Differential Processing/Degradation of Melanosomes by Epidermal Keratinocytes

University of Cincinnati

Date: 1/6/2011

I, Jody P. Ebanks , hereby submit this original work as part of the requirements for the degree of Doctor of Philosophy in Pharmaceutical Sciences/Biopharmaceutics.

It is entitled: Differential Processing/Degradation of Melanosomes by Epidermal Keratinocytes

Student's name: Jody P. Ebanks

This work and its defense approved by:

Committee chair: R. Randall Wickett

Committee member: Tomohiro Hakozaki, PhD

Committee member: Raymond Boissy

Committee member: Pankaj Desai

Committee member: Ana Luisa Kadekaro

Committee member: Gerald Kasting

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Last Printed:1/13/2011 Document Of Defense Form

Differential Processing/Degradation of Melanosomes

by Epidermal Keratinocytes

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 Division of Pharmaceutical Sciences of the James L. Winkle College of Pharmacy

Jody Patria Ebanks

2011

B.S. Biochemistry, Boston College, Chestnut Hill, MA, 2005

Committee Chair: R. Randall Wickett, Ph.D.

ABSTRACT

The synthesis and processing of melanosomes, the pigmented organelles of the follicular and interfollicular epidermis, is of major interest in the field of cutaneous biology. Additionally, clarification of the biological and cellular processes of cutaneous pigmentation has several therapeutic and cosmetic based applications, including the alleviation of skin hyperpigmentation.

Modification of skin complexion coloration has traditionally been accomplished by inhibition of the rate limiting enzyme of melanogenesis, tyrosinase, or attenuation of melanosome transfer from melanocytes to keratinocytes. The post transfer modification of pigmented melanosomes, the main focus of this dissertation project, provides an attractive and distinct avenue of modulating skin pigmentation. There is currently limited information on how epidermal keratinocyte process recipient melanosomes during terminal differentiation. Furthermore, the variability of melanosomal degradation seen between light and dark skin remains to be clearly established. Therefore, we have developed a novel model system to investigate the degradation of isolated melanosomes by cultured human keratinocytes. Fluorescently labeled and isolated melanosomes, cultured in the presence of light and dark skin derived keratinocyte cultures, were assessed for degradation. The extent of degradation has been qualitatively assessed, using transmission electron microscopy and indirect immunofluorescence with confocal microscopy, and quantitatively assessed using flow cytometry analysis. Within 48 hours of melanosome incorporation, indirect immunofluorescence and confocal microscopy images suggest that light derived keratinocytes may have accelerated melanosome degradation compared to dark keratinocytes. This time dependent decrease in fluorescence was then quantitatively analyzed using flow cytometry analysis. Consistent with the results of the confocal analysis, over a 48 hour time frame, light keratinocytes appear to degrade melanosomes more efficiently than the

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dark skin derived keratinocytes, P=0.039. This methodology offers a novel mechanism to address the differential ability of light and dark keratinocytes to degrade melanosomes.

To further delineate the process of melanosome degradation between light and dark skin, we focused on hydrolytic enzymes that have been implicated in epidermal differentiation and potentially melanosome degradation. To investigate this, we performed preliminary microarray analysis on suprabasal epidermis derived from light and dark skin, by laser capture microscopy

(LCM). Data analysis of the microarray experimentation showed over-expression of various hydrolytic enzyme genes in the suprabasal epidermal layers, when comparing light to dark skin.

Western blot analysis performed to confirm the expression pattern of hydrolytic enzymes from either light or dark skin derived epidermal lysates, demonstrated that cathepsin L2 was reproducibly upregulated in light skin, P=0.048. In addition, immunofluorescence analysis of cathepsin L2 in light and dark foreskin cryosections confirmed this differential expression and demonstrated that this enzyme was expressed throughout the epidermal layers. Biochemical analysis of cathepsin L2 activity in the two complexion types confirms an elevated enzyme activity in light compared to dark skin complexion samples, 1.75 fold higher activity in light skin compared to dark skin, P=0.03. Taken together these results confirm the differential expression of the acid hydrolase cathepsin L2 in light and dark skin at the gene and protein level. These results may have identified a specific acid hydrolase that may play a role in melanosome degradation and pigment processing.

ACKNOWLEDGEMENTS

In everything I do, I give God thanks for the gifts of talent, persistence, knowledge, and faith that He and only He is capable of endowing unto me.

I express the upmost appreciation and love to my Mother and Father for their continual support and belief in me. The sacrifices you have made for me will always be appreciated. I thank my

Dad for instilling in me, the value of hard work and dedication. To my Mom, you are the major driving force for me to work harder, your love for me has been steadfast and my love for you cannot be overstated.

To my sisters Paige and Alicia you both amaze me at how loving, smart and supportive you continually are. You girls have consistently motivated me to be a good role model and to be more loving. I am proud to be your sister and I cannot wait to attend your graduations! To my brother Patrick, you bring a smile to my face in even the most difficult of situations. I love you more than you will ever know.

To my family, I thank you all for supporting me along this journey. I especially thank Debbie and Romeash for their constant words of encouragement and persistent support, you have both constantly believed in me and I thank you for this.

I would like to extend my deepest appreciation to Dr. Raymond Boissy for being my mentor for the past five years. Your insight and critical evaluation has been invaluable. I thank you for the vi

opportunity to be a part of the Boissy laboratory and allowing me to develop as a student and as a scientist. Special thanks to Dr. R. Randall Wickett and Dr. Gerald Kasting for accepting me into the Cosmetic Science program and supporting me every step of the way throughout this journey. Dr. Wickett, you have shared your exceptional knowledge in cosmetic and skin sciences, laying the foundation for my work. Dr. Kasting, I thank you for challenging me to think critically, a fundamental skill required for success in our field. To Dr. Pankaj Desai, I appreciate your feedback and help throughout this dissertation! To Dr. Tomohiro Hakozaki, I thank you for giving me the chance to work on this research project. Your feedback, critical thoughts, and consistent help were vital to the development of this project. To Dr. Ana Luisa Kadekaro, I cannot explain how much appreciation I have for everything you have done for me for the past several years. Your support and consistent willingness to help me was invaluable, your scientific advice was indispensable, and your kindness unfailing.

I would be remiss not to mention the institutions that have provided me with the education and preparation to accomplish this degree. First, I am indebted to Procter and Gamble for providing the support and resources to make this work possible. To the University of Cincinnati, College of

Pharmacy/College of Medicine, and to the Graduate School as a whole, I send my appreciation for offering the curriculum and resources necessary for me to complete this degree. My deepest appreciation goes out to Boston College for providing the foundation of my education and teaching me to think critically and how to work diligently.

To the BGSA, in particular Amber Evans, Adeola Adeyemo, Tiffany Forde, Vanessa Saunders,

Ritch Hall, Teisha Murray, Fedoria Rugless, Sakinah Davis, Eme Amba and of course Dr.

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Bradford Mallory. You guys have become my family over the last few years and I am grateful for every single one of you and for everything we have accomplished together.

To my friends in the College of Pharmacy, Jennifer Karr, Rania Ibrahim, Jennifer Davis, Shoná

Burkes, Allison Rush, Rachna Gajjar, Amit Kulkarni, Kelly Smith, Divya Samineni, Ganesh

Moorthi, Terri Lacount, Shruthi Vaidhyanathan, Poonam Chopra, Sarah Ibrahim, John Anneken,

Ganesh Mugundu, Ned Berry, Nirmal Bhide, Nimitha Dave, Nicole Roenker, Courtney Huff,

Guarav Tolia, Cheryl Minges. You have all made up a very dynamic group that has helped me in so many ways. To Laura and Greg Temming, you guys have been extremely good friends and a constant support system; I will never forget that! Thanks to Lola Kelly-Smalls for your help as a mentor and friend. Special thanks to the staff in the College of Pharmacy, including but not limited to, Marcia Silver, Donna Taylor and Paula Shaw, I thank you for your assistance and friendly conversations. Dr. Carol Caperelli I thank you for all your help with enzyme kinetics and biochemical assays, I believe Michaelis and Menten would be pleased. I also send my appreciation to Dr. Marty Visscher for all her words of encouragement over the years.

Special thanks to Sandy Schwemberger and Andrew Osterburg for all your help with the flow cytometer. Thank you to Birgit Ehmer for all your help with the various microscopy techniques.

Your expertise was of great assistance and I appreciate everything you have done to assist me along this journey. I would also like to say a big thank you to Amy Albin for her assistance with the TEM.

I am grateful to the wonderful people in the members of the UC Department of Dermatology. I could not have picked a better group of people to work with and eat cake with every month. Dr.

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Zalfa Abdel-Malek, Dr. Diya Mutasim, Dr. Viki Swope, Renny Starner, Dr. Yuki Hashimoto,

Dr. Elodie LePape, Amy Roberto, Meghan Lee, Nancy Olson, Jennifer Yang, Josh Jameson,

Jared Swope, Christie Alexander, Priya Srivastava, Heather Henry, and Nicole Mosby. I also appreciate all the technical help from, and friendly conversations with, the dermatopathology ladies: Cindy Dewar, Jade Williams, and Sue Schaefer. Finally, Anne von Koschembahr thanks for being such a sweet friend over the last few months. I expect big things from you.

To the many wonderful and down-to-earth dermatologists: Dr. Mary Jo Kerns it was a pleasure working with you, Dr. Kerith Spicknall thanks for all your assistance especially with my various questions, Dr. Dave Crowe it was great working with you, I really appreciate you treating me as a friend and your help with my dermatologic questions, and of course Dr. Farah Abdulla for being my sister from another mother. You have been such a great friend and your advice has truly gotten me through the last few years!

To the members of the Boissy laboratory I send my deepest appreciation, for all the kindness and support. To Amy Koshoffer, I honestly and truly consider you and your family as part of my family. I would not be where I am today without your help and your kindness. I especially thank you for your great insight into the experimentation and your advice not only about science, but about life as a whole. It has been a great honor to be your colleague. To Adnan Mir and Howard

Epstein it was a pleasure to work with you guys. To Erika Osterholzer, Katherine Schmidt and

Allison Chalupa, it has been a great pleasure to work with you all; it is amazing how talented you girls are at such a young age. I am so proud of the work you all have accomplished and I cannot

wait to see how many things you all have in store for the future. To Bradley King, it has been a great pleasure to work with you for the last few months, I thank you for all the help you have given me with my work and I expect great things from you in medical school. To Gordon Shott, thank you for all the help, you really made the lab work flow a bit smoother.

To the Harneys, I cannot possibly thank you enough for your support and love over the last several years. I particularly appreciate you guys giving me a place to stay when I came back to

MA, the visits to the “yellow submarine” in Orlando, and of course Nana’s Pizzelles! Especially to Timothy Harney Jr., you pushing me to find the strength in myself to finish this degree was an invaluable experience.

To my closest friends: Angela Conant, you have been a pillar of strength for me since day one and even though we have been apart for so long, you have been a constant support system for me. To Douglas Ciarfella, it is hard to explain how important your friendship has been to me, I honestly don’t think I could have made it here without your advice and support. To the ladies of

Boston College Mod 22a: Katie Cronin, Danielle Hedderson, Laura Belden, Anne McLaughlin,

Christine Mulligan and Kara Brielmann/Sarah Burnie –Wishart included. You girls have been some of the greatest friends anyone could ever hope for. Every single one of you has been an inspiration to me in so many ways.

To the most beautiful island in the world JAMAICA, I hope I have made my country proud.

Dedication: This dissertation is dedicated to my mother, Amaui Ebanks, and father, Patrick Ebanks Sr., for their never-ending love and support.

TABLE OF CONTENTS

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ABSTRACT iii-iv ACKNOWLEDGEMENTS vi-x DEDICATION xi TABLE OF CONTENTS 1-4 LIST OF TABLES 5 LIST OF FIGURES 6-7 LIST OF ABBREVIATIONS 8-12 CHAPTER 1: INTRODUCTION 13 1.1. Human Skin Anatomy 14 1.1.1. Dermis and Hypodermis 15 1.1.2. Epidermis 16 1.1.2.1. Stratum Basale 17 1.1.2.2. Stratum Spinosum 18 1.1.2.3. Stratum Granulosum 18 1.1.2.4. Stratum Corneum 19 1.1.3. Pigmentary System of the Epidermis 20 1.1.3.1. Melanocytes and Skin Pigmentation 21 1.1.3.2. Melanosome Biogenesis 21 1.1.3.3. Mechanisms of Melanosome Transfer and Degradation 23 1.2. Lysosomes and Lysosome Related Organelles 23 1.2.1. Melanosomes as Specialized Lysosomes 24 1.2.2. Epidermal Lysosomal Hydrolases 25 1.2.2.1. Cysteine Cathepsins 26 1.2.2.1.1. Cathepsins L2 (Cathepsin V) 27 1.2.2.1.1.1. Cathepsin L2 Structure and Catalysis 28 1.2.2.1.1.2. Cathepsin L2 Inhibition 29 1.2.2.1.2. Cathepsins B 30 1.2.2.1.2.1. Cathepsin B Structure and Catalysis 31 1.2.2.2. Cathepsin D 32 1.2.2.3. β-3-glycosyltransferase-like 33 1.2.2.3.1. β-3-glycosyltransferase-like Structure and Catalysis 34 1.2.2.4. Prostatic Acid Phosphatase 35 1.2.2.5. β-glucuronidase 36 1.2.2.5.1. β-glucuronidase Structure and Catalysis 37 CHAPTER 2: HYPOTHESES AND SPECIFIC AIMS 40 CHAPTER 3: OBJECTIVES 43 CHAPTER 4: EPIDERMAL KERATINOCYTES FROM LIGHT VERSUS DARK SKIN EXHIBIT DIFFERENTIAL DEGRADATION OF MELANOSOMES 45 4.1. Abstract 46 4.2. Introduction 47

4.3. Results 49 4.3.1. Distinct Racial Skin Sources Demonstrate Variable Melanosome Degradation 49 4.3.2. Fluorochrome Labeling of Melanogenic Cells and Melanosome Isolation 50 4.3.3. Keratinocytes Uptake and Degrade Isolated Melanosomes 51 4.3.4. Degradation of Fluorochrome Labeled Melanosomes by Human Keratinocytes in an In Vitro Model System 52 4.3.5. Quantitative Analysis of Melanosome Degradation by Human Keratinocytes in an In Vitro Model System 54 4.4. Discussion 56 4.5. Materials and Methods 60 4.5.1. Cell Culture 60 4.5.2. Melanosome Isolation/Purification 60 4.5.3. Electron Microscopy 60 4.5.4. Pulse Incorporation of Melanosomes by Keratinocyte Samples 61 4.5.5. Assessing Keratinocyte Rate of Division 61 4.5.6. Indirect Immunofluorescence of Keratinocytes Using Confocal Microscopy 61 4.5.7. Flow Cytometry 61 4.5.8. Description of Statistics 61 4.6. Conflict of Interest 62 4.7. Acknowledgements 62 4.8. Figures 63 4.9. Supplementary Figures 70 4.10. Supplementary Materials and Methods 74 4.10.1. Cell Culture 74 4.10.2. Melanosome Isolation/Purification 75 4.10.3. Electron Microscopy 76 4.10.4. Pulse Incorporation of Melanosomes by Keratinocyte Samples 77 4.10.5. Assessing Keratinocyte Rate of Division 78 4.10.6. Indirect Immunofluorescence of Keratinocytes Using Confocal Microscopy 78 4.10.7. Flow Cytometry 79 CHAPTER 5: HYDROLYTIC ENZYMES OF THE INTERFOLLICULAR EPIDERMIS DIFFER IN EXPRESSION AND CORRELATE WITH THE PHENOTYPIC DIFFERENCE OBSERVED BETWEEN LIGHT AND DARK SKIN 81 5.1. Summary 82 5.2. Introduction 84 5.3. Materials and Methods 85 5.3.1. Cryosectioning of Human Foreskin Tissue for Indirect Immunofluorescence 85 5.3.2. Indirect Immunofluorescence of Skin Cryosections 86 5.3.3. Laser Capture Microscopy and Microarray Analysis 86 5.3.4. Isolation of Human Epidermis 87

5.3.5. Immunoblotting 87 5.3.6. Biochemical Assay 88 5.3.7. Immunogold Electron Microscopy 89 5.3.8. Statistical Analysis 89 5.4. Results 89 5.4.1. Light and Dark Skin Differentially Express Lysosomal Hydrolases in Suprabasal Epidermis 89 5.4.2. Immunoblot Analysis of Lysosomal Hydrolases in Epidermal Tissue 90 5.4.3. Indirect Immunofluorescence of Lysosomal Hydrolases in Epidermal Cryosections 91 5.4.4. Immunogold Labeling of Cathepsin L2 in Light and Dark Epidermal Tissue 92 5.4.5. Biochemical Analysis of Cathepsin L2 Activity 93 5.5. Discussion 94 5.6. Acknowledgments 96 5.7. Figures and Tables 97 CHAPTER 6: MECHANISMS REGULATING SKIN PIGMENATION: THE RISE AND FALL OF COMPLEXION COLORATION - A REVIEW 106 6.1. Abstract 107 6.2. Introduction 107 6.3. Transcriptional Regulation of Melanogenic Enzymes 110 6.4. Post-translational Modification of Melanogenic Enzymes 112 6.5. Attenuation of Tyrosinase and Related Melanogenic Enzymes Catalytic Activity 114 6.5.1. Hydroquinone 114 6.5.2. Monobenzylether 116 6.5.3. Arbutin and Deoxyarbutin 116 6.5.4. Mequinol 117 6.5.5. N-Acetyl-4-S-Cysteaminylphenol 118 6.5.6. Kojic Acid 119 6.5.7. Azelaic Acid 119 6.5.8. Gentisic Acid 120 6.5.9. Flavonoid-like Agents 120 6.5.10. Aloesin 121 6.5.11. Hydroxystilbene 122 6.5.12. Licorice Extract 123 6.5.13. Antioxidants and Redox Agents 124 6.5.14. L-Ascorbic Acid and Magnesium-L-Ascorbyl-2-Phosphate 124 6.5.15. Alpha Tocopherol and Alpha Tocopherol Ferulate 125 6.6. Interruption of Melanosome Transfer 126 6.6.1. Centaureidin and Methylophiopogonanone B 126 6.6.2. Niacinamide 127 6.6.3. PAR-2 Inhibitors 128 6.6.4. Lectins and Neoglycoproteins 130

6.7. Acceleration of Epidermal Turnover and Desquamation 131 6.7.1. α-Hydroxyacids 131 6.7.2. Salicylic Acid 132 6.7.3. Linoleic Acid 132 6.7.4. Retinoids 133 6.8. Conclusions 134 CHAPTER 7: CONCLUSIONS AND FUTURE RECOMMENDATIONS 135 7.1. Conclusions 136 7.2. Future Recommendations 137 REFERENCES 139 APPENDIX 175 AI. Supplemental Data to Chapter 4 176 AI.1. Structure of CFDA-SE 176 AII. Supplemental Data to Chapter 5 177 AII.1. Differentiation and Immunoblot Analysis of Keratinocytes 177 AII.2. Immunoblot Analysis of Cath D in Undifferentiated and Differentiated KC Lysates 179 AII.3. IIF of Acid Hydrolases in Epidermal Cryosections 179 AII.4. Supplemental Analyses and Calculations for Biochemical Assay 184 AII.4.1. Generating AMC Standard Curve 184 AII.4.2. Determining Substrate Concentration for Biochemical Assay 185 AII.4.3. Calculating Specific Activity 186

LIST OF TABLES

Table Page 5.1. Concentrations and dilutions of components utilized for western blot analysis. 97 5.2. Microarray analysis of suprabasal epidermis from light and dark skin. 98 5.3. Immunogold labeling of cathepsin L2 in light and dark skin. 99 A.1. Velocity of light and dark epidermal sample cleavage of ZLR-AMC. 186 A.2. Specific activity of light and dark epidermal samples. 187

LIST OF FIGURES

Figure Page 1.1. Schematic diagram detailing skin anatomy. 15 1.2. Various stages of melanosome maturation. 22 1.3. Schematic diagram of human cathepsin L2. 29 1.4. Schematic diagram of Cathepsin B. 32 1.5. Stereo ribbon schematic of β-GLU monomer . 38 4.1. Transmission electron micrographs of biopsies from light and dark skin. 63 4.2. SKMEL-188 labeled with CFDA-SE and MEL-5. 64 4.3. Incorporation and degradation of fluorescently labeled melanosomes by light and dark skin keratinocytes. 65 4.4. Keratinocytes incorporate and degrade fluorescently labeled SKMEL-188 melanosomes. 67 4.5. Ratio of baseline to 48 hour fluorescent intensities quantitatively analyzes melanosome degradation. 68 4.6. Keratinocytes incorporate and degrade fluorescently labeled normal human melanosomes. 69 4.S1. Transmission electron micrographs of SKMEL-188 melanosomes. 70 4.S2. Ultrastructure of melanosomes after 48 hours in keratinocyte cultures. 71 4.S3. Melanosomal perinuclear cap formation in the keratinocyte cytoplasm. 72 4.S4. Proliferation rate of light and dark keratinocytes with incorporated melanosomes. 73 5.1. Immunoblot analysis of key acid hydrolases completed in four light skin and dark skin derived epidermal lysates. 100 5.2. Indirect immunofluorescence of four distinct dark skin and light skin cryosections. 102 5.3. Immunogold electron microscopy of Cath L2 in epidermis of light and dark skin. 104

6.1. Process of melanogenesis within epidermal melanosomes. 109

A1. Chemical structure of the succinimidyl ester of carboxyfluorescein diacetate,

used to form a fluorescent dye-protein adducts with melanosomes. 176

A2. Differential expression of involucrin and p63 in differentiated KC lysates. 178

A3. Cath D expression in undifferentiated and differentiated KC lysates. 179

A.4. IIF of Cath L2 in four distinct DS and LS Samples . 180

A.5. IIF of Cath B in four distinct DS and LS Samples. 181

A.6. IIF of Cath D in three DS and four LS Samples. 182

A.7. IIF of B3GTL in two distinct DS and LS Samples. 183

A.8. Standard curve of various AMC concentrations versus RFU 184

LIST OF ABBREVIATIONS

α-MSH α-melanocyte stimulating hormone

α-TOC Alpha tocopherol

α-TF Tocopherol ferulate

β-GLU β-glucuronidase

AA Antibiotic/antimycotic

AHA α-hydroxyacids

AHP aminohydroxyphenylalanine

AP Acid phosphatase

ACPP Prostatic acid phosphatase

Arg Arginine

ASA L-Ascorbic Acid

Asn Asparagine

Asp Aspartic Acid

AZA Azelaic acid

B3GTL β-3-glycosyltransferase-like (also B3GALTL)

BBI Bowman-Birk protease inhibitor

BFGF Basic fibroblast growth factor

BPE Bovine pituitary extract

BSA Bovine serum albumin

Cath B Cathepsin B

Cath D Cathepsin D

Cath L Cathepsin L

Cath L2 Cathepsin L2

CFDA Carboxyfluorescein diacetate,

CFDA-SE Carboxyfluorescein diacetate, succinimidyl ester

CHS Chediak-Higashi syndrome

CREB cAMP-responsive element binding protein

CSC Constitutive skin color

Cys Cysteine

DCT/TYRP2 Dopachrome tautomerase/tyrosinase related protein-2

DA Deoxyarbutin

DHI 5,6-dihydroxyindole

DHICA 5,6-dihydroxyindole-2-carboxylic acid

DIC Differential interference contrast

DKC Dark keratinocyte

DMEM Dulbecco’s modified eagle medium

DOPA 3,4-dihydroxyphenylalanine

DS Dark skin

ECL Enhanced chemiluminescence

ECM Extracellular matrix

ER Endoplasmic reticulum

ERK extracellular-signal related kinase

FACS Fluorescence activated cell sorting

FBS Fetal bovine serum

FSC Facultative skin color

GA Glycolic acid

GERL Golgi-endoplasmic reticulum-lysosome

Glu Glutamic Acid

GSK3b glycogen synthase kinase-3b

HEPES N-2-Hydroxyethylpiperazine-N’-2-ethanesulfonic acid

His Histidine

HPS Hermansky-Pudlak syndrome

HQ Hydroquinone

IEM Immunogold electron microscopy

IIF Indirect immunofluorescence

IL-6 Interleukin-6

IRB Institutional review board

KA Kojic acid

KC Keratinocyte

KHG Keratohyalin granules

Kojyl-APPA 5-[(3-aminopropyl)phosphinooxy]-2-(hydroxymethyl)-4H-pyran-4-one

LA Lactic acid

LAMP-1 Lysosome associated membrane protein -1

LAMP-2 Lysosome associated membrane protein -2

LCM Laser capture microscopy

LKC Light keratinocyte

LPA Lysophosphatidic acid

LRO Lysosome-related organelle

LS Light skin

M6P Mannose-6-phosphate

MPRs Mannose-6-phosphate receptor

MAP Magnesium-L-ascorbyl-2-phosphate

MAPK Mitogen-activated protein kinase

MBEH Mono benzyl ether of hydroquinone

MC1R Melanocortin 1 receptor

Md X Median X

MG Methyl ester of gentisic acid

MITF Microphthalmia-associated transcription factor

MOPB Methylophiopogonanone B

NAD Nicotinamide adenine dinucleotide

NADP Nicotinamide adenine dinucleotide phosphate

NCAP N-acetyl-4-s-cysteaminylphenol

NGS Normal goat serum

NHM Normal human melanocyte

PAX 3 Paired-box homeotic gene

PBS Phosphate buffered saline

PE R-Phycoerythrin

PIH Post inflammatory hyperpigmentation

PF Paraformaldehyde

PKA Protein kinase A

PNS Post nuclear supernatant

PTCA pyrrole-2,3,5 tricarboxylic acid

PUVA 8-methoxy psoralen UVA

PVDF Polyvinylidene fluoride

RER Rough endoplasmic reticulum

RFU Relative fluorescence units

ROS Reactive oxygen species

RSK Ribosomal S6 kinase

RT Room temperature

SA Salicylic acid

SB Stratum basale

SC Stratum corneum

SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis

SG Stratum granulosum

SS Stratum spinosum

STI Soybean trypsin inhibitor

TEM Transmission electron microscopy

TG Transglutaminase

TGF-β1 Transforming growth factor-β1

TGN Trans-Golgi network

TM Transmembrane

TM-PAP Transmembrane prostatic acid phosphatase

TPA 12-O-tetradecanoylphorbol-13-acetate

TSRs Thrombospondin type 1 repeats

Tyr Tyrosine

TYRP1 Tyrosinase-related protein 1

TYRP2 Tyrosinase-related protein 2

UVR Ultraviolet radiation

V-ATPase Vacuolar ATPase

CHAPTER 1

INTRODUCTION

1. INTRODUCTION

Since all living single cell or a higher order organisms have some form of a limiting membrane that defines its place in space, it is thought provoking to say that they all have a form of ‘skin’.

