by which ET-1 exerts its effect following UV has yet to be elucidated in human MCs.

There is substantial evidence that the signaling pathway of the ETBR enhances melanoma tumor progression, by altering tumor-host interactions, promoting angiogenesis and inflammation

(Jamal and Schneider, 2002; Spinella et al., 2007), which are mediated through ET-1 and ET-3

(Saldana-Caboverde and Kos, 2010). These effects are enhanced by over-expression of ETBR by melanoma tumor cells (Spinella et al., 2007; Tang et al., 2008), and melanoma metastases, thus suggesting that ETBR may be used as a melanoma progression marker (Demunter et al., 2001).

Together, this places the endothelin signaling pathway as a critical player in the various aspects of MC biology: during embryonic development, and maintenance of MC in the epidermis

(protection from UV-induced DNA damage and mutagenesis) on one hand, and transformation and progression to melanoma on the other. Here, we focus on the protective role of ET-1 in normal human MCs exposed to UV.

1.6. UV activation of the MAP kinases

The longevity of MCs (for decades) in the epidermis and their resistance to apoptosis (due to high levels of the pro-survival Bcl-2 protein) (Plettenberg et al., 1995; Nys and Agostinis, 2012) make them vulnerable to malignant transformation by repetitive exposure to solar UV. MC survival can be a double-edged sword, since resistance of transformed MCs to apoptosis represents a major obstacle in melanoma therapy, and survival of MCs with DNA damage increases mutagenesis that can initiate melanoma. Maintenance of genomic stability involves repair of UV-induced DNA damage as well as initiating anti-tumor barriers, like cell-cycle arrest and senescence (Sinha and Häder, 2002; Nakanishi et al., 2009). An important factor that plays a major role in these processes is the tumor suppressor, p53 (Latonen and Laiho, 2005). The extent of DNA damage, which is UV-dose dependent (Hauser et al., 2006), along with the coordination of different players involved in UV response ultimately determines the MC’s fate and its genomic stability (López-Camarillo et al., 2012).

Exposure to UV activates different signaling pathways in a time, dose and wavelength specific manner (Bender et al., 1997; Ibuki and Goto, 2002). An important and early signaling pathway activated in response to UV involves the mitogen activated protein (MAP) kinases (Tada et al.,

2002; Bode and Dong, 2003; Wagner and Nebreda, 2009). The MAP kinases consist of extracellular signal-related kinases (ERK), p38 MAPK and c-Jun NH2-terminal protein kinase

(JNK). The ERK cascade is usually activated by mitogenic stimuli, and mediates cell proliferation and survival (Tada et al., 2002; Roux and Blenis, 2004). As previously mentioned,

ET-1 signaling has been shown to induce activation of ERK1/2 (Imokawa et al., 1996; Yada et al., 2001; Tada et al., 2002; Yogi et al., 2007; Chen et al., 2009) JNK and p38 are the stress-

32 induced MAP kinases that are known to be activated in response to UV (Figure 1.6) (Weston and

Davis, 2002; Wagner and Nebreda, 2009).

Figure 1.6. Activation of the MAP kinases JNK and p38 signaling pathways. MAP kinase pathways are activated by environmental stressors, growth factors and inflammatory cytokines. MAP3K and MAP2K family members are the upstream activators of JNK and p38. Once activated, JNK and p38 target

34 a variety of downstream targets, including transcription factors and other targets, which help to determine various biological responses, such as cell proliferation and survival. Modified from Wagner and Nebreda, 2009

Activation of both JNK and p38 are considered an early event following UV radiation (Kyriakis and Avruch, 1996; Tada et al., 2002). This activation occurs by different upstream activators, such as the MAP2K and MAP3K family members, which is outlined in Figure 1.6. Once activated, these kinases are able to mediate their effects by phosphorylating specific residues on target substrates, like transcription factors (c-Jun, p53, ATF-2, NF-κB) and other protein kinases, and participate in regulating apoptosis (Weston and Davis, 2002; Wada and Penninger, 2004;

Dhanasekaran and Reddy, 2008; Wagner and Nebreda, 2009; López-Camarillo et al, 2012).

Although JNK and p38 can be activated concurrently by the same stimuli, it has been shown that they can mediate very different outcomes that antagonize each other. Cultured murine KCs lacking p38α have increased baseline activation of ERK, and increased JNK activation following

UV exposure (Kim et al., 2008), which may serve as a compensatory mechanism. Murine embryonic fibroblasts (MEFs) lacking functional JNK are unable to proliferate, but selective pharmacological inhibition of p38α/β reverses this effect (Wada et al., 2008). This suggests that

JNK and p38 may have an antagonistic relationship with each other, and that this cross-talk allows for their activity to be regulated in a variety of different cellular responses.

The JNK protein kinases are encoded by three genes, MAPK 8 (encoding JNK1), MAPK9

(encoding JNK2) and MAPK10 (encoding JNK3), that ultimately are spliced to form up to 10 known isoforms (Wagner and Nebreda, 2009). Mice that lack JNK1 or JNK2 have defects in T cell development (Wagner and Nebreda, 2009). Interestingly, when treated with the carcinogen

7,12-dimethylbenz(a)anthracene (DMBA) and the tumor promoter 12-O-tetradecanoylphorbol-

13-acetate (TPA), JNK1-/- mice develop numerous skin papillomas (She et al., 2002), whereas

35 mice lacking JNK2 are resistant to tumorigenesis (Chen et al., 2001). This suggests that different forms of JNK have opposing roles in TPA-induced skin cancer formation.

Activation of JNK has been linked to apoptosis (both pro- and anti-) and cell proliferation (Liu and Lin, 2005; Bivik and Ollinger, 2008; Dhanasekaran and Reddy, 2008; Wagner and Nebreda,

2009). In the literature, chronic activation of JNK is associated with apoptosis, while transient

JNK activation enhances cell survival (Chen et al., 1996b; Christmann et al., 2007; Wu et al.,

2014). JNK1 and JNK2 knockout MEFs are resistant to UV-induced apoptosis, due to their inability to release cytochrome c (Tournier et al., 2000). In normal human MCs, JNK activation has been found to have a pro-apoptotic function following UV radiation, by regulating lysosomal membrane permeabilization (Bivik and Ollinger, 2008). UV activation of JNK was linked to apoptosis through two events: 1. phosphorylation of the pro-apoptotic protein Bim, allowing it to dissociate from the pro-survival protein Mcl-1, and 2. lysosomal release of cathepsins B and D, two proteases that convert Bid to Bax (Bivik et al., 2006). Bim and Bax are able to then converge and insert themselves into the mitochondrial membrane, thus promoting cytochrome c release and ultimately apoptosis (Bivik and Ollinger, 2008). Pharmacological inhibition of JNK resulted in increased cell death and attenuated CPD repair following UV exposure (Christmann et al.,

2006, 2007). JNK suppresses apoptosis by phosphorylating and inhibiting Bad, a pro-apoptotic protein that inhibits the effect of the pro-survival protein, Bcl-xL (She et al., 2002b; Yu et al.,

2004). JNK functions in cell survival by phosphorylating various targets that either promote or inhibit apoptosis. JNK is known to activate members of the activator protein-1 (AP-1) family of transcription factors, like c-Fos, c-Jun, JunD and ATF-2 (Wagner and Nebreda, 2009).

Activation of these proteins is important for cell cycle and DNA damage repair. MEFs that are deficient in c-Fos have increased amounts of UV-induced photolesions, due to their inability to

36 re-synthesize XPF, a component of the NER pathway that is downregulated following UV exposure (Christmann et al., 2006, 2007). Lamb et al. (2003) found that JNK can phosphorylate the transcription factor JunD, which cooperates with NF-κB to promote survival, most likely by increasing pro-survival gene expression. The dual role of JNK in apoptotic and survival signaling pathways indicates that JNK function is complex, depending on the cell type, the type and extent of stimuli, and the isoform of activated JNK.

The p38 protein kinases are encoded by four genes (p38α, p38β, p38γ and p38δ) to generate four isoforms, with p38α having the highest expression (Wagner and Nebreda, 2009). Mice deficient in p38α are embryonic lethal by E12.5 (Adams et al., 2000), whereas targeted deletion of p38α in murine KCs showed no observable phenotype and, in vivo have a dampened inflammatory response following UV irradiation (Kim et al., 2008). Mice lacking p38δ have no obvious phenotype but are resistant to developing skin papillomas following the DMBA/TPA skin carcinogenesis protocol (Schindler et al., 2009).

Similar to JNK, p38 has been linked to both apoptotic and anti-apoptotic responses.

Pharmacological inhibition of p38 was shown to protect human epidermis from apoptosis and inflammation following UV irradiation (Hildesheim et al., 2004). Oral administration of the p38 inhibitor SB 242235 followed by UVB irradiation of murine back skin prevented expression of two pro-inflammatory cytokines, interleukin- 6 and interleukin-8, as well as reduced erythema

(Kim et al., 2005). p38 has been shown to promote apoptosis by phosphorylating the pro- apoptotic protein Bim (S65) (Cai et al., 2006). Activation of p38 has been linked to expression of genes that promote apoptosis (PMAIP1 [Noxa], CYCS [cytochrome c]), inflammation (AREG,

HBEGF) and cell cycle regulation (TP63, CDKN1A [p21], SFN [14-3-3σ]) (Mouchet et al.,

2010). Several of these effects are mediated through transcriptional activation of p53, a well-

37 known downstream target of p38 (Chen et al., 1996; Mouchet et al., 2010). Use of SB 202190 caused increased apoptosis in Jurkat cells exposed to UV (Nemoto et al., 1998). Melanoma cells are protected from UV-induced apoptosis by p38 activation, as treatment with SB 203580 promoted apoptosis through increased Fas expression and NF-κB activity (Ivanov and Ronai,

2000). Jenkins et al. (2011) demonstrated that activation of p38 was necessary to regulate p16 expression in human MCs following H2O2 treatment, thus reducing oxidative stress. p38 has been shown to phosphorylate a downstream substrate, MAPKAP2, leading to phosphorylation and nuclear export of Cdc25b, thus promoting cell cycle arrest at the G2 checkpoint (Bulavin et al., 2001; Manke et al., 2005). In MCs, p38 has been found to be important for melanogenesis

(Corre et al., 2004), activation and stabilization of p53 in response to UV and/or α-MSH treatment (Kadekaro et al., 2012), and the oxidative stress response (Kadekaro et al., 2012; Liu et al., 2012). In UV-irradiated human FBs, p38 activation has been found to be critical for removal of photolesions by NER via by its ability to regulate chromatin relaxation and recruitment of

NER factors to the lesion site (Zhao et al., 2008; Wang et al., 2013). Just like JNK, p38 is involved in both apoptotic and survival responses, depending on the stimuli and cell type, indicating the complexity of p38 function. It has been shown that irradiation of human MCs with

UV activates both JNK and p38, but not ERK 1/2 (Tada et al., 2002). The roles of JNK and p38 in UV-induced DNA repair and apoptosis have yet to be explored in human MCs.