(Freinkel and Woodley, 2001) In humans, the skin acts as an interface that conveys a personal identity as well as a state of health, as it is the most visible aspect of an individual’s phenotype.(Jablonski, 2004) In addition, as human skin extends to approximately 2 m2 and comprises approximately 6-10% of the total body mass, it offers an extensive boundary for the body to interact with the surrounding environment. (Tobin, 2006)(Walters and Michael, 2002)

1.1. Human Skin Anatomy

At the interface between the human body and the external environment lies the largest organ of the integumentary system, the skin. This organ serves a variety of purposes including, but not limited to, thermoregulation and photoprotection of the body.(Lin and Fisher, 2007) The skin is also the foremost protective barrier that shields the body from external insults including chemical, mechanical, and microbial assaults.(Tobin, 2006) The diverse properties of human skin make it a complex and sophisticated sensory system that responds to stimuli and provides a mechanism to safeguard, restore, and maintain the body’s homeostasis.(Slominski et al,

2008)(Slominski and Wortsman, 2000) A detailed schematic of the skin and its appendages can be found in Figure 1.1. The complexity of this multilayered organ is immense, including heterogeneous cell types with extracellular components, as well as appendages (including eccrine sweat glands, hair follicles, sebaceous glands, apocrine glands, nerve fibers, blood vessels, and others). (Haake, Scott and Holbrook, 2001)(Tobin, 2006) Traditionally, it is accepted that skin is

comprised of three major layers: the epidermis, dermis, and hypodermis (in order from the outermost layer to the innermost layer).

Figure 1.1. Schematic diagram detailing skin anatomy http://cosmetique.ch/skin-care/skin-cross-section.jpg .

1.1.1. Dermis and Hypodermis

The second major layer of the skin, known as the dermis, is a supportive connective tissue that provides thermoregulation, tensile strength, pliability/elasticity, and receives sensory stimuli.

The dermis is primarily composed of fibroblasts that produce fibrous and elastic connective tissue and “ground” substance. The dermal layer is also composed of an extracellular matrix

(ECM) component, sensory receptors, and neurovascular networks. However, the overall fine structure varies depending on the depth in the dermis. (Tobin, 2006)(Haake, Scott and Holbrook,

2001)(Kanitakis, 2002) The dermal architecture can be broken down into two layers, the superficial papillary dermis and the deeper reticular dermis. The papillary dermis accommodates mechanical stress on the skin by protruding upwards and interdigitating with epidermal rete ridges to increase contact surface area and adhesion. The reticular dermis is the more resilient subdivision of the dermis, which contains larger collagen bundles and mature elastic fibers. In addition, the reticular dermis displays a graded difference in the overall size and characteristics of its inherent fibrous connective tissue, providing a system of mechanical support. (Kanitakis,

2002)(Haake, Scott and Holbrook, 2001) At the transitional boundary between fibrous connective tissue of the reticular dermis and underlying fat containing subcutaneous layer, begins the hypodermis. The two regions are well integrated and contain a series of nerve and vascular networks, as well as protruding epidermal appendages.

The deepest layer of the skin is the hypodermis, a subcutaneous fatty tissue mainly composed of adipocytes. The hypodermis plays an important role in providing insulation, thermoregulation, an energy supply reserve, protection from injuries, and molding of bodily contours. (Haake, Scott and Holbrook, 2001)(Kanitakis, 2002)

1.1.2. Epidermis

The epidermis is a non-vascularized, stratified squamous cell epithelium that comprises the most external portion of the skin. Keratinocytes, the major epidermal cell type, make up 90-95% of the epidermis and exhibits a progressive differentiation, called cornification, from a basal to surface direction. (Haake, Scott and Holbrook, 2001) Keratinocytes are derived from the superficial

ectoderm of the implanted embryo, however the remaining cell types of the epidermis are of distinct embryonic origin. The remaining cellular constituents of the epidermis are the melanocytes (neural crest derived), Langerhans cells (bone marrow derived), and Merkel cells.

Melanocytes function to produce pigmented organelles that participate in skin pigmentation, as well as protect the skin from ultraviolet radiation (UVR). (Haake, Scott and Holbrook, 2001,

Holbrook, 1994)(Walters and Michael, 2002)(Tobin, 2006) Keratinocyte form a network with spatial and organizational specificity, which assembles into distinct cellular layers as the cells terminally differentiates. (Holbrook, 1994) Specifically, the epidermis is composed of four distinct cellular layers or “strata” that exist at varying stages of differentiation.

1.1.2.1. Stratum Basale

The inner most layer, termed the stratum basale (SB) or stratum germinativum, lies at the junction of the epidermis and dermis. Basal cells are columnar type cells that are laterally bordered by other basal cells and bordered above by spinous cells. The juxtaposition and relationship of cells imparts an apparent polarity to the cells. (Holbrook, 1994) Basal keratinocytes express keratin intermediate filaments K5 and K14 in fine bundles that offer a cytoskeletal network with enough flexibility to allow for cell division and migration. (Haake,

Scott and Holbrook, 2001) The basal cell layer is primarily composed of stem keratinocyte cells and transient amplifying cells that undergo a limited number of divisions before transitioning into suprabasal layers. The transient amplifying cells of suprabasal layers become post-mitotic differentiated cells in suprabasal layers. (Holbrook, 1994)(Haake, Scott and Holbrook, 2001)

1.1.2.2. Stratum Spinosum

The ‘spiny’ layer is so named for adhesion points (desmosomes) that interconnect juxtaposed cells to impart a spine like appearance during histological analysis. The cell layer includes 8-10 sheets of keratinocytes with limited cell division capabilities. (Haake, Scott and Holbrook,

2001)(Tobin, 2006) Cellular morphology of the cells in the stratum spinosum (SS) transitions from a polyhedral type to a more flattened spinous cell as they move towards terminal differentiation. Keratin intermediate filaments of the SS are concentrically located around the nucleus and are infused peripherally into desmosomes. While K5/K14 persists from the SB,

K1/K10 are newly synthesized in the SS and are characteristic of differentiating epidermal cells.

(Holbrook, 1994)(Haake, Scott and Holbrook, 2001)

1.1.2.3. Stratum Granulosum

The stratum granulosum (SG) layer is comprised of 2-3 cell layers that contain amorphous, electron dense keratohyalin granules (KHG). It is the keratohyalin granules that give the characteristic granular appearance of the SG. The KHG is comprised of keratin intermediate filaments, loricin, electron-dense protein, and profilaggrin. (Haake, Scott and Holbrook, 2001)

(Holbrook, 1994) Filaggrin is thought to help aggregation of keratin filaments. (Eckert, 1989)

(Dale, Resing and Lonsdale-Eccles, 1985)(Lynley and Dale, 1983)

Lamellar (membrane coating) granules first appear in the SS, but they are primarily present and active in the SG. These lamellar granules deliver precursor stratum corneum (SC) lipids into the

intracellular space. (Haake, Scott and Holbrook, 2001)(Eckert, 1989) The lamellar granule derived lipid sheets provide the barrier properties attributed with the epidermis. (Holbrook, 1994)

At the transition zone between the SG and the SC a drastic remodeling of the layer occurs. This remodeling involves degradative enzymes, of the granular cells, that degrade cellular organelles including the nucleus. (Haake, Scott and Holbrook, 2001) It is for this reason that the SG is considered the last of the viable epidermal layers before transitioning into the non-viable SC.

The self-renewing nature of normal epidermis results in a finely balanced system of formation/proliferation of lower epidermal layers and desquamation of corneocytes in the SC.

This constant rejuvenation process allows for proper epidermal homeostasis. (Candi, Schmidt and Melino, 2005)(Bouwstra and Ponec, 2006)

1.1.2.4. Stratum Corneum

The last stage involved in the differentiation of epidermal keratinocytes results in the structural and compositional changes of the cell to corneocytes in the SC. (Bouwstra and Ponec, 2006) It is the SC that is directly responsible for mechanical integrity and indirectly responsible for the barrier capability of the epidermis.(Norlén, 2006) The barrier properties of the SC are a result of the transglutaminase (TG) mediated cross-linking of structural lipids and proteins.(Tobin, 2006)

The traditional structure of the SC has been described by the ‘brick and mortar’ model in which the “bricks” are corneocytes rich in proteins which are surrounded by “mortar,” an extracellular lipid matrix.(Haake, Scott and Holbrook, 2001) This model has been refined over the years and is reviewed by both Norlén as well as Rawling.(Norlén, 2006)(Rawlings, 2010) The compacted

epidermal sheets are packed with keratin filaments, surrounded by a matrix of filaggrin and interconnected by corneodesmosomes.(Tobin, 2006) The SC assembly is however not a homogeneous structure and includes a transition from the “stratum compactum” to the “stratum disjunctum”. This transition involves an increase in TG mediated cross-linkage of proteins and augmentation in cornified envelope associated ceramides and fatty acids to assist in the conversion from a fragile to a more rigid corneocyte. (Rawlings, 2010) The corneodesmosomes are gradually degraded in the stratum disjunctum by enzymes released from the lamellar bodies.

(Micali et al, 2001) The cohesiveness of the corneocytes is also dependent on non-desmosomal glycoproteins that function as lectins and lipids. During the process of desquamation, the corneocytes are shed from the SC. (Igarashi et al, 2004)

The human SC contains approximately 15 – 26 corneocyte layers in which dark skin contains a significantly higher number compared to light skin. However, the SC is of comparable thickness in all races and thus dark skin may have an increased level of corneocyte cohesion and compaction. (Tobin, 2006)(Berardesca and Maibach, 2003)

1.1.3. Pigmentary System of the Epidermis

As previously stated, keratinocytes comprise ~90-95% of epidermal structure. Many other critical constituents are found in the epidermis including Langerhans cell, Merkel cells, and melanocytes, which perform important tasks in the epidermal cell layers. The highly regulated processes of the epidermal pigmentary system results in skin phenotype and complexion coloration.

1.1.3.1. Melanocytes and Skin Pigmentation

Melanocytes of the interfollicular epidermis are neural crest derived cells, which reside in the

SB. Melanocytes participate in a highly regulated process of melanogenesis to produce and transfer pigmented organelles to surrounding keratinocytes.(Goding, 2007) A major source of skin color in most humans is pigmentation due to melanin. In the various ethnic groups, the constitutive skin color (CSC) defines the amount of melanization in an individual’s epidermis, as determined by cellular genetic programs, in the absence of external stimuli such as ultraviolet radiation (UVR). CSC can be enhanced by various factors including the tanning response following UVR exposure. Facultative skin color (FSC) describes this increase in skin pigmentation above the CSC, as a result of various stimuli including UVR and hormones. The

FSC is reversible once the stimulation factor has been removed. (Quevedo and Holstein,

2006)(Quevedo et al, 1974)

1.1.3.2. Melanosome Biogenesis

Epidermal melanocytes synthesize a unique specialized intracellular organelle termed the melanosome, in which melanin pigment synthesis and storage is accomplished. (Dell'Angelica,

2003)(Marks and Seabra, 2001) The formation of this membrane bound organelle has been delineated into four distinct maturation stages dependent on several characteristics including morphology.(Seiji et al, 1963) Figure 1.2. depicts the stages of melanosome maturity in which

(left) an electron micrograph of MNT-1 melanoma cell displays the various stages of development. Similarly, (right) is a schematic diagram of the maturation of the melanosomes.

Figure 1.2. The various stages of melanosome maturation. (A modified representation from

Raposo et al.) (Raposo and Marks, 2002)

The early stage “pre-melanosome” is initially a non-pigmented electron-lucent vesicle containing rudimentary matrix filaments (stage I).(Wasmeier et al, 2008)(Jimbow et al, 2000) Theories about the origin of the these stage I melanosomes include the hypothesis that the organelle originates from the endoplasmic reticulum (ER), as supported by the presence of ER proteins in preparations of isolated pre-melanosomes.(Basrur et al, 2003) A distinct view is that the premelanosome originates from the late secretory and endocytic pathways.(Raposo et al,

2001)(Dell'Angelica, 2003) These theories may be attributed to the bipartite process of melanogenesis, by which structural components of the melanosome are shuttled from the ER and melanosomal regulatory glycoproteins are shuttled from the Golgi-apparatus. (Turner, Taylor and Tchen, 1975)(Boissy, Huizing and Gahl, 2006)(Ebanks, Wickett and Boissy, 2009) The second stage in the ontogenesis of the melanosome, involves the elongation of the organelle into an ovoid shape and the formation of parallel intralumenal striations. (Marks and Seabra, 2001)

The transition from stage I to II melanosomes is dependent on the presence of the protein pmel-

17, which is cleaved into fragments to form the melanosome internal matrix.(Hoashi et al, 2005)

This intralumenal matrix becomes the scaffold onto which melanin pigment is deposited. A distinguishing characteristic of the stage III melanosome is the deposition of electron dense melanin onto the fibrillar internal matrix.(Kushimoto et al, 2003) The melanin subsequently accumulates and fully obscures the intralumenal matrix (stage IV).

Melanin synthesis and deposition in the melanosome is dependent on pivotal glycoproteins.

Specific regulatory proteins and enzymes are trafficked to the melanosome microenvironment, which function in converting the amino acid tyrosine to melanin. These proteins are type 1 membrane glycoproteins that belong to the tyrosinase gene family: tyrosinase, tyrosinase-related protein I (Tyrp1) and dopachrome tautomerase/tyrosinase related protein-2 (Dct/Tyrp2). These regulatory proteins are transcribed in the rough endoplasmic reticulum (RER), shuttled and modified in the Golgi apparatus. The glycoproteins are then shuttled to the premelanosomes in

50 nm coated vesicles. (Boissy, Huizing and Gahl, 2006)

1.1.3.3. Mechanisms of Melanosome Transfer and Degradation

This portion of the introduction is included in each of the attached manuscripts.

1.2. Lysosomes and Lysosomal Related Organelles

Lysosomes are a class of cytoplasmically located membrane-bound organelles that participate in protein degradation. Lysosomes degrade endogenous macromolecules transported from

biosynthetic pathway, exogenous proteins delivered from the endocytic pathway, and proteins from the cytosol.(Dell'Angelica et al, 2000) This degradation is accomplished by the more than

50 resident acid hydrolases, which are within the lumen of the lysosome, and includes proteases, lipases, and glycosidases (Dell'Angelica et al, 2000)(de Duve, 1963)(Kornfeld and Mellman,

1989)(Hunziker and Geuze, 1996)(Kornfeld and Mellman, 1989)(de Duve, 1963) Lysosomes are also comprised of vastly glycosylated integral membrane proteins along with the residential acid hydrolases, but lack mannose-6-phosphate receptors (MPRs). Other lysosomal membrane proteins influence the transport of amino acids, ions and solutes across the lysosomal membrane to assist in the maintenance of a luminal acidic pH of 4.6-5.0. (Dell'Angelica et al,

2000)(Huizing et al, 2008)

1.2.1. Melanosomes as Specialized Lysosomes

Ample evidence exists to suggest a similar biogenetic pathway between melanosomes and lysosomes, membrane bound organelles that function as the major degradative compartments in the eukaryotic cell. Melanosomes are often referred to as lysosomal-related organelles (LROs) as both lysosomes and melanosomes share similar biosynthetic and structural properties, but have dissimilar composition, morphology, and/or function.(Orlow, 1995)(Novak and Swank, 1979)

(Huizing et al, 2008) Characteristics that LROs and lysosomes share include: a low (acidic) intralumenal pH, similar membrane proteins, and common formation pathway. Other features shared between melanosomes and lysosomes include the presence of transmembrane lysosomal proteins such as lysosomal-associated membrane proteins 1 and 2 (LAMP1, LAMP2), a number of lysosomal hydrolases, the ability to fuse with phagosomes, and availability of endocytic

tracers. (Dell'Angelica et al, 2000)(Orlow, 1995) In addition, Tabata et al. has recently published that both lysosomes and LROs are able to establish an acidic lumen by the presence of vacuolar

ATPase (V-ATPase), which pumps protons into the lumen. (Tabata et al, 2008)

More evidence of a relationship between melanosomes and lysosomes is provided by genetic diseases that affects both pigmentation (melanosomes) and lysosomes in many tissues, Chediak-

Higashi syndrome (CHS) and Hermansky-Pudlak syndrome (HPS). (Diment et al,

1995)(Dell'Angelica et al, 2000)

Using cell fractionation, Diment et al. have shown that melanosomes are the recipient of several lysosomal hydrolases (β-glucuronidase, β-galactosidase, β-hexosaminidase, cathepsins B and L) within the melanocyte. (Diment et al, 1995)(Orlow, 1995) In distinct studies, cytochemical staining completed to ascertain the activity of acid phosphatase showed a similar distribution of acid phosphatase within mammalian and avian melanocytes.(Seiji and Kikuchi, 1969)(Wolff and

Schreiner, 1971)(Boissy, Moellmann and Halaban, 1987)(Orlow, 1995) Furthermore, Novikoff et al. found a similar distribution between tyrosinase and acid phosphatase in the GERL (Golgi- endoplasmic reticulum-lysosome). (Orlow, 1995)(Novikoff, Albala and Biempica, 1968)

1.2.2. Epidermal Lysosomal Hydrolases

Lysosomal hydrolases can be divided in three main categories, which includes ester hydrolases, glycoside hydrolases, and peptide hydrolases. Ester hydrolases include enzymes that degrade carboxylic esters, phosphate esters, and sulphate esters (of interest to us is: prostatic acid phosphatase). Glycoside hydrolases modify both simple and complex glycosyl compounds (of

interest to us are -glucuronidase. We are also interested in an analogous glycosyl transferase enzyme, -3-glycosyltransferase-like. Peptide hydrolases can be divided into exopeptidases which remove terminal amino acids and endopeptidases which cleave internal components of the protein (of interest to us are Cathepsin D, L2). Cathepsin B, another enzyme of interest to us, exhibits both endo- and exopeptidase activity. (Mier and van den Hurk, 1975c)(Mier and van den Hurk, 1975a)(Mier and van den Hurk, 1975b)

In the lysosomal system, cathepsin D (Cath D) is believed to act synergistically with cysteine proteinases as well as exopeptidases in the process of protein degradation. (Kawada et al, 1997b)

1.2.2.1. Cysteine Cathepsins

An increase in the activity of lysosomal cysteine protease is observed and associated with the terminal differentiation of keratinocytes. In addition, it has been reported that lysosomal proteases are pivotal in the physiological processes that are not necessarily restricted to the lysosomal compartment. (Zeeuwen, Cheng and Schalkwijk, 2009)(Tanabe et al, 1991)(Kawada et al, 1997b) Cysteine cathepsins have a functional preference for slightly acidic to an acidic pH environment and reducing conditions. Hence, these enzymes have a preferential localization in endocytic pathway compartments. (Brix et al, 2008) Most cysteine cathepsins are synthesized as preproenzymes in the RER and are cleaved co-translationally by signal peptidase. The procathepsins are then shuttled through the Golgi to the trans-Golgi network (TGN) then to the late endosome by mannose-6-phosphate (M6P) mediated sorting (although shuttling independent of M6P is a possibility and is reviewed by Brix et al.. (Brix et al, 2008)) (Brix et al, 2008)(Mort and Buttle, 1997)(Von Figura, 1991)(Ishidoh and Kominami, 2002)(Mach, 2002)(Wiederanders,

Kaulmann and Schilling, 2003)(Collette et al, 2004)(Saftig, Storch and Braulke, 2005) In keratinocytes, in the presence of the appropriate signal, mature cysteine proteases are selectively trafficked in protease-loaded vesicles. (Brix et al, 2008) Lysosomal cysteine proteases share similar sequences and folds in their structure. The enzymes feature an L-domain with a prominent helix structure and R-domain with a β-barrel motif. The domains interact to form a planar interface with an associated ‘V’- fashioned cleft for the active site. (Turk, Turk and Turk,

2000) Cysteine cathepsins that we will be focusing on includes Cathepsin L2 (Cath L2) and

Cathepsin B (Cath B)

1.2.2.1.1. Cathepsin L2 (Cathepsin V)

Cath L2, also known as cathepsin V (Cath V), (EC 3.4.22.43) is a member of the papain family of cysteine proteases, a family of enzymes that participate in the turnover of intercellular proteases, prohormone activation, enzyme activation, tissue remodeling and other functionalities.

(Santamaría et al, 1998)(Berti and Storer, 1995)(Dickinson, 2002)(Zeeuwen, Cheng and

Schalkwijk, 2009) Cathepsin proteases are active at a low pH and participate in a variety of cellular processes, which upon deregulation can be linked to the propagation of diseases such as cancer, osteoporosis, asthma etc.(Somoza et al, 2000) The enzyme shares 77% proenzyme and

80% mature enzyme sequence identity with cathepsin L, a ubiquitously expressed enzyme. The similarities between Cath L2 and L sequence and genomic organization (Cath L is adjacent to the chromosomal region 9q22.2 of Cath L2) suggests that these two cathepsins diverged late in evolution. (Somoza et al, 2000)(Puzer et al, 2004)

Cath L2 expression is predominantly found in the thymus, testis; but is also present in the brain, corneal epithelium, and skin. (Brix et al, 2008) In association with the skin, Cath L2 has been documented to be expressed in melanocytes, primary keratinocytes, HaCaT keratinocytes, but not in fibroblasts. (Chen, Seiberg and Lin, 2006) Work completed by Bernard et al. established that the “stratum corneum thiol protease,” determined to be Cath L2, is expressed as a proenzyme in the lower epidermis and is activated in the upper layers during keratinocyte differentiation. (Bernard et al, 2003)

1.2.2.1.1.1. Cathepsin L2 Structure and Catalysis

Self-activation of pro-cathepsin L2 occurs at a low pH. This activation results in a 221 residue mature protein. The enzyme is comprised of 2 domains (Figure 1.3.). The first domains shows structural similarities to the R-domain of papain, which primarily consists of β-sheets. The second domain resembles the papain L-domain and contains 3 alpha helices. The active site is found at the interface of the R- and L- domains. (Somoza et al, 2000) When comparing Cath L2 to other cysteine proteases Cath S, K, and L, there was a striking conservation of sequences with the only major difference found in the 10-residue loop structure formed by residues 168A to 169.

In fact comparative analyses of the papain-like cysteine proteases display sequence and length variability in this portion of the structure. (Somoza et al, 2000) Furthermore, the S3 subsite of

Cath L2 is distinct from other cysteine proteases by displaying an Arg at position 70, which may play a role in the enzymes specificity. (Puzer et al, 2004)(Somoza et al, 2000)

Figure 1.3. Schematic diagram of human Cathepsin L2. Within the structure, α-helices are represented by coils and β-strands are represented by arrows. (Somoza et al, 2000)

The catalytic center of Cath L2 is formed by Cys25 and His159. Prior to the binding of the substrate, Cys25 and His159 exist in a thiolate-imidazolium ion pair configuration. (Somoza et al,

2000)(Polgar, 1974)(Lewis, Johnson and Shafer, 1976)(Lewis, Johnson and Shafer, 1981)

(Sluyterman and Wijdenes, 1976) The thiolate mediates the hydrolysis reaction by acting as a nucleophile and attacking the carbonyl carbon of the scissile peptide bond of the substrate. The histidine is multifunctional in substrate hydrolysis by both maintaining the cysteine in a deprotonated state and possibly functions as a general base to assist in the deacylation reaction.

(Somoza et al, 2000)(Storer and Menard, 1994)

1.2.2.1.1.2. Cathepsin L2 Inhibition

Regulation of cysteine protease activity is controlled by a balance between the active form and inhibition by associated protease inhibitors. Cystatins are small proteins that function to regulate

endogenous proteases by competitively and reversibly binding and inhibiting cysteine proteases.

(Zeeuwen, Cheng and Schalkwijk, 2009)(Turk, Turk and Turk, 1997) Cystatin M/E has restricted expression to the differentiating and cornifying layers of epidermis, sweat and sebaceous glands, and hair follicles. (Cheng et al, 2006)(Zeeuwen et al, 2002)(Zeeuwen et al, 2001) Additionally, cystatin M/E is reported to be a strong inhibitor of Cath L2. Immunohistochemical staining of human epidermis, completed by Cheng et al., displays a strong co-expression of Cath L2 and cystatin M/E in the SG of epidermis. (Cheng et al, 2006)

1.2.2.1.2. Cathepsin B

Similar to Cath L2, cathepsin B (Cath B) (EC 3.4.22.1) is a lysosomal cysteine protease. Cath B predominantly functions as a dipeptidyl carboxypeptidase at an acidic pH, but at pH above 5.5 the enzyme transitions to an endopeptidase functionality. (Stachowiak et al, 2004) As an endopeptidase, Cath B cleaves internal peptide bonds with a preference for bulky hydrophobic side chains (will also accept Arg) within the substrate. As an exopeptidase, the enzyme cleaves dipeptides form substrate’s C-terminus, a result of the enzymes occluding loop structure.(Mort and Buttle, 1997)

Cath B participates in protein processing and toxin degradation within the lysosome, but is also involved in cellular processes, outside the lysosome, including activation of prohormones and antigen processing. (Stachowiak et al, 2004)(Authier et al, 1999) Cath B is ubiquitously expressed and has been shown to play a key role in remodeling of the human dermal extracellular matrix, cellular apoptosis/senescence, and tumor cell invasion of the skin. (Lai et al,

2010)(Brix et al, 2008)(Büth et al, 2007)(Kawada et al, 1997a)(Maciewicz et al, 1990)

Büth et al. have proposed that Cath B is involved in the regeneration of wounded epidermis, which has been exemplified by regeneration of scratch wounded keratinocyte monolayers. The authors suggest that Cath B influences the spontaneous migration of HaCaT and normal KC by mediating proteolytic activity dependent extracellular matrix remodeling, which is attenuated by

Cath B specific inhibition. (Büth et al, 2007)(Büth et al, 2004)

1.2.2.1.2.1. Cathepsin B Structure and Catalysis

Cath B is a bilobal protein with both the active site and substrate-binding cleft located at the interface of the two protein domains. (Mort and Buttle, 1997) A schematic of Cath B structure can be found in Figure 1.4. A unique element to Cath B is the presence of an occluding loop, mentioned previously. The occluding loop is comprised of Ile105-Pro126 which impedes access to the catalytic pocket. (Stachowiak et al, 2004)(Illy et al, 1997)(Musil et al, 1991)

Figure 1.4. Schematic diagram of Cathepsin B. Structure obtained from Stachowiak et al. displays the occluding loop structure in red. The blue amino acids indicate the catalytic center and yellow displays the interface of the catalytic cleft. (Stachowiak et al, 2004)

1.2.2.2. Cathepsin D

Cath D (EC 3.4.23.5) is a ubiquitously expressed aspartic endopeptidase that is normally found in intracellular vesicular structures including the lysosome, endosome, and phagosomes. The enzyme is synthesized as a preproenzyme in the RER, followed by signaling peptide removal and targeting of the propeptide to the aforementioned vesicular structures. (Benes, Vetvicka and

Fusek, 2008)(Kopitar-Jerala and Turk, 2000) The inactive 52 kDa preproenzyme is cleaved into an active 48 kDa intermediate proenzyme and lastly processed into the mature form of 32-34 kDa (heavy chain) and 13-14 kDa (light chain).(Kawada et al, 1997b)(Benes, Vetvicka and

Fusek, 2008)(Egberts et al, 2004) Cath D is involved in the turnover of proteins within the

lysosome and also has been proposed to be involved in prohormone activation and antigen processing. (Kopitar-Jerala and Turk, 2000)(Diment, Leech and Stahl, 1988)(Pillai and Zull,

1986) As expected, elevated levels of Cath D is associated with tumor cell invasion and metastasis. (Egberts et al, 2004)

In the skin Cath D is involved in extracellular and intracellular catabolism. In addition Cath D is thought to be involved in epidermal differentiation with an increased expression and activity dependent on the stage of differentiation. (Egberts et al, 2004)(Horikoshi et al, 1998) Cath D is suggested to be involved in the process of TG mediated crosslinking of substrates. Furthermore,

Cath D is reported to be a ceramide binding protein and has a ceramide related enhancement of its proteolytic activity. (Egberts et al, 2004)(Heinrich et al, 1999)(Candi, Schmidt and Melino,

2005) In the hyperproliferative hyperkeratosis skin disease psoriasis, Cath D as well as Cath B and cathepsin L are shown to be activated to a variable extent compared to normal skin. Psoriasis is characterized by abnormal differentiation (abnormal SG and nuclei remain in psoriatic SC) with altered KIF expression and premature expression of involucrin and TG. Following 8- methoxy psoralen UVA therapy (PUVA), a treatment used to attenuate the psoriatic lesions, Cath

D, B, and L expression reverted to normal. (Kawada et al, 1997b)(Chen et al, 2000)(Egberts et al, 2004)

1.2.2.3. β-3-glycosyltransferase-like

β-3-glycosyltransferase-like (B3GTL or B3GALTL) or β-1,3-glucosyltransferase (EC. 2.4.1.-) is a member of glycosyltransferase enzymes, which play a role in many biological processes

including molecular trafficking, signal transduction, cell adhesion, endocytosis, and other functions. (Reis, 2008) The enzymes function mainly in the ER ad Golgi apparatus to attach sugar molecules to particular acceptor sites. (Reis, 2008) B3GTL modifies thrombospondin type

1 repeats (TSRs) of extracellular proteins by addition of glucose residues to O-linked saccharides forming Glcβ1,3Fucα1-O-linked disaccharides. This TSRs specific modification influences extracellular proteins that function in cell-cell and cell-matrix interactions as well as signaling.

(Heinonen et al, 2003)(Kozma et al, 2006) Specifically TSRs are found in extracellular signaling proteins that influence neuronal guidance, angiogenesis, and tissue remodeling during development. (Heinonen and Mäki, 2009)

B3GTL is expressed in a variety of normal tissues and is suggested to be influenced by tissue- specific regulation. (Heinonen and Mäki, 2009) A mutation in B3GTL gene causes Peter’s-plus syndrome. The translated protein lacks the catalytic domain a result of erroneous splicing due to a point mutation. Peter’s-plus syndrome has characteristic features including craniofacial malformations, abnormal development of the skeletal system, mental retardation, and eye defects. (Heinonen and Mäki, 2009)

1.2.2.3.1. β-3-glycosyltransferase-like Structure and Catalysis

B3GTL is a 498 residue protein that is originally thought to be comprised of a short cytoplasmic amino-terminal region (residues 1-4), a transmembrane domain (residues 5-28), a “stem” region

(residues 29-260) and lastly a catalytic domain (residues 261-498). The catalytic domain is

conserved with three aspartic acid residues at its core. (Heinonen et al, 2003) The structural features were later described differently, as an amino-terminal signaling peptide region that directs the enzyme to the ER (residues 1-28), followed by a large domain that may be involved in carbohydrate binding or recognition of substrate glycoprotein (residues 29-260). The carboxy- terminus catalytic domain (residues 261-498), is as previously described. (Heinonen and Mäki,

2009)

1.2.2.4. Prostatic Acid Phosphatase

Acid phosphatase (AP) (EC 3.1.3.2) encompasses a great portion of the total acid hydrolases found in epidermal tissue. (Mäkinen, 1985) AP is part of a class of enzymes that catalyzes the hydrolysis of phosphomonoesters. (Amlabu, Nok and Sallau, 2009) The presence of lysosomal and non-lysosomal AP has both been reported in the epidermis. Two AP rich regions have been documented in the epidermis, including the lysosomal and cytoplasmic regions. Furthermore, AP is highly concentrated in epidermal layers undergoing cellular decomposition such as the SG and

SC. (Mäkinen, 1985) AP has been reported to be associated with the internal membrane of the melanosomes at the site of melanin deposition and tyrosinase activity. (Wolff and Schreiner,

1971)

It should be noted that there are distinct acid phosphatases that exist in human tissues. The multiple molecular forms are traditionally revealed by isoelectric focusing. (Mäkinen, 1985)

Lysosomal AP and prostatic acid phosphatase (ACPP) are encoded by two distinct genes on chromosome 11 and 3, respectively. While the two enzymes display discernible functions, they

have similar physical properties and sensitivity to inhibition by tartrate. Additionally, the enzymes display immunological cross-reactivity. (Lee, Lee and Li, 1991) Furthermore, Mäkinen has reported an acid phosphatase that presents with ACPP-like biochemical properties in the epidermis of the skin. (Mäkinen, 1985)(Partanen, 2008) To add even more complexity, ACPP has a secretory and non-secretory form with different molecular weights and isoelectric points.

The presence of a cellular form is suggested, but the physiological substrate remains unknown.

(Quintero et al, 2007)(Veeramani et al, 2005) Quintero et al. have also recently reported that

ACPP has two splice variants including a secretory form and a type I transmembrane (TM) protein. The TM prostatic acid phosphatase (TM-PAP) is localized to the plasma membrane of the endosomal-lysosomal pathway and displays a wide expression pattern in many nonprostatic tissues.(Quintero et al, 2007)

Immunohistochemical staining completed by Partenen for ACPP showed positive staining in epidermal tissue as well as a positive expression of catalytically active ACPP-like phosphatase in the ‘epidermal keratohyalin layer’ of the skin. (Partanen, 2008) The mechanism of catalytically active ACPP involves the dephosphorylation of macromolecules, using the catalytic residues

His12 and Asp258, which are located at the cleft of the domains. (Hassan, Aijaz and Ahmad, 2010)

1.2.2.5 β-glucuronidase

β-glucuronidase (β-GLU) (E.C. 3.2.1.31) is an exoglycosidase that catalyzes the cleavage of β- glucuronic acid from glycosaminoglycans (GAGs) including chondroitin sulfate, dermatan

sulfate, heparin sulfate. Specifically, β-GLU catalyzes the removal of β-D-glucuronic acid residues from the non-reducing termini of GAGs. This is believed to be essential for the restructuring and turnover of those extracellular matrix constituents.(Shipley, Grubb and Sly,

1993)(Islam et al, 1999)(Jain et al, 1996) The enzyme is found in most tissues specifically in lysosomes and in microsomes (ER).(Antunes et al, 2010) Eukaryotic β-GLU has an optimal pH around 4.7-4.8 and under the tissues normal pH, only approximately 10% of the enzymes maximal activity is maintained. (Krahulec and Krahulcová, 2007)(Antunes et al, 2010)

In humans the deficiency of β-GLU results in mucopolysaccharidosis type VII (Sly Syndrome), a mucopolysaccharide storage disease that is the result of partially degraded GAGs that accumulate in the lysosomes of many tissues. (Islam et al, 1999)(Wong et al, 1998) It is also reported that β-GLU as well as other lysosomal enzymes are released into the synovial fluid of inflammatory joint diseases including rheumatoid arthritis. β-GLU may also be involved in the pathogenesis of other diseases in which extracellular β-GLU is associated with metastatic potential and the invasiveness of tumors . (Jain et al, 1996)(Antunes et al, 2010)

1.2.2.5.1. β-glucuronidase Structure and Catalysis

The structure of human β-GLU belongs to the Family 2 β-glycosidases. (Wong et al, 1998) β-

GLU is synthesized as an 80 kDa precursor glycoprotein that is subsequently processed into a 78 kDa monomer. β-GLU is a homotetramer with four identical monomer units. The human enzyme contains four Asn-linked glycosylation sites that are required for proper protein folding.

(Jain et al, 1996)(Chen et al, 2008) Details on the roles of glycosylation and phosphorylation of the functionality of β-GLU were completed by Shipley et al.. (Shipley, Grubb and Sly, 1993)

Figure 1.5. Stereo ribbon schematic of β-GLU monomer obtained from Jain et al.. Three distinct structural domains are depicted in green, blue and red with yellow representing a hairpin structure. Bonded atoms are shown to represent the oligosaccharide attached to Asn 173. (Jain et al, 1996)

Figure 1.5. obtained from Jain et al. shows the structural domains of each monomer. The red, C- terminal domain is comprised of a TIM-barrel motif which contains the enzyme’s active site.

The active site of the enzyme lies at a large cleft interface between two monomers. The region of the TIM-barrel in which the active site lies contains a high concentration of charged residues that may steer electronegative substrates such as hyaluronic acid. (Jain et al, 1996) The active site residues are proposed to be Glu540, Glu451, and Tyr504. (Islam et al, 1999) The two glutamic acid

residues participate in the enzymes catalysis, where Glu540 is believed to the nucleophile and

Glu451 is believed to be function as the acid/base catalyst. (Islam et al, 1999)(Wong et al, 1998)

While Tyr504 was found to be important for catalysis, yet its role remains unclear. (Islam et al,

1999)(Wong et al, 1998)

CHAPTER 2

HYPOTHESES AND SPECIFIC AIMS

2. HYPOTHESES AND SPECIFIC AIMS

Over the past few decades, skin research has transitioned from the concentration of specialists and dermatologists to a more widespread interest point of many research scientists. While there has been many avenues to research skin, the cells that participate in the function and differentiation of the epidermis has been a primary interest in cellular and molecular biology.(Darmon and Blumenberg, 1993) The complex and dynamic mechanisms involved in the pigmentary system of skin work in tandem to provide complexion coloration as well as protection from UVR (ultra violet radiation) induced damage. Due to extensive research, the mechanisms involved in melanosome biogenesis and transfer to recipient keratinocytes are being elucidated, while the biochemical processes involved in the degradation of melanosomes, by recipient keratinocytes, remain enigmatic.

To date, the principal methods of modulating skin pigmentation is accomplished via interruption of melanin biosynthesis or alternatively through the disruption or inhibition of melanosome transfer from melanocytes to keratinocytes. In addition, understanding and modifying the processes involved in melanosome transfer has been/is currently being investigated heavily. A distinct, and less explored pathway of interest is the processing of melanosomes subsequent to transfer, through melanosomal degradation. Hydrolytic enzymes have been implicated in the processing of melanosomes, but the mechanisms involved in pigment degradation following transfer to epidermal keratinocytes remain to be properly established. Furthermore, the distinction between pigment degradation of light versus dark skin has not been clearly defined.

Therefore, the central hypotheses to be tested are 1.) Epidermal keratinocytes are capable of degrading melanin and the ability to do so is relatively enhanced in light skin and dampened in

dark skin counterparts, 2.) A cohort of hydrolytic enzymes and other candidate molecules, as determined by previous microarray analysis, are distributed in a specific patterns within the interfollicular epidermis and may differ between and correlate with light versus dark skin.

For these reasons we aim to:

1. Investigate if the differential degradation of melanosomes within light skin as opposed to

dark skin is regulated by the cytoplasmic milieu of the keratinocyte.

2. To determine if keratinocytes can degrade melanosomes and if the ability to do so is related

to the hydrolytic content in the cell.

3. To evaluate if known hydrolytic enzymes are distributed in a specific pattern throughout the

interfollicular epidermis and if this pattern differs between and correlates with light versus

dark skin.

CHAPTER 3

OBJECTIVES

3. OBJECTIVES

The long term objective of this project is to elucidate the mechanism of pigment degradation to allow for therapeutic or cosmetic based modulation of these processes. Understanding the mechanisms of pigment degradation will have therapeutic implications such as alleviating hyperpigmentation, melasma and other related skin pigmentary disorders. Clarification of the enzymatic processes will also assist in cosmetic product development aimed at non-solar induced tanning or alternatively skin/hair lightening technology. Additionally, the development of a screening tool that assesses successful melanosomal degradation will be influential in testing overall product efficacy. Finally, formation of a mechanism to successfully quantify melanosome degradation will have implications in advancing the research of skin pigment degradation in various skin types.

CHAPTER 4

EPIDERMAL KERATINOCYTES FROM LIGHT VERSUS DARK SKIN

EXHIBIT DIFFERENTIAL DEGRADATION OF MELANOSOMES

4. EPIDERMAL KERATINOCYTES FROM LIGHT VERSUS DARK SKIN EX HIBIT

DIFFERENTIAL DEGRADATION OF MELANOSOMES (IN PRESS: JOURNAL OF

INVESTIGATIVE DERMATOLOGY)

4.1. Abstract

Modification of skin complexion coloration has traditionally been accomplished by interruption or attenuation of melanogenesis and/or melanosome transfer. Post-transfer modification of pigmented melanosomes provides an attractive and distinct avenue of modulating skin pigmentation. The processing of melanosomes during keratinocyte (KC) terminal differentiation and the degradative variability observed between light and dark skin remains enigmatic. To evaluate this, we developed a model system to investigate the loss of fluorescently labeled and isolated melanosomes by cultured human KCs. The extent of melanosome loss has been qualitatively assessed using transmission electron microscopy (TEM) and indirect immunofluorescence (IIF) with confocal microscopy, and quantitatively assessed using flow cytometry analysis. Results show that melanosomes are incorporated into the cytoplasm of both light and dark keratinocytes (LKCs and DKCs, respectively) and trafficked to a perinuclear region. Within 48 hours, confocal microscopy images suggest that LKCs display accelerated melanosome loss. This time dependent decrease in carboxyfluorescein diacetate (CFDA) fluorescence was then quantitatively analyzed using flow cytometry. Consistent with the results of the confocal analysis, over a 48 hour time frame, LKCs appear to lose melanosomes more efficiently than DKCs. These experiments demonstrate that melanosomes are more rapidly lost in KCs derived from light as opposed to dark skin.

4.2. Introduction

Skin complexion coloration is a consequence of several biochemical and cellular pathways that work concomitantly to provide pigmentation and protection from photo-induced carcinogenesis.

(Sturm, 2009)(Costin and Hearing, 2007) The synthesis and processing of melanosomes, pigment containing organelles of the follicular and interfollicular epidermis, are an integral part of hair pigmentation and skin complexion, respectively. While the mechanisms involved in the synthesis and transfer of melanosomes are being defined, the process of melanosomal degradation by recipient KCs remains enigmatic.

Currently, modulation of skin pigmentation is predominantly accomplished via interruption of melanin biosynthesis (Parvez et al, 2007)(Ebanks, Wickett and Boissy, 2009) or alternatively through the disruption or inhibition of melanosome transfer from melanocytes to KCs (Boissy,

2003), as exemplified by lectins or neoglycoproteins (Minwalla et al, 2001a) and PAR2 (Paine et al, 2001) . A distinct pathway of interest is the processing of melanosomes subsequent to transfer, through melanosomal degradation.

The degradation of the melanosomal organelle is multifaceted and displays variability between distinct racial skin sources. Dark skin derived melanosomes are approximately 0.8 µm, but in contrast light-skinned melanosomes are significantly smaller in size. (Szabo et al, 1969)(Konrad and Wolff, 1973)(Minwalla et al, 2001b) Another distinguishing characteristic between light and dark skin is the pattern and distribution of melanosomes in the cytoplasm of KCs. It has been documented that melanosomes in KCs of light skin are often distributed in membrane bound

clusters of approximately four to eight melanosomes, while melanosomes of dark skin are predominantly individually dispersed. This distinct pattern of melanosome distribution within the

KC appears to be governed by the KC milieu itself. (Yoshida et al, 2007)(Minwalla et al, 2001b,

Thong et al, 2003)

Skin type may also regulate the pattern of melanosome degradation. As KCs undergo terminal differentiation, melanosomes are completely degraded in the upper skin layers of light skin resulting in corneocytes devoid of melanosomes. Alternatively, some melanosomes remained intact and are present in the desquamating corneocytes of dark skin. It has been proposed that light skin melanosomes, which are primarily in membrane bound clusters, are degraded more efficiently during terminal differentiation of the epidermis. (Thong et al, 2003)

Integral to the understanding of pigment degradation is the establishment of a technique to qualitatively and quantitatively analyze this phenomenon. In this present study we have utilized

IIF in conjunction with confocal microscopy, as well as TEM and flow cytometry to demonstrate that melanosomes are more rapidly lost in LKCs (i.e. phototype I/ II) compared to DKCs (i.e. phototype V/VI).

4.3. Results

4.3.1. Distinct Racial Skin Sources Demonstrate Variable Melanosome Degradation

Differential degradation has been observed between light and dark skin sources. Transmission electron micrographs of skin taken from light and dark upper-arm skin biopsies, demonstrate an enhanced degradation/reduction of melanosomes in light skin samples (Figure 4.1.). Figure 4.1. displays a distinct distribution of melanosomes in light and dark skin samples. Both light and dark skin micrographs showed melanosomes in the lower epidermal layers, the stratum basale

(SB) and stratum spinosum (SS). Dark skin showed retention of melanosomes throughout the epidermis, with melanosomes still apparent in the upper skin layers including the stratum granulosum (SG) and stratum corneum (SC). However, light skin micrographs lacked melanosomes in the SG and SC, suggesting a differential processing of melanosomes between light and dark skin. This variability in melanosomal distribution in the epidermal layers of light and dark skin is of particular interest when investigating melanosomal processing and degradation.

While TEM is an excellent technique often used to investigate the degradation of melanosomes, it offers only a qualitative analysis of the biochemical process. It is for this reason that we have developed both a qualitative and an associated quantitative technique to analyze the process of melanosome loss.

4.3.2. Fluorochrome Labeling of Melanogenic Cells and Melanosome Isolation

SKMEL-188 was chosen to be the source of melanosomes because of its highly proliferative and melanogenic capabilities. The cultures offered a high yield of vastly pigmented melanosomes for isolation. The SKMEL-188 cultures were pre-labeled with CFDA, a resilient long-term tracking dye. CFDA is supplied as a non-fluorescent and colorless chemical that passively diffuses into the cell, where intracellular esterases cleave the molecule’s acetyl group. Upon cleavage, the probe becomes highly fluorescent and binds free amino residues of intracellular macromolecules.

(Minwalla et al, 2001a)(Murphy, Watt and Jones, 1992)(Le Poole et al, 1993) The structure of

CFDA can be found in appendix AI.1.. At a 5 µM concentration the dye proved to be highly fluorescent, resilient and displayed minimal associated cell death.

SKMEL-188 labeled with CFDA displayed a green fluorescent signal (Figure 4.2b.) compared to unlabeled cells, which showed minimal to no green signal (Figure 4.2f.). Similarly, melanoma cells labeled with tyrosinase-related protein 1 (TYRP-1) specific MEL-5 antibody produced a red signal (Figure 4.2c. and 4.2g.) an indication of the high melanogenic nature of the cell line. Co- localization analysis of both the CFDA and MEL-5 (Figure 4.2d.), as indicated by a yellow/orange signal, showed a good overlap of the two signals in SKMEL-188. It should be noted that after several passages, occasionally some cells would halt melanogenesis and appear unpigmented, thus labeling with CFDA but not MEL-5 (Figure 4.2d.). Yet, this was not a concern in the melanosome isolation procedure, as only the highly melanin laden stage III and IV melanosomes localize to the fraction of interest during sucrose density gradient centrifugation.

Overall, SKMEL-188 proved to be an excellent source of numerous mature melanosomes.

Mature stage III and IV melanosomes were isolated from the labeled SKMEL-188 cells using sucrose density gradient centrifugation. (Figure 4.S1a.) It should be noted that occasionally the melanosomes appeared to have an aberrant morphology, visually similar to an electron lucent halo. This peculiar structural characteristic of melanoma derived melanosomes has also been previously reported in the literature following microanalysis of melanomas and pigmented nevi.

(Szekeres, 1975) However, intact SKMEL-188 cells (Figure 4.S1b.) displayed the same morphology as the isolated melanosomes, suggesting that the homogenization and isolation procedures were not damaging to the melanosomes. Higher magnification images of isolated

SKMEL-188 melanosomes and melanosomes within intact SKMEL-188 cells (Figure S1c upper row and Figure S1c lower row, respectively) have been included to show the preservation of melanosome ultra-structure following the isolation procedure.

4.3.3. Keratinocytes Uptake and Lose Isolated Melanosomes

To investigate the degradative properties of normal human KCs, we studied the incorporation and degradation of melanosomes using electron microscopy. An 18 hour timeframe for uptake of isolated melanosomes was obtained from a literature value, as determined by Virador et al.