One major downstream target of both JNK and p38 is the tumor suppressor p53, the universal sensor of cellular stress and regulator of DNA damage response (Chen et al., 1996; Levin, 1997;

Bai and Zhu, 2006). The main function of p53 is to serve as a transcription factor that activates a variety of different target genes that play roles in cell cycle arrest, apoptosis, DNA damage repair and survival (Levin, 1997; Latonen and Laiho, 2005). Following cellular stress or mitogenic

38 signaling, JNK and p38 phosphorylate the transcriptional domain of p53, allowing stabilization and accumulation of the protein, as well as enhancing its transcriptional activity (Levin, 1997;

Fuchs et al., 1998; Latonen and Laiho, 2005; Jones et al., 2007; Wagner and Nebreda, 2009;

Gong et al., 2010; Kadekaro et al., 2012). It has been suggested that low doses of UV trigger p53 to upregulate expression of target genes involved in cell-cycle arrest and other protective responses, while higher doses of UV induce p53 target genes involved in apoptosis (Latonen and

Laiho, 2005; Gong et al., 2010). p53 is highly associated with its tumor suppressor role by its ability to control apoptosis through gene transcription of various targets involved in mitochondrial and death receptor mediated apoptotic pathways. JNK has been shown to specifically phosphorylate p53 or p73 (Fuchs et al.,

1998; Jones et al., 2007), thus promoting apoptosis through the expression of pro-apoptotic genes, like Bax (Miyashita and Reed, 1995) and PUMA (Nakano and Vousden, 2001). Other p53 target genes associated with mitochondria-mediated apoptosis include Noxa (Oda et al., 2000) and Apaf-1 (Soengas et al., 1999). Death receptors that are induced by p53 include Fas (Müller et al., 1998), KILLER (Wu et al., 1997) and PERP (Attardi et al., 2000). p53 has also been shown to contribute to apoptosis by repression gene expression of pro-survival genes like Bcl-2

(Haldar et al., 1994) and survivin (Hoffman et al., 2002), or interact directly with Bcl-2 and Bcl- xL to prevent their pro-survival functions at the mitochondria (Mihara et al., 2003).

Although p53 is strongly associated with apoptosis, it has also been found to participate in pro- survival responses, like cell cycle arrest, melanogenesis, DNA damage repair and oxidative stress. p53 participates in cell cycle arrest in the G1, S and G2 phases. One transcriptional target of p53 is p21, which functions as an inhibitor of cyclin-dependent kinases (CDKs) leading to cell arrest in G1 (el-Deiry et al., 1993), as well as S and G2 (Agarwal et al., 1995) phases. 14-3-3σ

39 and growth arrest and DNA damage inducible protein 45 (Gadd45) are two additional transcriptional p53 targets that participate in G2 cell cycle arrest following cellular stress

(Hermeking et al., 1997; Zhan et al., 1999). p53 activation has been linked to melanogenesis, by increasing expression of pro-opiomelanocortin (POMC) (Cui et al., 2007), the precursor for

ACTH and α-MSH; tyrosinase (Khlagatian et al., 2000); and TYRP1 (Nylander et al., 2000). p53 plays a role in the DNA damage response by contributing to DNA repair. UV-irradiated MEFs lacking p53 or GADD45 (a target gene of p53) have deficient repair of photolesions, while loss of p21 (another p53 target gene) has little effect on DNA repair (Smith et al., 2000). p53 has also been shown to recruit histone acetyltransferase (HAT) complexes to sites of DNA damage, facilitating chromatin relaxation needed for DNA repair (Rubbi and Milner, 2003). Activation of p53 occurs in a p38-dependent manner in MCs treated with α-MSH, and inhibition of p53 reduced the ability of α-MSH to increase expression of genes involved in BER (OGG1 and APE-

1/Ref-1), as well as increased DNA damage following UV and/or H2O2, and increased UV- induced oxidative stress (Kadekaro et al., 2012). Additionally, p53 has been shown to upregulate other genes involved in reducing oxidative stress, like Sestrin (Budanov et al., 2004), GPX (Tan et al., 1999), ALDH (Yoon et al., 2004) and TIGAR (Bensaad et al., 2006). Together, these findings reveal that p53 can also play a protective role, especially in MCs.

1.7. Significance of the project to melanoma prevention

Given the continuous rise in the incidence of melanoma and the role of UV in melanomagenesis, it is critical to gain better understanding of how MCs respond to UV, and the role of paracrine factors in the UV response. The evidence that paracrine factors play a critical role in ameliorating the photodamaging effects of UV suggests their significant role in promoting genomic stability of MCs and preventing their malignant transformation. Our study focuses on the effects of ET-1 and its signaling pathways on the DNA damaging effects of UV, mainly induction and repair of the DNA photoproducts CPDs, and apoptosis. UV activates the stress- activated MAP kinases JNK and p38, which have been shown to play a role in determining cell fate. We have investigated how UV and ET-1 regulate the activation of these two kinases in human MCs, and the role of these MAP kinases in the repair of DNA damage and inhibition of apoptosis. Understanding the DNA damage response of human MCs to UV is significant for identifying pathways that can be targeted for prevention of melanoma.

CHAPTER 2

ENDOTHELIN-1 PROTECTS HUMAN MELANOCYTES FROM UV-INDUCED DNA DAMAGE BY ACTIVATING P38 AND JNK SIGNALING PATHWAYS

2. Endothelin-1 Protects Human Melanocytes from UV-Induced DNA Damage by

Activating p38 and JNK Signaling Pathways

Anne M. von Koschembahr1,2, Renny J. Starner1, Viki B. Swope1, and Zalfa A. Abdel-Malek1*

1Department of Dermatology, University of Cincinnati College of Medicine, USA

2Cancer and Cell Biology, University of Cincinnati College of Medicine, USA

*correspondence:

Mailing address: Department of Dermatology

University of Cincinnati

231 Albert Sabin Way

Cincinnati, Ohio 45267

Tel. #: 513-558-6246

Fax #: 513-558-0198

Email: [email protected]

Presently being editing for re-submission to Experimental Dermatology

Manuscript modified for dissertation purposes

2.1. Abstract

Endothelin-1 is a paracrine factor with mitogenic, melanogenic and survival effects on human melanocytes, and its synthesis by keratinocytes is increased upon exposure to ultraviolet radiation (UV). We report that endothelin-1 reduces the generation of UV-induced DNA damage and enhances repair of DNA photoproducts in cultured melanocytes, and inhibits apoptosis in the absence of any increase in melanin content, proliferation or signaling via cAMP. Treatment with endothelin-1 activates the MAP kinases JNK and p38, evident by phosphorylation of their target, activating transcription factor-2 (ATF-2), and enhances their UV-induced phosphorylation.

These effects are dependent on increasing intracellular calcium mobilization, an important component of endothelin B receptor signaling pathway. Activation of both JNK and p38 is required for reducing DNA photoproducts. ATF-2 activation depends on JNK, but not p38, yet is not sufficient for reducing UV-induced DNA damage, suggesting the requirement for other JNK and p38 targets in this effect. We conclude that endothelin-1 is part of a paracrine network that restores genomic stability of human melanocytes by reducing the genotoxic effects of UV via activating JNK and p38.

2.1.1. Keywords: Endothelin-1, human melanocytes, ultraviolet radiation, p38, JNK

2.2. Introduction

Exposure to ultraviolet radiation (UV) is the main etiological factor for melanoma, the deadliest form of skin cancer that originates from human melanocytes (HM) (1, 2). There is compelling clinical data implicating a few severe sunburns during the childhood years with increased melanoma incidence (3-5). Experimental evidence for the role of UV in melanomagenesis is provided by neonatal transgenic HGF mice that develop melanomas following a single, severe dose of UV (6). Elucidating the response of melanocytes to UV is critical for identifying targets for melanoma prevention.

Melanocyte survival, proliferation and melanin synthesis are regulated by a complex network of paracrine factors whose synthesis is increased upon UV exposure of epidermal cells

(7-11). There is increasing evidence that some paracrine factors, such as interleukin-12 and

1,25(OH)2 vitamin D3, reduce the genotoxic effects of UV on epidermal cells by increasing repair of DNA photoproducts (12-14). Additionally, α-melanocortin (α-MSH), synthesized by keratinocytes and melanocytes, and the keratinocyte derived endothelin-1 (ET-1) increase the survival and enhance repair of DNA damage in UV-irradiated HM (8, 11, 15-18). The significance of ET-1 in maintaining melanocyte homeostasis was demonstrated by selective ablation of ET-1 in murine epidermis in vivo, resulting in reduced number of melanocytes and compromised repair of DNA damage following UV exposure (19).

Exposure of various cell types, including HM, to UV activates the stress-activated MAP kinases JNK and p38 (20-22). The effects of JNK and p38 are mediated by activation of downstream transcription factors involved in the stress response, including p53, ATF-2, AP-1 and NF-κB, as well as USF-1 and MITF, known to be activated in HM (20, 23-25). Activation of

JNK and p38 augments nucleotide excision repair (NER), the main repair pathway of DNA photoproducts, which is critical for prevention of UV-induced skin cancers, including melanoma

(26-28). The role of paracrine/autocrine factors in enhancing DNA repair strongly suggests their photoprotective effects against UV-induced genotoxicity in epidermal cells, including melanocytes. Signaling through the endothelin B receptor (ETBR), the predominant ET-1 receptor expressed on HM, has been shown to include activation of the MAP kinases ERK1/2,

JNK and p38 (29), although ET-1 treatment has only been shown to activate ERK 1/2 in HM

(21).

The photoprotective effects of ET-1 have been reported (11, 19), but the mechanism(s) for these effects on HM are not been fully understood. We have been investigating the intracellular mechanism(s) activated by ET-1, which modulates the UV response. Our results define the significance of ET-1 activated signaling pathways in reducing the deleterious effects of UV via activation of JNK and p38.

2.3. Materials and Methods

Melanocyte culture: Primary HM cultures were each established from a single neonatal foreskin and maintained as described (10). Lightly-pigmented HM were used in all experiments to minimize the effect of variability in melanin content on the UV response. To test the effects of

ET-1, HM were maintained for 2-3 days before treatment and for the duration of each experiment in deprivation medium lacking bovine pituitary extract (BPE), source of melanocortins that stimulate cAMP formation (10), and 12-O-tetradecanoyl-phorbol-13-acetate

(TPA), which activates protein kinase C (PKC) and down regulates endothelin signaling (17, 30).

The only melanocyte-specific mitogen included was basic fibroblast growth factor (bFGF), which maintained the survival of HM without stimulating proliferation.

Irradiation of HM by UV: Cells were irradiated with a source of UV that has peak emission at

313 nM, as previously described (31).

Quantitation of UV-induced cyclobutane pyrimidine dimers (CPD): Unless otherwise stated,

HM were treated every other day for 4 days with 0 (control) or 1 nM ET-1 prior to and immediately following exposure to 90 mJ/cm2 UV. Melanocytes were harvested and fixed in 1% paraformaldehyde and 70% ethanol immediately after (0 or 3 h) to measure induction of CPD, and 48 h post UV to measure repair of CPD. Cells were immunostained using TDM-2 clone antibody (Cosmo Bio USA, Carlsbad, CA, USA) and goat-anti-mouse Alexa 488 antibody (Life

Technologies, Carlsbad, CA, USA), and analyzed by flow cytometry, as previously described

(31). To determine the effects of MAP kinases on UV-induced CPD induction and repair, HM were treated as described above, with additional groups treated with the JNK inhibitor SP

600125 (15 μM), or the p38 inhibitor SB 203580 (25 μM) (EMD Millipore, Billerica, MA,

USA). At least triplicate samples were included/group, and 10,000 events analyzed/sample.