(Virador et al, 2002) Human KC cultures from light and dark skin showed successful incorporation of melanosomes and trafficking of the organelles to a perinuclear location.

To determine the optimal time period to evaluate the degradation of mature stage III and IV melanosomes by KCs, samples were processed by TEM at baseline, 8 h, 24 h, 48 h and 96 h post the 18 hour pulse incorporation and washing of non-incorporated melanosomes. Analysis of

TEM micrographs of 48 hours samples showed ultrastructural disruption of melanosomes, a sign of degradation and by 96 hours KCs displayed minimal melanosomes in culture. Hence, the experimental timeframe was selected to be from baseline to 48 hours. Figure 4.S2. depicts representative TEM micrographs of 48 h samples from (a) LKCs and (b) DKCs. Black arrows indicate melanosomes that have ultrastructural disruption including visible striations from the melanosome scaffolding and/or an amorphous shape. White arrows indicate intact mature melanosomes, mostly found in the DKCs after 48 h.

4.3.4. Loss of Fluorochrome Labeled Melanosomes by Human Keratinocytes in an In Vitro

Model System

Upon determination of the time frame for melanosome incorporation and time period to visualize melanosome degradation, we established a methodology to qualitatively assess the loss of melanosomes in LKCs and DKCs in vitro. To qualitatively analyze the incorporation and loss of melanosomes, we used isolated CFDA labeled melanosomes and incubated them in the presence of pre-plated LKCs and DKCs. Following an 18 hour pulse incorporation period, all non- incorporated melanosomes were rigorously washed with PBS and baseline samples were immediately processed for confocal microscopy. 48 hour samples were re-fed with KC media, until 48 hours post baseline, then fixed and similarly processed. IIF and confocal microscopy analysis of KCs at baseline and 48 hours were completed to assess the concentration and localization of melanosomes in LKCs compared to DKCs. Representative images can be found in Figure 4.3.. In combination with the CFDA labeling, MEL-5 was concurrently used as a melanosome specific label for TYRP-1, as described in the materials and methods section. At baseline, both LKCs (Figure 4.3a.-4.3d.) and DKCs (Figure 4.3e.-4.3h.) displayed incorporation

of isolated melanosomes and trafficking of melanosomes predominantly to a perinuclear location within the cytosol of the recipient KCs. Additionally, CFDA pre-labeling of the melanosomes displayed excellent co-localization with MEL-5, as exemplified by the yellow/orange signal seen in the merge column of Figure 4.3. (Figure 4.3d., 4.3h., 4.3p.). After 48 hours LKCs (Figure

4.3i.-4.3l.) displayed a dramatic reduction in melanosomes, with minimal CFDA or MEL-5 labeling. On the contrary, 48 hour DKCs (Figure 4.3m.-4.3p.) have retained melanosomes with vivid signal. Additionally, DKCs that retained melanosomes displayed a predominant perinuclear cluster of the organelles. This perinuclear melanosome cap formation is also shown in supplementary Figure 4.S3.. Visual image analysis demonstrated that the CFDA labeled melanosomes were avidly fluorescent in the confocal baseline images, confirming that the isolation procedure did not disrupt the CFDA fluorescent labeling. Moreover, the general CFDA labeling of melanosomes shows consistent co-expression with TYRP-1, offering an effective way to specifically visualize melanosomal location and reduction with time.

The distinction between melanosome reduction in LKCs compared to DKCs, suggests that light skin derived KCs may have accelerated melanosome degradation. Yet, it is important to note that the confocal images serve as a qualitative technique and does not simultaneously assess a large cell population. For these reasons we have applied the model system to flow cytometry, which both quantifies melanosome reduction and assess a larger population of sample cells.

Concerns of a differential proliferation rate between LKC and DKC cultures resulting in a differential ‘dilution’ rather than loss of melanosomes were addressed by looking at the rate of division between LKC and DKC cultures. LKC and DKC cultures were plated and counted at 53

baseline, 24, 48 hours post melanosome incorporation. (Figure 4.S4.) LKC and DKC lines showed similar rates of proliferation over the 48 h time frame of the experiment. This data suggests that both the racial lines of KCs show a similar rate of growth and the depletion of melanosomes seen is actually representative of differential melanosomal loss between LKCs and

DKCs.

4.3.5. Quantitative Analysis of Melanosome Loss by Human Keratinocytes in an In

Vitro Model System

The parameters of the qualitative analyses were then applied to flow cytometry, to offer a quantitative technique to delineate the process of melanosome degradation. The signal of cells incubated with melanosomes was assessed above the signal of negative control KCs without melanosomes. This was completed to reduce the acquisition of nonspecific background signal due to auto-fluorescence of normal cellular constituents.

To quantify the amount of melanosomes within the KC populations at baseline and 48 hours we evaluated the percent of cells positive for CFDA or PE (%CFDA or %PE) as well as the degree of positivity within that population, as indicated by the median X (Md X) values. The product of

%CFDA and Md X (or product of %PE and Md X) was used to ascertain the degree of positivity within the overall cell population, in which an elevated value was interpreted as a higher melanosome quantity per cell. Figure 4.4. graphically depicts the results of baseline and 48 hour readouts of fluorescence intensity, an indication of melanosome concentration in LKCs and

DKCs. Frequently, we found that the baseline incorporation of melanosomes varied between

both KC types without a unilateral correlation. Within 48 hours of incorporation there was a significant diminution of CFDA and PE fluorescence over time, an indicative of a reduction in melanosome quantity in both cell lines. In addition, we found that melanosome concentration, as indicated by CFDA (Figure 4.4a.), showed a similar trend of incorporation and loss compared to

TYRP-1 specific labeling PE (Figure 4.4b.). This indicates that the CFDA labeling offers a valid methodology to quantitate overall melanosome reduction. Consequently, subsequent experiments were completed using only CFDA labeling of melanosomes without double labeling with MEL-

5/PE.

As previously stated, the distinct racially derived KCs showed variable abilities to incorporate melanosomes. Hence, to appropriately analyze/normalize for this variability, the ratio of melanosomes at baseline to 48 hours was used to assess melanosome loss with time (Figure 4.5.).

Consistent with the results of the confocal analysis, over the 48 hour period, the LKCs appear to lose melanosomes more efficiently than the DKCs and displayed a greater reduction in intensity values over the experimental time frame. The experiment was repeated four times using four distinct LKC and DKC cell lines (Figure 4.5.). Experiments depicted in Figures 4.5a. and 4.5b. were completed using a similar number of melanosomes for both experiments, while Figure 4.5c. was completed with a slightly lower number of melanosomes and Figure 4.5d. with a slightly higher number of melanosomes. In all cases, we found the same trend in which DKCs retained an increased proportion of its initial incorporated melanosomes and LKCs displayed a comparatively enhanced rate of melanosome loss. Statistical analysis of all four analysis suggests that there is a statistically significant difference between LKC and DKC ability to lose melanosomes with 95% confidence, P = 0.039.

The experimental model system was then applied to normal human melanocyte derived melanosomes. The experiment was completed using the same model parameters but, alternatively, using duplicate samples per category. The result of the normal melanosome experiment was consistent with the SKMEL-188 melanosome experimentation. Yet, there was a slight distinction noted in the normal melanosome experiment, in which LKCs initially incorporated more of the pigmented organelles than the DKCs (Figure 4.6a.). The percentage of cells that incorporated the melanosomes were similar between the two KC subtypes but the amount of melanosomes per cell varied, with a somewhat higher Md X value achieved in LKCs.

However, the overall change from baseline to 48 hour values, an indication of melanosome loss, shows consistent results with prior experiments as LKCs displays an enhanced capability to lose melanosomes with time (Figure 4.6b.).

4.4. Discussion

Presently we have demonstrated through various experimental methods that melanosomes are more rapidly lost from recipient LKCs compared with DKCs. This degradation is accomplished at differential rates between light and dark skin, with an apparent enhancement in degradation in

LKCs. TEM analysis of light and dark epidermis displayed a dissimilar location of melanosomes in the corresponding cell layers of the two skin types. Although both racial groups showed a reduction in overall melanosome number, once the KCs transitioned from the SB to the SS and

SG, the light skin samples showed a more pronounced reduction. Furthermore, unlike light skin which preserved little to no melanosomes in the SG, dark skin samples often retained these melanosomes throughout all the cell layers including within the desquamating corneocytes. This

was a fundamental finding, suggesting a distinction between the ways that melanosomes are processed by recipient KCs in the two racial categories.

To fully understand and delineate how the process of melanosome disintegration is accomplished, we established a model system to evaluate the procedure. Melanosomes were isolated from SKMEL-188 cells, which were highly melanogenic and provided an overabundance of melanosomes. The melanosomes were fluorescently labeled with CFDA and

MEL-5/PE, offering a convenient tool to examine the presence or absence of melanosomes with various fluorescence-based analytical techniques. Baseline incorporation of isolated melanosomes by the KC cultures was easily assessed using confocal microscopy and flow cytometry. Confocal analysis of melanosome location in conjunction with DIC optics demonstrates that following incorporation of melanosomes, the organelles are shuttled to a perinuclear location in the cytoplasm of KC cultures. Within 48 hours of initial analysis, fluorescence image analysis suggests that DKCs display a higher preservation of melanosomes compared to LKC, which display a more efficient reduction in fluorescence.

Similar median fluorescence readouts from flow cytometry analysis of LKCs and DKCs were obtained and again, LKCs displayed a higher melanosomal loss over the experimental time span.

Flow cytometry analysis focused on two major parameters. The first parameter of interest is the percent of cells that are positive for melanosome internalization (%CFDA or %PE). The percent positive was critical to see the initial proportion of cells that successfully incorporated the melanosomes. The second parameter of interest is the median fluorescence intensity value, which

is an estimate of the quantity of melanosomes expressed per cell (Md X). The product of these two parameters was analyzed in each sample to indicate the degree of positivity and the overall quantity of melanosomes within the cell population. In four separate experiments utilizing

SKMEL-188 derived melanosomes and four separate LKC and DKC lines, LKCs consistently showed an increase rate of melanosomal loss. These results were recapitulated using normal human melanocyte melanosomes. Dissimilar to the SKMEL-188 melanosome experiment, normal human melanosomes were internalized more efficiently by LKC than in DKC samples.

However, in agreement with melanoma melanosome experiments, the change in normal melanosome concentration from baseline to 48 hours was more enhanced in LKC compared to

DKC cultures. Our ability to both qualitatively and quantitatively analyze melanosome loss has been successfully accomplished using the proposed model system.

Although it is very clear that there is a differential degradation of melanosomes, which seems to be governed by the genetic composition of the KC, the mechanism needs to be discerned. We speculate that these molecular regulators could be known or novel degradative enzymes previously suggested to be involved in epidermal differentiation and melanosome turnover. In agreement with the suggested mechanism, literature studies have looked at the presentation of melanin in epidermal skin sections from various racial origins. In experimentation completed by

Pathak and Stratton and Quevedo et al., the authors treated skin sections to locate melanin throughout the epidermis. They found that melanin was present throughout the epidermis, but its quantity reduced during its ascent through the various epidermal layers towards the SC. They also, suggest that melanosomes may lighten in color due to the intervention of lysosomal hydrolases or a transition from a more oxidized to reduced state. (Pathak and Stratton,

1969)(Quevedo et al, 1989)(Quevedo and Holstein, 2006) The present study utilizing identical melanosomal sources demonstrate that the keratinocyte milieu regulates, in part, the differential loss of melanosomes. This process may also be influenced by the nature of the donor melanosomes themselves. A smaller, less dense and more pheomelanotic melanosomes synthesized by light skin melanocytes may also exhibit more efficient melanosomal loss than larger, denser and more eumelanotic melanosome synthesized by dark skin melanocytes.

In conclusion, we have successfully developed a new strategy to study and elucidate the process of melanosome loss. The technique offers a novel approach to investigate how various skin types process melanosomes during KC terminal differentiation. Throughout our various analyses, we have found that KC cultures from light skin have an apparent increased rate of melanosome loss when compared to DKCs of a similar passage. This model system offers a distinct analytical methodology that may provide valuable information concerning the turnover of the melanosome in the skin and hair. We propose and explore that this distinction in melanosomal processing, is the result of an initial differential melanosomal loss in part. Whether this differential rate of melanosomal loss we have demonstrated in our in vitro model represents the entire process in vivo remains uncertain. An additional mechanism that could be at play is that the more heavily pigmented and numerous melanosomes, that frequently occur in DKCs, may ultimately saturate the degradative machinery and could lead to persistent melanosomes in the upper layers of the skin.

4.5. Materials and Methods

4.5.1. Cell Culture

Cell cultures were established from neonatal foreskin samples following standard procedures.

For details on the establishment of cell cultures see supplementary materials and methods section

4.10.

4.5.2. Melanosome Isolation/Purification

Stage III and IV mature melanosomes were isolated from SKMEL-188 melanoma cells or melanocyte cell lines. For details on the procedure to isolate melanosomes, see the supplementary materials and methods section 4.10.

4.5.3. Electron Microscopy

Cells were fixed with Karnovsky’s fixative, processed and embedded in epon resin according to standard procedures. TEM analysis of both Caucasian and African American upper-arm skin biopsies was performed as previously described (Yoshida et al, 2007). Details of the procedure for electron microscopy can be found in the supplementary materials and methods section 4.10.

4.5.4. Pulse Incorporation of Melanosomes by Keratinocyte Samples

The procedure for the fluorescent labeling and incorporation of isolated melanosomes can be found in the supplemental materials and methods section 4.10.

4.5.5. Assessing Keratinocyte Rate of Division

A cell counting procedure was employed to assess the KC rate of division. For details see supplemental materials and methods section 4.10.

4.5.6. Indirect Immunofluorescence of Keratinocytes Using Confocal Microscopy

IIF was used to analyze KC cultures following pulse incorporation of isolated melanosomes as well as SKMEL-188 control samples. A modified version of the procedure has been previously detailed (Smith et al, 2005). For further details see the supplementary materials and methods section 4.10.

4.5.7. Flow Cytometry

Following pulse incorporation of melanosomes, baseline KCs from light and dark derived sources were processed for flow cytometry. A detailed explanation of the procedure can be found in the Supplemental materials and methods section 4.10.

4.5.8. Description of Statistics

Experimental analysis of melanosome degradation by flow cytometry was completed four times independently. Comparison of the ratio of baseline to 48 h melanosome concentration, between 61

LKCs and DKCs was analyzed using a paired Student’s t-test (two-tailed, type-1). Statistical significance was accepted at P < 0.05.

4.6. Conflict of Interest

The authors state no conflict of interest.

4.7. Acknowledgements

We would like to thank Andrew Osterburg for his technical assistance with the flow cytometer as well as Bradley King and Gordon Shott for their assistance with the flow cytometry experiments.

This work was supported by a grant from The Procter and Gamble Company.

4.8. Figures

Figure 4.1. Transmission electron micrographs of biopsies from light skin (top row) and dark skin (bottom row).

Scale bar = 500 nm. LS – light skin, DS – dark skin, SB – stratum basale, SS – stratum spinosum, SG – stratum granulosum, SC – stratum corneum. (a) The SB of light skin shows multiple, membrane bound, melanosome clusters. The number of melanosomes is drastically reduced by the SS (b). The SG (c) and SC (d) of light skin samples displayed no distinguishable melanosomes. The SB of dark skin (e) displayed larger, singly distributed melanosomes, which reduced by the SS (f) but to a lesser extent than in light skin. The SG (g) and SC (h) of dark skin retained melanosomes (indicated by arrows) unlike light skin.

Figure 4.2. SKMEL-188 labeled with CFDA-SE and MEL-5.

SKMEL-188 cells were used as the source of mature stage III and IV melanosomes. SKMEL-

188 as seen in the differential interference contrast (DIC) images (a, e) were double labeled with

CFDA (b), an avidly fluorescent green cell tracing reagent, and MEL-5 (c), a melanosome specific TYRP-1 marker (red signal). The majority of the cells were found to be actively involved in melanogenesis as display by the yellow/orange signal (d) in the cytoplasm of the cell.

Cells that were labeled with MEL-5 show only a red signal (g,h) and showed little to no green

CFDA signal (f).

Figure 4.3. Incorporation and loss of fluorescently labeled melanosomes by LKCs and DKCs.

CFDA (green) and MEL-5 (red) labeling of melanosomes were completed in LKCs (passage 2) and DKCs (passage 3). (a-d) LKCs at baseline display incorporation and shuttling of melanosomes to a perinuclear location in the cytosol of the KCs. (e-h) Similarly, DKCs at baseline demonstrate incorporation and translocation of melanosomes to a perinuclear region. (i- l) LKCs, after 48 hours, show a reduction in melanosomal protein labeling, with both CFDA and

TYRP-1 labeling significantly reduced. (m-p) DKCs after 48 hours show retained melanosomal protein labeling compared to LKCs at 48 hours, with a distinct localization pattern compared to

dark baseline counterparts. Higher magnification images of boxed areas in (d,h,p) have been included to show the co-localization of CFDA and MEL-5.

(%CFDA)*(Md X) CFDA (%PE)*(Md X) PE 150 1750 1500 125 1250 100 1000

75 750 50 500 25 250 0 0 Baseline Baseline 48 Hour 48 Hour

Baseline Baseline 48 Hour 48 Hour (ArbitraryIntensity units) Intensity (ArbitraryIntensity units) DKC LKC DKC LKC DKC LKC DKC LKC ab

Figure 4.4. Keratinocytes incorporate and lose fluorescently labeled SKMEL-188 melanosomes.

LKCs (passage 2) and DKCs (passage 3) evaluated at baseline and 48 hours after melanosome incorporation period for (a) CFDA fluorescence intensity and (b) TYRP-1 expression (as PE fluorescence). Results demonstrate that initial incorporation of melanosomes was substantial in both LKC and DKC lines and within 48 hours there was significant reduction in fluorescence indicating melanosome loss quantitatively.

Figure 4.5. Ratio of baseline to 48 hour fluorescent intensities quantitatively analyzes melanosome loss.

To evaluate the reduction of melanosomes in the various KC cultures, the ratios of incorporated baseline values to the remaining 48 hour values were analyzed. Experiment was repeated four times using four distinct LKC and DKC cell lines. (a-b) Show repeated analysis in which LKCs display an overall higher reduction in melanosomes. When the melanosome number was slightly reduced (c) or increased (d) during pulse incorporation of the pigmented organelle, the same trend in melanosome loss was observed. Overall on a consistent basis, light skin derived keratinocytes seem to more efficiently lose melanosomes than DKCs. P=0.039. Parenthesis after the KC type indicates cell passage number.

(% CFDA)*(MdX) CFDA Ratio of melanosomes at baseline to 48 h 350 4.5 3.944 300 4.0 250 3.5 3.0 2.694 200 2.5 150 2.0 1.5 100 1.0 50 0.5 0 0.0 Baseline Baseline 48 Hour 48 Hour DKC LKC Intensity (ArbitraryIntensity Units) Intensity (Arbitrary units) DKC LKC DKC LKC Keratinocyte Cultures ab

Figure 4.6. Keratinocytes incorporate and lose fluorescently labeled normal human melanosomes.

(a) LKC and DKC incorporate dark skin derived human melanosomes and lose them over a 48 hour time period. Although, LKCs displayed an interestingly higher incorporation compared to

DKCs, the reduction from baseline to 48 hours was consistent with prior experiments. (b)

Analysis of the ratio values from baseline to 48 hours shows an enhanced rate of loss in LKC compared to DKC cultures.

4.9. Supplementary Figures

Figure 4.S1. Transmission electron micrographs of SKMEL-188 melanosomes.

Scale Bars = 1 µm. (a) TEM micrographs of pure stage III and IV SKMEL-188 melanosomes, isolated via sucrose density gradient centrifugation. (b) Melanosomes in intact SKMEL-188 show the same morphology as isolated melanosomes. (c) Higher magnification micrographs of isolated SKMEL-188 melanosomes (upper row) and melanosomes within intact SKMEL-188 cells (bottom row).

Figure 4.S2. Ultrastructure of melanosomes after 48 h in keratinocyte cultures.

Scale Bars = 1 µm. TEM micrographs of melanosomes in (a) LKCs and (b) DKCs. Black arrows indicate melanosomes with ultrastructural disruption and irregularly defined shape, often with visibility of the melanosome scaffolding network suggesting melanin disappearance. White arrows indicate melanosomes that retain an ultrastructure similar to isolated mature melanosomes. DKCs displayed more intact melanosomes after 48 h as indicated by white arrows.

Figure 4.S3. Melanosomal perinuclear cap formation in the keratinocyte cytoplasm.

Differential interference contrast images with CFDA (green) fluorescence overlay shows the melanosome perinuclear formation in (a) LKCs at baseline, (b) DKCs at baseline and (c) DKCs at 48 h post incorporation into the KC cytoplasm. N is the nucleus and C is the perinuclear cap.

Figure 4.S4. Proliferation rate of light and dark keratinocytes with incorporated melanosomes.

Both LKCs (passage 2) and DKCs (passage 2) show a similar rate of growth over the 48 hour time frame of the experiment, as noted by the similarities in the slopes between baseline and 24 h, as well as between 24 h and 48 h.

4.10. Supplementary Materials and Methods

4.10.1. Cell Culture

This study was approved by the institutional review board at the University of Cincinnati [#03-1-

23-1]. Primary cultures of melanocytes and KCs were propagated for experimentation.

Melanocyte and KC cultures were established from neonatal foreskins obtained from the

University Hospital at the University of Cincinnati or the Christ Hospital of Cincinnati, following routine circumcision. Foreskins were visually inspected for the light-most (i.e. phototype I / II) and dark-most (phototypes V/VI) tissue. Our previous work details quantitative analysis done, to confirm the expression of low and high melanin content and the correlation with light and dark skin, respectively.(Yang et al, 2000) Light and dark cell cultures were established using the same culture conditions. The foreskin tissue samples were processed by washing in an iodine/PBS solution, adipose tissue removed and skin pieces placed dermis side down on 25 units/mL dispase soaked gauze overnight at 4°C. The epidermis was then collected and incubated in 0.05% trypsin-EDTA then pelleted by centrifugation. Epidermal cells were liberated by rigorous pipeting in cell culture media and seeded into 25 cm2 tissue culture flasks.

Melanocytes were maintained in MCDB-153 media (Sigma-Aldrich Co., St. Louis, MO) supplemented with 4% fetal bovine serum (Fisher Scientific, Pittsburgh, PA), 1 µg/mL vitamin

E, 5 µg/mL Insulin, 0.6 ng/mL basic fibroblast growth factor (PeproTech Inc, Rocky Hill, NJ),

13 µg/mL bovine pituitary extract (Hammond Cell Tech, Windsor, CA), 8 nM 12-O- tetradecanoylphorbol-13-acetate (TPA) and 1% antibiotic/antimycotic (Fisher Scientific,

Pittsburgh, PA). All the above reagents were purchased from Sigma-Aldrich Co. unless otherwise stated. KC cultures were maintained in KC medium Epilife supplemented with HKGS-

V2 (Cascade Biologics Inc., Portland, OR). SKMEL-188 (a gift from V. Setaluri Ph.D.,

University of Wisconsin, Madison, WI) were maintained in DMEM media (Fisher Scientific,

Pittsburgh, PA) supplemented with 6% fetal bovine serum, 2 mM L-glutamine (Fisher Scientific,

Pittsburgh, PA), 1 mM sodium pyruvate (Invitrogen, Carlsbad, CA) and 1% antibiotic/antimycotic. The cell cultures were maintained in a humidified incubator with 95% air/5% CO2 at 37°C. Melanocyte media and SKMEL-188 media was routinely changed twice weekly, while KC media was changed three times weekly.

4.10.2. Melanosome Isolation/Purification

Melanosomes were isolated and purified using a modification of the methodology published by

Kushimoto et al.. (Kushimoto et al, 2001) Cells were grown to confluency and harvested using

0.05% trypsin-EDTA. The cells were then washed with 1% FBS and pelleted by centrifugation at

1500 rpm for 5 mins at 4°C. Cells were washed in homogenization buffer (0.25 M sucrose, 10 mM HEPES, 1 mM EDTA, 2% antibiotic/antimycotic, pH 7.2) and homogenized with 120 strokes of a Dounce glass homogenizer (Thermo Fisher Scientific, Rochester, NY). The homogenate was then centrifuged at 700 g for 1 min at 4°C to prepare the post nuclear supernatant. The post nuclear supernatant was collected and pelleted by centrifugation at 14000 g for 5 min at 4°C. The pelleted cell constituents were then re-suspended in HEPES buffer (10 mM

HEPES, 2% antibiotic/antimycotic, pH 7.2) and separated on a preformed stepwise sucrose density gradient and centrifuged at 368000g for 1h at 4°C in a swing-out rotor. The sucrose density gradients were comprised of 1 M, 1.2 M, 1.4 M, 1.6 M, 1.8 M, 2.0 M and 2.4 M fractions, sequentially layered with the highest density fraction on the bottom. The stage III and

IV melanosomes, which preferentially localized to the high density sucrose fraction, were collected. The isolated fraction was diluted 1:1 with HEPES buffer and centrifuged at 16,000

rpm for 10 mins at 4°C to separate the melanosomal pellet from the sucrose. Melanosome pellets were collected in Epilife media with 2% antibiotic/antimycotic. An aliquot of the melanosome suspension was then taken and compared to a 0.6 µm mean particle size polystyrene bead standard curve (Sigma-Aldrich Co., St. Louis, MO) prepared using a Malvern Zetasizer® Nano

(Malvern Instruments Ltd., Worcestershire, UK).