Determination of UV-induced apoptosis by Annexin V staining: Melanocytes were untreated or treated for 4 days with 1 nM ET-1 prior to, and 24 h following exposure to 90 mJ/cm2 UV.

Cells were harvested, stained for Annexin V-APC (BD Biosciences, San Jose, CA, USA) and propridium iodide (PI), and analyzed by flow cytometry as previously described (11). The effect of MAP kinase inhibition on UV-induced apoptosis was determined by using the MAP kinase inhibitors, as described in the quantification of CPD. At least triplicate samples were included/group, and 10,000 events analyzed/sample.

Western Blot analysis: Activation of JNK, p38 and ATF-2 was determined by Western blotting.

Melanocytes were maintained for 2 days in deprivation medium, and then in unsupplemented

MCDB 153 for 18 h prior to, and following UV irradiation. Melanocytes were irradiated with 0 or 90 mJ/cm2 UV, and immediately treated with 0 or 1 nM ET-1. To determine the role of ET-1 signaling pathways in activation of JNK, p38 and/or ATF-2, HM were treated with 20 μM

BAPTA/AM (EMD Millipore), a potent calcium chelator, or 100 µM 2-APB (Cayman

Chemicals, Ann Arbor, MI, USA), an inhibitor of IP3, or 5 μM Bisindolylmaleimide I (EMD

Millipore), a pan PKC inhibitor, immediately following UV and/or ET-1 treatment. To determine the effects of the JNK inhibitor SP 600125 (15 μM) and the p38 inhibitor SB 203580 (25 μM),

HM were treated for 18 h prior to, and immediately after UV and/or ET-1 treatment with either inhibitor. Proteins were extracted 30 and 60 min post-treatment using RIPA buffer with protease and phosphatase inhibitors (Sigma-Aldrich, Saint Louis, MO, USA). Antibodies against phospho-ATF-2 (T69/71), phospho-p38 (T180/Y182), phospho-JNK (T183/Y185), phospho-

ERK1/2 (T202/Y204), phospho-MAPKAP2 (T222), phospho-MARCKS (S152/156), and the

48 respective total protein were purchased from Cell Signaling Technology (Danvers, MA, USA).

Phospho-c-Jun (S63) and total c-Jun antibodies were obtained from Cell Signaling Technology and Santa-Cruz Biotechnology (Santa Cruz, CA, USA), respectively. Tubulin and actin were detected as loading control, using HRP-conjugated antibodies purchased from Abcam

(Cambridge, MA, USA) and Santa Cruz Biotechnology, respectively. Membranes were incubated with the appropriate horseradish peroxidase-conjugated anti-rabbit or anti-mouse immunoglobin G (EMD Millipore) and detected by chemiluminescence. Detected bands were quantified by densitometry. The results of the 30 min time point are included in most figures since similar results were obtained at the 30 or 60 min time points. cAMP measurement: Melanocytes were maintained in deprivation media for 2 days, then treated with 0 (control) or 1 nM ET-1, or 10 µM forskolin, a known direct activator of adenylate cyclase, for 45 min. The levels of cAMP were determined in triplicate wells/group using a 125I- labeled radioimmunoassay kit (Perkin Elmer, Waltham, MA, U.S.A.), as previously described

(32-33).

Melanin content assay: HM were maintained in deprivation medium for 2 days, and then treated with 1 nM ET-1 every other day for a total of 4 days. Total melanin was determined as previously described (32).

Cell cycle analysis: Melanocytes were maintained in deprivation medium for 3 days, then treated with 0 or 1 nM ET-1, and harvested at 0, 6, 24, 48, and 72 h thereafter. Two hours before each time point, 10 μM bromodeoxyuridine (BrdU; Sigma-Aldrich) was added to each dish.

Melanocytes were harvested and fixed, permeabilized with 2N HCl + 0.5% Triton X-100, neutralized in 0.1 M sodium tetraborate + 0.5% Tween-20, incubated in blocking buffer, then

49 stained with mouse-anti-BrdU (Cell Signaling Technology) and goat-anti-mouse Alexa 488 antibodies (Life Technologies). Melanocytes were suspended in PBS containing 0.110 mg/ml

RNase and 6.85 µg/ml PI, and analyzed by flow cytometry. At least 10,000 events were analyzed/sample, with each group in at least triplicate. To determine the synergistic effect of ET-

1 and α-MSH on the cell cycle of HM, cells in deprivation medium were pre-treated with 10 nM

α-MSH, with or without 1 nM ET-1, every other day for 4 days prior to and immediately following UV irradiation. Melanocytes were harvested at 0, 24, 32 and 48 h post UV, stained and analyzed as described above.

Determination of melanocyte proliferation: To determine the effect of ET-1 on cell proliferation in deprivation medium, cells were treated with 0 or 1 nM ET-1 every other day for a total of four days. Cells were counted at the time of plating and 4 days thereafter, with each group plated in triplicate. To demonstrate that ET-1 signaling synergizes with cAMP signaling to drive HM proliferation, cells were treated with 0 or 1 nM ET-1 in deprivation medium containing BPE (source of α-MSH) every two days for a total of six days. Cells were counted at

2, 4 and 6 days, with triplicate dishes included in each group.

Gene expression analysis by quantitative real-time PCR (qRT-PCR): To investigate the transcriptional regulation of XPA and cyclin D1 by ET-1 and UV, melanocytes were plated and maintained for 3 days in deprivation medium. MC1R expression was also detected as a positive control, as ET-1 treatment upregulates MC1R expression (18). Melanocytes were irradiated with

90 mJ/cm2 UV, and then treated immediately with 0 or 1 nM ET-1. Cells were harvested at 8 and

24 h following UV and/or ET-1 treatment, pelleted and snap frozen in liquid nitrogen, before storage at -80°C. Total RNA was extracted using the RNeasy Mini Kit combined with the

RNase-free DNase system (Qiagen, Valencia, CA, USA). First-strand cDNA was synthesized

50 using 2.5 μg of total RNA using the SuperScript VILO Synthesis Kit (Life Technologies), and quantitative real-time reverse-transcriptase–PCR was performed using human XPA, cyclin D1,

MC1R and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) RT2 qPCR Primers and RT2

SYBR Green/Fluorescein PCR Master Mix (Qiagen). Amplification was performed using the

BIO-RAD iCycler iQ system (BioRad, Hercules, CA, USA). XPA, cyclin D1 and MC1R expression levels were normalization to GAPDH (34). Biological triplicates were processed individually and analyzed in technical triplicate. Expression levels were normalized to mean expression in control samples.

Statistical analysis: Data were statistically analyzed in GraphPad PRISM using one-way

ANOVA followed by the Student-Newman-Keuls post-hoc for multiple comparisons of sample means. Values were considered significant at p<0.05.

2.4. Results

ET-1 reduces DNA damage in UV-irradiated HM

The dose of UV 90 mJ/cm2, which induces significant CPD and ≤25% increase in apoptosis, was used in all experiments. Melanocytes established from different donors showed similar trends in response to UV and/or ET-1. In order to determine the appropriate treatment time with 1nM ET-1, HM were pre-treated for 0, 1, 2, or 4 days prior to, and immediately after

UV. Significant reduction in CPD induction at 0 h was only evident in the group treated with ET-

1 for 4 days, and repair (48 h post-UV) increased significantly with all different treatments with

ET-1, but was maximal in the group pre-treated for 4 days (Figure 2.S1A). To determine the kinetics of CPD in HM treated for 4 days prior to UV, the levels of CPD were compared at 0, 8,

24 and 48 h post UV and/or ET-1. Endothelin-1 significantly reduced the induction of CPD (at 0 h) and increased repair of CPD at 24 and 48 h, as compared to the respective UV groups (Figure

2.S1B). To determine the appropriate dose of ET-1, HM were treated with either 0.1 or 1 nM

ET-1 for 4 days prior to, and after UV exposure. There was a dose-dependent response of ET-on

CPD repair at 48 h post-UV, with only 1 nM ET-1 significantly reducing induction of CPD at the

0 h time point compared to UV 0 h (Figure 2.S2). Since pre-treatment for 4 days with 1 nM ET-1 caused significant reduction in CPD induction and maximal increase in CPD repair, this treatment protocol was used for measuring CPD in subsequent experiments. Treatment with 1 nM ET-1 for 4 days prior to UV exposure significantly reduced the induction of CPD, as compared to the UV group at 3 h, and further decreased the levels of CPD by 48 h (Figure 2.1A).

Endothelin-1 caused a marked reduction of apoptosis, consistent with our previous report on the survival effects of ET-1 on HM (Figure 2.1B)(11). The effect of ET-1 on CPD was independent of activation of the cAMP pathway, since 1 nM ET-1 had no effect on intracellular cAMP levels

(Figure 2.S3). Additionally, the effect of ET-1 on CPD was evident on HM expressing loss-of- function MC1R, by reduced induction and enhanced repair of CPD (Figure 2.S4). In all these experiments, there was no significant repair of CPD in the UV only groups 48 h post-irradiation.

The effects of ET-1 on DNA damage and apoptosis were independent of increased pigmentation, since 4 day treatment with 1 nM ET-1 did not cause any change in total melanin content (Figure 2.S5). Moreover, increased HM survival in response to 1 nM ET-1 was not due to HM proliferation, since no changes in the cell cycle or cell numbers were observed (Figure

2.S6A and B). Melanocytes showed an increase in proliferation with multiple treatments with 1 nM ET-1 in the presence of BPE, indicating that the signaling pathways of ET-1 and cAMP synergize together to drive HM proliferation (Figure 2.S6C). To validate these findings, HM were pre-treated for 4 days with 10 nM α-MSH, in the absence or presence of 1 nM ET-1, and immediately following UV irradiation (Table A1). Cell cycle analysis demonstrated that combined treatment with ET-1 and α-MSH increased the percent of HM in S phase at 24, as compared to treatment with α-MSH alone (control group). Similar results were observed at 32 h.

No significant difference was observed in all of the UV-irradiated groups, indicating that while

ET-1 and α-MSH treatment increased cell cycle progression in the absence of UV, it did not rescue HM from UV-induced cell cycle arrest (see Appendix AI, Table A1).

Treatment with ET-1 activates JNK and p38

We have reported that UV irradiation of HM induces the phosphorylation of the stress- activated MAP kinases p38 and JNK (21, 22). We found that treatment of HM for 30 or 60 min with 1 nM ET-1, in the absence of UV, markedly increased JNK and p38 phosphorylation

(Figure 2.S7). The effect of UV was remarkably less that of ET-1 (Figure 2.S7), and the

53 combined effect of ET-1 and UV exceeded that of ET-1 alone. UV activated the JNK and p38 target ATF-2 by inducing its phosphorylation on Thr 69 and 71 (Figure 2.S7) (35-37). Treatment with 1 nM ET-1 for 30 or 60 min profoundly stimulated the phosphorylation of ATF-2 compared to control, and augmented its phosphorylation by UV (Figure 2.S7). Using different HM strains, a similar trend in the effects of UV and ET-1 on phospho JNK, p38 and ATF-2 was observed

(Figures 2.2, 2.3, 2.S10). Exposure to UV had no effect, while treatment with ET-1 for 30 min robustly increased ERK1/2 phosphorylation (Figure 2.S8), as reported previously (21). Since UV did not affect ERK activity, we focused on the regulation of JNK and p38.

ET-1 induced activation of JNK and p38 is dependent on the Ca2+ signaling pathway

We investigated the significance of intracellular Ca2+ and the downstream kinase PKC, the two main signaling pathways for ETBR, on JNK and p38 activation (21, 38-40). We tested the effects of BAPTA/AM, a Ca2+ chelator; 2-APB, an inhibitor of inositol-(1, 4, 5)-triphosphate

(IP3), and Bisindolylmaleimide I, a pan PKC inhibitor. Treatment with 20 µM BAPTA/AM had a profound inhibitory effect on the ET-1-induced phosphorylation of JNK, ATF-2, and p38 activation (Figure 2.2A). To ensure that activation of JNK, p38 and ATF-2 was dependent on IP3 stimulated Ca2+ release from ET-1 signaling, the effects of Ca2+ inhibition (BAPTA/AM) were validated using 2-APB. Treatment with 100 μM 2-APB significantly inhibited ET-1 activation of

JNK, p38 and ATF-2 (Figure 2.2B). Additionally, PKC is another downstream kinase activated by ET-1 signaling, through DAG and Ca2+. To verify if PKC was involved in the activation of

JNK, p38 or ATF-2, 5 μM Bisindolylmaleimide I (a pan PKC inhibitor was used) was used.

Inhibition of PKC had no effect on UV or ET-1 activation of JNK, p38 and ATF-2 (Figure 2.2C).

To validate that the working concentration of PKC was sufficient for inhibiting PKC activity, phosphorylated MARCKS, a well-known downstream target of PKC, was investigated.

Significant inhibition of MARCKS phosphorylation was detected when HM were treated in the presence of 5 μM Bisindolylmaleimide I (Figure 2.S9C)(41).

JNK activation is required for ET-1-induced ATF-2 phosphorylation

Using pharmacological inhibitors of JNK and p38, we determined the significance of these kinases in mediating the effects of ET-1 on ATF-2 activation. Treatment with the JNK inhibitor SP 600125 (15 μM) reduced the levels of phospho-JNK and attenuated the phosphorylation of ATF-2 in HM exposed to ET-1 and/or UV (Figure 2.3A). Treatment with the p38 inhibitor SB 203580 (25 μM) inhibited the phosphorylation of p38, but not ATF-2, by UV and/or ET-1 (Figure 2.3B). The above concentrations of JNK and p38 inhibitors effectively inhibited the respective kinase activity, evidenced by abrogation of phosphorylation of the JNK substrate c-Jun and the p38 substrate MAPKAP2 by ET-1 (Figure 2.S9A and B). Treatment with the p38 inhibitor SB 203580 markedly increased the ET-1-induced JNK phosphorylation, however, treatment with the JNK inhibitor SP 600125 did not substantially increase the ET-1- induced phosphorylation of p38 (Figure 2.S10). These results suggest that the persistent phosphorylation of ATF-2 in ET-1 treated HM, despite inhibition of p38, is due to JNK activation.

Activation of both JNK and p38 mediate the reduction of CPD by ET-1, while JNK promotes survival

Inhibition of JNK had no effect on CPD levels in UV-irradiated HM at 3 or 48 h post UV in the absence of ET-1, yet abolished the effects of ET-1 on CPD repair (Figure 2.4A). Similarly, inhibition of p38 had no significant effect on the levels of CPD in UV-irradiated HM at 3 or 48 h, but totally inhibited the effect of ET-1 on CPD induction and repair (Figure 2.4B). Inhibition

55 of JNK phosphorylation increased UV-induced apoptosis by about 2.5 fold, and while the partially abrogates the inhibitory effect of ET-1 on UV-induced cell death (Figure 2.4C).

However, inhibition of p38 activation did not significantly alter UV-induced apoptosis or its reduction by ET-1 treatment (Figure 2.4D).

ET-1 has no effect on regulating gene expression of XPA or cyclin D1

In order to determine if ATF-2 plays a role in mediating the protective effects of ET-1,

HM were transfected with lentiviral vectors expressing shRNA against ATF-2. These attempts to silence ATF-2 in HM were unsuccessful, as the cells were unable to survive following transfection. Since ET-1 has been shown to impact DNA repair and cell cycle progression, the

ATF-2 gene targets XPA and cyclin D1 were selected for gene expression analysis using quantitative real-time reverse transcriptase-PCR (qRT-PCR). Results showed that UV irradiation severely downregulated mRNA expression of XPA or cyclin D1 at both 8 and 24 h, whereas treatment with 1 nM ET-1 had no effect on gene expression levels (see Appendix AI, Figure A2).

Previous studies have shown that UV irradiation downregulates (42), while 10 nM ET-1 treatment upregulates MC1R gene expression (18). Under our experimental conditions, 1 nM ET-

1 had no effect to increase MC1R, while UV exposure significantly reduced its gene expression

(see, Appendix AI, Figure A2).

2.5. Discussion

Understanding the DNA damage response of HM to UV, a major causative factor for malignant transformation of melanocytes (1, 2), is significant for identifying pathways that can be targeted for melanoma prevention. It is established that paracrine factors synthesized by epidermal keratinocytes regulate HM homeostasis (7-11, 13, 15, 18, 19). Of particular interest to us are the paracrine factors α-MSH and ET-1. We have reported that beside their melanogenic and mitogenic effects, α-MSH and ET-1 protect HM from UV-induced DNA damage and apoptosis (11, 17, 18, 43). The synthesis of ET-1 and α-MSH in the epidermis are increased upon

UV exposure, which underscores their physiological significance in the response of melanocytes to UV (8, 9, 43). In the present study, we investigated the effects of ET-1 and its signaling pathways on the DNA damaging effects of UV, mainly induction and repair of the DNA photoproduct, CPD, and apoptosis.

We found that ET-1 reduces the generation of CPD and enhances their repair, and inhibits apoptosis in UV-irradiated HM (Figures 2.1, 2.4, 2.S1, 2.S2, 2.S4). These effects were not dependent on increased melanin content or proliferation (Figures 2.S5, 2.S6). Reduction of

CPD in response to ET-1 was evident in HM expressing loss-of-function MC1R, suggesting that activation of the ETBR signaling pathway is sufficient for reducing CPD, independently of the cAMP pathway (Figure 2.S4). It was not surprising that 1 nM ET-1 did not stimulate HM proliferation in the deprivation medium used (Figures 2.S6A, 2.S6B), since mitogenic stimulation of HM requires activation of multiple signaling pathways, mainly PKC and cAMP, and unlike α-MSH, 1 nM ET-1 had no effect on cAMP formation (Figure 2.S3) (17, 18, 44). HM treated with ET-1 in the presence of BPE (source of melanocortins) increased proliferation, evident by increasing cell number over time (Figure 2.S6C). To validate that ET-1 and α-MSH

57 can synergize to drive HM proliferation (17-18), cell cycle analysis was performed and demonstrated that ET-1 and α-MSH increased entry into S phase at 24 and 32 h, but the combination did not rescue cells from UV-induced cell cycle arrest (see Appendix AI, Table

A1). We attribute the persistence of cell cycle arrest to DNA damage that was not fully repaired by 48 h following UV exposure.

The stress response to UV involves activation of the stress-activated MAP kinases p38 and JNK by dual phosphorylation (20), and subsequently, these kinases activate many targets, including the transcription factors ATF-2, p53, and USF-1 (20, 22, 35, 45). We previously reported that irradiation of HM with UV causes phosphorylation of the MAP kinases JNK and p38, but not ERK1/2 (21, 22). Here, we show that treatment of UV-irradiated or non-irradiated

HM with ET-1 caused robust phosphorylation of p38 and JNK, and that this activation was enhanced when were treated with ET-1 immediately following UV (Figures 2.2, 2.3, 2.S7,

2.S10). We detected phosphorylation of ATF-2, a known MAP kinase target involved in cellular stress responses, to validate activation of JNK and p38 (35-37). To our knowledge, we are the first to report that ATF-2 is activated in HM in response to UV or ET-1, and that its UV-induced activation is enhanced following ET-1 treatment (Figure 2.2, 2.3, 2.S7). We confirmed that while

1 nM ET-1 caused robust phosphorylation of ERK1/2, irradiation with 90 mJ/cm2 UV had no effect, (Figure 2.S8), in support of previous data suggesting that ERK1/2 contributes to HM proliferation in response to combined mitogenic signals (21). These results indicate that ET-1 enhances the activation of the UV stress response and participates in the maintenance of HM homeostasis.

The endothelin B receptor, the predominant ET-1 receptor expressed in HM, is a Gq protein-coupled receptor that rapidly activates phospholipase C-β (PLC-β), which hydrolyzes the

58 membrane-bound phosphatidylinositol-4’, 5’-biphosphate (PIP2) to diacylglycerol (DAG) and

2+ IP3 (18, 39, 40). Generation of IP3 stimulates the release of Ca from intracellular stores, and together with DAG, leads to activated PKC. Our data show that UV and ET-1-induced activation of JNK, p38 and ATF-2 occur through a Ca2+-dependent, PKC-independent signaling pathway

(Figure 2.2). Activation of JNK via a Ca2+-dependent, PKC-independent pathway that has been previously reported (48).

We utilized the pharmacological inhibitors of JNK and p38 to determine the role of the activated JNK and p38 in mediating the inhibitory effects of ET-1 on CPD and apoptosis in UV- irradiated HM. We found that activation of JNK and p38 by ET-1 is critical for reduction of UV- induced CPD (Figure 2.4A and B). While phosphorylation of JNK or p38 by ET-1 occurs within minutes, reduction of CPD requires prolonged treatment with ET-1, as shown in Figure 2.S1A, which might be required to sustain phosphorylation of JNK and p38, in order to activate downstream targets leading to activation of DNA repair pathways, mainly NER. Only inhibition of JNK, not p38, markedly reduced ATF-2 phosphorylation, suggesting that JNK is the primary kinase for ATF-2 in HM, and that p38 promotes DNA repair in HM by activating other transcription factors, such as p53 (Figure 2.3A and B)(22). Our results suggest that ATF-2 activation is not sufficient for enhancing CPD repair by ET-1, since inhibition of p38 compromised the effect of ET-1 on CPD repair despite ATF-2 phosphorylation. Other mechanisms by which p38 may enhance repair of CPD include chromatin remodeling, and inhibition of DDB2, a protein that facilitates the assembly of NER factors to initiate repair of

DNA photoproducts (27). Further studies are underway to identify other downstream targets of

JNK and p38 that are critical for the protective effects of ET-1 against UV-induced DNA damage and apoptosis.

Our data show that activation of JNK promoted survival of UV-irradiated HM (Figure

2.4C). In the presence of ET-1, JNK inhibition partially abrogated the inhibitory effect of ET-1 on UV-induced apoptosis, suggesting that JNK activation does not account for the entire protective effect of ET-1 on the survival of UV-exposed HM. Activation of other important pro- survival targets, such as Akt, by the ET-1/ETBR signaling pathway (11) may promote survival in the absence of JNK activation, serving as a possible compensatory mechanism. The anti- apoptotic effect of JNK is consistent with previous results showing that inhibition of JNK promoted UV-induced apoptosis and attenuated CPD repair in mouse embryonic fibroblasts (26).