4.10.3. Electron Microscopy

Following melanosome pulse incorporation, samples were fixed using half strength Karnovsky’s fixative (Karnovsky, 1965) in 0.2 M sodium cacodylate buffer at pH 7.2 for 30 mins. Cells were then washed with cacodylate buffer 3 x 5 mins and treated with 1% osmium tetroxide/1.5% potassium ferrocyanide (Karnovsky, 1971) for 30 mins. Cells were then washed en bloc with

0.5% uranyl acetate in 75% ethanol for 30 mins, gradually dehydrated in a graded series of ethanol and embedded in Eponate 12. Culture areas were randomly selected, cut from the epon cast and mounted onto epon pegs and sectioned into 90 nm ultrathin sections with a RMC MT

6000-XL ultramicrotome. Images were taken using a Joel JEM-1230 transmission electron microscope fitted with an AMT Advantage Plus 2K X 2K digital camera. (Joel Ltd., Tokyo,

Japan)

For TEM analysis of melanosomes isolated from SKMEL-188 cells, pellets of the isolated melanosomes were processed through the post-fixation step using osmium tetroxide and potassium ferrocyanide. The melanosome pellets were then embedded in a 2% agar solution prepared by heating the agar in cacodylate buffer. The warm agar solution was poured onto a

pre-cooled microscope slide and allowed to solidify. The pellet was then cut into small cubes and embedded into epon pegs and sectioned into 90 nm ultrathin sections.

TEM analysis of biopsies taken from either Caucasian or African American upper-arm skin was performed as described by Yoshida et al. (Yoshida et al, 2007)

4.10.4. Pulse Incorporation of Melanosomes by Keratinocyte Samples

SKMEL-188 cell were labeled with the succinimidyl ester of carboxyfluorescein diacetate

(CFDA). 5 µM CFDA in PBS was used to label SKMEL-188 for 30 mins at room temperature

(RT), as recommended by the manufacturer. Cells were then re-fed with DMEM media and stored overnight at 37°C in a 5% CO2 / 95% air atmosphere incubator. Labeled mature melanosomes were isolated from the SKMEL-188 lines and were dosed in the presence of pre- plated light and dark derived passage 2 or 3 KCs in 8-well Lab-Tek™ chamber slides (Thermo

Fisher Scientific, Rochester, NY) (coated with 1% coating matrix (Invitrogen, Carlsbad, CA)) for confocal experimentation. 25 cm2 tissue culture flasks were used for flow cytometry experiments. KC cultures were plated at 2 x 104 cells per well for IIF and 2 x 105 cells per 25 cm2 flask for flow cytometry. Isolated melanosomes were added at the same concentration per sample. Melanosomes were plated in excess, ranging from 1000 to 2000 melanosomes/KC plated. Following an 18 hour pulse period, to allow KCs to incorporate the isolated melanosomes, KCs were rigorously washed three times with 1X PBS to remove non- incorporated melanosomes. Baseline flasks were processed immediately and 48 hour flasks were re-fed with KC growth media until 48 hours post wash, and then processed for either IIF or flow cytometry analysis. In experiments using melanocyte cultures as the source of melanosomes, the cells were labeled and utilized in a similar manner as SKMEL-188 cells.

4.10.5. Assessing Keratinocyte Rate of Division

Distinct LKC and DKC cultures were plated at 2 x 105 KCs per 25 cm2 flask. Duplicate samples were plated for baseline, 24h and 48h time points for analysis in both the LKC and DKC culture lines. Samples were then pulsed with isolated melanosomes as described above. At baseline, 24 h and 48 h the appropriate samples were harvested by trypsinization. Flasks were washed once with 1X PBS and the rinses collected in the same 15 mL centrifuge tubes. Samples were then vortexed until homogeneous and counted on a Z1 Coulter® Particle Counter (Beckman Coulter

Inc., Mississauga, ON, Canada)

4.10.6. Indirect Immunofluorescence of Keratinocytes Using Confocal Microscopy

Following pulse incorporation baseline KCs/SKMEL-188 control samples were fixed with 3% paraformaldehyde for 30 mins and stored in PBS at 4°C. Forty-eight hours post wash; the remaining samples were fixed with 3% paraformaldehyde. Samples were then washed with PBS

3 X 5 mins, permeabilized with 100% HPLC-grade methanol for 3 mins at RT, washed with PBS

3 X 5 mins and blocked for 30 mins with 1% normal goat serum (NGS) in 5% bovine serum albumin (BSA) in PBS. The appropriate wells were then incubated with the primary antibody, mouse monoclonal antiserum to tyrosinase-related protein 1 (TYRP-1), MEL-5 (1:100)

(Covance Research Products Inc., Emeryville, CA) in 1% BSA in PBS and control wells were incubated in solely 1% BSA in PBS for 1 hour at RT. All sample wells were then washed with

PBS 3 X 5 mins and incubated with the secondary antibody, Alexa Fluor® 546 goat anti-mouse

IgG (1:250) (Molecular Probes, Eugene, OR) in 1% BSA in PBS for 45 mins at RT. Slides were washed with PBS 3 X 5 mins then mounted and coverslipped with Gel Mount (Biomeda Corp.,

Foster City, CA), an aqueous base mounting media that contains anti-fading agents to preserve

the sample fluorescence. Slides were then allowed to dry and edges sealed with clear nail varnish. Images were digitally photographed on a Zeiss LSM 510 Confocal Microscope (Carl

Zeiss, Thornwood, NY) using the same image parameter settings between sample wells. Images were then processed on the LSM 510 Image Browser Software (Carl Zeiss MicroImaging Inc.,

Thornwood, NY).

4.10.7. Flow Cytometry

For flow cytometry, cells were plated at a concentration of 2 x 105 cells/flask 24 h before the addition of melanosomes to the cultures. SKMEL-188 or normal human melanocyte derived melanosomes were labeled with 5 μM CFDA, before isolation and plating. Preliminary experiments included a supplementary labeling step, where the KCs were permeabilize and then double labeled with MEL-5/ R-phycoerythrin (PE) goat anti-mouse IgG before fixation and flow analysis. Again, MEL-5 labeling was analyzed in conjunction with the CFDA labeled melanosomes.

The procedure for flow analysis is as follows. At baseline and 48 hours KCs were washed with

1X PBS pH 7.4 and incubated using 2 mL of 0.05% trypsin-EDTA for 10-15 mins at 37°C and harvested. Cells were collected in 15 mL centrifuge tubes (Corning Life Sciences, Corning, NY) containing 100 µL FBS and centrifuged at 2000 rpm for 5 mins at 4°C. Supernants were aspirated and cells permeabilized with 200 µL of 1X BD FACSTM permeabilizing solution (BD

Biosciences, Franklin Lakes, NJ) for 10 mins at RT. Cells were pelleted at 2000 rpm for 5 mins at 4°C, supernatant aspirated and then blocked with 20% NGS in 4% FBS in PBS solution for 30 mins at RT. The appropriate tubes were incubated with MEL-5 diluted in 4% FBS in PBS

(1:100) and control tubes were incubated in 4% FBS in PBS solution for 45 mins at RT with shaking. Cells were centrifuged at 2000 rpm for 5 mins at 4°C, supernatant aspirated and rinsed

1X with PBS and re-pelleted. Appropriate cells were incubated with 200 µL of secondary antibody R-phycoerythrin goat anti-mouse IgG (1:20) (PE) (Molecular Probes, Eugene, OR) diluted in 4% FBS in PBS solution for 30 mins at 37°C. Cells were then washed in PBS 2 X 5 mins and fixed in 1 mL of 1% paraformaldehyde in 1X PBS. Samples were analyzed on a

COULTER® EPICS® XL™ Flow Cytometer (Beckman Coulter, Fullerton, CA). CFDA and PE were excited with the 488 nm line of an argon-ion laser, until 10,000 gated events were observed per sample. The emission spectra of CFDA and PE were collected using a 525 bandpass filter and a 575 bandpass filter, respectively.

Since analysis of the CFDA and PE data showed similar trends in melanosome degradation with time, subsequent experimentation was performed using solely CFDA pre-labeled melanosomes without secondary labeling with PE. Cells were analyzed in an identical manner to the flow cytometry methodology detailed above, up to the permeabilization step with 1X BD FACSTM permeabilizing solution. Following this step with cells were immediately fixed with 1 mL of 1% paraformaldehyde and analyzed by the flow cytometer.

The majority of flow experiments were completed with triplicate samples per experimental condition. However, the flow experiment that used normal human melanosomes was completed in duplicate samples due to plethoric number of melanosomes necessary to complete the experiment.

CHAPTER 5

HYDROLYTIC ENZYMES OF THE INTERFOLLICULAR EPIDERMIS DIFFER IN

EXPRESSION AND CORRELATE WITH THE PHENOTYPIC DIFFERENCE OBSERVED

BETWEEN LIGHT AND DARK SKIN

5. HYDROLYTIC ENZYMES OF THE INTERFOLLICULAR EPIDERMIS DIFFER IN

EXPRESSION AND CORRELATE WITH THE PHENOTYPIC DIFFERENCE

OBSERVED BETWEEN LIGHT AND DARK SKIN

5.1. Summary

Background Degradation of melanosomes by light skin (LS) appears to occur more rapidly than in dark skin (DS). Hydrolytic enzymes known to reside and be expressed in a differential pattern within the interfollicular epidermis are implicated in playing a role in epidermal differentiation and potentially melanosome degradation.

Objectives To evaluate the differential expression of hydrolytic enzymes that may correlate with the physiological and phenotypic differences seen between DS and LS.

Methods Hydrolytic enzyme expression was confirmed by microarray analysis of suprabasal epidermal layers from LS and DS. Specific lysosomal hydrolases identified by microarray analysis, were analyzed by indirect immunofluorescence (IIF) and immunoblot analysis.

Immunogold electron microscopy (IEM) was completed to visualize the cellular expression of the hydrolytic enzyme cathepsin L2 (Cath L2) and biochemical assay was performed to ascertain

Cath L2 activity.

Results Immunoblotting of light and dark epidermal lysates, demonstrated that both prostatic acid phosphatase (ACPP) and Cath L2 were reproducibly upregulated in DS and LS, respectively. IIF and IEM analyses of Cath L2 in tissue confirmed this differential expression.

Biochemical analysis of Cath L2 in light and dark epidermal lysates displays an increased activity of Cath L2 in LS samples.

Conclusion The results of this study confirm a differential expression of ACPP and Cath L2 in

DS and LS at the gene and protein level. Additionally, Cath L2 displays an increased activity in

LS derived epidermal lysates. These results have identified a specific acid hydrolase that may play a role in melanosome degradation and pigment processing.

What’s already known about this topic?

Cath L2 has been reported to be differentially expressed in LS and DS and it has been suggested that the enzyme may play a role in SC turnover. (Chen, Seiberg and Lin, 2006)(Zeeuwen, Cheng and Schalkwijk, 2009)(Zeeuwen et al, 2006)

What does this study add?

The association of Cath L2 with melanosomes and the potential of the enzyme to be involved in pigment processing has been explored.

5.2. Introduction

Skin is the major interface of the body and the surrounding environment. The complex ultrastructure of the organ, a result of finely regulated keratinization, differentiation, and cornification process, forms a unique and dynamic barrier.(Candi et al, 2008) Although, the procedure of terminal differentiation shares similarities, there are distinctions between skins of different racial origins. The most notable distinction between LS and DS is the degree of pigmentation. A major determinant in skin colour is the process of melanosome degradation and pigment processing during keratinocyte terminal differentiation. (Thong et al, 2003)(Ebanks,

Wickett and Boissy, 2009) It is generally accepted that racial skin types also determines the pattern of melanosome degradation. The processing of melanosomes in LS and DS is distinct, in that melanosomes are completely degraded and are absent from the corneocytes of LS, but intact melanosomes remain in the desquamating corneocytes of DS. (Thong et al, 2003)

For decades, the process of successful degradation of melanosomes has remained elusive. Yet, theories do exist surrounding a potential mechanism. Both optical and ultrastructural assessments have described the disintegration of melanosomes in lysosomal compartments. Furthermore, histochemical studies and biochemical analysis have suggested the involvement of acid hydrolases in melanosome degradation.(Borovansky and Elleder, 2003)(Ohtaki and Seiji,

1971)(Saito and Seiji, 1973) The role of the melanosome as a lysosome related organelle and the association of melanosomes with acid hydrolases have also been explored. (Ohtaki and Seiji,

1971)(Olson, Nordquist and Everett, 1970)(Wolff and Schreiner, 1971) In addition, literature

studies have investigated the association of acid phosphatase with melanosome complexes localized within keratinocytes.

For these reasons, we have investigated acid hydrolases of LS and DS epidermis, previously implicated in epidermal terminal differentiation, to assess their expression and possible involvement in melanosome degradation. The associated differences observed between the two racial skin types were also explored. Several candidate acid hydrolases, identified by microarray analysis, were investigated to establish if they are differentially expressed in DS and LS.

5.3. Materials and Methods

5.3.1. Cryosectioning of Human Foreskin Tissue for Indirect Immunofluorescence

LS and DS derived neonatal foreskins were collected from the University Hospital at the

University of Cincinnati as well as the Christ Hospital of Cincinnati (Cincinnati, OH, U.S.A.), following routine circumcision. Biopsies of foreskin tissues were washed with phosphate buffered saline (PBS) and embedded in OCT compound (Tissue-Tek®-Sakura, Torrance, CA,

U.S.A.) to prevent dehydration of sample edges, and then allowed to solidify at -20°C. Skin sample were stored at -80°C until they were sectioned (5µm) with a -20°C Leica CM1800 cryostat microtome (Leica Microsystems, Richmond, IL, U.S.A.), and collected on Poly-L-

Lysine charged slides (Thermo Fisher Scientific, Rochester, NY, U.S.A.). Slides were stored at

-80°C until IIF.

5.3.2. Indirect Immunofluorescence of Skin Cryosections

Slides were equilibrated at room temperature (RT) for approximately 1 hr. Samples were incubated in 3% paraformaldehyde (PF) for 10-15 mins, washed 3 x 5 mins with PBS and incubated with HPLC-grade methanol for 3 mins at RT. Samples were again washed with PBS and blocked with 1% normal goat serum (NGS) in 5% bovine serum albumin (BSA) in PBS, for

30 mins. The appropriate samples were labeled with primary antibody in 1% BSA for 1 hr, while control wells were incubated in solely 1% BSA. The samples were again washed with PBS and incubated with the appropriate specie specific secondary antibody conjugated to Alexa Fluor®

546 (Molecular Probes, Eugene, OR, U.S.A.) in 1% BSA for 45 mins at RT. Slides were washed with PBS and mounted with Gel Mount (Biomeda Corp., Foster City, CA, U.S.A.), an aqueous mounting media that preserves sample fluorescence with anti-fading agents. Slides were allowed to dry and the coverslip edges were sealed using a clear nail varnish. Images were taken on a

Zeiss Axioplan microscope with AxioCam-MR digital camera (Carl Zeiss MicroImaging Inc.,

Thornwood, NY, U.S.A.)

5.3.3. Laser Capture Microscopy and Microarray Analysis

8 μm thick foreskin sections were collected and stored at -80°C until use (≤ 1 week). Slides were prepared for LCM using an Arcturus HistoGene Frozen Section LCM Staining Kit. Suprabasal areas of foreskin cryosections from three dark and three light foreskins were microdissected and captured using the Arcturus Veritas Microdissection System. RNA was extracted using a

Picopure RNA Isolation Kit and purity and quantity was assessed using an Agilent 2100

Bioanalyzer (Santa Clara, CA, U.S.A.). All samples were within the usable range (total RNA

>5ng/sample and 260/280 purity ratio of >2.0). RNA samples were amplified and biotin-labeled using the OvationTM Biotin System (NuGEN, San Carlos, CA, U.S.A.), then hybridized to

Affymetrix HG-U133 plus 2.0 Array chips (human genome). Data were extracted from the chips by the BioMedical Informatics Core Facility at The Cincinnati Children’s Hospital Research

Foundation (Cincinnati, OH, U.S.A.). Each sample was normalized to an internal chip control.

Two experimental groups (LS and DS) were formed, each with three samples, which contained signals above background. For each gene (i.e., signal) the LS group was compared to the DS experimental group. Data were compiled from all genes that exhibited a two-fold or greater difference in the ratio of one pigmentary group compared to the other. All kits were purchased from Molecular Devices (Arcturus Molecular Devices, Sunnyvale, CA, U.S.A.), unless otherwise stated.

5.3.4. Isolation of Human Epidermis

LS and DS derived foreskin samples were processed by washing with an iodine/PBS solution and adipose tissue removed. Skin samples were cut into pieces and placed dermis side down on a

25 units/mL dispase soaked gauze overnight at 4°C. The following day, epidermal sheets were collected and washed 1 X with PBS and stored at -80°C until use.

5.3.5. Immunoblotting

For epidermal lysate preparation, epidermal sheets were sonicated on ice 3 x 20 secs in RIPA buffer (150 mM sodium chloride, 1% NP-40, 0.5% deoxycholate, 0.1% sodium dodecyl sulfate,

50mM Tris pH 8.0, and CompleteTM protease inhibitors (Roche, Indianapolis, IN, U.S.A.)). Table

5.1. details protein concentrations, antibodies used. Equal amounts of protein, as determined by a

BCA assay (Roche, Indianapolis, IN, U.S.A.), were loaded onto sodium dodecyl sulfate polyacrylamide gels (SDS-PAGE) and transferred to a polyvinylidene fluoride (PVDF) membrane. Following protein transfer, the membranes were blocked using 5% milk in TBST.

Membranes were probed (Table 5.1.). Immunoreactivity was determined by an enhanced chemiluminescence reaction (ECL, Perkin Elmer, Waltham, MA, U.S.A.). Densitometry was completed using Alphaease software (Alpha Innotech Corp., San Leandro, CA, U.S.A.). Protein expression was calculated using the ratio of sample band intensity relative to loading control intensity.

5.3.6. Biochemical Assay

Fluorometric assay of Cath L2 activity was analyzed in assay buffer (25mM Sodium Acetate,

0.1M NaCl, 5mM DTT, pH 5.5). 50µM Z-Leu-Arg-AMC (MP Biomedicals LLC, Solon, OH,

U.S.A.) was used as the substrate. For sample preparation, epidermal sheets were snap-frozen in liquid nitrogen for 20 secs then homogenized with a Potter-Elvehjem micro tissue homogenizer

(teflon/glass) (Thermo Fisher Pierce, Rockford, IL, U.S.A.). Samples underwent two 20 secs liquid nitrogen freeze thaws and stored at -80°C until use. Epidermal samples were mildly sonicated on ice for 1 min, homogenate centrifuged at 13,000 g, 10 min 4°C, and supernatant collected. Protein concentrations were determined by BCA assay and utilized at 10µg in 100µL of assay buffer. Activity was monitored as an increase in fluorescence from substrate hydrolysis at (excitation 360 nm, emission 460 nm) in a POLARstar OPTIMA plate reader (BMG Labtech,

Cary, NC, U.S.A.) every 30 secs for 15 mins at 37°C.

5.3.7. Immunogold electron microscopy

The transmission electron microscopy (TEM) procedure for embedding and sectioning of biopsies taken from either Caucasian or African American upper-arm skin, was performed as described by Yoshida et al..(Yoshida et al, 2007) Samples were processed for IEM using a modified procedure of that described by Orlow et al.(Orlow et al, 1993). The modifications are as follows. Samples were collected on formvar coated nickel grids. The primary antibody used was Cath L2 (1:10) (R&D Systems, Minneapolis, MN). Washed samples were incubated with a

1:15 solution gold-conjugated (goat) anti-mouse (20 nm Au, Ted Pella Inc., Redding, CA, USA).

Finally, samples were incubated with 2% glutaraldehyde for 15 mins and 2% aqueous osmium tetraoxide for 15 mins. Samples were dried overnight, and stained with Reynolds lead citrate for

15 mins at RT. Images were captured on a Joel JEM-1230 TEM. (Joel Ltd., Tokyo, Japan)

5.3.8. Statistical Analysis

Data were analyzed using the Student’s t-test (two-tailed, type-2) assuming equal sample variance. Statistical significance was accepted at P < 0.05.

5.4. Results

5.4.1. Light and dark skin differentially express lysosomal hydrolases in suprabasal epidermis

To explore the differential profile of gene products that may play a role in melanosome degradation, microarray analysis was performed on the suprabasal cell layers of LS and DS derived neonatal foreskin obtained by LCM. Known hydrolytic enzymes were identified during

the analysis and Table 5.2. lists the lysosomal enzymes and regulators that displayed a differential expression between the two complexion types. Lysosomal hydrolases elevated in the suprabasal layers of LS included B3GTL, Cath B, Cath L2 and those that are elevated in DS included ACPP, Cath D and β-GLU with the magnitude indicated in the fold change column

(Table 5.2.). These target enzymes were subsequently assessed at the protein level using both

WB and IIF.

5.4.2. Immunoblot analysis of lysosomal hydrolases in epidermal tissue

As a validation study to the microarray analysis, comparative proteomic analysis was completed by WB. Immunoblotting was completed to assess the expression of enzymes of interest in four

LS and four DS derived epidermal lysates. The concentration of protein used, antibody dilutions and loading control concentrations are detailed in Table 5.1.. The first lysosomal enzyme of interest, ACPP, demonstrated relatively higher protein levels in all four DS samples compared to

LS samples, P=0.048 (Figure 5.1a.). This observation is consistent with the results of the microarray analysis previously described. In contrast, B3GTL (Figure 5.1b.) and β-GLU (Figure

5.1c.) showed inconstant levels between all epidermal lysate samples regardless of skin complexion. Mature Cath D expression (33 kDa) (Figure 5.1e.) in the LS epidermal lysates appeared to be reduced compared to DS lysates, as predicted by microarray analysis, P=0.015.

The intermediate form of Cath D (46 kDa) (Figure 5.1e.) alone, however, was inconsistent and showed no obvious trend. Yet, if considered as a precursor to the mature Cath D, it demonstrates an overall increased expression of mature Cath D in DS compared to LS. Intriguingly, WB results of Cath D from differentiated and non-differentiated keratinocyte lysates from LS and DS

displayed variable expression, a contrary result to the Cath D expression in epidermal lysates

(unpublished data, which has been included in appendix section AII.2.).

Continued analysis of hydrolytic enzymes showed that Cath B (Figure 5.1d.) displayed variable expression of the pro-enzyme in both complexion lysates. Results of the protein expression of the mature form of Cath B were inconclusive. This result is consistent with the results of IIF, described below, but contrary to the results of the microarray data.

The most dramatic distinction in acid hydrolase expression between LS and DS was observed in

Cath L2 probed samples. LS lysates demonstrated higher levels relative to DS, P=0.048. (Figure

5.1f.)

5.4.3. Indirect immunofluorescence of lysosomal hydrolases in epidermal cryosections

Cryosections of epidermal biopsies taken from LS and DS were used for comparative IIF analysis of lysosomal hydrolases. Many LS and DS donor samples were used to compare the expression pattern of epidermal hydrolases. Figure 5.2. shows the expression of Cath L2 in images taken from four discrete DS samples (Figure 5.2a.-5.2d.) and similarly four LS samples

(Figure 5.2e.-5.2h.). LS samples consistently expressed a higher level of Cath L2 than DS samples. Furthermore, the elevated level of Cath L2 in LS was observed throughout all the epidermal layers including in the stratum corneum (SC). DS samples demonstrated a modest immunoreactivity to Cath L2 in all samples.

Cath B (Figure 5.2i.-5.2p.) displayed variable expression from sample to sample without regard for the pigmentation status of the skin utilized. However, most samples showed a high staining

pattern of Cath B in the SC of the epidermis. IIF of Cath D (Figure 5.2q.-5.2w.), similar to Cath

B, showed variable protein expression from sample to sample in both LS and DS. Furthermore, the expression pattern of Cath D varied between the stratum granulosum (SG) (i.e. Figure 5.2q. or 5.2u.) and the SC (i.e. Figure 5.2s. or 5.2t.).

There was low expression of B3GTL in both complexion samples (Figure 5.2x.-5.2ae.); however there was a slightly higher staining intensity apparent in two LS samples (Figure 5.2ab.-5.2ac.).

The remaining LS samples (Figure 5.2ad.-5.2ae.) displayed a similar staining intensity to DS specimens. However, it should be noted that B3GTL analysis was completed in two rather than four distinct samples with sample 1and 2 being from one specimen and sample 3 and 4 from a different specimen. In both LS (Figure 5.2ab.-5.2ae.) and DS (Figure 5.2x.-5.2aa.) samples

B3GTL displayed minimal expression in the SC.

5.4.4. Immunogold labeling of cathepsin L2 in light and dark epidermal tissue

To delineate the localization and function of Cath L2 at the ultrastructural level, we performed immunogold labeling of ultrathin sections of LS and DS. TEM micrographs of each epidermal cell layer were captured and analyzed for labeling of Cath L2 as indicated by gold particles. The number of gold particles were more abundant in the cytoplasm of LS as opposed to DS, as shown in Figure 5.3. Occasionally these cytoplasmic gold particles were associated with the edge of the melanosome in the basal area and vesicles with melanin debris in the SG zone, as indicated by arrows. The number of particles were quantified and detailed in Table 5.3.. LS samples again contained a higher intensity of immunolabeling for Cath L2 in all cell layers compared to DS. 92

Furthermore the expression of Cath L2 followed a gradient like progression with increasing levels as the keratinocytes transitioned from a basal to terminally differentiated cells. DS samples contained a very low expression of Cath L2 in the cytoplasm or nucleus of keratinocytes within the stratum basale (SB) and stratum spinosum (SS). For IEM analysis we took the ratio the number of gold particles (in the cytoplasm plus the nucleus) between the two skin types as detailed in Table 5.3.. When comparing the number gold particles in corresponding light to dark skin layers, we found a 4.2 fold higher expression in the SB, 8.1 fold higher in the SS, and 9.4 fold enhancement in labeling for Cath L2 in the stratum granulosum (SG). Similarly, we found a

4.2 fold higher intensity in the SG in conjunction with the stratum corneum (SC) and 2.1 fold increase in the SC only of LS compared to DS.

5.4.5. Biochemical analysis of cathepsin L2 activity

From the compilation of the previous experimental results, ACPP, mature Cath D and Cath L2 were observed to be differentially expressed between the distinct skin types. Cath L2 displayed the highest dissimilarity in protein expression level between the two skin phenotypes and was consequently analyzed for its activity in epidermal samples. Biochemical analysis was completed using ZLR-AMC as the fluorogenic substrate. Preliminary work was used to determine the appropriate substrate concentration (50µM) for the experimentation (supplemental analyses and calculations completed for the biochemical assay can be found in appendix section AII.4.).

Analysis of the specific activity in the lysates of three LS and three DS derived epidermal tissue was completed. The results indicate a specific activity in DS samples of 0.413 nmol/min*mg compared to 0.722 nmol/min*mg in LS samples. A 1.75 fold higher Cath L2 activity was observed in LS above that of DS samples, P=0.03.

5.5. Discussion

Lysosomal hydrolases have been reported to function in areas that are not necessarily restricted to the lysosome. The process by which lysosomal hydrolases are released and activated during epidermal cornification as well as the processes involved in the degradation of cellular organelles is not fully established. (Zeeuwen et al, 2006)(Turk et al, 2002) We have identified and verified the expression of acid hydrolase that display divergent expression in the suprabasal layers of skin with light (Type I/II) or dark (Type IV/V) complexion colouration. We investigated the differential expression of these enzymes, found to be critical for proper epidermal terminal differentiation, to determine a correlation with the differential pigment processing observed in

LS and DS. Cath L2, a member of the class of papain like cysteine proteinase, was initially found to be over-expressed in LS relative to DS by microarray analysis. Furthermore, cysteine cathepsins are reported to participate in proteolytic cascades, where a protease activates one or several other proteases to result in the cleavage of a proteinaceous substrate. (Brix et al,

2008)(Kobayashi et al, 1991)(Kostoulas et al, 1999)(Sloane et al, 2006) Cath L2 has been characterized in the process of epidermal homeostasis with prospective roles in corneodesmosome degradation and SC turnover.(Zeeuwen et al, 2006) Here we observe through

WB and IIF that Cath L2 shows a higher expression in LS compared to dark samples.

Furthermore, IEM analysis shows that Cath L2 occasionally localizes with melanosomes within the cytoplasm of the keratinocytes.

Acid phosphatase has been shown to be associated with the melanosome and is speculated to be involved in the organelle’s degradation. (Wolff and Schreiner, 1971)(Nakagawa et al, 1984) It has also been previously reported that an acid phosphatase with prostatic acid phosphatase-like

biochemical properties is expressed in the skin.(Mäkinen, 1985)(Partanen, 2008) In addition, histochemical staining of ACPP in human epidermis completed by Partenen displayed successful staining within the cell layers.(Partanen, 2008) We have shown that ACPP is differentially expressed in LS and DS, with an upregulation in DS by both WB and microarray analysis. In the case of ACPP, whether keratinocytes amplify the expression of hydrolytic enzymes in order to ultimately degrade a more resilient/ resistant melanosome compared to a more easily degradable light skin derived melanosome warrants further investigation.

β-GLU, B3GTL, and Cath B were all found to be present in the epidermis, yet all displayed variable expression in all specimens investigated. The aspartyl proteinase Cath D, similar to Cath

L2, is postulated to be involved in squame separation or regulation of the enzyme transglutaminase. (Zeeuwen, Cheng and Schalkwijk, 2009)(Igarashi et al, 2004) By our analysis,

Cath D IIF showed variability in staining, which is inconsistent with both the WB and microarray that displayed an over-expression in DS. IIF of the enzyme was observed in either the

SG or SC with an inter-sample variability that was not associated with skin ethnicity. This may be a result of staining of the inactive enzyme precursors alongside staining with the active mature enzyme.

Taken together, we observe that there are complexion coloration associated distinctions in the presence of lysosomal hydrolases in the epidermis, specifically in reference to ACPP, Cath D, and Cath L2.

5.6. Acknowledgments

We would like to thank Bradley King and Gordon Shott for their technical assistance with the IIF and biochemical assay experiments. This work was supported by a grant from The Procter and

Gamble Company.

5.7. Figures and Tables

Table 5.1. Concentrations and dilutions of components utilized for western blot analysis.

[Protein] %SDS [Primary] [Secondary] Expected MW Loading Control Prostatic acid ACPP 50µg 10% 1:100 GαM1:1000 52 kDa GAPDH 1:1000/GαR-HRP phosphatase 1:1000 (Lifespan Biosciences) (Santa Cruz Biotech) β-3-glycosyltransferase- B3GTL 50µg 12% 1:100 GαR 1:1000 57kDa GAPDH 1:1000/GαR-HRP like 1:2500 (Sigma-Aldrich Co.) β-glucuronidase β-GLU 50µg 10% 1:100 GαM1:1000 75 kDa GAPDH 1:1000/GαR-HRP (Lifespan Biosciences) 1:1000 Cathepsin B Cath B 60µg 10% 1:200 GαR 1:5000 pro (37-43kDa) GAPDH-HRP 1:1000 (US Biologicals) mature (25kDa) Cathepsin D Cath D 30µg 10% 1:200 DαG 1:1000 immature(52- GAPDH-HRP 1:5000 (Santa Cruz Biotech) 60kDa) interm (46kDa) mature (33kDa) Cathepsin L2 Cath L2 30µg 10% 1:500 GαM 1:2000 28kDa GAPDH 1:1000/GαR-HRP (R&D Systems) 1:2000

Table 5.2. Microarray analysis of suprabasal epidermis from light and dark skin.

Gene Products Fold Change

Elevated in Upper epidermis of Light Skin Compared to DS Beta 3-glycosyltransferase-like 7.539 Cathepsin B 3.545

Cathepsin L2 3.047

Elevated in Upper epidermis of Dark Skin Compared to LS Acid phosphatase, prostate 3.86 Cathepsin D (lysosomal aspartyl protease) 4.26 Glucuronidase, beta 4.9

Table 5.3. Immunogold labeling of Cath L2 in light and dark skin.

Skin Cell average # of gold particles # ratio light to Phenotype Layer micrographs dark cytoplasm cytoplasm nucleus + nucleus Light Skin SB 34.8 2.7 37.4 n=9 4.2 – SB Light Skin SS 42.8 26.3 69.1 n=12 8.1 – SS Light Skin SG 64.7 37.3 102.0 n=3 9.4 – SG Light Skin SG + SC 120.4 9.1 129.5 n=11 4.2 – SG+SC Light Skin SC 239.2 --- 239.2 n=11 2.1 – SC Dark Skin SB 8.1 0.9 8.9 n=19 Dark Skin SS 6.5 2.0 8.5 n=15 Dark Skin SG 10.8 --- 10.8 n=5 Dark Skin SG + SC 30.7 --- 30.7 n=3 Dark Skin SC 114.9 --- 114.9 n=8

Figure 5.1. Immunoblot analysis of key acid hydrolases was completed in four LS and DS derived epidermal lysates.

(a) ACPP band intensities (relative to GAPDH) were found to be significantly lower in light skin lysates than that observed in DS, P=0.048. B3GTL (b), β-GLU (c), pro Cath B (d), and intermediate

Cath D ((e) upper row) displayed inter-sample variability with no obvious over-expression in either

LS or DS. Mature Cath D ((e) middle row) was expressed at a higher concentration in DS in

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comparison to LS epidermal lysates, P= 0.015. (f) Cath L2 band intensities were apparent in LS specimens, while DS samples showed minimal immunoreactivity, P=0.048.

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Figure 5.2. Indirect immunofluorescence of four distinct dark skin (DS) and light skin (LS) cryosections.

Cath L2 expression in dark skin (a-d) displayed a low staining intensity compared to LS samples (e-h). The expression of Cath L2 was apparent in all cell layers of the LS epidermis. Cath B showed variable expression in all DS (i-l) and LS specimens (m-p), however,

Cath B was most apparent in the SC of all labeled samples. Similar to Cath B, Cath D (q-w) displayed variable staining from sample to sample regardless of skin pigmentary phenotype. Cath D staining showed the highest staining in either the SG (q,r,u,v) or SC (s,t,w)

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of the epidermis in comparison to the remaining cell layers. B3GTL displayed a modest expression in all DS samples (x-aa), which is recapitulated in some light skin samples (ad, ae). Yet, a few light sample showed a slightly higher level of B3GTL (ab, ac) than the remaining samples.

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Figure 5.3. IEM of Cath L2 in epidermis of LS and DS.

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Localization of Cath L2 with IEM in the SB of (a) LS and (b) DS and in the SG of (d) LS and (e) DS. Negative control of (c) SB and

(f) SG in LS. Arrows indicate gold particles associated with melanosomes or melanosomal debris. (*) indicates the basement membrane . SC is the stratum corneum and KH are keratohyalin granules. Scale bar = 500nm.

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CHAPTER 6

MECHANISMS REGULATING SKIN PIGMENATION: THE RISE AND FALL OF COMPLEXION COLLORATION - A REVIEW

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6. MECHANISMS REGULATING SKIN PIGMENTATION: THE RISE AND FALL OF

COMPLEXION COLLORATION - A REVIEW(Ebanks, Wickett and Boissy, 2009)

6.1. Abstract

Skin pigmentary abnormalities are seen as aesthetically unfavorable and have led to the development of cosmetic and therapeutic treatment modalities of varying efficacy. Hence, several putative depigmenting agents aimed at modulating skin pigmentation are currently being researched or sold in commercially available products. In this review we will discuss the regulation of processes that control skin complexion coloration. This includes direct inhibition of tyrosinase and related melanogenic enzymes, regulation of melanocyte homeostasis, alteration of constitutive and facultative pigmentation and down-regulation of melanosome transfer to the keratinocytes. These various processes, in the complex mechanism of skin pigmentation, can be regulated individually or concomitantly to alter complexion coloration and thus ameliorate skin complexion diseases.

6.2. Introduction

The acquirement of an aesthetically pleasing skin pigmentary appearance has been a primary focus of many cosmetic and therapeutic based industries. As a result, several treatment modalities are being investigated for their efficacy in treating skin hyperpigmentary lesions. This review will detail many of the current skin depigmenting agents and treatment approaches that are currently being employed to combat skin pigmentary disorders.

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Melanocytes, the pigment producing cells of the follicular and interfollicular epidermis, produce a specialized lysosomal related organelle termed the melanosome. Within the melanosome, biopolymers of the pigment melanin are synthesized to give hair and skin, as well as other tissue, its color. This melanin synthesis involves a bipartite process in which structural proteins are exported from the endoplasmic reticulum and fuse with melanosome-specific regulatory glycoproteins released in coated vesicles from the Golgi-apparatus. Melanin synthesis ensues subsequent to the sorting and trafficking of these proteins to the melanosome. (Turner, Taylor and Tchen, 1975)(Boissy, Huizing and Gahl, 2006) Each melanocyte resides in the basal epithelial layer and, by virtue of its dendrites, interacts with approximately 36 keratinocytes to transfer melanosomes and protect the skin from photo-induced carcinogenesis. Furthermore, the amount and type of melanin produced and transferred to the keratinocytes with subsequent incorporation, aggregation and degradation influences skin complexion coloration. (Boissy,

2003) Hyperpigmentary disorders of the skin such as melasma, age spots or solar lentigo can result from the overproduction and accumulation of melanin. (Urabe, Nakayama and Hori,

1998)(Virador et al, 2001) As such, depigmenting agents can have potent effects by acting on one or more steps in the melanogenic pathway, melanosome transfer or post-transfer pigment processing and degradation.

Tyrosinase, the rate limiting enzyme of the melanogenic pathway, is a copper containing glycoprotein of approximately 60-70 kDa and a common target for therapeutic agents intended to alleviate cutaneous hyperpigmentation. (Parvez et al, 2006)(Ando et al, 2007) The biosynthesis of the two major forms of melanin, black/brown eumelanin and yellow/red pheomelanin is initially catalyzed by tyrosinase. Specifically, the enzyme catalyzes the hydroxylation of the

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monophenol L-tyrosine to the o-diphenol 3,4-dihydroxyphenylalanine (DOPA) and the oxidation of DOPA to the o-quinone DOPAquinone (Fig. 1). (Sturm, Teasdale and Box, 2001)(Garcia-

Borron and Solano, 2002) Several depigmenting agents modulate skin pigmentation by influencing the transcription and activity of tyrosinase as well as related melanogenic enzymes tyrosinase related protein-1 (TYRP-1), tyrosinase related protein-2 (TYRP-2) and/or peroxidase.

(Briganti, Camera and Picardo, 2003)

HO COOH COOH HO COOH NH2 NH 2 NH 2 HO S HO N Tyrosine H 2N Cysteinen e or S Glutathioneion COOH Tyrosinase 1,4 -Benzothiazinyl-alaninei CysteinylDOPAOP A Tyrosinase

HO COOH COOH O NH 2 NH 2 PHEOMELANIN HO O

PA DOPA (Dihydroxyphenylalanine) DOPAquinone

DOPAchrome tautomerase (TYRP-2)

O HO HO + CO HO 2 HO N N COOH HO N COOH H H DHICA DOPAchrome DHI

DHICA oxidase (TYRP-1) Tyrosinase

O O

O N COOH O N H H Indole-5,6-quinone-carboxylicaar acid Indole-5,6-quinone

EUMELANIN

Figure 6.1. Process of melanogenesis within epidermal melanosomes. Tyrosinase, the rate limiting enzyme of melanogenesis, catalyzes the hydroxylation of L-tyrosine to DOPA and the oxidation of DOPA to DOPAquinone. If cysteine or glutathione is present, it reacts with

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DOPAquinone to produce cysteinylDOPA and the benzothiazine derivatives of pheomelanin. As cysteine is diminished, DOPAquinone cyclizes into DOPAchrome. TYRP-2 catalyzes the tautomerization of DOPAchrome to 5,6-dihydroxyindole-2-carboxylic acid (DHICA), which is later oxidized to DHICA-melanin subunits. The oxidation of DHICA to eumelanin is thought to be catalyzed by TYRP-1. In the absence of TYRP-2 the carboxylic acid moiety of DOPAchrome is spontaneously lost to form 5,6-dihydroxyindole (DHI). DHICA in conjunction with DHI comprise subunits of eumelanin. (Yamaguchi, Brenner and Hearing, 2007)(Ito, 2003)

6.3. Transcriptional Regulation of Melanogenic Enzymes

The transcriptional level is the first stage by which the expression of tyrosinase and related melanogenic enzymes may be modulated. Influential in this process, the microphthalmia- associated transcription factor (MITF) is a basic helix-loop-helix leucine zipper transcription factor that regulates melanocyte cellular differentiation as well as the transcription of melanogenic enzymes (tyrosinase, TYRP1 and TYRP2) and melanosome structural proteins

(MART-1 and PMEL17). (Shibahara et al, 2001)(Tachibana et al, 1996)(Levy, Khaled and

Fisher, 2006) Mutations in the MITF gene are associated with auditory and pigmentary abnormalities of Waardenburg syndrome type IIA. (Takeda and Shibahara, 2006)(Steingrimsson,

Copeland and Jenkins, 2004)

The MITF-M isoform, with the promoter most proximally located upstream to the common exon sequences, is exclusively expressed in melanocytes and is believed to bind the M box regulatory element and transactivate the promoter of tyrosinase, TYRP-1 and TYRP-2, as well as other genes. (Ando et al, 2007)(Briganti, Camera and Picardo, 2003)(Shibahara et al, 2001)(Levy,

Khaled and Fisher, 2006)(Lin et al, 2002)(Yasumoto et al, 1994) It is believed that upon stimulatory binding of α-melanocyte stimulating hormone (α-MSH) to the melanocortin 1

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receptor (MC1R), adenyl cyclase is activated and cAMP produced. cAMP then activates the protein kinase A (PKA) pathway to phosphorylate cAMP-responsive element binding protein

(CREB) transcription factors, which mediates MITF-M promoter activation to induce melanogenesis. MITF is also regulated at the transcriptional level by interleukin-6 (IL-6) and

Wnt Signaling pathway. Furthermore, MITF is post-transcriptionally regulated by phosphorylation via ribosomal S6 kinase (RSK), glycogen synthase kinase-3b (GSK3b), p38 stress signaling and mitogen-activated protein kinase (MAPK) pathways by currently undefined mechanisms/pathways.(Levy, Khaled and Fisher, 2006)(Lin et al, 2002)(Solano et al, 2006)(Saha et al, 2006) As MITF is considered an important regulator of melanogenesis, manipulation of the aforementioned signaling pathways may have potential therapeutic use.

Transforming growth factor-β1 (TGF-β1) is a cytokine that plays a role in cell differentiation, proliferation and apoptosis, in addition to the inhibition of pigmentation. TGF-β1 is believed to mediate the down-regulation of the MITF promoter activity, reducing the production of tyrosinase, TYRP-1, TYRP-2 and MITF protein levels. (Solano et al, 2006)(Kim, Park and Park,

2004) Yang et al. have shown that TGF-β1 inhibits the expression of paired-box homeotic gene

(PAX 3), a transcription factor and key regulator of MITF in melanocytes. (Yang et al, 2008)

Kim et al. have also demonstrated that TGF-β1 influences the extracellular-signal related kinase

(ERK) pathway and down-regulates MITF as well as melanogenic enzyme production. (Solano et al, 2006)(Kim, Park and Park, 2004)(Englaro et al, 1998)(Kim et al, 2006) Similarly, ERK activation by sphingosine-1-phosphate, C2-ceramide and sphingosylphosphorylcholine has also been reported by Kim et al., which the authors hypothesize, may play an important role in the inhibition of melanogenesis. (Kim et al, 2006)(Kim et al, 2003b)(Kim et al, 2002) ERK

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activation, in those cases, is thought to result in phosphorylation of MITF and its subsequent ubiquitination and degradation. (Kim et al, 2006)(Hemesath et al, 1998)(Wu et al, 2000)(Xu et al, 2000) In a distinct study, the authors demonstrated that lysophosphatidic acid (LPA), a serum phospholipid released by activated platelets, mediates the reduction of MITF promoter activity as well as MITF and tyrosinase protein and melanin synthesis in Mel-Ab cells, an immortalized mouse melanocyte line. (Kim et al, 2004)

In addition to the process of melanization, MITF also regulates melanocyte differentiation, development, and survival.(Widlund and Fisher, 2003) Pertaining to survival, MITF regulates the anti-apoptotic molecule Bcl-2 as well as additional survival genes.(McGill et al, 2002)(Buscà et al, 2005) It has recently been demonstrated that melanocytes deficient in MITF expression are compromised in their resistance to UV-induced apoptosis. (Hornyak et al, 2009) Therefore, caution is warranted when attempting to decrease skin pigmentation by down-regulating MITF since melanocyte death may be a consequence.

6.4. Post-Translational Modification of Melanogenic Enzymes

A major post-translational modification of melanogenic enzymes is the attachment of N-linked glycans to asparagine residues in Asn-X-Ser/Thr motifs (where X is not Pro), during the polypeptides translocation in the ER. This glycosylation is critical for the proper maturation of tyrosinase.(Briganti, Camera and Picardo, 2003)(Negroiu et al, 1999)(Branza-Nichita et al, 2000)

A detailed review of the processes involved in the N-glycosylation of melanogenic enzymes has been published by Branza-Nichita et al.. (Branza-Nichita et al, 2000) Inhibition of proper N- glycan processing of melanogenic enzymes can result in improper polypeptide folding and in

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turn inhibition of melanogenesis, as they facilitate association with lectin-chaperones. Treatment with various agents that inhibit N-glycosylation can result in the down-regulation of melanosomal enzyme activity and reduced melanosomal maturation. (Garcia-Borron and Solano,

2002)(Briganti, Camera and Picardo, 2003)(Branza-Nichita et al, 2000)(Imokawa and Mishima,

1984)

Studies completed by Mishima and Imokawa using tunicamycin and glucosamine, specific inhibitors of lipid carrier-dependent glycosylation of protein, resulted in decreased pigmentation and ultrastructural as well as biochemical aberrations in melanogenic compartments of treated

B16 melanoma cells. In addition, electron microscopic analysis showed melanosomes with internal structural irregularities and pigment loss. (Briganti, Camera and Picardo, 2003)(Mishima and Imokawa, 1983) In a more recent study, Terao et al. tested a novel compound, BMY-28565, that inhibited melanogenesis by depressing tyrosinase activity with no impact on tyrosinase mRNA levels in B16 melanoma cells. As other active derivatives of the compound cause an increase in protein glycosylation in B16 melanoma cells, the authors hypothesize that the test compound inhibited tyrosinase by modifying the sugar moieties of the enzyme. (Terao et al,

1992)(Land, Ramsden and Riley, 2006) In a distinct study by Choi et al., treatment of HM3KO melanoma cells with deoxynojirimycin, a α-glucosidase inhibitor that disrupts early ER N-glycan processing, and deoxymannojirimycin, an inhibitor of α-1,2-mannosidase which are thought to be responsible for late glycan processing, showed inhibition of glycosylation, transportation of tyrosinase to the melanosome and melanin synthesis. (Choi et al, 2007) Other factors explored for their ability to modulate tyrosinase glycosylation include calcium D-pantetheine-S-sulfonate

(Franchi et al, 2000), ferritin (Maresca et al, 2006) and glutathione (Imokawa, 1989). Glutathione

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induced inhibition of tyrosinase glycosylation, blocks the maturation and transfer of tyrosinase from GERL (Golgi-endoplasmic reticulum-lysosome) - coated vesicles to the pre-melanosome.