Inhibition of p38 did not alter the survival effect of ET-1 (Figure 2.4D). It is conceivable that despite inhibition of p38 activity, other forms of DNA damage that compromise HM survival might be efficiently repaired, or that survival pathways were activated by alternative mechanisms. One mechanism by which JNK promotes repair of CPD and cell survival may involve activation of FOXO transcription factors, known to activate genes that are involved in

DNA repair and cell cycle regulation (46, 47). These genes include Gadd45, or code for antioxidant enzymes, such as manganese superoxide dismutase and catalase. Recently, FOXO1 was reported to interact directly with JNK, and that this interaction is critical for the transcriptional activity of FOXO1 and for its role in repair of DNA damage (47). Collectively, these results reveal the significance of p38 and JNK in maintaining genomic stability of HM by modulating their response to UV, which includes activation of a variety of transcription factors and regulation of various substrates that impact DNA repair and survival.

As previously mentioned, JNK and p38 mediate their effects by phosphorylating various downstream targets. ATF-2 has been shown to function as a transcription factor following mitogenic stimulation or cellular stress in a variety of different cell types (37-39). Silencing

60 experiments for ATF-2 were unsuccessful in HM, as the cells were unable to survive following transfection. qRT-PCR analysis of the potential ATF-2 target genes XPA and cyclin D1 were significantly reduced by UV, but immediate treatment with ET-1 had no effect to increase their expression levels (see Appendix AI, Figure A2), despite activation of ATF-2 (Figures 2.2, 2.3,

2.S7, 2.S8). This suggests that ATF-2 may regulate different genes in different cell types in the response to different stimuli (49-50). Since the protective effects of ET-1 required 4 days pre- treatment, this would suggest that changes in gene expression may also require additional treatment with ET-1, allowing for comparable conditions between measuring DNA damage repair, apoptosis, and gene expression levels. The role of ATF-2 in the UV response of HM remains to be investigated.

Crosstalk between p38 and JNK has been reported (20). We observed that inhibition of p38 by SB 203580 robustly increased ET-1 induced phosphorylation of JNK, but inhibition of

JNK by SP 600125 did not markedly affect phosphorylation of p38 (Figure 2.S10). Our results suggest that both JNK and p38 are required for ET-1 induced enhancement of CPD repair, and neither kinase seems dispensable for this effect. JNK is critical for survival following UV exposure, but plays a significantly lesser role in the survival effect of ET-1 (Figure 2.4C).

Inhibition of p38 resulted in increased activation of JNK, but this did not appear to enhance the effect of ET-1 mediated survival (Figures 2.4D, 2.S10A). Even in the absence of UV, ET-1 had a profound effect on JNK and p38 phosphorylation (Figures 2.2, 2.3, 2.S7, 2.S10). Given that the extent and duration of JNK and p38 activation determine their downstream effects, we conclude that ET-1, which interacted synergistically with UV to activate JNK and p38, modifies the response of HM to UV.

In this study, we investigated the role of ET-1 in the acute response of normal HM to UV.

Our data, which are summarized in Figure 2.S11 (see also Appendix AI, Figure A3), clearly show that ET-1 is photoprotective for HM, including HM that express LOF MC1R, and suggest that ET-1 can restore genomic stability of melanocytes by reducing the burden of UV-induced

DNA damage. These data, combined with our previous study (11), demonstrate the significance of ET-1 in preventing mutagenesis and malignant transformation. Our results provide further support for the role of paracrine factors, whose synthesis is stimulated by UV in the epidermis, in reducing the genotoxic effects of UV on melanocytes.

2.6. Acknowledgements: Supported in part by R01 ES017561 for ZAM and T32 ES007250 for

AMK and ZAM. We thank Drs. Ana Luisa Kadekaro, David Plas, and William Miller for valuable discussions; Drs. Ying Xia, William Miller, Ze’ev Ronai and Eric Lau for generously providing reagents; Drs. Philip Hexley and George Babcock for assistance in flow cytometry analysis; Kevin McFarland and Dorothy Supp for assistance in qRT-PCR sample processing and analysis; Joshua Jameson, Jared Swope, and David Abad for their technical assistance.

2.7. Authors’ Contributions

AMK, RK, VS, ZAM performed the research

AMK, VS, ZAM designed the research study

ZAM contributed essential reagents or tools

AMK, RK, VS, ZAM analyzed the data

AMK, ZAM wrote the paper

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2.9. Figures

Figure 2.1. ET-1 protects HM from UV-induced DNA damage and apoptosis. In (A), CPD induction and repair were determined at 3 h and 48 h post UV in HM that were untreated or treated with ET-1, and irradiated with 90 mJ/cm2 UV. Each data point is the mean of 3 h ± SEM of 12 independent experiments performed in 8 different HM strains. *, p < 0.001, statistically different from UV 3 h and 48 h; #, p < 0.001, statistically different from ET-1 + UV 3 h. In (B), apoptosis was measured by Annexin V-APC staining of HM that were treated as previously described. Data are expressed as percent increase in apoptosis above the respective control

(untreated or ET-treated HM). Each data point is the mean ± SEM of 9 independent experiments performed in 6 different HM strains. *, p < 0.001, statistically different from UV group.

Figure 2.2. Role of intracellular Ca2+ and PKC signaling pathways in mediating the activation of JNK and p38 and the downstream target ATF-2 by UV and ET-1. Western blot analysis was performed using HM that were UV-irradiated with 90mJ/cm2 and immediately treated with 0 or 1 nM ET-1 in the absence or presence of the Ca+2 chelator BAPTA/AM (20

μM) in (A), the IP3 inhibitor 2-APB (100 μM) in (B), or the pan PKC inhibitor

Bisindolylmaleimide I (5 μM) in (C). Results presented represent 30 min time point detection.

Actin served as the loading control for all total proteins. The numbers represent the densitometric values normalized by actin as a percentage of the experimental control. (BAPTA) =

BAPTA/AM, (2-APB) = 2-Aminoethoxydiphenyl borate and (Bis) = Bisindolylmaleimide I.

Figure 2.3. Significance of JNK and p38 for activation of ATF-2 in response to UV and ET-

1. Western blot analysis was performed using HM that were treated or untreated with the JNK inhibitor SP 600125 (15 μM) in (A) or the p38 inhibitor SB 203580 (25 μM) in (B), with or without exposure to UV, in the absence or presence of ET-1, as previously described in Materials and Methods. Results presented represent 30 min time point detection. Actin served as the loading control for all proteins. The numbers represent the densitometric values normalized by actin as a percentage of the experimental control. (SP) = SP 600125 and (SB) = SB 203580.

Figure 2.4. Effect of JNK or p38 inhibition on UV-induced CPD and apoptosis. CPD induction and repair were determined in HM that were pretreated with 0 or the JNK inhibitor

(SP) in (A) or the p38 inhibitor (SB) in (B), in the absence or presence of 1 nM ET-1 as previously described. Each data point is the mean percent of UV 3 h ± SEM of 3 independent experiments using 3 different HM strains. In (A) *, p < 0.01, statistically different from UV 3 h and 48 h; #, p < 0.05, statistically different from ET-1 + UV + SP 48 h. In (B), *, p < 0.001, statistically different from UV 3 and 48 h. #, p < 0.001, statistically different from ET-1 + UV +

SB at the respective time point. The role of JNK and p38 in modulating the apoptotic effect of

UV was determined by Annexin V staining at 24 h post UV. In (C), HM were treated with SP

600125 or in (D), with SB 203580. The data are expressed as increase in apoptosis above the respective unirradiated control (untreated, ET-1, inhibitor, inhibitor +ET-1). Each data point is the mean ± SEM of at least triplicate samples/group. In (C) *, p < 0.001, statistically different from UV or UV + SP; #, p < 0.001, statistically different from ET-1 + UV + SP. In (D), *, p <

0.001, statistically different from UV or UV + SB. Similar results were obtained in three independent experiments using three different HM strains.

2.10. Supplemental Figures

Figure 2.S1. Effect of duration of ET-1 pre-treatment on CPD induction and repair, and the kinetics of CPD repair in ET-1 treated HM. In (A), HM were treated with 0 or 1 nM ET-1 for 0, 1, 2 or 4 days prior to and immediately following irradiation with 90 mJ/cm2 UV. Cells were harvested at 0 h and 48 h, and analyzed as previously described. Each data point is the mean percent of UV 0 h ± SEM of at least triplicate samples/group. **, p < 0.001, statistically different compared to UV 0 h and 48 h; #, p < 0.05, statistically different compared to 0.1 nM

ET-1 + UV 48 h; ^^, p < 0.01, statistically different compared to 1 nM ET-1 + UV 0h. In (B),

HM were treated with 0 or 1 nM ET-1 every 48 h for a total of 4 days prior to and immediately following UV exposure. Cells were harvested at 0 h, 8 h, 24 h and 48 h, and analyzed as previously described. Each data point is the mean percent of UV 0 h ± SEM of at least triplicate samples/group. #, p < 0.001, statistically different compared to UV 0 h and 8 h; ^^, p < 0.05, statistically different compared to UV 48 h; *, p < 0.001, statistically different compared to UV time point; ^, p < 0.05, statistically different compared to ET-1 + UV 24 h.

Figure 2.S2. Dose-dependent response of ET-1 on CPD induction and repair. HM were treated with 0, 0.1 nM or 1 nM ET-1 every 48 h for a total of 4 days prior to and immediately following UV 90 mJ/cm2 exposure. Cells were harvested at 0 h and 48 h, and analyzed as previously described. Each data point is the mean percent of UV 0 h ± SEM of at least triplicate samples/group. **, p < 0.001, statistically different compared to UV 0 h and 48 h; ^, p < 0.05, statistically different compared to 1 nM ET-1 no pre-treatment + UV 0 h; *, p < 0.01, statistically different compared to UV 48 h; #, p < 0.001, statistically different compared to UV 0 h and 8 h;

^^, p < 0.05, statistically different compared to UV 48 h.

Figure 2.S3. Effect of 1 nM ET-1 on cAMP production. HM were maintained in deprivation media for 2 days, as previously described. Cells were treated with 0 (control) or 1 nM ET-1 for

45 min. Treatment with 10 μM forskolin served as a positive control. Each data point is the mean percent of control ± SEM of 2 independent experiments with triplicate samples/group. *, p <

0.001, statistically different compared to control or 1 nM ET-1.

Figure 2.S4. Effect of ET-1 on CPD induction and repair in HM expressing LOF MC1R.

HM (MC1R genotype R151C/R160W) were treated with 0 or 1 nM ET-1, 10 nM α-MSH or 10

μM forskolin every 48 h for a total of 4 days prior to and immediately following UV 90 mJ/cm2 exposure. Cells were harvested at 0 h and 48 h, and analyzed as previously described. Each data point is the mean percent of UV 0 h ± SEM of 2 independent experiments with at least triplicate samples/group. *, p < 0.001, statistically different compared to UV 0 h and 48 h; ^, p < 0.01, statistically different compared to ET-1 + UV 0 h or For + UV 0 h; **, p < 0.05, statistically different compared to UV 0 h.

Figure 2.S5. Effect of ET-1 on HM pigmentation. Total melanin content of four lightly- pigmented HM strains derived from individual donors was determined and is presented as mean

μg/106 cells ± SEM of triplicate samples/group.