Yet, other mechanisms of action proposed for glutathione include (A) the direct inactivation of tyrosinase by chelating copper within the enzyme’s active site, (B) mediating the transition from eumelanogenesis to pheomelanogenesis, as glutathione participates in the conversion of dopaquinone to pheomelanin, (C) antioxidant properties that quench free radicals and peroxides that induce melanin formation and (D) modulating the depigmenting capabilities of melanocytotoxic agents. (Solano et al, 2006)(Villarama and Maibach, 2005)

6.5. Attenuation of Tyrosinase and Related Melanogenic Enzymes Catalytic Activity

6.5.1. Hydroquinone

Hydroquinone (1,4-dihydroxybenzene, HQ) has been the gold standard for treating hyperpigmentation for more than 50 years and has been successfully used to treat melanosis. The compound can be found in wheat, tea, berries, beer and coffee, but is detoxified within the liver into inert compounds.(Parvez et al, 2006)(Nordlund, Grimes and Ortonne, 2006) Hydroquinone is a phenolic compound and depigmenting agent that mainly exerts its effect on melanocytes with active tyrosinase. As HQ dependent melanogenic inhibition requires the presence of active tyrosinase, it is therefore not useful in altering the color of melanin that is previously present in the dermis and epidermis. (Halder and Nordlund, 2006)

The structural similarity between HQ and melanogenic precursors enables HQ’s interaction with tyrosinase. This interaction mediates HQ’s inhibition of tyrosinase by binding histidines or copper at the active site of the enzyme. Additionally, HQ induced generation of reactive oxygen

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species and quinones leads to the oxidative damage of membrane lipids and proteins such as tyrosinase. Hydroquinone is also thought to inhibit pigmentation by depleting glutathione, modifying melanosome formation or reducing DNA and RNA synthesis with concomitant melanosome degradation and melanocyte destruction. (Briganti, Camera and Picardo,

2003)(Solano et al, 2006)(Draelos, 2007)(Picardo and Carrera, 2007)

Traditional hydroquinone formulations contain other constituents that promote a synergistic effect. A popular formulation is comprised of hydroquinone and a corticosteroid to reduce inflammation, along with tretinoin, shown to reduce the atrophy associated with the corticosteroid and remove pigmentation by increasing keratinocyte turnover and the penetration of hydroquinone. (Picardo and Carrera, 2007)(Badreshia-Bansal and Draelos, 2007)

Due to the risks of side effects such as exogenous ochronosis and permanent depigmentation following long term use, hydroquinone has been banned by the European Committee (24th Dir

2000/6/EC) and formulations have been withdrawn from cosmetics and are available only through prescription.(Briganti, Camera and Picardo, 2003)(Solano et al, 2006)(Picardo and

Carrera, 2007)

Other phenolic compounds have been evaluated for their depigmenting capabilities. In fact the chemical structures of several phenolic compounds have been investigated to delineate structure related tyrosinase inhibitory activity. It has been suggested that having a hydroxyl group para to an electron donator group is required for a compound to be an effective alternative substrate for tyrosinase.(Briganti, Camera and Picardo, 2003) Distinct structure-activity based analysis done

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by Ni-Komatsu et al. on quinolines, which contain a 4-substituted amino group with a tertiary amine side chain, shows significant inhibitory effect. Yet these quinolines, such as chloroquine, were not reported to influence the enzymatic activity of tyrosinase, but rather the intracellular trafficking of tyrosinase related proteins and lysosome associated membrane protein -1 (LAMP-

1).(Ni-Komatsu et al, 2008)

6.5.2. Monobenzylether

The mono benzyl ether of hydroquinone (MBEH) is a related compound that is metabolized within the cell to form a quinone species that interacts with and results in permanent depigmentation, even at areas distant from the site of application. MBEH can destroy melanocytes and should not be used to treat post-inflammatory hyperpigmentation or melasma.

MBEH therapy is appropriate for generalized depigmentation in the treatment of patients with vitiligo unresponsive to repigmentation therapy. (Briganti, Camera and Picardo, 2003)(Solano et al, 2006)(Halder and Nordlund, 2006) Proposed mechanisms of action for MBEH are both cytotoxicity to melanocytes as a result of free radical formation and competitive inhibition of tyrosinase activity. (Parvez et al, 2006)

6.5.3. Arbutin and Deoxyarbutin

Arbutin (hydroquinone-O-beta-D-glucopyranoside), a derivative of hydroquinone, is a botanically derived compound found in cranberries, blueberries, wheat and pears. (Parvez et al,

2006)(Badreshia-Bansal and Draelos, 2007) Arbutin is used as an effective treatment of hyperpigmentary disorders, and displays less melanocyte cytotoxicity than hydroquinone. The compound inhibits melanogenesis by competitively and reversibly binding tyrosinase without

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influencing the mRNA transcription of tyrosinase. It also inhibits the maturation of melanosomes, possibly by its reported influence on DHICA polymerase activity and Pmel-17 protein. The mild effect of arbutin is attributed to the controlled release of hydroquinone as a result of in-vivo cleavage of the glycosidic bond. Higher concentrations of arbutin are more efficacious than lower concentrations, but may cause paradoxical hyperpigmentation. (Solano et al, 2006)(Imokawa, 1989)(Draelos, 2007)(Badreshia-Bansal and Draelos, 2007)(Chakraborty et al, 1998) Deoxyarbutin (dA), a synthetic form of arbutin synthesized without the hydroxyl moiety, provides a promising treatment for reducing skin hyperpigmentation.(Picardo and

Carrera, 2007) dA shows reversible inhibition of tyrosinase activity with associated skin lightening in both a hairless guinea pig model system and in human skin. The reversibility of dA’s impact on skin pigmentation suggests that the compound does not permanently destroy melanocytes. (Solano et al, 2006)(Chawla et al, 2008)(Boissy, Visscher and DeLong, 2005) In addition to the reported efficacy, Hamed et al. have found that dA is less cytotoxic/cytostatic than HQ in treatment of cultured human melanocytes.(Hamed et al, 2006) Chawla et al. have reported that dA and associated second-generation derivatives, dose-dependently inhibit tyrosine hydroxylation and DOPAoxidase activity of tyrosinase. This may be attributed to the chemical structure of dA, as the deoxysugars may increase skin penetration and binding affinity for tyrosinase. (Chawla et al, 2008)(Solano et al, 2006)

6.5.4. Mequinol

Mequinol (Hydroquinone monomethyl ether, 4-hydroxyanisole, para-hydroxy-methoxy benzene), another derivative of hydroquinone, is enzymatically oxidized by tyrosinase to produce melanocytotoxic quinones. The formation of quinones results in pigment cell destruction

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and skin depigmentation.(Briganti, Camera and Picardo, 2003)(Petit and Pierard, 2003) The combination of 0.01% tretinoin with mequinol has been reported to inhibit melanin production and has been shown to be effective and safe in the treatment of solar lentigenes and related hyperpigmentation. (Halder and Nordlund, 2006)(Picardo and Carrera, 2007)(Colby et al,

2003)(Keeling et al, 2008)

6.5.5. N-Acetyl-4-S-Cysteaminylphenol

N-Acetyl-4-S-Cysteaminylphenol (NCAP) is a phenolic thioether that has been used in the treatment of epidermal hyperpigmentation such as melasma and also in anti-melanoma studies.

NCAP is structurally similar to tyrosine and acts specifically in melanin-synthesizing cells as an alternative tyrosinase substrate. (Halder and Nordlund, 2006)(Picardo and Carrera, 2007)

(Jimbow, 1991) Ferguson et al. suggests that NCAP may undergo oxidation by tyrosinase to form a reactive o-quinone that is capable of alkylating thiol groups of essential enzymes, which may interfere with cell growth and proliferation. (Ferguson et al, 2005) In a 12 patient study of the efficacy of NCAP completed by Jimbow, the author describes an 8% complete loss and 66% marked improvement in visible changes of melanoderma after 2 to 4 weeks of topical application. The author attributes this depigmentation to a decrease in the number of functional melanocytes and the number of melanosomes transferred to keratinocytes. NCAP is suggested to be a depigmenting agent that is less irritating and more stable than HQ, with specificity for melanin-synthesizing cells. (Jimbow, 1991)

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6.5.6. Kojic Acid

Kojic Acid (5-hydroxy-2-hydroxymethyl-4H-pyran-4-one, KA) is a naturally occurring hydrophilic fungal metabolite obtained from species of Acetobacter, Aspergillus and Penicillium.

(Halder and Nordlund, 2006)(Draelos, 2007) Kojic acid is believed to inactivate tyrosinase by chelating copper atoms as well as suppressing the tautomerization of dopachrome to

DHICA.(Picardo and Carrera, 2007) Although KA is a popular treatment for melasma, it is associated with sensitization, contact dermatitis and erythema.(Badreshia-Bansal and Draelos,

2007) A distinct, more stable derivative of kojic acid synthesized by Kim et al., 5-[(3- aminopropyl)phosphinooxy]-2-(hydroxymethyl)-4H-pyran-4-one (Kojyl-APPA), showed increased skin penetration and pigment lightening efficacy in melanoma and normal human melanocytes. (Solano et al, 2006)(Kim et al, 2003a)

6.5.7. Azelaic Acid

Azelaic acid (1,7-heptanedicarboxyilic acid, AZA) is a naturally occurring non-toxic straight chain, saturated dicarboxylic acid derived from Pityrosporum ovale. (Petit and Pierard,

2003)(Kim and Uyama, 2005)(Parvez et al, 2007) AZA appears to selectively influence the mechanism of hyperactive and abnormal melanocytes, but minimally influences normal skin pigmentation, freckles, nevi and senile lentigenes.(Parvez et al, 2006)(Briganti, Camera and

Picardo, 2003)(Grimes, 1995) AZA’s antiproliferative and cytotoxic effect may be mediated by the inhibition of DNA synthesis and mitochondrial oxidoreductase activity. The compound is also able to bind amino and carboxyl groups and may prevent the interaction of tyrosine in the active site of tyrosinase and thus function as a competitive inhibitor. Although not all authors are in agreement with the therapeutic efficacy of AZA, it has been reported to be effective in

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treatment of melasma and acne. (Briganti, Camera and Picardo, 2003)(Grimes, 1995) A 24 week multicenter, controlled, double blind clinical trial of 329 women completed by Baliña et al. compared the efficacy of a 20% AZA cream to a 4% HQ cream in treating melasma. The authors reported no significant difference between the results where ~65% of the patients treated with

AZA were reported to achieve good to excellent results compared to ~73% of HQ treated patients.(Baliña and Graupe, 1991) 20% azelaic acid seems to be well tolerated in treated patients with no systemic side effects, but some local cutaneous irritation, a burning sensation, mild erythema, scaling and pruritus that subsided 2 to 4 weeks post treatment. (Grimes, 1995)

(Fitton and Goa, 1991)

6.5.8. Gentisic Acid

The methyl ester of gentisic acid (2,5-dihydroxybenzoic acid, MG) is a natural derivative of

Gentianas root with the capacity to inhibit tyrosinase. MG can act as a pro-drug that releases HQ which subsequently inhibits tyrosinase. Yet, methyl gentisate is less cytotoxic and mutagenic than HQ. (Briganti, Camera and Picardo, 2003)(Solano et al, 2006)(Picardo and Carrera, 2007)

6.5.9. Flavonoid-like Agents

Flavonoids are natural plant polyphenols found in leaves, bark and flowers. 4000 members have been identified to date, all benzo-γ-pyrane derivatives are comprised of phenolic and pyrane rings. These polyphenolic compounds are known to have anti-inflammatory, antiviral, antioxidant and anticarcinogenic properties. (Solano et al, 2006)(Picardo and Carrera, 2007)(Kim and Uyama, 2005) The flavonoids may also have ROS scavenging properties and the ability to chelate metals at the active site of metalloenzymes. Flavonoids may have hypopigmenting

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capabilities by directly inhibiting tyrosinase activity at distal portions of the melanogenic pathway. (Solano et al, 2006)Structure-function analysis of flavonoids suggests that flavonoids with an α-keto group show potent tyrosinase inhibition due to the similarity between the dihydroxyphenyl group of DOPA and the α-keto containing flavonoids. (Kim and Uyama, 2005)

Similar analysis completed by Kubo et al. suggests that flavonoids have the capability to chelate copper in tyrosinase’s active site as long as the 3-hydroxygroup is free. (Parvez et al, 2007)

(Kubo et al, 2000) A comprehensive review on the properties of plant polyphenols has been published by Kim et al. (Kim and Uyama, 2005) Flavonoids that will be reviewed here include aloesin, hydroxystilbene derivates and licorice extracts.

6.5.10. Aloesin

Aloesin is a natural hydroxymethyl chromone compound derived from aloe vera plants.(Draelos,

2007) It competitively inhibits the function of tyrosinase by inhibiting the hydroxylation of tyrosine to DOPA and oxidation of DOPA to dopaquinone. (Picardo and Carrera, 2007) Studies completed by Jones et al. on normal human melanocytes treated with aloesin, showed a dose dependent decrease in tyrosinase activity.(Jones et al, 2002) The hydrophilic nature of the compound reduces the skin penetration of aloesin. Hence, combination treatment of aloesin with arbutin has been studied by Jin et al. to assess the synergistic effects on tyrosinase activity. The authors report that the two adhere to different mechanisms of action where aloesin exhibits noncompetitive inhibition while arbutin inhibits competitively.(Draelos, 2007)(Parvez et al,

2007)(Jones et al, 2002)(Zhu and Gao, 2008)(Jin et al, 1999) Testing of aloesin revealed no cytotoxicity, which makes it a good alternative to HQ.(Picardo and Carrera, 2007)

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6.5.11. Hydroxystilbene

One of the more efficient pigment lightening flavonoid subcategories is the hydroxystilbene compounds, that are derived from natural products found in oriental herbal medicines. There are more than 30 stilbene and stilbene glycosides with a structural skeleton comprised of two aromatic rings linked by an ethylene bridge.(Solano et al, 2006)(Kim et al, 2004) Commonly studied hydroxystilbene products include resveratrol (3,4’,5 trihydroxystilbene), its isomer oxyresveratrol and methoxylated or glycosylated analogs piceid-glucoside, rhapontigenin and rhaponticin. (Solano et al, 2006)(Fremont, 2000) The number and position of hydroxyl substituents of hydroxystilbene compounds seem to play an important role on the inhibition of tyrosinase activity.(Kim et al, 2002) However, glycosylated hydroxystilbene compounds such as piceid, the glycoside of resveratrol at position 3, exhibit decreased tyrosinase inhibition compared to the parent compound.(Kim et al, 2004) In all cases, tyrosinase inhibition is reversible and in turn requires a high concentration of hydroxylated stilbenes within melanocytes. (Solano et al, 2006) Resveratrol, a commonly studied hydroxystilbene, is found in red wine and displays free radical scavenging, anticancer and anti-inflammatory activities.(Briganti, Camera and Picardo, 2003) Some data attribute resveratrol depigmenting affect to its ability to reduce Mitf and tyrosinase promoter activities in B16 mouse melanoma cells.(Lin et al, 2002)(Solano et al, 2006) However, other contradictory results suggest that resveratrol treated normal human melanocytes (NHM) display steady-state tyrosinase RNA and, as such, regulation of tyrosinase transcription does not influence its depigmentation.(Newton et al, 2007) Additionally, further analysis of the resveratrol treated NHM displayed ER-retained immature tyrosinase, suggesting disrupted trafficking of tyrosinase in the GERL and elevated

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proteolytic degradation. (Newton et al, 2007) The resveratrol analog oxyresveratrol is a stronger inhibitor than resveratrol. Oxyresveratrol has been described to have a potent dose-dependent non-competitive inhibition on L-tyrosine oxidation by tyrosinase from mushroom and murine melanoma B-16, without suppression of tyrosinase synthesis or expression. (Kim et al, 2002)

The hydroxystilbene products still require more analysis to properly elucidate their pigment lightening effect and mechanism(s) of action.

6.5.12. Licorice Extract

Licorice extract is obtained from the root of Glycyrrhia Glabra Linneva and has been used in traditional Chinese medicine.(Parvez et al, 2006)(Solano et al, 2006)(Halder and Nordlund,

2006) The main component of the hydrophobic fraction of licorice is glabridin. Glabridin has been shown to prevent UVB-induced pigmentation and to inhibit tyrosinase activity, superoxide anion production and cyclo-oxygenase activity. This suggests an influence of glabridin extract on both melanogenesis and inflammation of the skin. (Halder and Nordlund, 2006)(Picardo and

Carrera, 2007) Other active agents of licorice extracts include liquiritin and isoliquertin.

Liquiritin is a glycoside containing flavonoid that induces skin lightening by dispersing melanin.(Draelos, 2007) Licorice extracts also influence pigmentation by removing epidermal melanin, inhibiting the biosynthesis of melanin and inhibiting the activity of tyrosinase in a dose- dependent manner. Licorice extract has been tested in the treatment of melasma with good results and very mild irritation.(Picardo and Carrera, 2007)(Badreshia-Bansal and Draelos, 2007)

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6.5.13. Antioxidants and Redox Agents

Ultraviolet radiation influences the proliferation of melanocytes and the production and secretion of paracrine and autocrine factors that stimulate melanogenesis. UV radiation can also produce reactive oxygen species (ROS) in the skin that may induce melanogenesis, DNA damage, melanocyte proliferation and/or apoptosis.(Yamakoshi et al, 2003)(Hachiya et al, 2004) The skin contains a number of antioxidants that can be depleted by UV exposure and cause oxidative damage. The application of topical antioxidants has the capacity to prevent oxidative damage to the skin.(Berson, 2008) Redox agents capable of scavenging ROS generated in the skin can inhibit second messengers that may stimulate melanogenesis. Redox agents can also influence skin pigmentation by interacting with copper at the active site of tyrosinase or with o-quinones to impede the oxidative polymerization of melanin intermediates. (Briganti, Camera and Picardo,

2003)(Karg et al, 1993) Here we will review vitamin C, vitamin E and later we will review the vitamin B derivative niacinamide.

6.5.14. L-Ascorbic Acid and Magnesium-L-Ascorbyl-2-Phosphate

L-Ascorbic Acid (vitamin C, AsA), obtained from citrus fruits and leafy green vegetables, is a water soluble vitamin and the most plentiful antioxidant in human skin.(Badreshia-Bansal and

Draelos, 2007)(Farris, 2005) AsA interferes with melanin synthesis by reducing oxidized dopaquinone, interrupting DHICA oxidation and interacting with copper ions at the active site of tyrosinase. AsA acts as an ROS scavenger by donating electrons to neutralize free radicals found in the aqueous compartment of the cell. (Briganti, Camera and Picardo, 2003)(Farris, 2005)

Unfortunately, AsA is highly unstable and is rapidly oxidized and decomposed in aqueous solutions. The hydrophilic nature of AsA also limits its skin penetration, unless the stratum

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corneum barrier is disrupted. (Briganti, Camera and Picardo, 2003)(Badreshia-Bansal and

Draelos, 2007) The more stable ascorbate ester, magnesium-L-ascorbyl-2-phosphate (MAP) is more lipophilic and has a greater permeation through the stratum corneum.(Picardo and Carrera,

2007)(Badreshia-Bansal and Draelos, 2007) MAP is then hydrolyzed by phosphatases in the skin to AsA and demonstrates the reducing capabilities of AsA.(Kameyama et al, 1996) In related skin lightening studies, Hakozaki et al. have used ultrasound to increase the transepidermal penetration and efficacy of the vitamin C derivative, ascorbyl glucoside and niacinamide.(Hakozaki et al, 2006)

6.5.15. Alpha Tocopherol and Alpha Tocopherol Ferulate

Vitamin E (Alpha Tocopherol, α-Toc) is a lipophilic antioxidant in the body, derived from cereal, vegetables, vegetable oil and nuts. (Badreshia-Bansal and Draelos, 2007) α-Toc is known to inhibit the oxidative attack of free and membrane bound unsaturated fatty acid and interferes with the lipid peroxidation of melanocyte membranes. The vitamin is also able to scavenge free radicals including superoxide anions, hydroxyl radicals and singlet molecular oxygen. It can also act as a humectant. The depigmenting effect of α-Toc can further be attributed to an increase in intracellular glutathione and the inhibition of tyrosinase. (Badreshia-Bansal and Draelos, 2007)

(Shimizu et al, 2001)(Fukuzawa and Gebicki, 1983)(Choi and Berson, 2006) α-Tocopherol ferulate (α-TF) is a related compound, in which α-tocopherol is linked by an ester bond to ferulic acid, an antioxidant that stabilizes α-Toc. (Briganti, Camera and Picardo, 2003) The presence of another antioxidant ferulic acid, causes a rapid regeneration of α-Toc and maintains a long lasting antioxidant effect. A study completed investigating the biochemical effect of α-TF in human melanoma cells suggests the whitening effect is due to tyrosinase inhibition at the post-

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transcriptional level, possibly by an unidentified secondary molecule.(Picardo and Carrera,

2007)(Funasaka et al, 1999)

6.6. Interruption of Melanosome Transfer

Regulation of cutaneous pigmentation is dependent on several processes beyond melanin synthesis within the melanosome. The efficiency of melanosomal transfer from melanocytes to keratinocytes, followed by melanosome processing in the recipient keratinocytes plays a critical role in skin pigmentation. Without successful transfer of melanosomes to keratinocytes, the skin can appear essentially unpigmented.(Minwalla et al, 2001a) Treatment modalities aimed at inhibiting melanosome transfer may influence and modulate skin pigmentation.

6.6.1. Centaureidin and Methylophiopogonanone B

The first step of melanosome transfer from melanocytes to surrounding keratinocytes is successful melanocytic dendrite formation and extension towards surrounding keratinocytes. The extension of melanocytic dendrites requires the reorganization of the melanocyte cytoskeletal elements such as actin filaments and microtubules.(Ito, Kanamaru and Tada, 2006a) Small

GTPases Rho, Rac and Cdc42 play a pivotal role in cell morphology and dendrite formation.

Specifically, Rac stimulates membrane ruffling and lamellipodia formation, Rho activates dendrite retraction and Cdc42 mediates filopodia and peripheral actin microspike formation. (Ito,

Kanamaru and Tada, 2006a)(Ito, Kanamaru and Tada, 2006b)(Ito, Kanamaru and Tada, 2007)

Ito et al. has shown that treatment of melanocyte and keratinocyte co-cultures with methylophiopogonanone B (5,7-dihydroxy-6,8-dimethyl-3-(4-methoxybenzyl)chroman-4-one,

MOPB), an agent reported to activate Rho and induce microtubule disorganization and tubule

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depolymerization, appeared to reduce melanosome transfer. (Ito, Kanamaru and Tada,

2006b)(Ito, Kanamaru and Tada, 2007) The authors also showed that treatment with 1µM MOPB did not influence melanin synthesis or the expression of melanogenic enzymes. MOPB appeared to induce a reversible dendrite retraction and transfer inhibition without associated cytotoxicity

(tested up to 72 hours).(Ito, Kanamaru and Tada, 2006b) Centaureidin (5,7,3’-trihydroxy-3,6,4’- trimethoxyflavone), a flavonoid glucoside derived from yarrow, also reduces melanosomal transfer to keratinocytes. Centaureidin is believed to directly or indirectly activate Rho, leading to melanocyte dendrite retraction without influencing melanogenic enzyme expression or melanin synthesis.(Ito, Kanamaru and Tada, 2006a)(Lin et al, 2008) More analysis is needed to confirm the applicability of MOPB and centaureidin as skin lightening agents.

6.6.2. Niacinamide

Niacinamide (vitamin B3, nicotinamide, 3-pyridinecarboxamide) is a biologically active form of niacin found in many root vegetables as well as in yeast.(Zhu and Gao, 2008)(Hakozaki et al,

2002) Physiologically, niacinamide functions as a precursor to the co-factors nicotinamide adenine dinucleotide (NAD) and nicotinamide adenine dinucleotide phosphate (NADP). Along with their reduced forms NADH and NADPH, these enzymes participate in numerous enzymatic reactions and also act as antioxidants. (Zhu and Gao, 2008)(Hakozaki et al, 2002) Niacinamide has several proposed medicinal applications in the skin including anti-inflammation, prevention of photoimmunosuppression and increased intercellular lipid synthesis.(Greatens et al, 2005)

Niacinamide’s role as a co-enzyme precursor may explain the multiple roles it has in skin, but this is not clearly defined.(Hakozaki et al, 2006)(Bissett et al, 2004) Topical niacinamide is described to have several benefits on aging skin including but not limited to improved barrier

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function, improved appearance of photoaged facial skin (including texture, hyperpigmentation, redness, fine lines and wrinkles) and reduced sebum production. (Hakozaki et al, 2002)(Bissett et al, 2004)(Bissett, Oblong and Berge, 2005)(Bissett, 2002)(Bissett et al, 2003) Additionally, niacinamide is believed to influence cutaneous pigmentation by down-regulating transfer of melanosomes from the melanocytes to the keratinocytes. (Solano et al, 2006)(Petit and Pierard,

2003)(Hakozaki et al, 2002) Studies completed by Hakozaki et al. suggest that niacinamide has no effect on tyrosinase activity, melanin synthesis or melanocyte number in a monolayer culture system. Alternatively, the authors found that niacinamide down-regulated the number of melanosomes transferred from melanocytes to keratinocytes by 35 to 68% in a co-culture model system. (Hakozaki et al, 2002) The actual process by which niacinamide down-regulates melanosome transfer remains to be properly established.

6.6.3. PAR-2 Inhibitors

PAR-2 belongs to a family of transmembrane G-protein coupled receptors (PAR1-PAR4) that are proteolytically activated by serine proteases. Specifically, the serine proteases (including trypsin or mast cell tryptase) cleave the extracellular amino terminal domain exposing a newly created N-terminus tethered ligand that undergoes a conformational change, binds and subsequently activates the receptors.(Van Den Bossche, Naeyaert and Lambert, 2006)(Seiberg et al, 2000a) Within the epidermis, PAR-2 is expressed in keratinocytes, but not in melanocytes and is involved in the regulation of skin pigmentation through melanocyte-keratinocyte interactions.

(Seiberg et al, 2000a)(Lin et al, 2008)(Derian, Eckardt and Andrade-Gordon, 1997)(Marthinuss,

Andrade-Gordon and Seiberg, 1995)(Seiberg et al, 2000b) Studies completed on keratinocyte

PAR-2 indicate that it may influence melanosome incorporation and phagocytosis by

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keratinocytes and play a regulatory role in skin pigmentation. Therefore, modulation of PAR-2 activity augments or decreases melanosome transfer and in turn skin pigmentation. (Seiberg et al,

2000a)(Seiberg et al, 2000b)(Babiarz-Magee et al, 2004)(Sharlow et al, 2000)(Paine et al, 2001)

PAR-2 activation can be achieved by synthetic peptides corresponding to the sequence of the N- terminal ligand. The peptide of mouse PAR-2 cleavage sequence SLIGRL is an equipotent activator of mouse and human PAR-2 receptor in comparison to the human tethered ligand

SLIGKV. (Lin et al, 2008) Interestingly, PAR-2 expression and induction by ultraviolet irradiation is dependent on skin type, with a higher overall expression and induction in darker skin individuals. (Van Den Bossche, Naeyaert and Lambert, 2006)(Babiarz-Magee et al,

2004)(Scott et al, 2001) Activation of PAR-2 enhances melanosome transfer, while inhibition of

PAR-2 by serine protease inhibitors can result in reduced melanosomal transfer and distribution.

Work published by Seiberg et al. and Paine et al. has shown that this inhibition leads to a dose- dependent lightening of skin pigmentation. (Seiberg et al, 2000b)(Babiarz-Magee et al,

2004)(Paine et al, 2001)Work completed by Seiberg et al. examining the effect of serine protease inhibitor RWJ-50353 on epidermal equivalents shows an accumulation of melanosomes in melanocytes, with an increase in early stage melanosomes compared to untreated controls. They hypothesize that the keratinocytes inability to receive the presented melanosomes leads to an accumulation of melanosomes in the melanocytic dendrite and a concomitant negative feedback mechanism that slows pigment production. The authors additionally showed that Yucatan swine skin treated with RWJ-50353 for an 8 week period demonstrated a dose dependent, reversible skin lightening effect. (Seiberg et al, 2000b)

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“Natural” therapies derived from soybeans have been explored for their safety and efficacy as depigmentation treatments. Two protein proteinase inhibitors have been isolated from soybean seeds Kunitz-type trypsin inhibitor (soybean trypsin inhibitor, STI) and the Bowman-Birk protease inhibitor (BBI). STI and BBI are found in the seeds of soybeans, but not in the other regions of the plant. STI inhibits trypsin proteolysis by forming a stable stoichiometric complex and BBI inhibits trypsin and chymotrypsin at separate reactive sites. (Paine et al, 2001)(Birk,

1985) Work completed by Paine et al. has shown that soymilk and soybean extract reduces pigmentation in dark skinned Yucatan swine treated for an 8 week period. The authors suggest that the soymilk inhibits PAR-2 activation in the skin and results in skin depigmentation.

Moreover, the authors suggest that STI and BBI inhibit PAR-2 activation, causes a reduction in keratinocyte phagocytosis and a reduction in resulting skin pigmentation. (Paine et al, 2001)

Soymilk also contains other constituents that may induce skin lightening such as trace amounts of free fatty acids and their acyl CoA esters that can inhibit trypsin and may participate in PAR-2 inhibition. Soybeans contain isoflavones, which are antioxidants that may reduce tyrosinase’s

DOPA oxidase activity. In addition, soybeans contain phospholipids which the authors suggest may assist in the epidermal delivery of STI and BBI without the assistance of liposomes, when soy milk is used as a topical treatment. (Badreshia-Bansal and Draelos, 2007)(Paine et al, 2001)

6.6.4. Lectins and Neoglycoproteins

Cellular recognition between melanocytes and keratinocytes is an important event involved in melanosome transfer. (Minwalla et al, 2001a) Lectins and neoglycoproteins have been explored as candidates that are involved in this phenomenon, because of their influence in cellular processes including intracellular trafficking, endocytosis and cell-cell recognition. (Minwalla et

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al, 2001a)(Greatens et al, 2005) Interestingly, Minwalla et al. have demonstrated a role for melanocyte and keratinocyte membrane glycosylated residues in the process of receptor- mediated endocytosis to facilitate melanosome transfer. The authors suggest that lectins and neoglycoproteins play an inhibitory role in this process. (Solano et al, 2006)(Minwalla et al,

2001a) Specifically, plasma membrane lectins and their glycoconjugates are thought to interrupt melanocyte and keratinocyte contact and interaction, by binding their specific plasma membrane receptors, inhibiting melanosome transfer. (Brenner and Hearing, 2008) This inhibition is reversible and is shown to be enhanced by the presence of niacinamide.(Greatens et al, 2005)

6.7. Acceleration of Epidermal Turnover and Desquamation

The capability of a compound to accelerate the turnover of epidermal layers and/or disperse melanin pigment can result in skin lightening. Chemical agents used to exfoliate the skin, stimulates the removal of pigmented upper layer keratinocytes to lighten skin. (Briganti, Camera and Picardo, 2003)(Zhu and Zhang, 2006)

6.7.1. α-Hydroxyacids

α-Hydroxyacids (AHA) are weak organic acids found in fruits, plants and milk sugars. (Bowe and Shalita, 2008) For centuries, α-hydroxyacids have been one of the most commonly utilized peeling agents used to treat dry skin, acne, actinic damage and to improve skin color/texture.

(Badreshia-Bansal and Draelos, 2007) AHAs are also reported to effectively treat pigmentary lesions such as solar lentigenes, melasma and post inflammatory hyperpigmentation (PIH). At low concentrations AHAs promotes exfoliation by decreasing corneocyte cohesion and stimulating new growth in the basal layer, while at higher concentrations AHAs promote

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epidermolysis and dispersed basal layer melanin. The accelerated desquamation of the stratum corneum by AHAs is complemented by a direct inhibition of tyrosinase, without influencing mRNA or protein expression. (Badreshia-Bansal and Draelos, 2007)(Zhu and Zhang,

2006)(Bowe and Shalita, 2008)

Lactic acid (LA) and glycolic acid (GA) are AHAs derived from sour milk and sugarcane juice, respectively. (Zhu and Zhang, 2006) In a study investigating the histological differences between

Japanese subjects treated for 6 weeks with 40% AHA, either glycolic, lactic or acetic acids,

Yamamoto et al. reported an increase in epidermal thickness, decreased melanin deposition and up-regulated collagen levels. The authors also suggest that the AHAs induced a remodeling of the epidermis with accelerated desquamation.(Yamamoto et al, 2006)

6.7.2. Salicylic Acid

Salicylic Acid (SA) is a beta-hydroxy acid found in willow bark and sweet birch. It is also a phytohormone, a plant product that acts similar to a hormone and regulates cell growth and differentiation. SA functions as a desquamating agent that penetrates and dissolves the intercellular matrix of the stratum corneum.(Picardo and Carrera, 2007)(Bowe and Shalita, 2008)

6.7.3. Linoleic Acid

Unsaturated fatty acids including oleic acid (C18:1), linoleic acid (C18:2) or α-linolenic acid

(C18:3) suppresses melanogenesis and tyrosinase activity, while saturated fatty acids such as palmitic acid (C16:0) or stearic acid (C18:0) increases it.(Ando et al, 1999) Linoleic acid reduces the activity of tyrosinase in melanocytes, while mRNA levels remain unchanged. (Badreshia-

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Bansal and Draelos, 2007) No evidence of change in TYRP-1 and TYRP-2 protein levels suggest that fatty acids selectively target tyrosinase. This may influence the enzyme’s degradation via a physiologic proteasome-dependent mechanism, altering the tyrosinase protein content in hyperactive melanocytes. (Briganti, Camera and Picardo, 2003)(Halaban et al, 1997) Linoleic acid also influences skin pigmentation by stimulating epidermal turnover and increased desquamation of melanin pigment from the epidermis. (Badreshia-Bansal and Draelos, 2007)

Studies completed to assess the skin lightening capabilities of unsaturated fatty acids, linoleic acid or α-linoleic acid, on UV induced hyperpigmentation of brown guinea pig skin, showed an efficient lightening effect.(Ando et al, 1998) It is thought that the unsaturated bonds of these molecules can be easily peroxidized, which in combination with an increase in epidermal turnover, correlate with an inhibitory effect on melanogenesis in vivo. (Badreshia-Bansal and

Draelos, 2007)(Ando et al, 1998)

6.7.4. Retinoids

Retinoids are a common treatment option used to ameliorate acne, photodamage and PIH. The mechanism of action likely involves the inhibition of tyrosinase, the dispersion of keratinocyte pigmented granules, reduction in pigment transfer and a reduction in corneocyte cohesion with an associated acceleration of epidermal turnover. (Solano et al, 2006)(Badreshia-Bansal and

Draelos, 2007)(Nair et al, 1993) Tretinoin (all-trans retinoic acid) is a derivative of vitamin A that is thought to have an inhibitory effect on tyrosinase transcription.(Berardesca et al, 2008)

Tretinoin is reported to be effective in treating melasma, with some associated side effects including erythema, peeling at the site of application and PIH. (Solano et al, 2006) Tretinoin is also used in combination in topical creams, such as a formulation proposed by Kligman and

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Willis containing 5% HQ, 0.1% tretinoin and 0.1% dexamethasone. Tretinoin in this formulation acts as a stimulant of epidermal turnover and pigment reduction via epidermopoieses, an antioxidant to reduce the oxidation of HQ and a mild irritant to enhance the epidermal penetration of HQ. (Zhu and Zhang, 2006)

6.8. Conclusions

Great advances have been made in understanding the cellular and biochemical mechanisms in pigment biology and the processes underlying skin pigmentation. This has led to the development of various skin lightening agents to reduce skin hyperpigmentation. While several agents target the rate limiting enzyme of melanogenesis tyrosinase, there has been an increased interest in alternative hypopigmenting mechanisms. Yet, as addressed by Lei et al., there is a need for a standardized and streamlined protocol to screen melanogenic regulatory compounds, to simplify the difficult task of product comparison.(Lei et al, 2002) Also, as the number of putative depigmenting agents grows there is an increased need for studies to clarify product efficacy, cytotoxicity, topical skin penetration, stability and safety. To add to the complexity, it may be more advantageous to test compounds together to address the synergistic effects on skin lightening, particularly when the active components influence distinct steps of melanogenesis.(Briganti, Camera and Picardo, 2003) While it is clear that great progress has been made in the study of skin lightening, it is even more apparent that there is a great deal of work still left to be done.

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CHAPTER 7

CONCLUSIONS AND FUTURE RECOMMENDATIONS

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7.1. Conclusions

Presently, through various analytical techniques we have demonstrated that the degradation of melanosomes is accomplished at a differential rate between light and dark skin. This differential rate suggests a distinction in how the melanosomes are processed with an apparent enhancement of processing by light skin derived KCs. Qualitative analysis using confocal microscopy showed that LKCs and DKCs were able to incorporate fluorescently labeled melanosomes that were shuttled to a perinuclear location within the cytoplasm of the keratinocytes. After 48 hours it is apparent that LKCs, comparative to DKCs, were devoid of pigmentation as indicated by a reduction in fluorescent labeling (CFDA and MEL-5). Quantitative assessment of melanosome degradation by flow cytometry reinforces the findings from the qualitative analysis. Specifically, flow analysis showed that DKC degrades the melanosomes at a slower rate than LKCs over a 48 hour time frame. This information was gathered from four separate experiments utilizing four distinct LKC and DKC lines. The ratio comparing baseline to 48 hour SKMEL-188 derived melanosome values in DKCs to LKCs were 2.663:3.404, 2.585:3.401, 1.398:1.806, and

3.121:4.735. These values indicate that DKCs retained a higher concentration of melanosomes over a 48 hour experimental period, again suggesting an enhanced rate of degradation in LKCs.

To further delineate how melanosome disintegration occurs we explored acid hydrolases that are implicated in the process of epidermal cornification and terminal differentiation. Candidate enzymes were identified using microarray analysis of suprabasal LS and DS captured by LCM.

ACPP was found to be over-expressed in DS at the gene level, by microarray analysis, and protein level, by WB analysis. We hypothesized that the enzyme was amplified in DS to degrade a more resilient/resistant melanosome in DS compared to a more easily degradable LS derived

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melanosomes, but this warrants further investigation. A major finding of our research displayed that Cath L2 was over-expressed in LS relative to DS in both microarray and WB analyses. IIF of Cath L2 expression in cryosections of LS and DS epidermis showed a dramatic over- expression in LS. In concurrence with the aforementioned results, biochemical analysis of Cath

L2 activity showed a 1.75 fold higher activity level in LS compared to DS samples, P=0.03.

Finally, to elucidate the associations of Cath L2 with melanosome processing, we completed

IEM analysis. IEM showed that Cath L2 occasionally localized with melanosomes in the cytoplasm of the KC. Furthermore, when comparing the number of gold particles, we found a much higher expression level in LS, specifically 4.2 fold in the SB, 8.1 fold in the SS, 9.4 fold in the SG, and 2.1 fold in the SC; all higher in LS compared to DS. Taken together, we have established a model system to study melanosome disintegration, which seems to be enhanced in

LS, and have identified potential components that may be influential in the process of melanosome degradation.

7.2. Future Recommendations

Application of the both the qualitative and quantitative model systems to CFDA/Mel-5

labeled pheomelanosomes to observe the persistence or degradation of pheomelanosomes by

LKC and DKC. This can be compared to the analyses that have already been completed

using SKMEL-188 derived eumelanosomes.

Determine by various biochemical analysis, the activity of other candidate acid hydrolases,

shown here to be differentially expressed in light skin and dark skin, including Cath D and

ACPP.

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Using enzyme inhibitors selective for specific enzymes, such as Cath V, and specific classes

of enzymes, such as cysteine proteases, to modulate their expression in KC cultures. These

LKC and DKC cultures would subsequently be utilized in our model system to see the

influence on pigment processing compared to an untreated control.

Testing the influence of acid hydrolases, by selectively inhibiting candidate enzymes, and

assessing the influence on melanosome degradation in a model system that closely resembles

in vivo circumstances. Organotypic cultures derived from skin biopsies or xenograft of

human skin on immunodeficient (SCID) mice can be treated with lysosomal hydrolase

specific inhibitors to clarify the role of these enzymes in melanosome degradation, in a

model system that recapitulates in vivo conditions.

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APPENDIX

175

Appendix I: Supplemental Data to Chapter 4.

AI.1. Structure of CFDA-SE

Figure A1. Chemical structure of the succinimidyl ester of carboxyfluorescein diacetate (CFDA-

SE), used to form a fluorescent dye-protein adducts with melanosomes. This structure was obtained from the manufacturer’s (Invitrogen) product description. (Invitrogen, 2006)

176

Appendix II: Supplemental Data to Chapter 5.

AII.1. Differentiation and Immunoblot Analysis of Keratinocytes

The expression of the acid hydrolase, Cath D, in both undifferentiated and differentiated KC cultures was explored. Later experimentation focused on usage of epidermal lysates rather than undifferentiated and differentiated KCs. As previously mentioned in chapter 5.4.2., Cath D expression in KC lysates were intriguingly different than that in epidermal lysates. Ultimately it was determined that using epidermal lysates to investigate the expression of the various acid hydrolases would more easily re-capitulate an in vivo like expression. Furthermore, the potential variability between the differentiation of distinct KC cell lines was negated using epidermal lysates

Differentiation of KC was accomplished by supplementing the KC media Epilife with 1.2mM

Ca2+ for a duration of 4 days. Prior to testing for Cath D expression, the cell lines were evaluated for specific antibodies, indicative of the state of differentiation. It is known that there are cues that assist in the initiation of epidermal KC stratification. The gene for the transcription factor p63, which is a homolog for p53, is expressed in progenitor (basal) layer of the epidermis. p63 is instrumental in the molecular switch towards differentiation and has also been implicated in functions that maintain the stem properties of basal KCs. (Mack, Anand and Maytin, 2005)

(Candi et al, 2008) Here, the reduction in p63 expression indicates the transition from a more stem like (basal) KC to a differentiated KC. Simultaneous analysis of involucrin, a differentiation marker, was completed. The expression of involucrin was expected and observed to increase in KC cultures supplemented with 1.2mM Ca2+ . This information is depicted in the

177

WB of Figure A2. Human melanocytes, which are terminally differentiated unlike basal KC of the epidermis displayed no expression of p63. Similarly, as melanocytes were not expected to express involucrin, a marker for KC stratification the cell lines not display expression of involucrin in the WB. Human epidermal lysates showed a slight p63 expression, indicative of basal keratinocytes, but a strong involucrin expression.

Figure A2. Differential expression of involucrin and p63 in differentiated KC lysates.

178

AII.2. Immunoblot Analysis of Cath D in Undifferentiated and Differentiated KC Lysates

Figure A3. Cath D expression in undifferentiated and differentiated KC lysates.

AII.3. IIF of Acid Hydrolases in Epidermal Cryosection

An expanded version of Figure 5.2. can be found in the subsequent figures. Acid hydrolase expression and location in epidermal cryosections was assessed by multiple images of each sample cryosection, which have been compiled here. 179

Dark Foreskin Samples Light Foreskin Samples

Dark Foreskin Sample 1 _Cath L2 1:20 and GαM-546 1:200 Light Foreskin Sample 1 _Cath L2 1:20 and GαM-546 1:200

Dark Foreskin Sample 2_ Cath L2 1:20 and GαM-546 1:200 Light Foreskin Sample 2 _Cath L2 1:20 and GαM-546 1:200

Dark Foreskin Sample 3_ Cath L2 1:20 and GαM-546 1:200 Light Foreskin Sample 3 _Cath L2 1:20 and GαM-546 1:200

Dark Foreskin Sample 4_ Cath L2 1:20 and GαM-546 1:200 Light Foreskin Sample 4 _Cath L2 1:20 and GαM-546 1:200

Figure A.4. IIF of Cath L2 in four distinct DS and LS Samples

180 Dark Foreskin Samples Light Foreskin Samples

Dark Foreskin Sample 1 _ Cath B 1:100 and GαM-546 1:200 Light Foreskin Sample 1 _Cath B 1:100 _GαM-546 1:200

Dark Foreskin Sample 2 _ Cath B 1:100 and GαM-546 1:200 Light Foreskin Sample 2 _Cath B 1:100 _GαM-546 1:200

Dark Foreskin Sample 3 _ Cath B 1:100 and GαM-546 1:200 Light Foreskin Sample 3 _Cath B 1:100 _GαM-546 1:200

Dark Foreskin Sample 4 _ Cath B 1:100 and GαM-546 1:200 Light Foreskin Sample 4 _Cath B 1:100 _GαM-546 1:200

Figure A.5. IIF of Cath B in four distinct DS and LS Samples

181 Dark Foreskin Samples Light Foreskin Samples

Dark Foreskin Sample 1 _Cath D 1:100 and GαM-546 1:200 Light Foreskin Sample 1 _Cath D_ 1:100 _GαM-546 1:200

Dark Foreskin Sample 2_ Cath D 1:100 and GαM-546 1:200 Light Foreskin Sample 2 _Cath D_ 1:100 _GαM-546 1:200

Dark Foreskin Sample 3 _ Cath D_ 1:100 and GαM-546 1:200 Light Foreskin Sample 3 _Cath D_ 1:100 _GαM-546 1:200

Light Foreskin Sample 4 _Cath D_ 1:100 _GαM-546 1:200

Figure A.6. IIF of Cath D in three DS and four LS Samples

182 Dark Foreskin Samples Light Foreskin Samples

Dark Foreskin Sample 1 _ B3GTL 1:25 and GαR-5461:200 Light Foreskin Sample 1 _ B3GTL 1:25 and GαR-5461:200

Dark Foreskin Sample 1A _ B3GTL 1:25 and GαR-5461:200 Light Foreskin Sample 1A _B3GTL 1:25 and GαR-5461:200

Dark Foreskin Sample 2 _ B3GTL 1:25 and GαR-5461:200 Light Foreskin Sample 2_ B3GTL 1:25 and GαR-5461:200

Dark Foreskin Sample 2A_ B3GTL 1:25 and GαR-5461:200 Light Foreskin Sample 2A _ B3GTL 1:25 and GαR-5461:200

Figure A.7. IIF of B3GTL in two distinct DS and LS Samples

183

AII.4. Supplemental Analyses and Calculations for Biochemical Assay

AII.4.1. Generating AMC Standard Curve

A 7-amino-4-methylcoumarin (AMC; Sigma-Aldrich Co., St. Louis, MO) standard was generated to be used as a reference standard for the biochemical assay. The AMC standard was used for two reasons, first to indicate the highest readout in relative fluorescence units (RFU) at a given AMC concentration and secondly as a conversion factor needed to convert from RFUs to a more commonly utilized unit of measure. The AMC reference standard was created by diluting

AMC powder in dimethyl sulfoxide (DMSO) to create a 2 mM stock. This 2mM stock was used to create a 20 µM solution in assay buffer (components detailed in materials and methods section

5.3.6.) The AMC was then serially diluted with assay buffer to create concentrations utilized in the assay: 10 µM, 5 µM, 2.5 µM, 1.25 µM, 0.625 µM, 0.3125 µM. The standard curve graph generated for the biochemical assay can be found in Figure A.8..

μM AMC vs RFU 62500

50000 37500 y = 5617.7x + 1178.3

RFU 25000 R² = 0.9974 12500 0

0 2.5 5 7.5 10 12.5 μM AMC

Figure A.8. Standard curve of various AMC concentrations versus RFU.

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AII.4.2. Determining Substrate Concentration for Biochemical Assay

To determine the appropriate substrate concentration to utilize in the biochemical assay of Cath

L2 in skin biological samples, various substrate (ZLR-AMC) concentrations were tested. The substrate concentrations tested were 5 µM,6 µM,8 µM,11 µM,18 µM and 50 µM in the presence of 1µg/mL recombinant human Cath L2 (rhCathL2; R&D Systems, Minneapolis, MN). The substrates were assessed using the buffer, pH, and temperature detailed previously in the materials and methods section 5.3.6.. The readouts were obtained as RFU of the released AMC product, which were then converted to a reading of velocity in RFU/min. A Lineweaver-Burke plot of 1/substrate versus 1/velocity (1/[S] vs. 1/[V]) was completed to determine the Michaelis-

Menten constant (Km) and the enzyme’s maximum rate (Vmax) of reaction. The Km was determined to be the negative inverse of the x-intercept and Vmax was the inverse of the y- intercept. From the experimentally determined Km, the appropriate substrate concentration,

50µM, was calculated at (10 x Km) to ensure saturated substrate conditions. The RFU value corresponding to 50µM substrate was then compared to the RFU value of 50µM AMC generated from the AMC standard curve. It was confirmed that the RFU values corresponding to the release of AMC from 50µM ZLR-AMC substrate (in the presence of 1µg/mL rhCathV) was below 5% of corresponding RFU readout from the 50µM AMC value of the standard curve, to ensure linear kinetics. 50µM ZLR-AMC was used in the biochemical assay as the suitable substrate concentration.

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AII.4.3. Calculating Specific Activity

Slope of the AMC standard curve. ൌ ͷ͸ͳ͹Ǥ͹ Ɋ

ൈ ൌ ͷ͸ͳ͹Ǥ͹ Slopes of the linear portion of sample plots in RFU (AMC released by substrate cleavage minus blank values) versus time (min). This velocity calculation indicates the product formed per unit time.

Table A.1. Velocity of light and dark epidermal sample cleavage of ZLR-AMC Dark Epidermal Sample Readouts Light Epidermal Sample Readouts ͳ ൌ ͳͶͷǤͷʹ ͳ ൌ ͵ͺͲǤ͹͹ ʹ ൌ ʹͻʹǤͶͲ ʹ ൌ ͶͷͷǤ͸ͺ ͵ ൌ ʹͷͺǤͳͳ ͵ ൌ ͵ͺͲǤͻͻ

The readout from the samples were then converted using the slope of the standard curve. ൌ

ൌ ൈ ൈ ൌ ൈ The specific activity was then calculated to determine the amount of product produced per mg protein per unit time (min). The amount of protein per sample was calculated by a protein content assay (as described in materials and methods section 5.3.5.) and utilized at a final concentration of 0.1µg/µl. Ɋ ൌ ൈ ൈ ͲǤͳɊ

Ɋ ͳ ͳͲͲͲɊ ൌ ൈ ൈ ൈ ൈ ͲǤͳɊ ͳͲͲͲɊ

ൌ ͲǤͳ ൈ 186

Table A.2. Specific activity of light and dark epidermal samples. Specific Activity (Dark Epidermal Samples) Specific Activity (Light Epidermal Samples) ͳ ൌ ͲǤʹͷͻ ͳ ൌ ͲǤ͸͹ͺ ൈ ൈ ʹ ൌ ͲǤͶͷͻ ʹ ൌ ͲǤͺͳͳ ൈ ൈ ͵ ൌ ͲǤͷʹͲ ͵ ൌ ͲǤ͸͹ͺ ൈ ൈ

Average = 0.413 ܖܕܗܔ Average = 0.722 ܖܕܗܔ ܕܑܖൈܕ܏ ܕܑܖൈܕ܏

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