Figure 2.S6. Effect of ET-1 on HM proliferation. To determine if ET-1 treatment impacted cell proliferation, BrdU analysis was performed as described in Material and Methods. In (A),

Cells were irradiated and then treated with 0 or 1 nM ET-1 in deprivation medium. Samples were harvested at 0, 6, 24, 48 and 72 h following UV and/or ET-1 as described. Each data point is the mean percentage of cells in G0/G1, S or G2/M phases ± SEM of 2 independent experiments with at least triplicate samples/group. To validate that the ET-1 treatment protocol used for CPD and apoptosis analysis did not impact cell proliferation, melanocytes were treated with 1 nM ET-1 in derivation medium and counted. In (B), cells were treated every two days for a total of four days with 0 or 1 nM ET-1 in deprivation medium. Samples were counted at day 0 and day 4 (n = triplicate samples/group). Each data point is the mean number of cells ± SEM. To demonstrate that ET-1 synergizes with α-MSH to drive melanocyte proliferation, melanocytes were plated and treated in media containing BPE (source of melanocortins). In (C), HM were treated every two days for a total of six days with 0 or 1 nM ET-1 in medium containing BPE (but lacking

TPA). Samples were counted at day 2, day 4 and day 6 (n = triplicate samples/group). Each data point is the mean number of cells ± SEM. *, p < 0.001, statistically different to Control 6 d; ^, p

< 0.01, statistically different to ET-1 4 d.

Figure 2.S7. Activation of the MAP kinases JNK and p38 and their downstream target

ATF-2 by UV and ET-1. In (A), Western blot analysis was performed using HM that were treated as described in Materials and Methods. Actin served as the loading control for all

86 proteins. Results presented represent 30 and 60 min time point detections. (B-D) represent densitometry of JNK, p38, and ATF-2, respectively, whereby the ratio of phospho/total protein is calculated after normalizing each band to the loading control.

Figure 2.S8. Effect of UV and ET-1 on ERK activation. Western blot analysis was performed using HM that were treated as described in Materials and Methods. The upper tubulin bands served as the loading control for phosphorylated and total ERK. The lower tubulin bands served as the loading control for phosphorylated and total ATF-2. Results presented represent 30 min time point detection.

Figure 2.S9. Verification of efficacy of JNK, p38 and PKC pharmacological inhibitors. To determine efficacy of the pharmacological inhibitors for JNK, p38 and PKC, specific targets were detected by Western blot analysis. In (A), phosphorylated and total c-Jun, a known substrate of JNK, was detected in HM treated as previously described in Figure 3A. In (B), phosphorylated and total MAPKAP2, a known substrate of p38, was detected in HM treated as previously described in Figure 3B. In (C), phosphorylated and total MARCKS, a known

89 substrate of PKC, was detected in HM treated as previously described in Figure 2C. Results for

(A), (B) and (C) presented represent 30 min time point detection.

Figure 2.S10. Reciprocal effects of MAP kinase inhibition on activation of JNK and p38.

(A) Western blot analysis was performed to determine the effect of p38 inhibition on JNK activation, as previously described in Figure 3B. (B) Western blot analysis was performed to determine the effect of JNK inhibition on p38 activation, as previously described in Figure 3A.

Results for (A) and (B) represent 30 min time point detection.

Figure 2.S11. Schematic representation of the effects of ET-1 on the UV response of HM.

Exposure of human skin to UV increases ET-1 synthesis by epidermal keratinocytes and induces

DNA damage and apoptosis. In HM, UV activates the stress-activated MAP kinases JNK and p38, and their target substrate ATF-2. ET-1 is known to bind and activate ETBR expressed on

2+ the cell surface of HM, leading to increased intracellular Ca concentration (from IP3 generation), and activation of PKC. The Ca2+ signaling pathway of ETBR leads to activation of

JNK and p38, and as we have previously reported, of ERK1/2, known to be involved in mitogenic signaling . Phosphorylation of JNK by ET-1 results in profound activation of ATF-2.

Activation of JNK and p38 by ET-1 reduces the induction and enhances the repair of UV- induced DNA damage, and JNK promotes HM survival. Collectively, these results strongly suggest that ET-1, via activating JNK and p38, protects HM from the genotoxic effects of UV and promotes their genomic stability.

CHAPTER 3

CONCLUSION

3. Conclusion: Major Findings, Summary, Future Directions and Significance

3.1. Major Findings

1. ET-1 is an important paracrine factor for MCs that plays a critical role in the UV

response by reducing generation and enhancing repair of CPD, and reducing apoptosis in

UV-irradiated human MCs, and these effects contribute to the maintenance of MC

genomic stability.

2. The effects of ET-1 are mediated by activation of the MAP kinases JNK and p38.

3. Increased intracellular Ca2+ release, an important signaling pathway for the activated

ETBR, is critical for JNK and p38 activation.

4. JNK and p38 activation are necessary for mediating the protective effects of ET-1 on

reducing generation and enhancing repair of CPD. JNK activation seems to be necessary,

but not sufficient, for mediating the survival effect of ET-1.

5. The effects of JNK and p38 on CPDs are independent of stimulation of melanin synthesis

or proliferation.

6. While the mitogenic and melanogenic effects of ET-1 require the synergistic interaction

with other signaling pathways, mainly cAMP, the effects of ET-1 on DNA photoproducts

and survival do not require activation of the cAMP signaling pathway.

7. ET-1 activates ATF-2, a downstream target of JNK and p38, in human MCs, mainly

through JNK activation. The role of ATF-2 in the UV response remains to be elucidated

in human MCs.

8. ET-1 compensates partially for LOF MC1R in UV-irradiated human MCs, since effects

of ET-1 on reducing CPD and increasing survival are independent of melanin synthesis.

3.2. Summary

This study is the first to elucidate, in detail, the mechanism by which ET-1 mediates its photoprotective effects on UV-irradiated human MCs through the activation of the MAP kinases

JNK and p38. UV radiation is considered the major cause of skin cancer, including malignant melanoma, the most fatal form (Elwood and Jopson, 1997; Jhappan et al., 2003; Miller and

Mihm, 2006). MCs are unique in that they are resistant to apoptosis, yet they live for decades in the epidermis (Quevedo et al., 1969; Plettenberg et al.,1995). Since MCs have a low proliferative capacity in vivo (Quevedo et al., 1969) and are routinely exposed to UV, maintaining their genomic stability is critical for prolonged cutaneous photoprotection and prevention of skin cancer, mainly melanoma. Understanding the molecular mechanisms activated in normal human

MCs that reduce the genotoxic effects of UV and prevent malignant transformation is critical for reducing the incidence of melanoma. This thesis provides new insight into the role that ET-1, a

KC-derived paracrine factor, plays in protecting human MCs from the photodamaging effects of

UV via activation of the stress-induced MAP kinases JNK and p38.

The first described effects of ET-1 on human MCs were stimulation of proliferation and melanogenesis (Imokawa et al., 1992; Hara et al., 1995; Imokawa et al., 1995, 1996; Tada et al.,

1998). Later, ET-1 was reported to have survival effects on MCs, by inhibiting the apoptotic effects of UV (Kadekaro et al., 2005). Here, we report that ET-1 reduces UV-induced CPDs by inhibiting their induction, as well as enhancing their repair (Figures 2.1A, 2.S1, 2.S2). Our results identified, in detail, the duration of treatment with ET-1 that is required for reduction of

CPD and the kinetics of CPD repair in ET-1 treated MCs (Figures 2.S1, 2.S2). MCs that express

LOF MC1R have increased vulnerability to UV due to compromised DNA repair (Hauser et al.,

2006; Kadekaro et al., 2010). ET-1 treatment in MCs with LOF MC1R had reduced induction

96 and enhanced repair of UV-induced CPD (Figure 2.S4), indicating that ET-1 helps to compensate for the loss of functional MC1R and that the effects of ET-1 are not mediated by increased cAMP production, but by increased Ca2+ mobilization. These findings strongly suggest that the survival effects of ET-1 on MCs are due to reduction of the genotoxic effects of UV, and that ET-1 plays a significant role in the prevention of mutagenesis that results from unrepaired

DNA damage. Accordingly, we conclude that ET-1 participates in the protection against malignant transformation of normal human MCs to melanoma. This highlights ET-1 as a component of a paracrine network that plays an important role in maintaining the genomic stability of MCs.

Increased melanogenesis in response to UV serves as a major photoprotective mechanism

(Gilchrest and Eller, 1999; Eller and Gilchrest, 2000; Brenner and Hearing, 2008). Under the experimental conditions we used, treatment with 1 nM ET-1 did not increase melanin content

(Figure 2.S5), yet reduced UV-induced CPD. This suggests that ET-1 activates DNA repair pathways, independently of increasing melanin content. The increase in the survival of UV- irradiated MCs in response to ET-1 treatment is due to inhibition of apoptosis, as determined by

Annexin V staining (Figure 2.1B), and is not due to increased MC proliferation. Under our experimental conditions, 1 nM ET-1 had no mitogenic effect on MCs, since our results showed no increase in the percentage of cells in S phase or in cell number in response to ET-1 treatment

(Figure 2.S6A and B). Therefore, unlike the mitogenic and melanogenic effects of ET-1 that require synergistic interaction with the cAMP pathway (Swope et al., 1995; Tada et al., 1998;

Halaban, 2000), signaling of ETBR is sufficient for the effects of ET-1 on CPD induction and repair, and MC survival. This is further supported by the effects of ET-1 on MC with LOF

MC1R, suggesting that ET-1 can partially compensate for the comprised DNA repair in these

97 cells. Our results with ET-1, together with previously reported effects of other paracrine factors, such as α-MSH, NGF and vitamin D3, on the extent of UV-induced DNA damage and apoptosis in MCs underscore the significance of the epidermal paracrine network in reducing the deleterious effects of UV and protecting MCs from mutagenesis (Zhai et al., 1996; Tada et al.,

1998; Böhm et al., 2005; Dixon et al., 2005; Kadekaro et al., 2005, 2010; Dixon et al., 2011;

Kadekaro et al., 2012; Swope et al., 2014).

One of the major signal transduction pathways activated in response to UV is the MAP kinase pathway (Weston and Davis, 2002; Bode and Dong, 2003; Wagner and Nebreda, 2009; López-

Camarillo et al., 2012). Activation of JNK and p38 occurs following various external stimuli, like UV, while ERK activation occurs following mitogenic stimulation (Tada et al., 2002; Roux and Blenis, 2004; Berridge, 2012). Tada et al. (2002) demonstrated in human MCs that UV- irradiation alone activated the stress-activated MAP kinases, namely JNK and p38, while it had no effect on activation of ERK. Treatment with 0.1 nM ET-1 was shown to activate ERK but no apparent effect on activating JNK or p38 was noted under the experimental conditions used

(Tada et al., 2002). Our results demonstrate that 1 nM ET-1 alone, a concentration that is 10 fold higher than that used by Tada et al., (2002), has a significant effect on the activation of JNK and p38, and that ET-1 treatment immediately following UV exposure results in further enhanced activation of these two kinases (Figures 2.2, 2.3, 2.S7). Since ET-1 activates JNK and p38, it is important to identify the downstream substrates of these MAP kinases that can potentially be targeted for melanoma prevention.

The effects of ET-1 on MCs are mediated by activating multiple signaling pathways (Saldana-

Caboverde and Kos, 2010; Rosanò et al., 2013), mainly through intracellular Ca2+ mobilization and PKC (Imokawa et al., 1992, 1995, 1996). These pathways have been shown to mediate the

98 effects of ET-1 on MC proliferation and melanogenesis, including activation of ERK 1/2 and the downstream targets p90rsk and CREB (Tada et al., 2002). Using pharmacological inhibitors of

2+ Ca (BAPTA/AM), IP3 (2-APB) and PKC (Bisindolylmaleimide I) revealed that increasing intracellular Ca2+, but not PKC, is required for activation of JNK and p38 by ET-1 (Figure 2.2).

Since PKC does not play a role in activating JNK and p38 in human MCs, it is critical to identify which upstream kinases are activated in response to increased Ca2+ mobilization, allowing them to target and phosphorylate JNK and p38.

JNK and p38 play important roles in the signal transduction pathways that coordinate cellular responses to stress, such as cell proliferation and survival by phosphorylating downstream transcription factors (Wagner and Nebreda, 2009). ATF-2 is considered an early response transcription factor following various cellular stressors, including UV, and is a target of JNK and p38 (Hayakawa et al., 2004; Bhoumik et al., 2007; Lopez-Bergami et al., 2010; Lau and Ronai,

2012). Our results are the first to reveal in human MCs that UV and ET-1 activate ATF-2, and that this activation is enhanced when MCs are treated with ET-1 immediately following UV exposure (Figures 2.2, 2.3, 2.S7). Here, we showed that JNK is the major contributor to ATF-2 activation in human MCs following UV exposure or ET-1 treatment, while p38 had no effect on

ATF-2 phosphorylation (Figure 2.3). ATF-2 activation did not seem to be sufficient for mediating the effects of ET-1 on CPD in UV-irradiated MC, as inhibition of p38 abrogated the effects of ET-1 on CPD induction and repair (Figure 2.4B), despite activation of ATF-2.

JNK and p38 have been reported to be involved in UV-induced NER. Activation of JNK increases the expression of XPF following UV exposure and enhances CPD repair in MEFs

(Christmann et al., 2006, 2007). p38 activation in UV-irradiated human FBs has been shown to enhance NER by augmenting CPD recognition and regulating chromatin relaxation (Zhao et al.,

2008; Wang et al., 2013). Pharmacological inhibitor studies of JNK or p38 reveal that ET-1 mediates its protective effects to inhibit CPD formation and promote DNA repair through the activation of JNK and p38 (Figure 2.4 A and B). Our observation that inhibition of p38 compromises the effect of ET-1 on CPD repair without impacting ATF-2 phosphorylation suggests that ATF-2 activation is not sufficient for enhancing CPD repair by ET-1 (Figure 2.4B), suggesting the involvement of other transcription factors, such as p53, which has a known role in

DNA repair and apoptosis and whose activation is p38-dependent in UV-irradiated HM (Latonen and Laiho, 2005; Kadekaro et al., 2012). These findings identify JNK and p38 as important players in mediating the protective effects of ET-1 against UV exposure in human MCs, particularly their involvement in DNA repair.

Activation of JNK and p38 have been found to be both pro- and anti-apoptotic, indicating their complex roles in the response to UV (Chen et al., 1996; Wada and Penninger, 2004; Liu and Lin,

2005; Bivik and Ollinger, 2008; Dhanasekaran and Reddy, 2008; Wagner and Nebreda, 2009). In regards to cell survival, as determined by UV-induced apoptosis, JNK plays an role in MC cell fate (pro-survival) while p38 appears to have no effect on UV-induced apoptosis (Figure 2.4C and D). Although JNK promotes survival in UV-irradiated MCs, our results suggest that JNK participates in, but is not sufficient for the survival effects of ET-1 on human MCs (Figure 2.4C).

Our results suggest that JNK and p38 affect different targets, which explains their differential effects on survival. This may be due to activation of other targets that promote survival, such as

Akt, by ET-1/ETBR signaling (Kadekaro et al., 2005), which may serve as a compensatory mechanism when JNK is inhibited. These findings identify that JNK promotes survival in UV- irradiated MCs, but may not be the major player in mediating the pro-survival effects of ET-1 on

UV-induced apoptosis in human MCs.

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3.3. Significance of the study to MC biology and melanoma

UV is now recognized as the major etiologic factor for melanoma, therefore it is important to elucidate how MCs respond to UV. An important component of the UV response in the skin is increased synthesis of paracrine factors. ET-1 is an important element of the cutaneous paracrine network, as previous studies have illustrated protective effects of ET-1 in MCs exposed to UV.

Here, our study focuses specifically on the role of ET-1, in the absence of other protective factors

(i.e, α-MSH, SCF, vitamin D3, etc), allowing us to precisely elucidate the signaling mechanism activated by ET-1 and ETBR in response to UV. The fact that ET-1 was able to convey photoprotection in human MCs demonstrates the importance of the ET-1 signaling pathway, especially since ET-1 treatment reduced DNA damage in MC expressing LOF MC1R, whose defects result in decreased pigmentation, increased DNA damage and increased ROS generation.

This suggests that ET-1/ETBR may be a significant pathway used by MCs to repair UV-induced

DNA damage and promote their survival. The significance of the ET-1 pathway in conveying photoprotection occurs through the activation the MAP kinases, specifically JNK and p38, whose activation is important for enhancing DNA damage repair in UV-irradiated MCs. Our results underscore the significance of these MAP kinases in the response of human MCs to UV exposure. The results hereby presented support the role of ET-1 in counteracting the genotoxic effects of UV on human MCs. Understanding the DNA damage response of MCs to UV, and how this pathway can be modulated by ET-1, is significant for melanoma prevention, as it provides new targets for prevention strategies.

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3.4. Future Directions

To fully understand the role of ET-1 in UV-irradiated human MCs, it will be necessary to determine the downstream targets of the stress-activated MAP kinases JNK and p38 that allow

ET-1 to convey its photoprotective effects. Detecting the transcription factor(s) that is/are activated in response to ET-1, as well as critical target genes, will allow for complete elucidation of the signal transduction pathway activated by UV and enhanced by ET-1. p53 is a known target of JNK and p38, and has been shown to play a protective role by enhancing DNA damage repair and reducing oxidative stress (Tan et al., 1999; Smith et al., 2000; Budanov et al., 2004; Yoon et al., 2004; Bensaad et al., 2006; Kadekaro et al., 2012) as well as enhances melanogenesis

(Khlagatian et al., 2000; Nylander et al., 2000; Cui et al., 2007), which supports the significance of p53 in human MCs and suggests that future studies include p53 in future investigations to understand the DNA repair mechanisms activated by ET-1 through the MAP kinases JNK and p38. Other potential transcription factor targets for JNK and p38 include the early response genes c-Fos and c-Jun (Wagner and Nebreda, 2009). As previously mentioned, loss of c-Fos in MEFs results in deficient repair of UV-induced photolesions (Christmann et al., 2006, 2007), making this an interesting target to possibly investigate. JNK has also been shown to activate the

Forkhead box O (FOXO) transcription factors, which function to regulate cell proliferation,

DNA repair and survival (Lam et al., 2006; Huang and Tindall, 2007; Klagge et al., 2011; Ju et al., 2014). Ju et al. (2014) reported that FOXO1 and JNK interact directly, and that this interaction is important for FOXO1’s transcriptional activity, as well as its role in DNA damage repair. Whole transcriptome shotgun sequencing (RNA-sequencing aka RNA-seq) of MCs treated with UV and/or ET-1 would also allow for identification of critical players that are involved in the DNA damage response and repair. Since ET-1 activates JNK and p38, it is

102 important to identify the downstream substrates of these MAP kinases that can potentially be targeted for melanoma prevention.

JNK and p38 are activated by numerous upstream kinases, namely MAP3K and MAP2K, following external environmental stress (Weston and David, 2002; Wagner and Nebreda, 2009).

Although we were able to show the significance of the Ca2+-dependent mechanism by ET-1, it remains to be determined which kinase(s) is/are responsible for their activation. A potential protein kinase to be investigated in the activation of JNK and p38 would be apoptosis signal- regulating kinase-1 (ASK1), a MAP3K which has been shown to be an upstream activator of both JNK and p38 signaling pathways following stress responses (Ichijo et al., 1997; Miyazono et al., 2001; Matsukawa et al., 2004). It has been shown in vitro that activation of ASK1 occurs following UV irradiation of human corneal epithelial cells (Wang and Lu, 2007). Another upstream protein kinase to possibly investigate is Ca2+/calmodulin dependent kinase (CaM-K), which has been shown to activate JNK and p38, as well as ATF-2 (Enslen et al., 1996), and has also been shown to be activated following UV exposure (Wright et al., 1997). Determining the kinase(s) responsible for activating JNK and p38 would help to chart the signaling pathway, in order better understand the regulation of their activation and activity.

Given the complex paracrine network that is activated in the skin by UV (Chakraborty et al.,

1996; Gilchrest et al., 1996; Yamaguchi et al., 2007; Berridge, 2012), it is important to determine if other paracrine factors (i.e., α-MSH, NGF, vitamin D3) that have signaling pathways different from ET-1, are also capable of activating the MAP kinases JNK and p38. This will help determine if JNK and p38 are globally involved in enhancing DNA repair and regulating survival of MCs.

103

Since ET-1 was found to reduce formation of CPD following UV irradiation, it would be interesting to consider the effects of ET-1 on chromatin remodeling. This process transmits DNA damage signals within cells, facilitates access of repair pathway proteins to DNA damage sites, and is mediated by enzymatic complexes, like SWI/SNF, that destabilize histone-DNA associations in an ATP-dependent manner (Halliday et al., 2009). It has been reported that three components of the NER pathway, namely RPA, XPA and XPC, stimulate the remodeling activity of the SWI/SNF complex (Hara and Sancar, 2002), allowing for the excision and removal of 6,4-

PP lesions but appeared to have no effect on CPD (Hara and Sancar, 2003). Zhao et al. (2009) demonstrated that knockdown of Brg1, the ATPase subunit of the SWI/SNF complex, prevented repair of CPDs, but not 6,4-PPs in UV-irradiated FBs. Due to these conflicting studies, it is important to thoroughly investigate the role of SWI/SNF complex in NER. Studying this complex and chromatin remodeling in the UV response of human MCs would allow us to understand how paracrine factors may impact the DNA repair process.

While UV is the main etiological factor for melanoma, there is also evidence that oxidative stress, which can be UV dependent or independent, plays a significant role in melanoma development (Prota, 2000; Song et al., 2009; Kadekaro et al., 2012; Mitra et al., 2012; Denat et al., 2014; Meierjohann, 2014; Panzella et al., 2014). As previously mentioned, UV exposure induces oxidative stress in human MCs due to ROS production (Kadekaro et al., 2005; Song et al., 2009). ET-1 plays a critical role in the UV-induced oxidative stress response of human MCs.

Kadekaro et al. (2005) reported that ET-1 profoundly reduces UV-generated ROS, as detected by

H2O2 formation. As of yet, there have been no additional reports investigating the role of ET-1 in oxidative stress (antioxidant response, repair of oxidative DNA damage, etc.), suggesting future studies that should be performed. Additionally, both JNK and p38 have been found to be

104 activated and involved in the oxidative stress response, including following UV exposure

(Iordanov and Magun, 1999; Katiyar et al., 2001; Aggeli et al., 2006; Gutiérrez-Uzquiza et al.,

2012; Kadekaro et al., 2012). It would be interesting to elucidate the activation and signaling pathways of JNK and p38 in human MCs exposed to UV or H2O2 that have been treated with the antioxidant N-acetylcysteine (NAC). Pharmacological inhibition of p38 and JNK would allow us to understand the role JNK and p38 play in the oxidative stress response.

Melanoma is the most serious type of skin cancer, as it is an aggressive disease that is very challenging to treat. With its incidence on the rise, there is a need to find clinical applications that protect epidermal cells from the detrimental effects of UV by inhibiting photocarcinogenesis. It is known that complex signaling pathways are activated in human MCs in response to UV exposure, and the importance of the paracrine signaling network to modulate this response via enhancing DNA damage repair and reducing of ROS generation, thus promoting genomic stability and MC survival. The two major paracrine factors that have been shown to promote MC survival following UV exposure are α-MSH and ET-1. One strategy for melanoma prevention has been to test analogs of α-MSH, which activate the MC1R signaling pathway, leading to increased melanin synthesis, enhanced DNA repair, and reduction of apoptosis (Abdel-Malek et al., 2006; Barnetson et al., 2006), but only in individuals who express functional MC1R. Our finding sheds light on the signaling mechanism that promotes photoprotective effects of ET-1 to reduce DNA damage, and that this protection occurs even in

MC that express LOF MC1R, as ET-1 conveys its effects through ETBR. Hyter et al. (2013) demonstrated the significance of ET-1 in DNA damage repair and MC survival following UV, and additional in vivo studies of the effects of ET-1 should be performed by using MC1R-/- mice to validate these findings. Development of topical applications (similar to that of the α-MSH

105 analogs) may serve as a prophylaxis to convey protection against UV exposure, especially for individuals who lack the ability to response to α-MSH. The challenge with developing ET-1 therapeutics is that expression of ETBR is expressed in a variety of different cells types and is not selectively expressed in human MCs.

Our study demonstrates that ET-1 protects MC against UV by promoting DNA repair through the activation of p38 and JNK. Additionally, JNK activation participates, but is not sufficient, for the effect of ET-1 to inhibit apoptosis in UV-irradiated MCs. Using mouse models where various isoforms of JNK and/or p38 are knocked out would allow for comparison of in vitro and in vivo results, as well as determine the specific roles of the various MAP kinase isoforms in promoting

DNA damage repair and MC survival. These kinases may function as potential therapeutic targets that can be manipulated to promote DNA repair, however they may prove difficult to develop. UV mediated signal transduction pathways are complex. The cell type, dose of UV and the activation of other signaling pathways, as well as the level and duration of kinase activation can all influence the how these kinases function to control various cellular responses. Given that underlying molecular mechanisms are complex, inhibiting these pathways presents a real challenge. Pharmacological inhibitors have been used to look at the effects of JNK or p38 both in vitro and in vivo. There are significant limitations to performing silencing experiments in vitro, due to the various isoforms expressed of JNK and p38. Genomic and proteomic strategies will help identify new MAP kinase substrates and regulators, while molecular signatures found using microarray data and sequencing should identify new biomarkers and potential therapeutic targets.

Additionally, the use of mouse models, in combination with human genetic and pharmacological approaches will help better define the specific roles of JNK and p38.

106

Based on our findings, ET-1 is able to promote DNA damage repair in UV-irradiated human

MCs via activation of the MAP kinase signaling pathways, specifically through JNK and p38. In regards to survival, p38 appears to have no effect, while it is still unclear what contribution JNK plays in promoting the anti-apoptotic effect of ET-1 following UV exposure in MCs. This suggests that ET-1 may mediate MC survival through the activation of a different signaling pathway. A major signaling pathway involved in promoting cell survival is the PI3K/Akt signaling pathway. Kadekaro et al. (2005) reported that treatment with ET-1 in UV-irradiated human MCs activated this pathway, by increasing activation of Akt. This activation was shown to phosphorylate the pro-apoptotic protein Bad (Kadekaro et al., 2005), thus inhibiting its ability to bind to other pro-survival Bcl-2 family members like Bcl-2 and Bcl-xL. Akt has also been shown to phosphorylate other promoters of apoptosis, like caspase 9 (Cardone et al., 1998). This suggests that ET-1 may be able to promote survival by regulating the caspase signaling cascade, and recommends that future studies look at this effect in human MCs exposed to UV.

Additionally, Akt has been shown to phosphorylate the FOXO family of transcription factors, specifically FOXO3, inhibiting its ability to increase expression of pro-apoptotic proteins

(Brunet et al., 1999; Dijkers et al., 2002; Cifarelli et al., 2012; Zhang et al., 2012).

Phosphorylation of FOXO1 by JNK has been shown to have a protective role in promoting DNA damage repair (Huang and Tindall, 2007; Ju et al., 2014). These roles of the FOXO genes may be isotype and stimuli dependent in response to different kinases. This suggests that future studies look at the regulation of FOXO family members (i.e., which kinases activate or inhibit these transcription factors) and how ET-1 influences their expression of target genes. Finally, the

PI3K/Akt signaling pathway has been shown to be involved in NF-κB signaling, particularly by phosphorylating the NF- κB inhibitor IKK (Kane et al., 1999; Romashkova and Makarov, 1999),

107 allowing NF-κB to translocate to the nucleus and promote survival, by regulating expression of pro-apoptotic genes, like Bcl-xL and capsase inhibitors (Barkett et al., 1999; Karin and Lin,

2002). ET-1 Additionally, ET-1 can also activate the MAP kinase ERK 1/2, which can promote

MC proliferation and survival (Imokawa et al., 1996; Tada et al., 2002). One downstream target of ERK 1/2 is microphthalmia associated transcription factor (MITF), a transcription factor that is a master regulator of melanocyte function and survival (Veis et al., 1993; McGill et al., 2002;

Levy et al., 2006), and has been linked to expression of Bcl-2 and other pro-survival genes, both in vitro and in vivo (McGill et al., 2002; Kadekaro et al., 2005; Sato-Jin et al., 2008; Hornyak et al., 2009; Hartman and Czyz, 2014). Hornyak et al. (2009) demonstrated that in vivo MITF expression inhibits UV-induced apoptosis independently of melanin synthesis. Kadekaro et al.

(2005) showed that treatment with 0.1 nM ET-1 activated MITF, while Bcl-2 expression remained unchanged, both in the absence and presence of UV. This suggests that the role of ERK

1/2 in activating MITF and the regulation of Bcl-2 expression in promoting ET-1 mediated survival following UV exposure should be elucidated under similar experimental conditions used in this study, including the use of pharmacological inhibitors against ERK and MITF.

Activation of the MAP kinases JNK and p38 are necessary to promote the ET-1 mediated effect to enhance DNA damage repair in UV-irradiated MCs. In regards to ET-1 promoted survival, the

MAP kinases do not play a clear role in mediating this effect. This suggests that ET-1 promotes two different mechanisms, where activation of the DNA damage repair pathway is independent of the pro-survival pathway. The implication of specifically inhibiting the DNA repair pathway would cause a significant amount of mutagenesis, due to inability to repair DNA photoproducts.

Accumulation of these DNA lesions in a pro-survival state would significantly contribute to cellular transformation and potential oncogenesis. Inhibition of survival would significantly kill

108 cells, and activation of the DNA repair pathway may not compensate for that effect. This indicates that both arms of the ET-1 signaling mechanism are equally important in mediating genomic stability and maintaining survival in MCs exposed to UV. ET-1 is a ubiquitously expressed protein and plays multiple roles in normal physiology (Rubanyi and Polokoff, 1994).

Broad inhibition of ET-1 would have significant consequences in various physiologic systems, whereas dermal inhibition of ET-1 would result in a significant decrease in survival of MCs, as demonstrated in vivo by Hyter et al. (2013). Additionally, antagonists of ETBR have also resulted in decreased survival in MCs (Kadekaro et al., 2005; Hyter et al., 2013) and in melanoma cells (Lahav et al., 1999; Bagnato et al., 2003), both in vitro and in vivo. This study demonstrates the significance of ET-1 in MC maintenance post-UV, and suggests the possibility of therapeutic manipulation of these pathways. However, it is not currently understood how all paracrine factors produced in the epidermis work together to establish the full range of receptor- mediated cellular responses in response to UV. This suggests that additional studies are necessary to further elucidate the epidermal events that occur in response to UV. Understanding how paracrine factors work as photoprotectors to reduce DNA damage and prevent cellular transformation of epidermal MCs is vital for improved treatment of UV-induced skin cancers.

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APPENDIX

150

AI: Additional Data for Chapter 2.

Table A1. Synergistic effect of ET-1 and α-MSH on melanocyte proliferation

HM were maintained in deprivation medium for at least 3 days and were treated with 10 nM α-

MSH (control), with or without 1 nM ET-1, every other day for 4 days prior to irradiation with 0 or 90 mJ/cm2 UV, and immediately thereafter for 0, 24, 32 or 48 h. Samples were stained for

BrdU incorporation and analyzed for cell cycle. Data presented are the mean percent of HM in

G0/G1, S or G2/M phases (triplicate samples/group).

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diately with 0 0 with diately

1; #, p < 0.05, statistically statistically 0.05, < p #, 1;

Melanocytes were were and Melanocytes plated

1 and UV. UV. and 1

. The data was GAPDH a data as The . using normalized

genes by genes

1, statistically different compared to Control to or different ET compared statistically 1,

MC1R

and

1; ***, p < 0.00 < p ***, 1;

cyclin D1 cyclin

, ,

each gene were normalized to control and mean relative expression levels are presented presented were expression levels relative normalized are gene mean and each control to

XPA

^, p < 0.01, statistically different compared to Control to different ET compared or statistically 0.01, < p ^,

, as described in Materials and Materials in as , Methods described

PCR

- SEM.

1. Total RNA was isolated 8 and 24 h after treatment, and equal amounts of RNA from each group and were treatment, 24 from equalgroup amounts and 8 RNA Total 1. each isolated RNA of after h was

Figure A2. Regulation of Figure imme irradiated Cells treated and mJ/cm2 90 with days. 2 UV deprivation in for were maintained media nM 1 ET or qRT by analyzed for Levels gene. housekeeping change fold as Control to ET or compared different

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Figure A3. Revised summary of study findings.

(A) is a schematic representation of the activation of the MAP kinases p38 and JNK, and their downstream target ATF-2, by UV and ET-1. UV exposure activates the MAP kinases p38 and

JNK via upstream MAP3K and MAP2K, outlined in Figure 1.6. ET-1 binds to ETBR, signaling through the Gαq protein, activating PLC-β to cleave PIP2. This cleavage results in generation of

IP3 and the membrane bound DAG. IP3 binds to specific receptors on the surface of the ER, promoting endogenous Ca2+ release. Together, DAG and Ca2+ can activate PKC. JNK and p38 are activated in a Ca2+-dependent, PKC-independent pathway. Activation of JNK is the major contributor to ATF-2 activation in human MCs in response to UV and/or ET-1.

(B) is a summary of results on DNA damage (induction and repair), survival and ATF-2 activation in human MCs following UV exposure and/or ET-1 treatment. Pharmacological inhibitor studies were also performed to determine the role of JNK and p38 in DNA damage and survival following exposure to UV, in the absence or presence of ET-1.

MCs. Exposure of human skin to UV induces DNA damage and apoptosis, as well as increases the synthesis of ET-1 by epidermal keratinocytes. Through the signaling pathway discussed in part (A), UV and ET-1 are able to activate the MAP kinases JNK and p38. Pharmacological inhibitor studies, as described in part (B), revealed that both JNK and p38 participate in the DNA damage repair mediated by ET-1. Whereas p38 had no effect on survival, JNK appears to have a role in UV-mediated apoptosis, but plays a minor role in mediating the protective effect of ET-1 on MC survival.

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